Stem-Cell Nanoengineering [1 ed.] 9781118540671, 9781118540619

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Stem-Cell Nanoengineering

Stem-Cell Nanoengineering

Editors

Hossein Baharvand and Nasser Aghdami

Copyright © 2015 by Wiley-Blackwell. All rights reserved Published by John Wiley & Sons, Inc., Hoboken, New Jersey Published simultaneously in Canada No part of this publication may be reproduced, stored in a retrieval system, or transmitted in any form or by any means, electronic, mechanical, photocopying, recording, scanning, or otherwise, except as permitted under Section 107 or 108 of the 1976 United States Copyright Act, without either the prior written permission of the Publisher, or authorization through payment of the appropriate per-copy fee to the Copyright Clearance Center, Inc., 222 Rosewood Drive, Danvers, MA 01923, (978) 750-8400, fax (978) 750-4470, or on the web at www.copyright.com. Requests to the Publisher for permission should be addressed to the Permissions Department, John Wiley & Sons, Inc., 111 River Street, Hoboken, NJ 07030, (201) 748-6011, fax (201) 748-6008, or online at http://www.wiley.com/go/permissions. Limit of Liability/Disclaimer of Warranty: While the publisher and author have used their best efforts in preparing this book, they make no representations or warranties with respect to the accuracy or completeness of the contents of this book and specifically disclaim any implied warranties of merchantability or fitness for a particular purpose. No warranty may be created or extended by sales representatives or written sales materials. The advice and strategies contained herein may not be suitable for your situation. You should consult with a professional where appropriate. Neither the publisher nor author shall be liable for any loss of profit or any other commercial damages, including but not limited to special, incidental, consequential, or other damages. For general information on our other products and services or for technical support, please contact our Customer Care Department within the United States at (800) 762-2974, outside the United States at (317) 572-3993 or fax (317) 572-4002. Wiley also publishes its books in a variety of electronic formats. Some content that appears in print may not be available in electronic formats. For more information about Wiley products, visit our web site at www.wiley.com. Library of Congress Cataloging-in-Publication Data: Stem cell nanoengineering / edited by Hossein Baharvand and Nasser Aghdami.   p. cm.   Includes bibliographical references and index.   ISBN 978-1-118-54061-9 (cloth)   I.  Baharvand, Hossein, editor.  II.  Aghdami, Nasser, 1971– , editor.  [DNLM: 1. Stem Cells. 2. Cell Engineering. 3. Nanostructures. 4. Nanotechnology. QU 325]  QH588.S83  616.02ʹ774–dc23 2014032059 Cover image: Stem-Cell 7980051 © Luismmolina /iStockphoto Molecule 12500167 © Cosmin4000 /iStockphoto Cluster-modified © Alwayncooper /iStockphoto

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To the memory of Dr. Saeid Kazemi Ashtiani, a wonderful colleague, a great stem cell biologist, and an inspirational advocate for stem cell research in Iran. To our professors, teachers, students, and families.

Contents

About the Editors Contributors Preface

ix xi xvii

Part 1  An Introduction to Stem Cells

1

  1 Adult Stem Cells Andreas Nussler and Sahar Olsadat Sajadian

3

  2 Pluripotent Stem Cells Hossein Azizi, Akbar Hajizadeh Moghaddam, and Thomas Skutella

25

  3 Interactions of Stem Cells and Components of the Extracellular Matrix Anna K. Blakney, Julie J. Antetomaso, Winnie W. Leung, and Deok-Ho Kim

35

  4 Regenerative Medicine and Cell Therapy: Past, Present, and Future Hooman Sadri-Ardekani and Anthony Atala

47

Part 2  An Introduction to Nanotechnology

67

  5 Principles of Nanotechnology Jerzy Leszczynski

69

  6 Stem-Cell Nanoengineering: Explorations in a Rapidly Moving Field Abhalaxmi Singh and Sanjeeb K. Sahoo

87

Part 3 Nanostructures for Stem-Cell Engineering – Engineering Approach97   7 Nanopatterned Surfaces for Stem-Cell Engineering Waleed Ahmed El-Said, Tae-Hyung Kim, Ki-Bum Lee, and Jeong-Woo Choi

99

  8 Biomimetic Nanostructures by Electrospinning and Electrospraying Elham Vatankhah, Molamma P. Prabhakaran, and Seeram Ramakrishna

123

  9 Nanoparticles for Stem-Cell Engineering Esmaiel Jabbari

143

10 Toxicology of Nanobiomaterials Shahin Bonakdar and Omid Mashinchian

171

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Contents

Part 4 Control of Stem-Cell Fate by Engineering of Microenvironment

185

11 Stem-Cell Responses to Surface Nanotopographies Peng-Yuan Wang and Wei-Bor Tsai

187

12 Control of Mesenchymal Stem-Cell Fate by Engineering the Nanoenvironment Habib Nikukar, Stuart Reid, Mathis O. Riehle, Adam S.G. Curtis, and Matthew J. Dalby

205

13 Delivery of Molecules and Genes/Small Interfering RNA into Stem Cells by Nanoengineering Mohsen Ashjari

Part 5 Nanotissue Engineering – Biological Approach along with Differentiation

223

243

14 Expansion of Stem Cells by Nanotissue Engineering Amir Mellati and Hu Zhang

245

15 Nanotissue Engineering of Neural Cells Sasan Jalili-Firoozinezhad, Fahimeh Mirakhori, and Hossein Baharvand

265

16 Nanotechnology and Cardiovascular Tissue Engineering Savneet Kaur and Upasana Rishiraj

285

17 Nanotissue Engineering of Musculoskeletal Cells Mohamadreza Baghaban Eslaminejad, Leila Taghiyar, and Fatemeh Safari

299

18 Nanotissue Engineering of Skin Cells Daisy M. Ramos, Aditi Subramanian, Aja Aravamudhan, Matthew Harmon, Roshan James, Namdev B. Shelke, and Sangamesh G. Kumbar

315

19 High-Throughput Screening of Stem Cell Self-Renewal and Differentiation on Nanomaterials327 Perry T. Yin, Tae-Hyung Kim, Jeong-Woo Choi, and Ki-Bum Lee

Part 6  Nanotechnology in Stem-Cell Imaging

345

20 Nanotechnology for Cellular Imaging Miroslaw Janowski, P. Walczak, and J.W.M. Bulte

347

Part 7  Nanotissue Engineering and Clinical Applications

363

21 Advancing Translational Nanotechnology to Clinical Application Michelle Griffin, Shima Salmasi, Naghmeh Naderi, Peter E. Butler, and Alexander M. Seifalian

365

22 Stem-Cell Nanoengineering from Bench to Bed Omid Mashinchian, Shahin Bonakdar, Shahriar Sharifi, and Morteza Mahmoudi

381

Index

397

About the Editors

Born in 1972, Hossein Baharvand received his PhD degree from Khwarizmi University (formerly Tarbiat Moallem University), Tehran, Iran in 2004 in the field of cell and developmental biology. He joined the Royan Institute when he was a MSc student in 1995. He became professor of Stem Cells and Developmental Biology in 2012, director of the Royan Institute for Stem Cell Biology and Technology from 2002 to the present, and head of the Department of Developmental Biology at the University of Science and Culture from 2006 to the present. He established the first mouse and human embryonic stem cells in Iran (2003). Since then his team has generated several mouse- and human-induced pluripotent stem cells (2008). This has enabled them to pursue many avenues of research into translational research and regenerative medicine. He has published 197 international and 98 national peer-reviewed papers, as well as six chapters in international books, seven books in Persian, and translated eight English text books into Persian. He was editor of four international books between 2009 and 2014, which were published by Springer and John Wiley & Sons, Inc. in the USA. He is editorial board member of six international journals and has been an invited speaker at several meetings. He has been the recipient of 26 international and national prizes, including: 10th (2004), 12th (2006), and 17th (2012) annual Razi research award on medical science, which is hosted by the Iran Ministry of Health and Medical Education; 26th Khwarizmi International Award (2013), which is hosted by the Iran Ministry of Science, Research and Technology; 27th annual book of the year of the Islamic Republic of Iran (2010); Dr. Hadavi’s Award of the Academy of Medical Sciences of Iran in 2010 and 2014; and winner of the Islamic Educational, Scientific, and Cultural Organization (ISESCO) Prize for research in the field of Biology (2010). Moreover, he was introduced as prominent Professor in the third term of Allameh Tabatabaei’s Award hosted by Iran Vice Presidency for Science and Technology, Presidency and National Elite Foundation (2014). He is the winner of the United Nations Educational, Scientific and Cultural Organization’s (UNESCO) Equatorial Guinea international prize (2014) for research in life sciences due to improving the quality of human life through his stem-cell research, with numerous applications in regenerative medicine. Dr. Nasser Aghdami earned a MD degree from Urumieh Medical University (Iran) in 1998 and a PhD in immunology from Tarbiat Modares University (Iran) in 2004. He joined the Hematology, Oncology and BMT Research Center in 2000 as a Principal Investigator, and became assistant professor at the Royan Institute for Stem Cell Biology and Technology in 2006. Dr. Aghdami has published over 40 international papers and is a member of the scientific board of the Royan Institute for Stem Cell Biology and Technology. He established the Department of Regenerative Medicine at the Royan Institute in 2010, and is the head of the department. He is the co-editor of three international books in the field of stem-cell and regenerative medicine. He received the Young Investigator Award from the International

ix

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About the Editors

Society for Cell Therapy in 2012. Dr. Aghdami’s research interests lie in the field of stem cell biology and regenerative medicine, and he is the principal investigator of over 50 clinical trials in cell therapy. His efforts in regenerative medicine and its related standards have led to numerous applications in cell therapy. Currently he is the chief executive officer of Cell Tech Pharmed Company. This achievement, as a tremendous honor, has enabled him to realize his goal of converting widespread research to reality.

Contributors

Nasser Aghdami Department of Regenerative Medicine and Department of Stem Cells and Developmental Biology at Cell Science Research Center Royan Institute for Stem Cell Biology and Technology, ACECR, Tehran, Iran Julie J. Antetomaso Department of Bioengineering University of Washington Seattle, WA, USA Aja Aravamudhan Institute for Regenerative Engineering, Raymond and Beverly Sackler Center for Biological, Physical and Engineering Sciences, and Department of Orthopaedic Surgery University of Connecticut Health Center Farmington, CT, USA Mohsen Ashjari Institute of Nanoscience and Nanotechnology University of Kashan Kashan, Iran Department of Stem Cells and Developmental Biology at Cell Science Research Center Royan Institute for Stem Cell Biology and Technology, ACECR, Tehran, Iran

Anthony Atala Wake Forest Institute of Regenerative Medicine (WFIRM) Wake Forest University School of Medicine Winston-Salem, NC, USA Department of Urology Wake Forest University School of Medicine Winston-Salem, NC, USA Hossein Azizi Institute for Anatomy and Cell Biology Department of Neuroanatomy, Medical Faculty University of Heidelberg Heidelberg, Germany Amol University of Special Modern Technologies Amol, Iran Department of Stem Cells and Developmental Biology at Cell Science Research Center Royan Institute for Stem Cell Biology and Technology, ACECR, Tehran, Iran Hossein Baharvand Department of Stem Cells and Developmental Biology at Cell Science Research Center Royan Institute for Stem Cell Biology and Technology, ACECR, Tehran, Iran

xi

xii

Contributors

Anna K. Blakney Department of Bioengineering University of Washington Seattle, WA, USA Shahin Bonakdar National Cell Bank Pasteur Institute of Iran Tehran, Iran J.W.M. Bulte Russell H. Morgan Department of Radiology and Radiological Science, Division of MR Research; Cellular Imaging Section and Vascular Biology Program, Institute for Cell Engineering; Departments of Oncology, Biomedical Engineering and Chemical and Biomolecular Engineering The Johns Hopkins University School of Medicine Baltimore, MD, USA Peter E. Butler UCL Centre for Nanotechnology and Regenerative Medicine Division of Surgery and Interventional Science University College London London, UK Department of Plastic and Reconstructive Surgery Royal Free London NHS Foundation Trust Hospital London, UK Jeong-Woo Choi Department of Chemical and Biomolecular Engineering Sogang University Seoul, Republic of Korea Adam S.G. Curtis Centre for Cell Engineering Institute for Molecular, Cell and Systems Biology College of Medical, Veterinary and

Life Sciences University of Glasgow Glasgow, UK Matthew J. Dalby Centre for Cell Engineering Institute for Molecular, Cell and Systems Biology College of Medical, Veterinary and Life Sciences University of Glasgow Glasgow, UK Waleed Ahmed El-Said Department of Chemistry Faculty of Science Assiut University Assiut, Egypt Mohamadreza Baghaban Eslaminejad Department of Stem Cell and Developmental Biology at Cell Science Research Center Royan Institute for Stem Cell Biology and Technology, ACECR, Tehran, Iran Sasan Jalili-Firoozinezhad Department of Stem Cells and Developmental Biology at Cell Science Research Center Royan Institute for Stem Cell Biology and Technology, ACECR, Tehran, Iran Departments of Surgery and of Biomedicine University Hospital Basel University of Basel Basel, Switzerland Michelle Griffin UCL Centre for Nanotechnology and Regenerative Medicine Division of Surgery and Interventional Science University College London London, UK

Contributors

Matthew Harmon Institute for Regenerative Engineering, Raymond and Beverly Sackler Center for Biological, Physical and Engineering Sciences, and Departments of Orthopaedic Surgery and Materials Science and Engineering University of Connecticut Health Center Farmington, CT, USA Esmaiel Jabbari Biomimetic Materials and Tissue Engineering Laboratories Department of Chemical Engineering University of South Carolina Columbia, SC, USA Roshan James Institute for Regenerative Engineering, Raymond and Beverly Sackler Center for Biological, Physical and Engineering Sciences, and Department of Orthopaedic Surgery University of Connecticut Health Center Farmington, CT, USA Miroslaw Janowski Russell H. Morgan Department of Radiology and Radiological Science, Division of MR Research and Cellular Imaging Section and Vascular Biology Program, Institute for Cell Engineering The Johns Hopkins University School of Medicine Baltimore, MD, USA

xiii

Deok-Ho Kim Department of Bioengineering, Center for Cardiovasular Biology and Institute of Stem Cell and Regenerative Medicine University of Washington Seattle, WA, USA Tae-Hyung Kim Department of Chemical and Biomolecular Engineering Sogang University Seoul, Republic of Korea Department of Chemistry and Chemical Biology Rutgers, The State University of New Jersey Piscataway, NJ, USA Sangamesh G. Kumbar Institute for Regenerative Engineering, Raymond and Beverly Sackler Center for Biological, Physical and Engineering Sciences, Departments of Orthopaedic Surgery and Chemical, Materials and Biomolecular Engineering University of Connecticut Health Center Farmington, CT, USA Ki-Bum Lee Department of Chemistry and Chemical Biology and Department of Biomedical Engineering Rutgers, The State University of New Jersey Piscataway, NJ, USA

Departments of NeuroRepair and Neurosurgery Mossakowski Medical Research Centre Polish Academy of Sciences Warsaw, Poland

Jerzy Leszczynski Interdisciplinary Center for Nanotoxicity Department of Chemistry and Biochemistry Jackson State University Jackson, MI, USA

Savneet Kaur School of Biotechnology Gautam Buddha University Uttar Pradesh, India

Winnie W. Leung Department of Bioengineering University of Washington Seattle, WA, USA

xiv

Contributors

Morteza Mahmoudi Department of Nanotechnology and Nanotechnology Research Center Faculty of Pharmacy Tehran University of Medical Sciences Tehran, Iran Division of Cardiovascular Medicine, Stanford University School of Medicine Stanford, CA, USA

Habib Nikukar Centre for Cell Engineering Institute for Molecular, Cell and Systems Biology College of Medical, Veterinary and Life Sciences University of Glasgow Glasgow, UK Shahid Sadoughi University of Medical Sciences and Health Services Yazd, Iran

Omid Mashinchian Department of Medical Nanotechnology School of Advanced Technologies in Medicine Tehran University of Medical Sciences Tehran, Iran

Andreas Nussler Department of Traumatology Eberhard Karls Universität Tübingen BG Clinic Tübingen Tübingen, Germany

Amir Mellati School of Chemical Engineering The University of Adelaide Adelaide, South Australia, Australia

Molamma P. Prabhakaran Center for Nanofibers and Nanotechnology Nanoscience and Nanotechnology Initiative Faculty of Engineering National University of Singapore Singapore

Fahimeh Mirakhori Department of Stem Cells and Developmental Biology at Cell Science Research Center Royan Institute for Stem Cell Biology and Technology, ACECR, Tehran, Iran School of Biology, College of Science University of Tehran Tehran, Iran Akbar Hajizadeh Moghaddam Amol University of Special Modern Technologies Amol, Iran Naghmeh Naderi UCL Centre for Nanotechnology and Regenerative Medicine Division of Surgery and Interventional Science University College London London, UK

Seeram Ramakrishna Center for Nanofibers and Nanotechnology, Nanoscience and Nanotechnology Initiative, Department of Mechanical Engineering Faculty of Engineering National University of Singapore Singapore Daisy M. Ramos Institute for Regenerative Engineering, Raymond and Beverly Sackler Center for Biological, Physical and Engineering Sciences, Departments of Orthopaedic Surgery and Materials Science and Engineering University of Connecticut Health Center Farmington, CT, USA Stuart Reid SUPA, Thin Film Centre University of the West of Scotland Paisley, UK

Contributors

Mathis O. Riehle Centre for Cell Engineering Institute for Molecular, Cell and Systems Biology College of Medical, Veterinary and Life Sciences University of Glasgow Glasgow, UK Upasana Rishiraj School of Biotechnology Gautam Buddha University Uttar Pradesh, India Hooman Sadri-Ardekani Wake Forest Institute of Regenerative Medicine (WFIRM) Wake Forest University School of Medicine Winston-Salem, NC, USA Reproductive Biotechnology Research Center Avicenna Research Institute, ACECR, Tehran, Iran Department of Urology Wake Forest University School of Medicine Winston-Salem, NC, USA Fatemeh Safari Department of Stem Cell and Developmental Biology at Cell Science Research Center Royan Institute for Stem Cell Biology and Technology, ACECR, Tehran, Iran Sanjeeb K. Sahoo Laboratory of Nanomedicine Institute of Life Sciences Bhubaneswar, Odisha, India Sahar Olsadat Sajadian Department of Traumatology Eberhard Karls Universität Tübingen BG Clinic Tübingen Tübingen, Germany Shima Salmasi UCL Centre for Nanotechnology and Regenerative Medicine

xv

Division of Surgery and Interventional Science University College London London, UK Alexander M. Seifalian UCL Centre for Nanotechnology and Regenerative Medicine Division of Surgery and Interventional Science University College London London, UK Department of Plastic and Reconstructive Surgery Royal Free London NHS Foundation Trust Hospital London, UK Namdev B. Shelke Institute for Regenerative Engineering, Raymond and Beverly Sackler Center for Biological, Physical and Engineering Sciences, Department of Orthopaedic Surgery University of Connecticut Health Center Farmington, CT, USA Shahriar Sharifi Department of Biomedical Engineering University Medical Center Groningen Groningen, The Netherlands Abhalaxmi Singh Laboratory of Nanomedicine Institute of Life Sciences Bhubaneswar, Odisha, India Thomas Skutella Institute for Anatomy and Cell Biology Department of Neuroanatomy, Medical Faculty University of Heidelberg Heidelberg, Germany Aditi Subramanian Institute for Regenerative Engineering, Raymond and Beverly Sackler Center for Biological, Physical and Engineering Sciences,

xvi

Contributors

Departments of Orthopaedic Surgery and Biomedical Engineering University of Connecticut Health Center Farmington, CT, USA Leila Taghiyar Department of Stem Cell and Developmental Biology at Cell Science Research Center Royan Institute for Stem Cell Biology and Technology, ACECR, Tehran, Iran

The Johns Hopkins University School of Medicine Baltimore, MD, USA Department of Radiology Faculty of Medical Sciences University of Warmia and Mazury Olsztyn, Poland Peng-Yuan Wang Industrial Research Institute Swinburne (IRIS)

Wei-Bor Tsai Department of Chemical Engineering National Taiwan University Taipei, Taiwan

Department of Chemistry and Biotechnology, Faculty of Science, Engineering and Technology, Hawthorn, Victoria, Australia

Elham Vatankhah Center for Nanofibers and Nanotechnology Nanoscience and Nanotechnology Initiative Faculty of Engineering National University of Singapore Singapore

CSIRO Manufacturing Flagship, Clayton, Victoria, Australia

P. Walczak Russell H. Morgan Department of Radiology and Radiological Science, Division of MR Research and Cellular Imaging Section and Vascular Biology Program, Institute for Cell Engineering

Perry T. Yin Department of Biomedical Engineering Rutgers, The State University of New Jersey Piscataway, NJ, USA Hu Zhang School of Chemical Engineering The University of Adelaide Adelaide, South Australia, Australia

Preface

Nanobiotechnology is a fast growing area of research that aims to create nanodevices, nanoparticles, and nanoscale development in the field of stem-cell and tissue-engineering based therapies. Concepts and discoveries from this field, along with stem cell research, provide exciting opportunities of using stem cells for regeneration of tissues and organs and to address the challenges of disease therapeutics. In this book, Stem-Cell Nanoengineering, we aim to provide the premier source of reviews in nanotechnology approaches towards stem cells and tissue engineering and regenerative medicine, which will serve as a textbook. This book overviews the fast moving field of stem cells, discusses challenges to the field that can be addressed through nanotechnology, provides information on stem cells, principles of nanobiotechnology, manufacture of nanoscaffolds, nanotissue engineering, changes in stem-cell-fate decisions by micro- and nanoengineering of the microenvironment, application of nanotechnology in stem-cell tracking and molecular imaging, and finally stem-cell nanoengineering from bench to bedside. The contributions to this book, all written by renowned experts in their respective disciplines, describe and explore various facets of this field. This book will be an especially valuable resource for biomedical and bioengineering researchers and clinicians. This book could useful for postgraduate students and advanced undergraduate students in cell biology, biochemistry, genetics, developmental biology, biomaterial, medical engineering, nanotechnology, and biomechanics; physicians; life science scientists; biomedical researchers; cell biologists; academics; surgeons; scientists and engineers in the field of regenerative medicine; clinicians and specialists; biotechnology and pharmaceutical industry professionals. We want to sincerely thank all authors that have contributed to this book for their devoted efforts and their excellent contributions. We hope that you, as a reader, will enjoy this book. We are also grateful to Drs. Hamid Gourabi, Abdolhossein Shahverdi, and Ahmad Vosough Dizaj for having faith in and supporting us throughout this project. We also wish to acknowledge the great support provided by many at John Wiley and Sons. A special thank you goes to our dedicated colleagues at Royan Institute for Stem Cell Biology and Technology, who, with their tireless commitment for stem-cell research and therapy, have become crucial factors in encouraging us to edit this book. We are grateful to Asma Ghodsi and Zahra Maghari for their assistance with collecting the chapters and in follow-up. Our thanks also to the many scientific colleagues and students, Sasan Jalili-Firoozinezhad, Mohammad Kazemi Ashtiani, Hossein Ghanian, Leila Montazeri, and Fahimeh Khayatan, and other members of the Cell Engineering program at the Royan Institute who stimulated our intrests, encouraged us, and supported our efforts. Hossein Baharvand and Nasser Aghdami

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Part 1

An Introduction to Stem Cells

Chapter 1

Adult Stem Cells Andreas Nussler and Sahar Olsadat Sajadian

Eberhard Karls Universität Tübingen, Department of Traumatology, Tübingen, Germany

Introduction Humans and many animals share the ability to regenerate missing parts of the body. Although humans are not able to replace missing parts of the body as a whole, like legs or hands for example, the human body is able to perpetually regenerate various tissues and blood. The “mysterious” cell type that enables the human body to perform this regeneration was discovered in the 1950s and subsequently named “stem cell” [1]. The first stem cells were discovered in the bone marrow. Therefore, at the beginning, stem cells were almost exclusively isolated from human bone marrow. Later on, the routine isolation and manipulation of bone marrow stem cells led to the development of a method of bone marrow transplantation for the treatment of blood diseases, such as leukemia, that is used all over the world today. Regardless of their origin, stem cells have the following specific properties: their division and self-renewal capacity over long periods of time, their lack of specification, and their ability to differentiate into specialized cell types. Over the past 50 years, many important discoveries have raised hope that stem cell research will achieve major breakthroughs in medicine. A short history of stem-cell research is presented in Table 1.1, and the different kinds of stem cells with their respective sources are displayed in Table 1.2.

Adult Stem Cells Adult stem cells (ASCs) are also called stromal cells. Their regenerative potential has been ­recognized several decades ago. For instance, it has been demonstrated that hematopoietic stem cells that are derived from adult tissues are able to generate every type of blood cell [2]. However, adult stem cells were thought to have a rather restricted potential for generating new tissue, but recent studies have changed this viewpoint. Recent observations have suggested that, in addition to the production of the derivatives of the blood system, stem cells from the bone marrow of the juvenile and adult organism can create muscle and neuron-like

Stem-Cell Nanoengineering, First Edition. Edited by Hossein Baharvand and Nasser Aghdami. © 2015 John Wiley & Sons, Inc. Published 2015 by John Wiley & Sons, Inc.

3

Table 1.1  Important events in the history of stem cell research Year

Event

Reference

1860–1920

Existence of stem cells inferred from the analysis of embryonic development and microscopy of bone marrow

http://www.stemcellnetwork.ca/index. php?page=stem-cell-timeline&hl=eng

1948–1958

Stem cell mechanisms deduced for sperm development and intestinal epithelium replacement

http://www.stemcellnetwork.ca/index. php?page=stem-cell-timeline&hl=eng

1956

First bone marrow transplants performed in human patients

[1]

1961

The existence and properties of transplantable stem cells in mouse bone marrow are established and the first colony methodology for counting these cells is introduced. This discovery paves the way for all current research on adult and embryonic stem cells

[49]

1968

Achievement of first allogeneic human bone marrow transplantations without lethal rejection reactions

[119]

1974

Embryonic cancer cells of mice and telomerase are shown to participate in the development of normal tissue

[120]

1978

Transplantable stem cells are discovered in human umbilical cord blood

http://www.stemcellnetwork.ca/index. php?page=stem-cell-timeline&hl=eng

1981

First embryonic stem cells derived from the inner cell mass of mouse blastocysts

[121]

1982–1986

First methodology for targeted genetic modification in embryonic stem cells developed

http://www.stemcellnetwork.ca/index. php?page=stem-cell-timeline&hl=eng

1992

Neural stem cells identified in the adult human brain

[122]

1994

First isolation of cancer stem cells from cancer cells

[53]

1995

First isolation of embryonic stem cell lines from primates

[123]

1998

First human embryonic stem cell line derived

[99]

2000

Retinal stem cells identified in mice

[124]

2001

Dermal stem cells identified in adult skin tissue

[125]

2003

Cancer stem cells identified in human brain tumor Rare human breast cancer stem cells identified

[126, 127]

2005

First evidence for human bone cancer stem cells

2006

First induced pluripotent stem cells (iPSCs) generated by reprogramming adult mouse skin cells.

[128]

2007

Mario Capecchi, Martin Evans and Oliver Smithies win the Nobel Prize for Physiology or Medicine because of their discoveries enabling modifications of the germline gene in mice Human colon cancer stem cells identified

http://www.stemcellnetwork.ca/index. php?page=stem-cell-timeline&hl=eng

2008

Sam Weiss is awarded the Gairdner Prize for the discovery of neural stem cells

http://www.stemcellnetwork.ca/index. php?page=stem-cell-timeline&hl=eng

2009

John Gurdon and Shinya Yamanaka win the Lasker Prize for their discoveries in the field of nuclear reprogramming. Yamanaka is also awarded the Gairdner Prize

http://www.stemcellnetwork.ca/index. php?page=stem-cell-timeline&hl=eng

2010

IPS cells created by transfection of m-RNA Adult cells reprogrammed directly to blood cells, cardiac muscle and neurons

[129, 130]

2011

Identification of multipotent human blood stem cells capable of generating all types of cells in the blood system

[131]

2012

John Gurdon and Shinya Yamanaka win the Nobel Prize for Physiology or Medicine for their finding that mature cells can be reprogrammed to a pluripotent state

http://www.stemcellnetwork.ca/index. php?page=stem-cell-timeline&hl=eng

Not yet

Laboratory

Induced pluripotent stem cells

Behave like embryonic stem cells Produce all types of cells Useful tool for drug development and modeling of diseases

Generation of skeletal Cartilage and bone repair, tissues: bone, Repair of blood cartilage, fat vessels after heart attack

Could provide patient-specific treatment Ability to produce almost all kinds of cells in the body

Can be easily extracted from the bone marrow of the patient Can be used for an efficient generation of the skeletal tissue in the human body

No major ethical concerns Various sources of MSCs and it cannot be established whether cells from different sources are identical or not.

No major ethical Expensive and difficult to concerns culture Need of feeder layer Possibility of tumor growth

No major ethical concerns

No major ethical concerns

Limited number of cells produced (1 in 104−5 ) Specific adult tissue with cell selection Difficult to identify and grow in the laboratory

Tissue-specific cell line Safety Immune privilege Lack of tumor growth No need for feeder layer

Limited number of cells Blood from the cord must be matched to the patient

Destruction of the human blastocyst Donation requires informed consent

Ethical concerns

Expensive and difficult to use Need for multiple growth factor Need for feeder layer Possible tumor formation Difficult to control their differentiation

Disadvantages and limitations

Produce all kinds of cells in the body. Immortality and capacity of eternal self-renewal

Advantages

Treatment of leukemia Can be frozen and and blood disorder stored

Skin and blood stem cells have been used for skin graft and bone marrow transplantation

Bone marrow

Tissue-specific Only produce the type of cells that belong to their own tissue

Mesenchymal stem cells

Adult tissue (blood, skin, bone, skeletal muscle, pancreas, liver)

Adult stem cells (tissue-specific stem cells)

Macular degeneration Generation of an entire organism consisting of hundreds of cell types Large source of excess blastocysts from IVF

Clinical trial

Generation of blood-specific cells

Blastocyte (early stage embryo; 0–2 weeks)

Embryonic stem cells

Characteristics

Umbilical cord Umbilical cord blood stem cells (hematopoietic stem cells) (tissue stem cells)

Source

Type of stem cells

Table 1.2  Sources, characteristics, advantages and disadvantages of different types of human stem cells

6

An Introduction to Stem Cells

cells in the brain [3].Therefore, if ASCs turn out to have the same potential as embryonic stem cells (ESC), some ethical issues concerning ESCs may be overcome. Hematopoietic stem cells (HSCs) were the first type of adult stem cells to be discovered, by Becker et al., in 1963 [4]. These cells reside in the bone marrow and are able to renew themselves and to differentiate [5]. Bone marrow contains at least three types of stem cells: HSCs, the first type discovered are able to differentiate into every type of blood cell. The second type discovered a few years later, is known as bone marrow stromal cells, ­mesenchymal stem cells or skeletal stem cells: they are a mixed cell population capable of generating bone, cartilage, fat and fibrous connective tissue [6, 7]. The third type is known as endothelial progenitor cells (EPCs), and they contain a unique population of peripheral blood mononuclear cells [8]. In the 1990s, Caplan popularized the term “mesenchymal stem cells (MSCs)” [9]; however, when publishing clinical studies of MSCs, some investigators still prefer not to refer to them as stem cells [10, 11]. The Mesenchymal and Tissue Stem Cell Committee of the International Society for Cellular Therapy (ISCT) proposed a uniform nomenclature for MSCs published in 2005 [12] according to the following guidelines: first, the MSCs must be plastic-adherent when they are maintained in standard culture conditions; second, they have to express ­surface markers, such as CD105, CD73, and CD90, without expressing the surface molecules CD45, CD34, CD14 or CD11b, CD79a or CD19, and HLA-DR; third, they have to differentiate into osteoblasts, adipocytes, and chondroblasts in vitro. In addition, it has been demonstrated that MSCs differentiate into a large variety of specialized MSCs, including myocytes, tendocytes, and ligament cells [13, 14]. The MSCs reside in different locations of the body, for example in bone marrow, around blood vessels as pericytes, in fat, skin, muscles, teeth and other locations [13–15]. Until recently, it had been believed that adult stem cells only differentiate into mature phenotypes of various cells, but within the MSC lineages, it has been demonstrated that chondrocytes are able to differentiate into osteoblasts and adipocytes and change their phenotype to osteoblasts. These findings have demonstrated the plasticity of MSCs [16–19].Adult stem cells are generally thought to be rare, difficult to isolate and to purify, and difficult to maintain in an undifferentiated state when grown in culture. Therefore, developing methods for upscaling adult stem cells outside the body is currently one of the priorities of stem-cell research targeted at further clinical application.

Adult Stem-Cell Plasticity Until recently, no researcher had seriously considered the possibility of adult stem cells generating the specialized cell types that are necessary for the formation of different tissues, either from the same embryonic germ layer or from a different germ layer. Recently, however, studies have demonstrated that blood stem cells (derived from the mesoderm) may be able to generate both skeletal muscle (also derived from the mesoderm) and neurons (derived from the ectoderm) [20, 21]. The term “plasticity” refers to the ability of a stem cell that is derived from one adult tissue to differentiate into a differentiated cell type from another tissue. This process is referred to either as “unorthodox differentiation” or as “trans-differentiation” [22, 23]. However, many examples have demonstrated that MSCs are able to differentiate into an endodermal phenotype, such as hepatocyte-like cells [24–26]. In addition, it has been demonstrated that MSCs differentiate into epithelial cells, such as retinal pigment epithelial cells [27, 28], sebaceous duct cells [29], skin epithelial cells [30], and tubucular epithelial cells in the kidney [17, 31]. According to the experiments that have been reported to date, adult stem cells may assume the characteristics of the cells that have developed from the primary germ

 1  Adult Stem Cells

7

layer. For example, many experiments on plasticity involve stem cells derived from bone marrow, which is considered to be of mesodermal origin. The bone-marrow stem cells may differentiate into another mesodermally derived tissue, such as skeletal muscle or cardiac muscle [20, 32–34].

Classification of Adult Stem Cells Adult stem cell niches are distributed throughout several regions of the body, including bone marrow, brain, fat, skeletal muscle, retina, liver, and skin. Some of these adult stemcell types are discussed in detail below. Many tissues that have specific ASC phenotypes have been identified so far. Some examples are mammary stem cells, intestinal stem cells, endothelial stem cells, olfactory stem cells [35], liver-derived stem cells, testicular stem cells, and dental pulp stem cells [8].

Bone Marrow-Derived Stem Cells Today, bone marrow is one of the most popular sources of stem cells. Bone marrow-derived stem cells (BMSCs) have a self-renewal potential and they are readily available through bone marrow biopsy. The BMSCs contain at least three different types: HSCs, MSCs and EPCs [8]. The HSCs and the MSCs are presented in more detail later, since they are considered to have the highest potential in therapeutic approaches. Hematopoetic stem cells (HSCs) are involved in the production of blood cells. This process is called hematopoiesis. They give rise to the entirety of the blood cells in the human body. Therefore, HSCs are a potential tool for curing blood diseases, for example leukemia or lymphoma, as well as blood disorders, for example anemia or immunodeficiencies. Mesenchymal stem cells (MSCs) are derived from bone marrow stromal progenitor cells and can form “mesenchyme,” a loose connective tissue. The MSCs can be isolated from HSCs by their capability to adhere to tissue culture plastic. They are able to form mesodermal and non-mesodermal tissues, such as bone, cartilage, tendon, adipose tissue, and muscle [36]. Endothelial progenitor cells (EPCs) include a unique population of peripheral-blood mononuclear cells derived from bone marrow that are involved in postnatal neovascularization during wound healing, limb ischemia, post-myocardial infarction syndrome, arteriosclerosis, and tumor development. Both HSCs and EPCs are derived from common a precursor called hemangioblast [37].

Adipose Tissue-Derived Stem Cells or Adipocyte-Derived Mesenchymal Stem Cells White adipose tissue is one of two types of adipose tissue found in mammals. The tasks of this tissue consist in saving energy and in acting as a thermal insulator. Adipose tissues originate from the mesodermal layer of embryos and can develop pre- as well as postnatally. Three different types of cells are present in adipose tissues: adipocytes, pre-adipocytes, and a heterogeneous population called stromal vesicular fraction. Adipose tissue-derived stem cells (ADSCs) are multipotent cells that are able to differentiate into other types of mesenchymal tissues, such as adipocytes, chondrocytes, myocytes, and osteoblasts [38]. Furthermore, they grow faster and are easier to culture in vitro over a long period of time. In the past decade, abundant evidence has been presented for the fact that the secretion of vascular endothelial growth factor by ADSCs leads to the healing of damaged tissue. Several studies have been presented indicating that adipocyte-derived mesenchymal stem cells (AD-MSCs) have the ability to differentiate to hepatocyte-like cells under specific conditions [39–41].

8

An Introduction to Stem Cells

Therefore, MSCs are expected to be an ideal source for transplantation or liver tissue engineering: however, the hepatic differentiation of MSCs is still insufficient for clinical application. But the proliferation and the differentiation capacity of ASCs also cause changes in their metabolic activity. These changes may, in turn, increase the risk of tumor formation [42].

Neural Stem Cells Neural stem cells (NSCs) differentiate into three major cell types: neurons, astrocytes, and oligodendrocytes [43]. The NSCs that have been identified in the ventricular zone of the brain include neuroblasts, precursor cells and astrocytes. They all express proteins like GFAP (glial fibrillary acidic protein) and the glycoprotein CD133, which permits the identification of this cell. Most NSCs of the lateral ventricles (ependymal cells) are quiescent and do not perform active division. The NSCs express nestin, which is a specific marker of neural precursors. Furthermore, NSCs of the hippocampus are ciliated and play an important role in the memory function of the brain. They express neuronal markers, such as NeuN, neuron specific enolase, and calbindin [44]. Neural stem cells are commonly cultured in vitro as so-called neurophases. In this case, they assume a free-floating cell cluster configuration. By using the neurophase in cell culture, NSCs are capable of differentiating into glial-like cells [45]. It is believed that they have the potential of curing brain disorders, such as anxiety, depression, memory deterioration, and some brain tumors [8]. Furthermore, there is some speculation that these cells may overcome the paralyzation caused by spinal cord injuries [46, 47].

Neural Crest Stem Cells A remnant of embryonic neural-crest stem cells has been identified in the hair follicles. Similar cells have also been found in the gastrointestinal tract, in the sciatic nerve, and in the spinal cord. Neural crest stem cells can differentiate into neurons, Schwann cells, myofibroblasts, chondrocytes, and melanocytes [48].

Hematopoietic Stem Cells Nearly 50 years ago, HSCs were identified by Till and McCullough [49]. The remarkable characteristics of these cells are their ability for continuous self-renewal in the bone marrow and their ability to differentiate to all types of blood cells (Figure 1.1). Hematopoietic stem cells normally reside in bone marrow, but under certain conditions they migrate through the blood in order to settle in other tissues. They are also present in fetal liver, the spleen, placenta blood, and in the umbilical cord. A decade ago, several studies have revealed that HSCs can give rise to a liver-like cell phenotype [23, 50]. One study has demonstrated that HSCs that are transplanted into an irradiated mouse evolve not only into various blood-cell types (from the mesoderm layer of the embryo), but also into epithelial cell phenotypes in the lung, gut (endoderm layer), and skin (ectoderm layer) [51]. If HSCs are truly multipotent, their potential for life-saving regenerative therapies may be considerably expanded in the future. There are several problems concerning the implementation of standardized HSC protocols. The identification and characterization of HSCs is difficult, as those with long-term replicating ability are rare and difficult to upscale. Furthermore, HSCs have multiple phenotypes and resemble other blood or bone-marrow cells, which makes them difficult to distinguish from each other [52].

 1  Adult Stem Cells

Natural killer (NK) cell

9

Neutrophil

Bone

T lymphocytes

Lymphoid progenitor cell

Basophil

Eosinophil B lymphocyte

Hematopoietic stem cell Multipotential stem cell

Monocyte/macrophage Myeloid progenitor cell

Red blood cells Bone (or cartilage) Stromal cell

Bone matrix

Stromal stem cell

Hematopoietic supportive stroma

Osteoblast

Marrow adipocyte

Lining cell

Blood vessel

Osteocyte Pre-osteoblast

Pericyte

Skeletal muscle stem cell?

Osteoclast

Platelets

Hepatocyte stem cell?

Adipocyte

Hematopoietic stem cell

Figure 1.1  Differentiation of hematopoietic and stromal cells. (See insert for color representation of the figure.) Table 1.3  Proposed cell-surface markers of undifferentiated hematopoietic stem cells Mouse

Human

CD34 low/−

CD34+

SCA-1+

CD59+

Thy1+/low

Thy1+

CD38+−

CD38low/−

C-kit

C-kit−/low

Lin

Lin−

+

−

Over the past years, different kinds of surface markers have been used to identify, i­solate, and purify HSCs derived from the bone marrow and the blood. Undifferentiated HSCs and hematopoietic progenitor cells express c-kit, CD34, and H-2 K. These cells often lack the Lin lineage marker completely, or only express it at a very low level (Lin−/ low ). Weissman and his collaborators have focused on surface protein markers of blood cells from mice and have identified the closest common markers for mouse and human HSCs. Moreover, it has been demonstrated that the cell surface markers can no longer be identified during cell development [53]. Indeed, for transplantation purposes, cells that express the surface proteins CD34+, Thy1+, and Lin− are the most likely to contain stem cells [54, 55] (Table 1.3). There are also different sources of HSCs, for example, bone marrow, peripheral blood, umbilical cord blood, and fetal hematopoietic system. In 1985, Perkins demonstrated that all major lineages of progenitor cells can be obtained from the bodies of mouse

10

An Introduction to Stem Cells

embryos, even without adding the hematopoietic growth factor [56]. Several scientists have demonstrated that, at earlier developmental stages, HSCs from different tissues have a great ability of self-replication, show different homing and surface characteristics, and are less likely to be rejected by the immune system. Therefore, they could be used for therapeutic transplantation [57]. Further studies have shown that there are two types of HSCs: long- and short-term HSCs. Long-term HSCs proliferate during their entire lifetime. It has been demonstrated that in young adult mice, 8–10% of these HSCs enter the cell cycle and divide every day. Short-term HSCs proliferate only for a limited time. Long-term HSCs have a higher telomerase activity than their short-term counterparts [58]. Active telomerase is a feature of dividing, undifferentiated cells and is also found in cancer cells. In mice, only 1 in every 10,000–15,000 bone-marrow cells is considered to be a long-term HSC [58].

Mesenchymal Stem Cells The existence of non-hematopoietic stem cells was suggested by Cohnheim in 1867 [59]. He claimed that bone marrow may be the source of fibroblasts with collagen fibers that are part of the normal wound-healing process [60]. In 1974, Friedenstein and his colleagues performed the first isolation of MSCs from bone marrow. They reported that the adherent cells (the non-adherent cells were HSCs that were removed 4 h after the cells had been seeded on plastic culture dishes) had a heterogeneous appearance, but that most of them were spindle-shaped and formed foci of two to four cells, which remained inactive for 2–4 days and then began to grow rapidly. They also demonstrated that these cells were able to differentiate into colonies that had some similarities to bone and cartilage. Further studies extended Friedenstein’s observations and demonstrated that these cells were multipotent and able to differentiate into osteoblasts, chondrocytes, adipocytes, and even into myoblasts. They are currently referred to as mesenchymal stem cells or marrow stromal cells [61].

Sources of MSCs Mesenchymal stem cells have the potential to differentiate into chondrocytes, osteoblasts, adipocytes, fibroblasts, marrow stromal cells, and other tissues of mesenchymal origin. The MSCs have various origins and the ability to regenerate specific cell types for several tissues, for example adipose tissue, periosteum synovial membrane, muscle, dermis blood, bone marrow, and teeth. Bone marrow stroma is considered to be the source of a large amount of multipotent cells that have access to various tissues via the blood circulation. It has been established that MSCs from bone marrow stroma are capable of differentiating into adipocytes, osteoblasts, chondrocytes, and also into hematopoiesis supporting stromal cells. The differentiation of MSCs into adipocytes, osteoblasts, and chondrocytes can be significantly increased by the use of specific differentiation cocktails [13, 17] (Figure 1.2). Although BMSCs are an option for stem-cell therapies, their use is still subject to some limitations: first, a bone marrow harvest is a painful procedure; second, although MSCs grow well under standard tissue culture conditions, ex vivo expansion is necessary due to the relatively low numbers of MSCs that are present in the harvested marrow. Therefore, and since obesity is becoming increasingly widespread in industrialized countries, adipose tissue has become an attractive alternative source of stem cells for clinical and nonclinical applications [62]. Moreover, adipose tissue yields a much higher amount of MSCs than bone

 1  Adult Stem Cells

Adipocyte, astrocyte, neuron, cardiomycyte, chondrocyte, hepatocyte, muscle, osteoblast Different embryonic tissue lineage, adipocyte, osteocyte, endothelial cell, neuron, chondrocyte

Bone marrow

11

Adipocyte, chondrocyte, muscle, osteoblast

Synovial membrane

Muscle

Adipocyte, chondrocyte, muscle, osteoblast

Teeth Adipocyte, muscle, chondrocyte, osteoblast

Adipocyte, chondrocyte, osteoblast

Dermis

Sources of adult mesenchymal stem cells

Blood

Pericyte

Adipocyte, osteoblast, fibroblast

Chondrocyte

Trabecular bone Adipose tissue

Periosteum Chondrocyte, osteoblast

Adipocyte, chondrocyte, muscle, osteoblast, stromal cell

Figure 1.2  Sources of adult mesenchymal stem cells and their potential of differentiation into multilineage cell types.

marrow [63]. The two following characteristics of ADSCs have caught the attention of ­scientists over the past few years: their immune privilege properties that are due to their lack of human leukocyte antigen (HLA) DR (a class II major histocompatibility antigen – MHC II) expression, which enable their use in therapeutic applications [64], as well as their ­suppression of the proliferation of activated allogeneic lymphocytes [62, 65]. Other studies, however, have demonstrated that these cells also possess immunosuppressive capacities, as MSCs can lead to tumor growth and cell transformation [66, 67]. These contradictory results demonstrate that further studies and protocols are necessary in order to identify the effects of ADSCs on tumor formation [66, 68]. Seeliger et al. have demonstrated that hepatocyte-like cells that are derived from adipose tissue offer a promising alternative approach to the treatment of urgent metabolic liver dysfunctions or for in vitro use. The researchers claim that hepatocyte-like cells have outstanding advantages as they maintain important metabolic functions and many enzymatic activities during cryopreservation, which renders them constantly available. As MSCs have an immunosuppressive capacity, the data obtained by Seeliger et al. suggest that these cells can be applied autologously without any lifelong immunosuppressive therapy being necessary [24].

Identification of MSCs Mesenchymal stem cells are often identified by their morphology or by their phenotype, if they have a fibroblast-like morphology. They can also be identified by several other means, such as their differentiation potential into colony forming units (CFUs), which underlines their proliferative capacity. In addition, the ability to adhere to plastic is another characteristic marker for MSCs. Phenotypically, MSCs express various markers, but unfortunately none of them is really specific to MSCs. However, it is generally agreed upon that adult human MSCs

12

An Introduction to Stem Cells

Table 1.4  Expression of profile surface markers of mesenchymal stem cells (MSCs) [13, 132, 133] Marker type

Markers expressed in MSCs

Markers not expressed in MSCs

Specific antigen

SH2, SH3, SH4, Stro-1, ACTA1

CD133

Hematopoietic markers

CD4, CD14, CD34, CD45, CD117

Cytokines and growth factors

IL-1α, 6, 7, 8, 11, 12, 14, and 15 LIF, SCF, GM-CSF, G-CSF

Cytokines and growth factor receptors

IL1R, IL3R, IL4R, IL6R, IL7R, LIFR, SCFR, G-CSFR, IFNγR, TNFR1,TNFR2, bFGFR, PDGFR, EGFR

IL-2R (CD25)

Adhesion molecules

Integrinsα1 (CD49a), α2 (CD49b), α3 (CD49c), aα (CD49e), β1 (CD29) β3 (CD61), β4 (CD104)

α4 (CD49d), αL (CD11a) Cβ2 (CD18)

Extracellular matrix molecules and receptors

ICAM-1 (CD54), ICAM-2 (CD102) VCAM-1 CD 106, ALCAM-1 (CD166), LFA3 (CD58), L-Selectin (CD62L), Endoglin (CD105), Hylaronate (CD44), CK18, CK19

ICAM-3 (CD50), E-Selectin (CD62E), P-Selection (CD62P), PECAM-1 (CD31), VWF, Cadherin5

Others

CD9, CD13, Thy-1 (CD90), HLA-ABC (MHC I) (Low)

HLLA-DR (MHC II)

do not express hematopoietic markers, such as CD45, CD34, CD14 or CD11 [69, 70]. In Table 1.4, different surface markers of mesenchymal stem cells are presented.

Immunosuppressive Properties of MSCs In 2006, it was reported that MSCs possess immunomodulatory properties [71, 72]. Based on these findings, it was speculated that MSCs may play a significant role in the maintenance of peripheral tolerance, transplantation tolerance, tumor evasion, and fetal maternal tolerance [73]. Various studies on human, baboon, and murine MSCs have confirmed the immunosuppressive characteristics of MSCs and have illustrated that these cells are able to suppress the activation and in vitro proliferation of T lymphocytes [74, 75]. In line with these findings, multipotent mesenchymal stromal cells have been intensively studied in regenerative medicine. Besides their effects on T cells, their immunosuppressive effects are attributed to the secretion of a soluble factor by MSCs. Mesenchymal stem cells have paracrine effects, such as immunomodulation, which occurs through the secretion of soluble mediators, like nitric oxide, cytokines (e.g. interleukin-6), transforming growth factor-β, human leukocyte antigen G5, and prostaglandin E2 [76, 77]. Moreover, MSCs from the bone marrow are in close contact with T and B cells and are able to regulate the immunological memory by setting various survival niches for plasma cells and memory T cells of the bone marrow. In addition, it has been demonstrated that MSCs modulate the function of B cells by inhibiting their proliferation [73]. Furthermore, there is some evidence that adult MSCs suppress the differentiation and functions of dendritic cells (DC) [78]. Since the immunosuppressive characteristics of BMSCs have been reported in vitro and  in  vivo, clinical trials on allogeneic transplantation through the reduction of the

 1  Adult Stem Cells

13

graft-versus-host disease (GVHD) in the recipient have been supported [62]. One possible explanation is the lack of expression of HLA-DR (MHC II) in MSCs and the suppression of the proliferation of activated allogeneic lymphocytes as demonstrated in ADSCs. It has been demonstrated that ADSCs promote engraftment and prevent or treat severe GVHD in allogeneic stem-cell transplantation in vitro and in vivo [62]. Recently, it has been speculated that the use of MSCs in diseases, such as alcoholic liver fibrosis, may suppress the inflammatory response [79]. If this assumption is correct, the use of MSCs could become a very innovative approach for the treatment of progressing liver fibrosis, since, so far, all routine clinical applications for treating this disease have been ineffective. However, several studies have suggested that the immunosuppressive effects of ASCs may support the growth of tumor cells [80, 81].

Aging During the aging process, the ability of the human body to regenerate tissues and organs decreases significantly. Some scientists have even stated that humans are as old as their ­tissue-specific adult stem cells [82, 83]. Aging is related to a functional decrease in various tissues and organs, for example impairment of the immune system, skin fragility, cardiac dysfunction, bone degeneration, and an increased risk of cancer development. Cellular senescence is a complex phenotype that leads to changes in the function and the replication capacity of the affected cells. Different protocols, culture conditions, and cell types lead to different kinds of senescence. However, in vitro, senescent cells generally exhibit a characteristically enlarged, flattened morphology [84]. Senescence is characterized by an irreversible arrest of G1 growth. This arrest leads to the repression of genes that induce the progression of the cell cycle, to the upregulation of cell-cycle inhibitors [85], to an increased global methylation [86], and to a reduced multipotency of ASCs [84].

Aging of ASCs and Possible Consequences Adult stem cells could be the perfect source for patient-specific cell therapy of a wide range of regeneration defects, including age-related diseases, such as acute bone and cartilage defects, various liver diseases, macular degeneration, acute myocardial ischemia (AMI), stroke, and amyotrophic lateral sclerosis (ALS). There exist numerous clinical trials with MSCs that have already entered phase III [87, 88]. There is increasing evidence, however, that ASCs change their function with increasing age. For example, aging leads to a loss of the self-renewing capacity of stem cells or to their incomplete differentiation into a new cell type [89, 90]. Aged MSCs are reported to be larger than younger ones. These old MSCs exhibit more podia, spread further and possess more actin stress fibers, a decreased number of CFUs, and a lower replicative lifespan [84, 91]. Aged cells exhibit important changes in their DNA, such as telomere shortening. When telomeres reach a certain length, cells stop dividing and enter into senescence. When the telomere has reached its minimal length, the lifespan of primary human cells is limited to 50–70 cell divisions. Age-related telomere shortening has been identified in osteoblasts, chondrocytes, and myocytes. A similar pattern has been detected in older MSCs [84, 92]. These findings prove that adult stem cells undergo the same aging process as primary cells. Further evidence of this process is provided by the observation that, in vitro, MSCs lose their telomeric repeats at the same rate as primary cells (30–120 bp/PD) [92]. It has to be noted, however, that the telomeres of osteoblasts and chondrocytes are longer than the telomeres of MSCs [93, 94].

14

An Introduction to Stem Cells

Our own studies with AD-MSCs have demonstrated an age-dependent decline in cell ­ roliferation as well as a decreased expression of pluripotent associated genes (NANOG, p OCT, and LIN28A). These changes are related to an impaired osteogenic differentiation due to age [95]. In the same line of evidence, we have been able to show that the reduction of the global methylation in AD-MSCs improves hepatic differentiation. These findings clearly demonstrate that the epigenetic modification of ASCs improves the differentiation potential of MSCs. The most interesting finding was that 5-azacytidine (AZA), a well-known DNA methyltransferase inhibitor, and BIX01294 (BIX), a histone deacetylase inhibitor, are equally able to improve the differentiation capacity of AD-MSCs towards hepatocyte-like cells. The authors have even been able to demonstrate the partial rejuvenation of old AD-MSCs [24]. The epigenetic modification of the genome apparently plays a significant role as a regulatory pathway in the control of stem-cell aging. These changes are mainly identified in DNA methylation and chromatin remodeling [96]. Increased global DNA methylation has been observed in many aged adult tissue and cell types [86]. Recently, it has been reported that ten-eleven translocation (TET) proteins mediate the conversion of 5-methyl cytosine (5mC) to 5-hydroymethyl-cytosine (5hmC), which plays a crucial role in DNA demethylation in embryonic stem cells [97]. Our own group has been able to demonstrate for the first time that TET proteins also mediate the conversion of 5mC to 5hmC in AD-MSCs. In donors with old AD-MSCs, we have observed a lower level of expression of 5hmC, together with a higher level of expression of 5mC. This finding suggests a DNA hypermethylation pattern in aged AD-MSCs [98]. Furthermore, it has been shown that the growth and the osteogenic differentiation potential of old adult AD-MSCs have been improved by the pretreatment with small molecules AZA and BIX. A gene expression analysis of pluripotent markers (NANOG, OCT-4, lIN28A, SOX2) has revealed age-dependent changes in AD-MSCs which suggest that an increase in age leads to a reduction of DNA demethylation activity. Moreover, it has been demonstrated that TET2 and TET3 gene expressions are increased by treatment with AZA and BIX. This suggests that TET2 and TET3 might be involved in the conversion of 5mC to 5hmC in AD-MSCs [99]. However, the specific role of TET2 and TET3 in the process of aging will require further investigation. In summary, these data suggest that a reduction of the DNA demethylation activity in AD-MSCs correlates with an increased donor age. Furthermore, the treatment of AD-MSCs with small molecules (AZA, BIX) is able to restore the growth and the differentiation potential of these cells.

Stem Cells in Regenerative Medicine and in vitro Application Regenerative medicine is a multidisciplinary field of research. It includes the use of biomaterials, growth factors, and stem cells for repairing, replacing, or regenerating damaged tissues and organs. Due to their self-renewal and differentiation capacity, stem cells are ideal candidates for an application in regenerative medicine [62]. However, many questions about the criteria that should define the application of stem cells in regenerative medicine still have to be answered. The first and foremost problem that needs to be solved consists in finding a possibility to access the millions or even billions of ASCs that can be found in every human body. Second, it has to be analyzed whether the stem cells can be harvested in an in vitro environment

 1  Adult Stem Cells

15

without any risk of transformation. Third, it has to be investigated whether these cells can be differentiated along multiple cell lineage pathways in a controllable and reproducible manner, or whether it is even better to use these cells in an undifferentiated state. The question of whether ASCs can be safely and effectively transplanted to an autologous or an allogeneic host also has to be answered. Finally, it has to be examined whether ASCs can be safely ­produced on a large scale according to Good Manufacturing Practice guidelines (GMP). Since James Thomson performed the first isolation of human ESCs, their potential of unlimited self-renewal and differentiation has led to numerous attempts to use them in drug discovery, disease modeling, and regenerative medicine [100]. The purpose of these attempts consists in differentiating human ESCs into hepatocytes [101, 102], cardiomycytes [103], neurons [104], and intestinal tissue [105]. Several pluripotent and multipotent stem cellbased therapeutics have entered clinical trials. Table 1.5 presents a few examples of therapeutics that have already been approved for clinical application. The most well-known example of the use of stem cell therapy is the transplantation of HSCs. This procedure has been practiced successfully for decades in order to treat serious hematological diseases. In the framework of this procedure, HSCs are injected directly into the blood of the recipient. This injection is called “bone marrow transplantation.” The stem cells recognize their pathway through “homing,” in which chemokines play a crucial role [106, 107]. Different types of studies have indicated the potential of mesenchymal and embryonic stem cells for undergoing similar homing into injured tissue [108–110]. However, there is still a debate as to whether stem cells should be systemically applied or brought to the site of the damage. Systemic application of MSCs in humans has shown beneficial effects in osteogenesis imperfecta and graft-versus-host disease [10, 111]. It is widely accepted that cell therapy alone is not sufficient for the regeneration of large tissue defects. Therefore, a combination of tissue engineering and the differentiated cells from MSCs seems to be required to trigger the healing of damaged tissue [112]. In fact, tissue engineering has already been used for the replacement of some tissues, for example skin, bone, or fat [112, 113] . Stem cell-based therapy, however, brings about several safety challenges that cannot be addressed by using standard analytical procedures. A particular difficulty is the ability to monitor biological cell distribution, since it may not be possible to distinguish the injected cells from the host cells. The ability to track the therapeutic cells enables the assessment of the risk of the formation of inappropriate ectopic tissue and of tumorigenesis. Moreover, the detection of misplaced or transformed cells may necessitate the development of methods for their removal. However, such a removal of cells is not technically possible at the moment. Furthermore, the delivery of a cell with an unlimited renewal potential and a capacity to differentiate into any human cell type brings about a huge safety concern that is not raised by any other type of treatment. The finding that undifferentiated stem cells that are injected into immunocompromised animals are capable of forming teratomas, emphasizes the importance of addressing this safety issue in the future, and underlines the caution that is necessary in the development of therapies based on the use of stem cells [114]. If cells contain genetic abnormalities, they could potentially develop into teratocarcinoma [115, 116], that is tumors composed of elements of teratomas together with persisting undifferentiated cells that are highly malignant [114]. Another safety issue that should be tackled is the immunogenicity of stem cells [117]. Although there are many reports confirming the immune privilege of human ESCs, any foreign cell that is introduced into a patient will be subjected to immune surveillance [118]. This evidence indicates that our understanding of stem cells is probably not yet good enough to completely evaluate the safety of these therapies in a comprehensive manner. Therefore, further research on this topic is necessary.

Stable ischemic stroke

Type 1 diabetes mellitus

Stargard’s disease, retinal pigment epithelial (RPE) derived from human ESCs Crohn’s Disease

Liver disease

Not completed Phase 1

Phase 2

Phases 1, 2

Completed

Phase 1/2 Suspended

Pilot investigation of stem cells in stroke patients

Human adult stem cells for the treatment of recently diagnosed type 1 diabetes mellitus (T1DM)

Advanced cell technology receives FDA clearance for the first clinical trial using embryonic stem cells (ESCs) to treat macular degeneration

Extended evaluation of PROCHYMAL [tm] adult human stem cells for treatment-resistant moderateto-severe Crohn’s Disease

A Phase 1/2 safety and tolerability dose-escalation study of autologous stem cells with patients suffering liver insufficiency

Spinal cord injury

Not completed Phase 1

Safety study of GRNOPC1

Condition

Study phase

Trials

15

20–65, M + F, chronic liver insufficiency, abnormal serum albumin and/or bilirubin and/or prothrombin time, no reception of a liver transplant, WHO performance score of less than 2, life expectancy of at least 3 months, written consent has to be given

Participation in Osiris Protocol 603, with a drop of at least 100 points in CDAI at day 28

Not published yet

12

98

12 + 35 yr, M + F, T1DM, at least 1 DM-related autoantibody, some beta cell function

 G0/G1 [22]. During apoptosis and necrosis, phosphatidylserine is translocated from the inner to the outer cell membrane. Annexin V, a calcium-dependent protein, can be used as a sensitive probe for phosphatidylserine detection. However, annexin V should be combined with another exclusion dye, such as PI, to enable better discrimination between types of cell death [23]. The presence of a nick from DNA fragmentation during apoptosis or necrosis can also be labeled by enzymes that include terminal transferase in the terminal deoxynucleotidyl transferase dUTP (deoxyuridine triphosphate) nick-end labeling (TUNEL) method, or DNA polymerase in the in situ nick-translation (ISNT) method. Enzymatic DNA labeling by the TUNEL assay on rat blastocytes has shown that cadmium selenide (CdSe) quantum dot (QD) nanoparticles induce apoptosis and inhibit cell proliferation [24]. Cysteine-aspartic proteases (caspases) are implemented as indicators of apoptosis processes. Activation of a caspase triggers a proteolytic cascade by activation of other caspases, which intensifies apoptotic signaling and consequently cell death [25]. Lao et al. have shown that fullerene nanoparticles can selectively enter oxidation-damaged endothelial cells rather than normal ones, preventing the apoptosis process by regulation of signaling events such as caspase 3 activation [26]. The viability or proliferation rate of cells exposed to nanomaterials can be relatively determined and compared with a control by the use of several reagents. In these assays, reduction of a dye as well as alteration in its absorption wavelength correlates with the ­metabolic activity of live cells. The same protocol as depicted in Figure 10.1 is performed for all  agents. Formazan dyes such as 3-[4, 5-dimethyl thiazol-2-y1]-2, 5-diphenyl tetrazolium  bromide (MTT) and 2,3-bis-(2-methoxy-4-nitro-5-sulfophenyl)-2H-tetrazolium-5-­ carboxanilide (XTT), as the most popular chromogenic products, have been mentioned in ISO 10993-5. Table  10.1 summarizes important reagents, their conversion by viable cells, and the detection procedure. Of note, optical properties of the materials may be altered at the nanoscale level with interference by cytotoxic dye reagents. For example, false readings of absorption spectra have been obtained for the MTT and neutral red (NR) assays of carbon nanosized materials [14]. A review of in vitro toxicity assessments on iron oxide nanoparticles has been published by Mahmoudi et al. [27]. Cell reproductive capacity can be examined using the colony formation assay within 1–3 weeks. This assay determines the ability of each cell to divide and establish a colony (at least 50 cells). Seeded cells that have been exposed to nanomaterials may lose their capacity to produce colonies. At the end of the test, colonies can be fixed and stained with reagents such as crystal violet for better observation [28]. Figure 10.2 summarizes four categories of cytotoxicity tests.

Assay reagent addition

Test compound addition

Cell seeding

Equilibration

Exposure

0–24 hours

1 hour–7days

Data recording

Assessment 10min–24hours

Figure 10.1  Time-scale representation for toxicity assessment. Table 10.1  Colorimetric assays for semi-quantitative analysis of cytotoxicity Reagent

Process by live cells

Dissolved in

Wavelength (nm)

MTT

Conversion of yellow water soluble tetrazolium salt to blue formazan

DMSO or isopropanol

570

XTT

Conversion of yellowish water soluble tetrazolium salt to orange formazan

Water or PBS

450–500

Crystal violet

Uptake of violet dye in less than 10 min by live cells

Acetic acid

570

Neutral red

Uptake of red dye and conversion in lysosome

Acetic acid-ethanol

540

Alamar blue

Conversion of nonfluorescence blue dye to red fluorescence

Water or PBS

Excitation: 530 Emission: 590

DMSO, dimethyl sulfoxide; MTT, 3-[4, 5-dimethyl thiazol-2-y1]-2, 5-diphenyl tetrazolium bromide; PBS, phosphate buffer solution; XTT, 2,3-bis-(2-methoxy-4-nitro-5-sulfophenyl)-2H-tetrazolium-5-carboxanilide.

Fluorescence imaging Confocal microscopy

LDH FDA/PI Crystal violet Trypan blue Hoechst AO/PI

Membrance integrity

Cell morphology

In vitro cytotoxicity assays

Optical microscopy Electron microscopy

ISNT assay Comet assay TUNNEL assay Cell cycle Genetic (Flowcytometry) indices DNA synthesis (Thymidine, brdU) Gene expression (c-Myc, p53, Bcl-2, caspase-3, ...)

ROS determination Organelle function Lysosome (Neutral red) Mitochondria (MTT, XTT, MTS, Alamar-blue) Intracellular esterases (Calcein AM)

Figure 10.2  In vitro cell toxicity/viability assays in four categories: genetic indices, membrane integrity, organelle function and cell morphology.

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Genotoxicity of Nanomaterials Several genes are recognized as modulator genes for cell growth or death, and include c-Myc, p53 and the Bcl-2 family [29]. Apoptosis induction of silver nanoparticles has been reported as a mitochondria-dependent mechanism, associated by generation of reactive oxygen species (ROS) and c-Jun N-terminal kinase (JNK) activation [30]. Translocation of JNK into mitochondria as a result of external stress leads to ROS generation [31, 32]. The p53 pathway as a mediator for cellular-stress responses is shown to be involved in DNA repair, senescence, and apoptosis [33]. Nanoparticles can be developed to target p53 for cell-cycle arrest regulation and growth suppression in tumor cells [34, 35]. Senzer and ­co-workers have transported p53 by using a liposomal nanodelivery complex to restore the normal tumor suppressor gene [36]. Apoptosis induction on cell cultures via the p53 pathway by zinc oxide [37], titanium oxide [38], nickel ferrite [39], silver [40], and tansferrin and selenium [41] nanoparticles has been reported. B-cell lymphoma 2, encoded by the Bcl-2 gene, and the Ras family of proteins play essential roles in cell growth and survival [42, 43]. Activation of JNK, c-Jun, p53, caspase-3 and NF-kappaB, and suppression of Bcl-2 protein have been observed for human umbilical vein endothelial cells (HUVECs) during exposure to silica nanoparticles [44]. Nanoparticles can be modified for efficient delivery of drugs or small interfering RNA (siRNA) into solid tumors to suppress c-Myc expression and activation of cell apoptosis [45, 46].

Immunotoxicity of Nanomaterials Preliminary in vitro assays for immunotoxicity of nanomaterials include red blood-cell destruction (hemolysis), platelet aggregation (thrombogenicity), complement activation, and attraction of macrophages (chemotaxis). Immunity stimulation of nanostructures is determined by evaluation of antigenicity, adjuvant properties, and inflammatory responses [47]. Stimulation of T-cell CD8/CD4 by polystyrene nanoparticles [48], inflammatory potential of TiO2 nanoparticles [49], and low degree of inflammation of carbon nanotubes in the mouse lung [50] have been reported. From another point of view, polymeric nanoparticles may be utilized for targeted delivery of anti-inflammatory drugs [51, 52]. In addition, phagocytosis and nitrogen oxide production by macrophages exposed to the nanostructures may be useful for studying interactions between the immune system and materials [53]. Nitric oxide (NO) levels can be determined as a criterion for the activation of macrophages against certain pathogens. Surface modifications with polyethylene glycol (PEGylation) or CD 47 are proposed routes for prevention of opsonization by macrophages [54, 55]. Phagocytosis is the process of foreign particle engulfment by cells. Digestion of ingested materials by lysosomes is facilitated by the formation of superoxide anion, hydrogen peroxide, and ROS. Therefore, increased ROS levels are correlated by cell activation that results from p ­ athogen destruction. It has been shown that TiO2 nanoparticles induce rapid ROS production in brain-resident macrophages (microglia) [56]. Cobalt and magnesium in the structure of engineered nanoparticles intensify the formation of oxidative stress [57]. Due to the large amount of variables in synthesis of nanomaterials and response of living systems, ­high-throughput screening is required. In one study, ROS production, mitochondrial depolarization, and plasma membrane permeability were analyzed, revealing lethal toxicity of ZnO in comparison with Pt, Ag, SiO2, Al2O3, and Au particles, which had sublethal toxicities [58].

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In some cases, such as diamond nanoparticles [59], there is no significant detectable ROS production, hence additional research is necessary to finalize all aspects of nonimmunotoxicity. For example, granulocyte/macrophage colony stimulating factors (GM-CSF), interleukin 4 (IL-4), and interferon gamma appear to be important cytokines for stimulation of macrophages and oxidative activity [60]. The concentration of ROS can be determined by using indicators such as 2′,7′-difluorescein diacetate (DFDA) for cerium oxide nanoparticles [61] and dichlorodihydrofluorescein diacetate (DCHFDA) for iron oxide nanoparticles [62],  which fluoresce when oxidized by ROS [63]. Respiratory immunotoxicity has been observed in rats administered zinc oxide nanoparticles by intratracheal instillation [64]. A  size-­dependent activation of the inflammatory response was reported for 15-nm-sized ­silver particles by the detection of TNF-α, MIP-2, and IL-1β [65]. The expression of inflammatory related genes such as IL-1, IL-6, TNF-α, iNOS, and COX-2 increased in ­cultured peritoneal macrophages that were induced by silica nanoparticles [66].

Carcinogenicity and Reproductive Toxicity Long-term studies are required in order to determine carcinogenic potential of nanomaterials, particularly those without known significant toxicities. Toxicity of single-walled carbon nanotubes (SWCNTs) and multiwalled carbon nanotubes (MWCNTs) have been studied extensively. However, due to the diverse physical properties of carbon-based materials and the possibility for proper functionalization with chemical groups, different biological outcomes have been reported. Injection of SWCNTs into the bloodstreams of mice revealed no evidence of toxicity over 4 months [67]. In another study, MWCNTs were present in the subpleural wall and subpleural macrophages after 6 h of inhalation [68]. DNA damage has been shown in embryonic stem cells exposed to MWCNTs [69]. In another experiment, the Comet (single-cell gel electrophoresis) assay was applied to human lymphocytes in order to determine the presence of any genotoxic effects of colloidal C60 fullerenes [70]. The genotoxicity assay on V79 cells exposed to small diameter SWCNTs (96 mg cm−2) after 3 h of incubation showed significant DNA damage [71]. The same results (genotoxicity) were observed for WIL2-NS cells exposed to 65 mg mL−1 TiO2 nanoparticles for 24 h [72]. Carbon black nanoparticles (14 nm diameter) at 100 mg mL−1 showed a significant increase in single-strand DNA breaks, but not double strand, after 3 h exposure to A549 cells [73]. These nanoparticles, at 75 mg mL−1, with the same scale (295 m2 g−1 specific surface area) weakly induced the frequency of mutation in FE1 MutaTMmouse lung epithelial cells. The results demonstrated a significant increase in DNA strand breaks and oxidized purines by the alkaline Comet assay, both with and without formamidopyrimidine glycosylase (FPG) [74]. Additionally, diverse nanoparticles from vehicle exhaust were assessed by the use of a specific tube in the breathing zone during bicycling. These nanoparticles influenced mononuclear blood cells and induced oxidative DNA base damage with no evidence of DNA strand breaks [75]. In another experiment a water-soluble semiconductor such as CdSe capped with a shell of zinc sulfide QDs was incubated with double strands of plasmid DNA. After the precipitation procedure and gel electrophoresis, exposure to the QDs led to DNA damage. DNA isolation from QDs at time 0, which was intended for the plasmid damage assay, demonstrated instantaneous modification when the QDs and targeted DNA were mixed [76]. The SWCNTs were prepared from CO and Fe(CO)5 as the iron-containing catalyst precursor. Specific analyses by NMAN 5040 (Manual of Analytical Methods, US National Institute for Occupational Safety and Health) and inductively coupled plasma atomic emission spectroscopy (ICP-AES) have shown that SWCNTs are comprised of 99.7% (wt) elemental

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Table 10.2  Important genotoxicity tests with diverse nanomaterials Material

Test procedure

Results

References

Particles from air

Alkaline Comet assay plus FPG-mononuclear blood cells

The sample of nanoparticles was positive in the Comet Assay with FPG in the air (high benzene levels)

[96]

Urban air particles

Alkaline Comet assay plus FPG-mononuclear blood cells DNA

DNA damage: positive

[97]

Carbon black

HPRT mutation assay-type II alveolar lung cells

HPRT mutation: positive

[98]

Zinc oxide

Chromosome aberration assay – CHO cells

Clastogen: positive

[81]

Magnetite nanoparticles

Micronuclei test-mouse polychromatic erythrocytes

Increase in micronuclei: positive

[99]

Silica-overcoated magnetic nanoparticles

Chromosome aberration assay – Chinese hamster lung fibroblasts

Increase in chromosome aberrations: negative

[100]

Single-walled carbon nanotubes

Ames Salmonella assay

Increase in mutation frequency: negative

[71]

Multiwalled carbon nanotubes

In vitro cytokinesis-block Micronucleus assay – MCF-7 cells

Increase in micronuclei Increase in centromeres

[101]

SiO2 nanoparticles

HPRT mutation assay

Induction of HPRT mutants

[79]

Gold nanoparticles

Plasmid DNA mixed with gold nanoparticles plus irradiationagarose gel electrophoresis test

Gold nanoparticles increased the single/double strand breaks of DNA by irradiation

[102]

CHO, Chinese hamster ovary; FPG, formamidopyrimidine glycosylase; HPRT, hypoxanthine guanine phosphoribosyltransferase.

carbon and 0.23% (wt) iron. Numerous techniques such as near-infrared and Raman spectroscopy confirmed that the majority of carbons in this structure have carbon nanotube morphology. Aortic mitochondrial DNA damage was detected after SWCNTs exposure, as demonstrated by quantitative polymerase chain reaction (qPCR) [77]. Consequently, the most frequently utilized test for DNA damage assessment is the Comet assay. In one study, 120 mg mL−1 of 99% pure SiO2 nanoparticles were exposed to WIL2-NS human B-cell lymphoblastoid cells for a 24-h period. The hypoxanthine guanine phosphoribosyltransferase (HPRT) mutation assay showed a significant increase in mutation frequency [78]. Table 10.2 lists important genotoxicity tests with diverse nanomaterials. The micronucleus assay is the second most regularly used genotoxicity test on nanobiomaterials after the Comet assay. The micronucleus assay has been used to analyze human B-cell lymphoblastoid cells exposed to SiO2 nanoparticles after vortexing and sonication. The assay with cytochalasin B indicated a significant dose-dependent increase in micronucleated binuclear cells with increasing particle doses [79]. Moreover, this assay was used to analyze CoCr nanoparticles that had been applied to human fibroblasts. Conformation of these particles was similar to the material that composed orthopedic joint-replacement prostheses. In this study, following a 12-h exposure, significant intensity in micronuclei was detected [80]. Chromosome aberrations in Chinese hamster ovary cells have been reported in 105 mg mL−1 of uncoated zinc oxide nanoparticles that were cultured in McCoy’s 5A medium [81]. Evaluation

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Table 10.3  Important recommendations for genotoxicity testing of nanomaterials Know what nanomaterial has been tested and in what form Recognize that nanomaterials are not all the same Consider uptake and distribution of the nanomaterial Take nanomaterials specific properties into account Use standardized methods Use in vivo studies to correlate in vitro results Learn about the mechanism of nanomaterials genotoxic effects Reprinted with permission from reference [69]. Copyright 2009 Elsevier.

of DNA damage biomarkers as a specific agent for preliminary screening of genotoxicity is another methodology for nanoparticulated materials. For example, p53 phosphorylation at serine 15 was increased in the A549 cell lines (adenocarcinoma human alveolar basal epithelial cells) when the cells were exposed to small-size carbon black nanoparticles (14 nm, printex 90), whereas no comparable results were detected for a larger size (260 nm) [73]. Table 10.3 lists recommendations for genotoxicity testing of nanomaterials. However, it should be stated that the same nanomaterial can produce varying outcomes with different genotoxicity assays. Reproductive toxicity in a fetus or offspring is considered as an important aspect of material biocompatibility. For example, injection of silica and TiO2 nanoparticles into pregnant mice have been shown to result in smaller sized fetuses. Transportation of nanoscale materials through the placental barrier leads to neurotoxicity in descendants [82].

Blood Compatibility of Nanomaterials Alteration in blood components such as plasma residual molecules and blood cells determines the extent of hemocompatibility of a nanomaterial. Assessments of thrombosis, coagulation, complement activation, or blood-cell malfunctions should be considered for interaction between nanomaterials and blood elements. Charge, hydrophobicity, and area of  the contact surface, as well as exposure duration and thermodynamic conditions, all influence the material–blood interaction. Nanostructures can be engineered as a carrier for both coagulant and anticoagulant factors. A quantitative measurement of hemoglobin is a simple way to find the extent of red blood-cell lysis. Ilinskaya and Dobrovolskaia have discussed interactions between nanoparticles and the coagulation system from the cellular, biochemical, and hydrodynamic points of view [83]. Different aspects can be considered for the role of nanoparticle physicochemical properties in interactions with blood components. For example, a higher degree of hemolysis and inflammation have been observed for smaller size polystyrene nanoparticles [84]. Minimal interaction with blood cells was noted for PEGylated negatively charged siRNA-loaded dextran nanogels [85]. Platelet adsorption was reduced by incorporation of Cu nanoparticles into hydroxyethyl methacrylate [86]. Dendrimers are synthesized as highly branched polymers with a nanosize spherical structure and numerous functional groups. They can be used independently or as a coating for other nanostructures in applications such as drug delivery for cancer therapy [87]. Duncan and Izzo have discussed biocompatibility and blood compatibility related to dendrimer chemistry in a comprehensive review [88]. Red blood cells, hematocrit, and hemoglobin are significantly higher for cationic and positively charged dendrimers (NH2 terminated) in comparison with anionic ones [89].

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Engineered nanostructures with the potential for immobilizing biomolecules (enzymes, antibodies, drugs, nucleic acids) or capturing harmful bioproducts such as viruses or venoms have been designed [90–92]. Hoshino et al. have stated that repulsive forces between negatively charged particles and cell-membrane phosphate groups minimizes toxicity [93]. Absorption of biomolecules may change the fate of the nanomaterial or affect the activation of a mechanism. Nanoparticle protein binding occurs almost instantaneously once the particle enters the biological medium and the physical properties of such a particle–protein complex often differ from those of the formulated particle [94]. These new properties can contribute to different biological responses and change nanoparticle biodistribution. Physical properties of nanosized materials, such as size, charge, and hydrophobicity, influence protein adsorption and protein corona formation, which suppress the intended performance [95].

Conclusions Among the useful and beneficial applications of nanoscience, nanomedicine has the greatest potential for improving quality of life. In order to reduce adverse health effects, the safety of nanobiomaterials should be verified by different screening tests. The complexity of both structure and function in biological systems as well as diversity of biomolecular interactions with nanostructures influences the predictability of the outcomes. Therefore, future research is necessary by using high-throughput screening (HTS) methods to provide a foundation for clinical trials. Noninvasive diagnostic methods that include imaging or laboratory t­ echniques must be developed to identify the fate of the nanobiomaterial in the body.

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[54] Moon JJ, B Huang and DJ Irvine (2012). Engineering nano- and microparticles to tune ­immunity. Advanced Materials 24: 3724–3746. [55] Yoo J-W, E Chambers and S Mitragotri (2010). Factors that control the circulation time of nanoparticles in blood: challenges, solutions and future prospects. Current Pharmaceutical Design 16: 2298–2307. [56] Long TC, N Saleh, RD Tilton, GV Lowry and B Veronesi (2006). Titanium dioxide (P25) produces reactive oxygen species in immortalized brain microglia (BV2): implications for ­ nanoparticle neurotoxicity. Environmental Science and Technology 40: 4346–4352. [57] Limbach LK, P Wick, P Manser, RN Grass, A Bruinink and WJ Stark (2007). Exposure of engineered nanoparticles to human lung epithelial cells: influence of chemical composition and catalytic activity on oxidative stress. Environmental Science and Technology 41: 4158–4163. [58] George S, T Xia, R Rallo, Y Zhao, Z Ji, S Lin, X Wang, H Zhang, B France, D Schoenfeld, et al. (2011). Use of a high-throughput screening approach coupled with in vivo zebrafish embryo screening to develop hazard ranking for engineered nanomaterials. ACS Nano 5: 1805–1817. [59] Schrand AM, H Huang, C Carlson, JJ Schlager, E Omacr Sawa, SM Hussain and L Dai (2006). Are diamond nanoparticles cytotoxic? The Journal of Physical Chemistry B 111: 2–7. [60] Gordon S (2003). Alternative activation of macrophages. Nature Reviews Immunology 3: 23–35. [61] Park E-J, J Choi, Y-K Park and K Park (2008). Oxidative stress induced by cerium oxide nanoparticles in cultured BEAS-2B cells. Toxicology 245: 90–100. [62] Naqvi S, M Samim, M Abdin, FJ Ahmed, A Maitra, C Prashant and AK Dinda (2010). Concentration-dependent toxicity of iron oxide nanoparticles mediated by increased oxidative stress. International Journal of Nanomedicine 5: 983–989. [63] Jones CF and DW Grainger (2009). In vitro assessments of nanomaterial toxicity. Advanced Drug Delivery Reviews 61: 438–456. [64] Liu H, D Yang, H Yang, H Zhang, W Zhang, Y Fang, Z Lin, L Tian, B Lin and J Yan (2013). Comparative study of respiratory tract immune toxicity induced by three sterilisation nanoparticles: Silver, zinc oxide and titanium dioxide. Journal of Hazardous Materials 15: 248–249, 478–486. [65] Carlson C, SM Hussain, AM Schrand, L K. Braydich-Stolle, KL Hess, RL Jones and JJ Schlager (2008). Unique cellular interaction of silver nanoparticles: size-dependent generation of reactive oxygen species. The Journal of Physical Chemistry B 112: 13608–13619. [66] Park E-J and K Park (2009). Oxidative stress and pro-inflammatory responses induced by silica nanoparticles in vivo and in vitro. Toxicology Letters 184: 18–25. [67] Schipper ML, N Nakayama-Ratchford, CR Davis, NWS Kam, P Chu, Z Liu, X Sun, H Dai and SS Gambhir (2008). A pilot toxicology study of single-walled carbon nanotubes in a small sample of mice. Nature Nanotechnology 3: 216–221. [68] Ryman-Rasmussen JP, MF Cesta, AR Brody, JK Shipley-Phillips, JI Everitt, EW Tewksbury, OR Moss, BA Wong, DE Dodd and ME Andersen (2009). Inhaled carbon nanotubes reach the subpleural tissue in mice. Nature nanotechnology 4: 747–751. [69] Landsiedel R, MD Kapp, M Schulz, K Wiench and F Oesch (2009). Genotoxicity investigations on nanomaterials: methods, preparation and characterization of test material, potential artifacts and limitations – many questions, some answers. Mutation Research/Reviews in Mutation Research 681: 241–258. [70] Dhawan A, JS Taurozzi, AK Pandey, W Shan, SM Miller, SA Hashsham and VV Tarabara (2006). Stable colloidal dispersions of C60 fullerenes in water: evidence for genotoxicity. Environmental Science and Technology 40: 7394–7401. [71] Kisin ER, AR Murray, MJ Keane, X-C Shi, D Schwegler-Berry, O Gorelik, S Arepalli, V Castranova, WE Wallace and VE Kagan (2007). Single-walled carbon nanotubes: geno-and cytotoxic effects in lung fibroblast V79 cells. Journal of Toxicology and Environmental Health, Part A 70: 2071–2079. [72] Wang JJ, BJ Sanderson and H Wang (2007). Cyto- and genotoxicity of ultrafine TiO2 particles in cultured human lymphoblastoid cells. Mutation Research/Genetic Toxicology and Environmental Mutagenesis 628: 99–106.

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Part 4

Control of Stem-Cell Fate by Engineering of Microenvironment

Chapter 11

Stem-Cell Responses to Surface Nanotopographies Peng-Yuan Wang1,2 and Wei-Bor Tsai3

Industrial Research Institute Swinburne (IRIS), and Department of Chemistry and Biotechnology, Faculty of Science, Engineering and Technology, Hawthorn, Victoria, Australia 2  CSIRO Manufacturing Flagship, Clayton, Victoria, Australia 3  Department of Chemical Engineering, National Taiwan University, Taipei, Taiwan 1 

Introduction Stem cells, undifferentiated cells with the ability to self-renew for producing more stem cells and with the potential for differentiation into specialized cells in response to appropriate signals [1], have attracted great attention in recent years due to their promise in regenerative medicine. Stem cells can be classified into three categories, based on the sources from which they are obtained and their differentiation ability: embryonic stem cells (ESCs), adult stem cells, and induced pluripotent stem cells (iPSCs) [2]. Among ESCs, the mesenchymal stem cell (MSCs), which can be harvested from most tissues with different characteristics (sizes and shapes) and populations [3], is the most studied. Since both controlled self-renewal and directional differentiation are requisites for the application of stem cells in regenerative medicine, knowledge regarding the maintenance of stem cells during cell expansion and directional differentiation of stem cells into a specific lineage is of great importance to clinical applications of stem-cell-based therapies. Stem cells, when removed from the in vivo stem-cell niche, often differentiate spontaneously in an inefficient and uncontrolled way during in vitro culture [1]. It is often thought that the stemcell niche, a microenvironment where stem cells are found, interacts with stem cells to ­regulate cell fate. Scadden described stem-cell niches as “specific anatomic locations that regulate how the stem cells participate in tissue generation, maintenance and repair” (abstract, pp. 1075–1079) in a review article in Nature [4]. The stem-cell niche contains both anatomic and functional cues and integrates the signals that mediate the balanced response of stem cells to the needs of organisms. Through an understanding of controlling the microenvironment surrounding stem cells, an important strategy of regenerative medicine is to restore the niche environment in order to control the fate of stem cells. The microenvironment surrounding stem cells includes biochemical cues (e.g., cytokines, cell–cell contacts, and cell–extracellular matrix interactions), topographic cues and mechanical induction [5]. The interplay of these signals regulates and directs the fate of stem cells. Although biochemical signals have drawn much attention in this research area, topology of the niche environment has been demonstrated to be an important mediator for regulating stem cells.

Stem-Cell Nanoengineering, First Edition. Edited by Hossein Baharvand and Nasser Aghdami. © 2015 John Wiley & Sons, Inc. Published 2015 by John Wiley & Sons, Inc.

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Control of Stem-Cell Fate by Engineering of Microenvironment

Mammal cells are organized into cooperative assemblies such as tissues and organs. Most of the cells are held together via cell–cell contacts or cell–extracellular matrix (ECM) via specific receptor-mediated interactions. The basic macromolecular components of the ECM include collagenous molecules, glycoproteins, elastins, hyaluronic acid, proteoglycans, and glycoaminoglycans [6]. The ECM also serves as a reservoir for harboring growth factors and cytokines. The ECM provides mechanical support for cellular anchorage, directs cell ­orientation, morphology and migration, and establishes the tissue microenvironment for controlling cell proliferation and differentiation. The ECM is enriched in nanotopology. For example, collagen, a major ECM protein in connective tissues such as bone, skin, tendon, and ligament, is assembled into 50–300-nm diameter fibrils, and then are further organized into larger cable-like collagen fibers around 1–4 μm in diameter [7]. Therefore, the base structure of the ECM is composed of nano- to microscale collagen fibers. Another example

(A)

Human anterior basement membrane

(B)

500 nm

(C)

500 nm

(D)

Porous silicon (p type)

500 nm

Human descemmet’s membrane

Electrospun gelatin fibrous mesh

25 µm

Figure 11.1  Scanning electron microscopy images: (A and B) the basement membrane of the human cornea, (C) artificial porous silicon substrate, and (D) artificial electrospun gelatin micro-fiber. Images A and B were extracted from Last et al. [8] by permission from Elsevier: Journal of Structural Biology.

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is the texture of the basement membrane (or basal lamina), which is a flexible, thin matlike structure in which epithelial cell sheets adhere. Inspection by scanning electron microscopy reveals that the mesh-like macaque anterior corneal basement membrane consists of interwound 24–183-nm diameter fibers, with nanopores ranging from 22 to 216 nm, occupying approximately 15% of the total surface area (Figure 11.1A and 1B) [8]. The complex surface topography governs not only the behavior of corneal epithelial cells, but also serves as a stem-cell niche for limbal epithelial stem cells and corneal stromal stem cells [9]. Topographic substratum mimicking the ECM feature is a powerful tool to assist researchers in understanding the physiological cell–substrate interaction. To this end, t­echnologies for fabrication of nanotopography are required. A summary of the fabrication techniques can be found in the literature [10–12]. For example, semiconductor industries have made a great effort in the development of techniques for the formation and manufacturing of micro- and nanoscale patterns on a silicon wafer, which provides cell biologists great tools to fabricate substrata with a defined surface nanotopography. Therefore, cell biologists can systematically explore the effects of the substrata topography on cell morphology and cell behavior, such as adhesion, spreading, attachment, growth, differentiation, and migration. The purpose of this chapter is to introduce the responses of stem cells to physical nanotopography, such as protrusion, pores, grooves/ridges, and fibers, comprising topographic features smaller than 1000 nm. The current theories regarding the topographic induction of stem-cell differentiation are also briefly covered. Although chemical patterns are also important for directing stem-cell functions [13], this topic is not discussed in this chapter.

Responses of Stem Cells to Nanoprotrusion Surfaces Cells inevitably encounter nanotopographies in vivo or in vitro. The natural ECM in many tissues contains topographical nanoprotrusions [14, 15]. On the other hand, artificial biomaterial surfaces usually contain at least nanoroughness since processing techniques for biomedical devices are rarely capable of producing a molecularly flat surface. A polymeric material could obtain surface nanoroughness easily by treatment with caustic (or hydrolytic) reagents [16, 17]. Surface roughness can be controlled by the strength of the erosive reagents and the time of incubation. Polymeric nanoprotrusion surfaces can be prepared by polymerdemixing. For example, mixtures of polystyrene and poly(4-bromostyrene) spontaneously undergo phase separation during spin casting, resulting in a nanotopographic surface [18]. By controlling the concentrations and the ratios of polystyrene and poly(4-bromostyrene), a variety of topographical forms, including pits, islands, or ribbons with varying height, depth, or distribution, can be fabricated. A nanopillared surface is also a popular model to investigate cell interactions with nanoprotrusions, since the technology for the fabrication of silicon nanopillared surfaces is well developed for optoelectronics application [19, 20]. Nanopillared surfaces can be fabricated by e-beam lithography, but low throughput is its drawback [21]. Alternatively, using a colloidal gold etch mask combined with reactive ion etching, silicon nanopillars can be produced [22]. A common cell response to a nanoprotrusion surface is reduced cell adhesion [23]. The inhibition in the adhesion of human skeletal stem cells is shown to increase with increasing nanopillar height, > 70 nm in height, on which the formation of focal adhesions is perturbed [24]. Similarly, Berry et al. showed that polymer demixed nanotopographical features reduced adhesion in bone-marrow-derived osteoprogenitor populations [25]. However, several cell types when cultured on surfaces with a lower protrusion ( 50 nm (−)

Human BMSCs

TiO2

30–300 nm

30 nm (+)

Rat BMSCs

Silicon

20–30 nm

0

89

Rat BMSCs

Silicon

Few nm to 1 µm

0

48

Silicon Rat BMSCs/ human BMSCs/ human ASCs

Few nm to 2 µm

(−)

49

Rat BMSCs

Silicon

Few nm to 230 nm

(−)

(−)

Osteogenic/ adipogenic (+)

46

Human BMSCs

Alumina

50 nm to 3 µm

(−)

0

Osteogenic (+)

35

Osteogenic (−)

42

(+)

Osteogenic (+)

45

< 30 nm (+); > 50 nm (−)

< 30 nm (+); > 50 nm (−)

44

Osteogenic: 30 nm (+)

29

40

BMSCs, bone-marrow mesenchymal stem cells; ASCs, adipocyte-derived mesenchymal stem cells; PMMA, polymethyl methacrylate; (+), positive effect; (−), negative effect; 0, no significant effect.

Several recent reviews have summarized the effects of engineered nanopits on the fates of mammalian cells [37, 38]. Dalby and his colleagues have carried out a series of studies on stem-cell responses to nanopits [39–42]. They used electron-beam lithography to precisely create defined nanopits (120 nm diameter, 100 nm deep) of different symmetry and with varying degrees of disorder on polymethylmethacrylate substrates. They found that osteogenesis of human MSCs is stimulated by disordered nanopits in the absence of chemical induction, with similar efficiency to the osteogenesis of cells cultured with osteogenic media [39]. They proposed that topographical modification could modulate c­ ellular differentiation in multipotent stem-cell populations [42] through the ERK/MAPK (extracellular signal-­ regulated kinase/mitogen activated protein kinase) signalling pathway via integrin-mediated cellular adhesion [43]. They showed that nanopit arrays disrupt c­ellular adhesion and spreading, which prevents osteogenesis indicated by the downregulation of several osteospecific genes (Ets and Stat1) in skeletal stem cells. The results imply that the formation of focal adhesions and the cytoskeleton has an impact on directing cell differentiation. The responses of stem cells to nanoporous topography vary with cell type/source, ­topographic feature, and material types. It is difficult to infer a general conclusion from the studies regarding nanoporous surfaces. Some results are contradictory to each other. For example, Park et al. showed that the adhesion, proliferation, and osteogenesis of rat MSCs on nanoporous TiO2 were reduced on a surface with pore size larger than 50 nm compared with a plane TiO2 surface or a surface with smaller pore size (~15 nm) [44], while some studies showed that larger pore sizes (79 and 150 nm) stimulate osteogenesis [29, 45]. The diversity in cell type/source, culture condition, material chemistry, and topographic feature may be the reason for the difficulty in making a conclusion on nanoporous effects. It should be noted that most of the porous studies focus only on the size of pores and porosity. However, the architecture

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Control of Stem-Cell Fate by Engineering of Microenvironment

Pore size gradient

1 µm

1 µm

1 µm

1 µm

300 nm

150 nm

1

1 µm

2

1 3

4

µm

100 µm

1 µm

1 µm

300 nm

2

1 3

4

µm

100 µm

2

3

4

µm

100 µm

Figure 11.2  Cell responses on pore-size gradients. Pore-size gradients were created on N-type Si wafers, so called n-type porous silicon gradients (n-pSi). The n-pSi with average surface pore sizes ranging from 5 to 230 nm and ridge roughness from 3 to 22 nm were fabricated using an electrochemical etching approach in a HF-based solution. These new surfaces allow stem cells to grow at least 1–2 months. Cell morphology of rat bone marrow-derived MSCs (rBMSCs) was affected on the n-pSi dependent on the pore size and nanoroughness between pores. Cells display a round shape at the high nanoroughness end but a spread-out shape at the low nanoroughness end of the gradients. The subsequent osteo- and adipogenic differentiations of rBMSCs were improved by the surface topographies dependent on the positions and local cell density along the gradients [46]. (See insert for color representation of the figure.)

of the spacing between pores may be different and affect cell responses. Due to the difference in the fabrication processes, some porous surfaces contain a flat interpore area, while some contain a rough area. Wang et al. showed that a porous Si surface with 200-nm pores with rough ridges (Ra ~10 nm) stimulated osteogenesis of rat MSCs [46]. Therefore, it might lead to a wrong conclusion on porous substrates without consideration of the topography of the interpore area. We believe that not only the averaged pore size, but also pore-to-pore distance, solid surface fraction, solid surface roughness, and overall ­surface roughness need to be considered for evaluating the effect of porous substrates on stem-cell behaviour [47]. In order to systematically investigate the porous effects on cell responses, a substrate with pore-size gradients is especially required for high-throughput screening in order to reduce the time and the material required for sample preparation [48]. More importantly, ­topographic gradients reduce the difference between discrete samples and individual experiments, and the cellular activity on different topographies can be compared at the same culture condition. By adjusting the parameters of the chemical or electrochemical etching process, silicon (Si) or

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aluminium porous substrates with a pore-size gradient can be readily obtained within a range of few nanometers to few microns [35, 49]. However, controlling the topographic features across wide-ranged feature sizes and the stability of the Si gradient formats during culture remain challenges to overcome. Recently, Wang et al. fabricated relatively stable porous Si gradients using n type Si wafer (n-pSi) [46]. This new surface allow stem cells to grow at least 1–2 months. The results showed that cell adhesion is extremely sensitive to the ­nanoroughness of the interpore space. Cell adhesion and spreading was inhibited at high nanoroughness end (small pore size), whilst cells were well attached at low nanoroughness end (large pore size, Figure 11.2). Differentiation of rBMSCs into osteogenic and adipogenic lineages showed that osteogenesis was dependent on both topography and local cell density whilst adipogenesis was dependent on topography only [46].

Responses of Stem Cells to Nanogrooved Surfaces Surfaces with nanogrooves (or nanograting, nanostripes) are frequently used to study cell– topography interactions. The methods for the fabrication of nanogrooved topographies include e-beam lithography, nanoimprint lithography, mechanical polishing, grinding, and laser ablation, which have been summarized in the literature [50, 51]. The general feature of cell responses to such a topographic feature is cellular alignment and elongation along the groove direction (Figure 11.3), a phenomenon called contact guidance. Cells have anisotropic morphology on the grooved surface, whereas cells are spread out isotropically on the flat control (Figure 11.3). (A)

900 nm width/100 nm depth

(B)

rBMSCs on nanogrooves (900/100)

(C)

Rat cardiomyocte on nanogrooves (450/350)

400 nm

(D)

2

4

6

8

µm

Flat control

25 µm

25 µm

2 µm

Figure 11.3  Typical cell morphology on a grooved surface. (A) A typical atomic force microscopy image of a nanogrooved surface. The feature size is 450 nm in width and 100 nm in depth (ridge-to-groove ratio 1:1). (B) Rat bone-marrow-derived mesenchymal stem cells (rBMSCs) and (C) rat cardiomyocyte are aligned with the direction of the nanogrooves. (D) The rBMSCs are spread out isotropically on the flat control. In (C) the cells on the grooved polyurethane surface could pull and deform the substrate, but still align with the direction of the grooves. In (B) vinculin staining shows that focal adhesions are mainly located at the two ends of the cells on the nanogrooves, whereas in (D) they are located around the cell periphery on the flat surface. (See insert for color representation of the figure.)

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Control of Stem-Cell Fate by Engineering of Microenvironment

Wang et al. showed that rat cardiomyocytes were highly aligned on nanogrooved polystyrene and polyurethane surfaces [52]. However, the cells had a better beating behavior on the polyurethane substrate, probably due to its softer compliance. Fujita et al. described the dynamic process of the adhesion of human MSCs on a nanogrooved polycarbonate surfaces (870-nm-wide ridge/670-nm-wide groove and 200 nm in depth) [53]. They found that cells extend filopodia that continuously probe the surface patterns during cell adhesion and spreading. Cell protrusions extending perpendicularly to the grooves tend to retract more rapidly than those extending in parallel with the grooves. The overall outcome is that cells extend and align along the grooves. Addition of actin disrupting agents such as cytochalasin D and latrunculin B leads to a loss of cytoskeleton organization, including α-SMA (smooth muscle actin), vimentin and α-tubulin, resulting in a rounded cell morphology on nanogrooved polydimethylsiloxane (PDMS) (600-nm-wide ridges/grooves of 600 nm depth) [54]. The degree of cell alignment on grooved topography depends on the dimension of the grooves, such as width and depth [55, 56]. Due to the ability to guide cell alignment, grooved architecture is of particular interest for use in tissues with anisotropic geometry, such as skeletal muscle [57], cardiac muscle [52], tendon [58], ligament [59], and nerve [60]. Although several reviews discuss the effect of nanogratings on mammalian cells [37, 50], few report a focus on stem cells. In this section, we mainly discuss the effects of nanogrooved surfaces with defined feature sizes (10–1000 nm) on stem cells, and some of the results are summarized in Table 11.2. Regarding the influence of nanogrooved topography on the induction of the differentiation of stem cells, cell lineages related to bone, adipose, muscle, and neurons are particularly attractive. Yim et al. showed that the induction of human MSCs toward neuronal lineage on PDMS substrates was enhanced on a nanograting surface (350 nm in width and 350 nm in depth) compared with plane and micrograting (1 μm/10 μm in width and 350 nm in depth) surfaces [61]. They showed that such a nanotopographic feature possesses better neurogenic induction ability than some chemical reagents such as retinoic acid. They found that the formation of focal adhesions is downregulated on the nanograting surfaces [62]. The expresTable 11.2  Stem-cell response on nanogrooved substrates

Cell type

Substrate material (surface coating)

Feature size

Attachment/ adhesion

Proliferation Differentiation

Human BMSCs PDMS (COL 1) 350 nm to 1 µm Human BMSCs PC (Fn)

670/870 nm

Human ESC line H9

PUA (gel/O2 plas)

350 nm

Human MSCs

PDMS (COL 1) 350 nm

Reference

Neurogenic (+)

61

Neurogenic (+)

64

Migration

53

(−)

62

FA; cytoskeleton

63

Human BMSCs PDMS/TCPS (COL 1)

350 nm/500 nm

Human ESCs

PE (O2 plas)

60–380 nm + µm

Rat BMSCs

PS (O2 plas)

450, 900 nm

0

0

Osteogenic (−); 91 adipogenic (+); myogenic (+)

200 nm to 2 µm

(−)

0

Osteogenic (+)

Human BMSCs PU

Cardiogenic (+) 90

65

BMSCs, bone-marrow mesenchymal stem cells; ESCs, embryonic stem cells; FA, focal adhesion; PC, polycarbonate; PDMS, polydimethylsiloxane; PE, polyethylene; PS, polystyrene; PUA, polyurethane acrylate; TCPS, tissue culture polystyrene; COL 1, type I collagen; Fn, fibronectin; O2 plas, oxygen plasma; gel, gelatin (+), positive effect; (−), negative effect; 0, no significant effect.

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sion of zyxin protein, a marker for mature focal adhesions, was decreased on 350-nm-grating PDMS, leading to smaller and more dynamic focal adhesions. Furthermore, the expression of integrin subunits α2, α6, αV, β2, β3, and β4 were also decreased on the nanograting surface compared with flat controls [63]. These results suggest that the traction force at focal adhesions on the nanogratings is decreased. Lee et al. also demonstrated direct differentiation of human ESCs into selective neurons on a nanogrooved poly(urethane acrylate) surface (350-nm-wide ridges/grooves of 500 nm depth) without any specific induction reagent [64]. The human ESCs on the nanogrooved surface expressed increased NeuroD1 (a neuronal marker) and decreased GATA6 (endoderm marker) and DCN (mesoderm markers) compared with controls. Tuj1 (immature neuronal marker), HuC/D (human neuronal protein) and MAP2 (mature neuronal marker) with well-structured neurite extension was immunostained positively, while the brachyury (mesoderm markers), Pdx1 (endoderm marker) and GFAP (intermediate filament proteins of mature astrocytes) was not found. The results indicate that on the nanogrooved surface human ESCs differentiate toward neuronal lineages, but not a glial lineage such as astrocytes. Watari et al. showed that nanogrooved surfaces (400 nm pitch of 300 nm depth) enhanced osteogenesis of human MSCs with respect to the expression of RUNX2 (runt-related transcription factor 2) and osteocalcin, and calcium deposition relative to cells cultured on 1400 or 4000 nm pitch or planar control [65]. Several biosignals, including Wnt and bone morphogenetic proteins (BMPs) signalling pathways, were identified as pivotal regulators in osteogenic differentiation. Watari et al. demonstrated that nanogroove cues enhance osteogenesis of human MSCs, likely via the BMP signalling pathway, since ID1 (a target of the BMP pathway) was enhanced on the 400-nm-pitch nanogrooved surfaces in the presence of osteogenic induction components, while TCF3 and AXIN2 (downstream genes of the Wnt pathway) were not significantly affected [65].

Responses of Stem Cells to Nanofibrous Structures Cell anisotropic spreading along fibrous substrates (contact guidance) was reported as early as in 1952 [68]. Recently, the investigation of cell responses to nanofibrous structure has been increased due to a wide application of the electrospinning technique. The structure of electrospun fibers and its fibrous membrane mimic the native collagen fibrils and native ECM topography (Figure 11.1D). The diameters of nanofibers can be modulated ranging from hundreds of nanometers to a few microns by controlling the electrospinning conditions, such as polymer type and concentration, solvent type, flow rate, and applied voltage [69]. Some researchers think that electrospun fibrous meshes display a three-dimensional topographic architecture since cells might migrate in the space between fibers. However, electrospun fibrous meshes are usually too compact for cells to penetrate. Furthermore, aligned nanofibers can be obtained using special collectors such as a rotating collector [70]. Aligned nanofibrous mesh is especially of interest in applications to study the differentiation of MSCs into differentiated lineages, such as neuron [69, 71], tendon [72, 73], ligament [74], and cardiac muscle/muscle (Table 11.3) [75]. In this section, the effect of nanofibrous mesh on stem cell responses is discussed. Aligned electrospun fiber is reported to promote the neurodifferentiation of stem cells. Lim et al. reported that aligned poly(caprolactone) (PCL) nanofibrous meshes (mean ­diameter 480 nm) enhanced the differentiation of rat neural stem cells in the presence of retinoic acid compared with random fibrous meshes or flat films [69]. Similarly, Wang et al. showed that neuronal differentiation (β-III-tubulin) and neurite outgrowth of human ESCsderived neural precursors were promoted on aligned silk fibroin nanofibrous meshes (mean fiber diameter 400 nm) compared with the random nanofibrous meshes and flat controls [76].

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Table 11.3  Stem-cell response on nanofibrous substrates Cell type Human BMSCs

Substrate material

Feature size

Attachment/ adhesion Proliferation Differentiation

Hydroxybutyl Median chitosan 400–600 nm (−)

Myogenic (+)

77

Neurogenic (+)

69

Tendogenic (+); Osteogenic (−)

72

Osteogenic (+)

92

Tendogenic; osteogenic; myogenic

73

Rat neural stem PCL (LMN) cells

260, 480, 930 nm

Human tendon stem cells

PLLA

median 430–450 nm

Human USSC

PCL/PLLA/ nHA

Did not mention (~few µm)

C3H10T1/2 cells

PS

655 nm

Rabbit BMSCs

Silk fibroin

800, 1200 nm

(+)

Ligament

74

Human BMSCs

PCL

270 nm

(−)

Neurogenic (+)

71

Human ESCs-derived NP

Tussah silk fibroin

400, 800 nm

400 nm (+)

Neurogenic (+)

76

Human CSSC

PEUU

165 nm

Keratocytes (+)

78

Rat BMSCs

PHBHHx

3, 5 µm

Adiopogenic (random: +); osteogenic (aligned: +)

70

2.8 µm

Cardiomyocyte (+) 75

PU Murine ESC-derived cardiomyocytes

(−)

Reference

(+)

BMSCs, bone-marrow mesenchymal stem cells; CSSC, corneal stromal stem cells; ESCs, embryonic stem cells; LMN, Laminin, nHA, nano-hydroxyapatite; NP, neural precursors; PCL, poly(caprolactone); PEUU, poly(etherurethane urea); PHBHHx, poly(3-hydroxybutyrate-co-3-hydroxyhexanoate); PLLA, poly(l-lactide acid); PS, polystyrene; PU, polyurethane; USSC, human cord blood-derived unrestricted somatic stem cells; (+), positive effect; (−), negative effect; 0, no significant effect.

Wang et al. reported that osteogenic differentiation of rat MSCs cultured on aligned poly(3-hydroxybutyrate-co-3-hydroxyhexanoate) fibrous meshes (mean fiber diameter 5.7 μm) were promoted, while the adipogenic differentiation was impaired [70]. The gene and protein analysis showed that PPAR-γ (peroxisome proliferator-activated receptor gamma) signalling pathway was reduced on the aligned fibres, which might contribute to the impaired adipogenesis and enhanced osteogenesis. The downregulation of PPAR-γ on the aligned fibres was related to phosphorylated activation of ERK without an increase in ERK expression. This study indicated that aligned fibres impaired adipogenesis but enhanced osteogenesis through the MAPK-dependent PPAR-γ signalling pathway. Dang et al. investigated the responses of human MSCs on aligned hydroxybutyl chitosan (HBC) electrospun nanofibrous meshes (median fiber diameter 436 nm) [77]. The cells and nuclei aligned and elongated along the fibers, and the cells formed an aligned cell sheet on the aligned fibrous mesh. Furthermore, when compared to genes expressed by human MSCs cultured on HBC films and TCPS, some of myogenic genes (collagen IV, desmin, Pax-3, Pax-7, myogenin) were apparently upregulated without chemical induction. Similarly,

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Yin  et  al. showed that human tendon stem cells cultured on aligned poly(l-lactide acid) (PLLA) nanofibrous mesh expressed significantly higher levels of integrin α1, α5 and β1 ­subunits, and myosin II B and the expression of tendon-specific genes (Eya 2, SCX, and COL 14) were significantly higher in cell culturing on aligned nanofibers compared with the cells cultured on randomly oriented nanofibers in both normal and osteogenic media [72]. Another study showed that aligned silk fibroin nanofibrous meshes enhanced the expression of ligament-related proteins in rabbit MSCs compared with random nanofibrous meshes [74]. Wu et al. reported that human corneal stromal stem cells cultured on aligned poly(ester urethane urea) nanofibrous meshes (mean fiber diameter 165 nm) expressed unique gene markers to keratocyte (KERA, ALDH, PTGDS, CHST6 and PDK4) together with uniform collagen fibrils and ordered matrix organization [78].

The Mechanism Governing Nanotopographical Control of Stem Cell Fate As we described in previous sections, stem cells can be differentiated into several specific ­lineages (such as osteoblasts [65] and neurons [61]) on nanotopographic surfaces without addition of specific biochemical signals. However, understanding of the fundamentals of nanotopographic influence on lineage commitment is still inadequate. One hypothesis is that cell responses are regulated via the formation of focal adhesions on nanotopographic surfaces. Cell attachment to the ECM is critical in regulating cell morphology, migratory behavior, growth, apoptosis, and differentiation. A large family of transmembrane receptors, ­integrins, is responsible for cell attachment to the ECM [79]. Integrins consist of a pair of heterodimeric α and β units with transmembrane-spanning domains and they bind ECM ligands such as fibronectin, laminin, and collagen. Integrins serve as the bridge connecting the extracellular matrix to intracellular cytoskeleton, which is a filament-like structure in the cytoplasm, composed of actin, myosin, actinin, and tropomyosin. The integrin–ECM contact points form specialized sites of adhesion called focal adhesions [80], which consist of aggregated integrins and other associated proteins, bridging the extracellular ligands and cytoplasmic stress fibers, organized contractile bundles of actin filaments. The other end of stress fibers can bind to a second focal adhesion or a meshwork of intermediate filaments that surrounds the nucleus. Stress fibers form and disappear in response to tension across a cell from focal adhesions. The ligand-bound integrins also activate many intracellular signaling pathways, such as the ERK/MAPK, signaling a cascade that is critical for cell survival and differentiation. The details regarding the integrin-mediated signaling pathway have been summarized in several reviews [81, 82]. A previous report suggested that nanotopographyinduced human MSC differentiation is modulated by the integrin-activated focal adhesion kinase [83]. The findings indicate that human MSCs sense and transduce nanotopographical signals through focal adhesions and actomyosin cytoskeleton contractility to induce differential gene expression. F-actin formation on nanogrooved surfaces appears on the top of grooves [84], indicating that the focal adhesions are formed along the groove. Furthermore, several studies show that cell nuclei become elongated along the direction of nanogrooves [84, 85]. These clues reflect the connectivity of cytoskeletal and nuclear structures. Maniotis et al. reported that living cells and nuclei are hard-wired such that a mechanical force on cell-surface integrins can instantly modulate the organization of molecular assemblies in the cytoplasm and nucleus [86]. Intermediate filaments are responsible for effectively mediating mechanical force transfer to

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Anisotropic cytoskeletal tension

Isotropic cytoskeletal tension

Cell-generated force Cadherin/integrin Cell/ECM/substrate

Figure 11.4  Schematic illustration of the concept of force isotropy applied to cells. This figure is extracted from Nava et al. [87] by permission from Hindawi Publishing Corporation: Journal of Biomedicine and Biotechnology.

the nucleus from the focal adhesions. The stretched nuclei on nano-grooved surfaces can be explained by the concept of “force isotropy” proposed by Nava et al. (Figure 11.4) [87]. The cytoskeletal tensional state depends on the balance between the intracellular actomyosin contractility and the reaction forces exerted by the underlying substrate. If cell adhesion-mediated traction forces, exerted by the cell on the underlying substrate, differ in magnitudes at different spatial orientations, then the cell nucleus tends to elongate (anisotropic cytoskeletal tension in Figure 11.4). On the contrary, the cell nucleus tends to remain a round shape if the traction forces remain similar at different orientation (isotropic cytoskeletal tension in Figure 11.4). The fact of nucleus deformation on nanotopographic surfaces provides a clue for s­ tem-cells– nanotopology interactions. The intranuclear environment contains structurally interconnected nuclear matrix elements that are responsible for modulation of gene ­expression [88]. Dang and Leong proposed that the stretching of nuclei in the cells on the nanotopographic surfaces may alter the internal matrix structure sufficiently enough to change the expression profile of genes [77]. However, direct evidence for the reprogamming of the gene expression profile in the cells cultured on nanotopographic substrates is still under investigation.

Conclusions More and more evidence indicates that surface nanotexture exerts significant impacts on directing stem-cell differentiation. However, this area remains less explored. We believe that with more knowledge regarding stem-cell–nanotopology interactions, researchers can design better strategies combining biochemical and topographic cues for tissue engineering and regenerative medicine.

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Chapter 12

Control of Mesenchymal Stem-Cell Fate by Engineering the Nanoenvironment Habib Nikukar1,2, Stuart Reid3, Mathis O. Riehle1, Adam S.G. Curtis1, and Matthew J. Dalby1

Centre for Cell Engineering, Institute for Molecular, Cell and Systems Biology, College of Medical, Veterinary and Life Sciences, University of Glasgow, Glasgow, UK 2  Shahid Sadoughi University of Medical Sciences and Health Services, Yazd, Iran 3  SUPA, Thin Film Centre, University of the West of Scotland, Paisley, UK 1 

Introduction The complexity of the human body and coordinated harmony of various parts of living organisms are reflections of more difficult integration at the cellular and molecular level. Thanks to modern microscopy and molecular approaches, biologists are finding understanding inside the smallest parts of living organs, the cells. Growth, development, and repair are continuously occurring to preserve health to the end of life. Medicine, as a science to conserve human health and treat diseases, is always looking for new methods to support us as we grow older. Through progress in the field of the very small (microand nanosciences), the possibility of finding new techniques for prevention and treatment of diseases may perhaps be found. The hope for use of cells directly for cell-based therapy, stem-cell therapy, gene therapy, and understanding cell control and signaling mechanisms will help us realize regenerative therapies. Use of autologous stem cells to make new tissues and organs to replace or treat those that have become damaged or diseased is a major tissue-engineering aim. In this chapter, the effects of nanoscale topography and mechanical stimulations on cells, with particular focus on human mesenchymal stem cells, to understand the potential usefulness of these nanoscale tools will be explored.

Bionanotechnology and Regenerative Medicine Development and availability of modern technology has facilitated discovery within the very small world at the micro- and nanoscale [1]. Desire to find new methods to maintain human health and treat the life-threatening diseases in medical sciences linked to nanotechnology, is defining “bionanotechnology.” The term “nanomedicine” sometimes is applied to the usage of high-performance nanotechnology for the improvement of medical methods and devices. The basics of these sciences focus on nanoscale, that is, (1–100) × 10−9 m materials such as

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synthetic nanotubes, nanoparticles, molecular engines, and also the study of cellular nanostructures such as pores, envelopes, chromosomes (DNA), etc. Health and lifestyle improvements have caused a progressive increase in old age groups around the world [2]. As a result, the body and its organs can suffer disease or damage and these global changes cause pressure to discover new methods for replacement of organs, joints, etc., to support the aging population. It is further clear that the numbers of allograft transplants are not sufficient and the adverse immune reactions with transplanted organs leading to the need for immunosuppressive drugs coupled to limitations in donors make regeneration of organs and/or engineered replacements very attractive [3]. Manipulation of cellular fates is one of the most important aspects of the biologists’ interests in developing new materials for tissue engineering. Use of these methods in cell therapy and replacement of body tissues or organs using the multipotent stem cells taken from the same patient is a particular focus. This scope of such innovation is constantly expanding as more is discovered about the usefulness of these adult stem cells. Engineering the tissues and making new organs are major drivers in regenerative medicine [4–6]. Making bone tissue [7, 8], joint [9–11], tendon [11], heart [12–14], bladder [15], pancreatic island cells, liver and hepatobilliary system [16, 17], kidney [18, 19], eyelayers [20], brain, and neural tissues [21], and many other tissues, are under research and at various stages of success/ clinical usage [22].

Mesenchymal Stem Cells Stem cells are capable of making one or several tissues and organs according to body needs, based on complex messages at different developmental and regenerative stages. The other characteristic aspect of stem cells is their ability to replicate, making a cell similar to themselves (self-renewal) and another daughter cell that will also be a stem cell (symmetrical self-renewal) or a specialized, progenitor cell (asymmetrical self-renewal). As well as pluripotent embryonic stem cells, there are multipotent adult stem cells throughout the body, residing in niches in many tissues and organs. They are supported by special microenvironments (lacuna or niches) surrounded by blood vessels, neurons, supportive cells, and soft tissue. This environment works to control quiescence, a way of preserving stem-cell numbers without DNA damage and self-renewal/differentiation based on tissue demand [23, 24]. It is hoped that their potential can be tapped to aid with transplantation, organ donation and rejection, malfunctioning organs and limbs, genetic diseases, cancer treatment, threatening infectious diseases, and many other medical issues. Mesenchymal stem cells (MSCs) are a type of adult stem cell, residing since early neonatal (cellular) phase, in special niches close to the red bone marrow and surrounded by neural, vascular, and bone tissue. The laboratory characteristics of MSCs are: (i) adherence to tissue plastic culture dishes under standard culture conditions; (ii) cell surface characterization (e.g., positive for CD73, CD90, and CD105, and negative for CD11b or CD14, CD34, CD45, CD79α or CD19, HLA-DR); and (iii) in vitro mesodermal differentiation [25, 26]. They are typically considered capable of differentiation to fibroblasts, osteoblasts, chondrocytes, and adipocytes [27, 28]. It has been shown that there is a strong correlation between changes in focal adhesion (FA) complex (larger, mature, highest in number) and MSC differentiation toward, for example osteoblasts and adipocytes, with MSCs with very few, small adhesions forming fat and MSCs with supermature (>5 μm in length) adhesions forming bone [29–33], as will be discussed later. This information on adhesions linked to differentiation potential of the MSCs could be used for tissue engineering, especially for bone, cartilage, and tendon tissue

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regeneration [34, 35]. Based on this finding, some types of simulation such as topographical, mechanical, chemical, and extra cellular matrix (ECM) stiffness changes have been introduced to show how the responses of MSCs could be controlled.

Cell Structure Throughout the cell cytoplasm is the cytoskeleton. The cytoskeletal architecture of cells consists of various protein dimers or polymers aiding in structure and shape, division, motility, and perhaps environmental sensing of the surrounding area and transferal of messages from outside the cell to the nucleus. The cytoskeleton is traditionally categorized as having three main fiber types: microtubules, microfilaments, and intermediate filaments. An interconnecting network of thin cytoskeletal fibers, the microtubules, helps support the cytoplasmic membrane. They are composed of tubulin and form a polarized meshwork to control protein and organelle passage throughout the cell cytoplasm. Actin microfilaments, composed of actin subunits, are important in cell movements by making cell protrusions and developing contractile forces through stress fiber formation and myosin interaction [36–38]. It seems that the cortical network of actin microfilaments in contact with membrane proteins is an important factor in cell motility based on cell adhesion and sensing of the surrounding area. Other actin microfilaments, again emanating from FA, make an internal network and make a connection with the nuclear envelope through linkers of nucleoskeleton and cytoskeleton complexes (LINC) [39]. Intermediate filaments are generally considered a more rigid type of cytoskeletal fiber compared to microfilaments and are involved in maintenance of cell shape and cell structural support. The filaments also integrate with the nucleus through LINC complexes [40, 41]. The nucleus, the largest and stiffest intracellular organelle, contains the genomic DNA and governs cell activities by transcription of genes and transfer of the resultant mRNA to the ribosomes for protein translation. The outer layer of the nucleus, the nuclear membrane, is a continuation of the endoplasmic reticulum that encloses all the chromosomes and has many highly specialized pores to control the entrance to the nucleus. Nuclear lamins, as the nucleoskeleton, make a network inside the nuclear membrane to protect the chromosomal assembly and preserve nuclear shape [42]. Lamins are type V intermediate filaments and are divided into two types: lamin A from the LMNA gene (lamin C is derived from alternative splicing of this gene) and lamin B from the LMNB1 and LMNB2 genes. The most common type of lamin is lamin A and cells without lamin A could survive but its genetic defect is the cause of laminopathies, a group of human diseases usually known by muscular dystrophy and cardiomyopathy. Lamin B gene knockout could be lethal for animal model embryos. It could be indicative for lethality of knockdown in human embryos as well. Lamin B is present in all eukaryotic cells but Lamin A/C involvement in the lamina is limited to differentiated cells [43–45].

Cells and Mechanical Forces The responses of the body after birth to various types of mechanical forces and stimuli are an important factor for adaptation and growth. The human body is exposed to various types of daily stimuli such as mechanical, electrical, shear stress, thermal, chemical, and acoustic waves. Cells could respond individually or in groups to these stimuli [46, 47]. Some adaptive responses are necessary for cell maintenance and some others could perhaps be detrimental. Back disorders due to work with vibrating equipment [48] and hypertrophied

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vasculature and heart muscle in response to hypertension [49] are potential detrimental mechanoresponsive effects. However, here the focus will be on positive mechanotransduction to allow control of cell fate in a desired manner [50]. After the discovery of stem cells, understanding the best way for manipulation, stimulation, or conduction of them has been a major scientific focus [51, 52]. Research on the effect of chemical, matrix stiffness, topography, electrical, magnetic, and mechanical stimuli on stem-cell behavior have been considered and some of these approaches will be described with a particular focus on topographical and mechanical stimulation [52–56]. It is noted that many different types of mechanical stimuli, in combination or individually, have been tried previously. These include stretching forces [57], compression [58], shear stress [59], and vibration [60, 61], with results demonstrating that type of stimulus, time-scale, and amplitude are all important [62–64].

Mechanotransduction For the purposes of this chapter, mechanical stimulation will be referred to as mechanotransduction, although it is acknowledged that mechanotransduction can be initiated by  a wide variety of stimuli that differentially regulate intracellular tension and cell morphology, which will be discussed in parallel (e.g., material surfaces). The molecular response of cells to any mechanical stimulus usually is in the form of altered protein production and changing phenotype. This occurs through a process called mechanotransduction, and this is a type of adaptation that is necessary for cellular life [65]. Briefly, the process starts by sensing changes in the external environment, transfer of this mechanical message by cytoskeleton to the nucleus evokes a chromosomal response to the stimulus, which will affect transcript to protein synthesis: this may be in the form of indirect (biochemical) or direct (mechanical) stimulation [47, 66]. Various mechanisms are involved in the mechanotransductive process. These responses start at the cellular level and are usually driven by cell–matrix interactions, adhesions, as a start point – or at least adhesions are a mechanism with particular interest in the field [67]. Indirect (biochemical) and direct (distortion of the nucleus) mechanotransductive events are covered later in this chapter. Both forms, at the nuclear level, will lead to different genes being transcribed in response to the changing mechanical environment, resulting in differential protein expression typical of different phenotypes, for example, osteocalcin and osteopontin in bone-cell differentiation [64, 66].

Focal Adhesion Complexes Without adhesion, tissue cells die through apoptotic mechanisms known as anoikiseven quicker than through lack of nutrients [68]. Attachment of cells to the surfaces is dependent to several factors: cell-surface proteins, such as integrins, cadherins, selectins, and immunoglobulins, to ECM characteristics, such as type and concentration of matrix proteins, to matrix stiffness and physical characteristics of the matrix [69]. Integrins are considered the most important cell-surface proteins in cell/interfacial research as they form the base of FA complexes (Figure 12.1) [70]. They are a heterodimer consisting of an α and a β subunit, each with three domains: an extracellular domain in contact with ECM, a transmembrane, and a cytoplasmic domain. Currently, about 18 varieties of α subunits and eight of β subunits and twenty-four separate combinations are known. It seems integrins are major transmitters of biomechanical signals from the ECM to the cell interior. The process

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F-actin

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α-actinin

Vinculin

FAK

Talin ə

β

Cell membrane

Integrins ECM

Biomaterial surface

Figure 12.1  A simplified overview of the focal adhesion. Focal adhesions are macromolecular structures that serve as mechanical linkages of the cell cytoskeleton (F-actin) to the extra cellular matrix (ECM), and as biochemical signaling hubs involved with the transmission of external mechanical forces through numerous signaling proteins that interact at sites of integrin binding and clustering. Reproduced from [141] with permission from Elsevier.

of activation starts with pairing of β and α integrins. By the connection of integrins to the ECM proteins, several signaling pathways inside the cell will be activated, and more than 100 proteins share this complex for transmission of signal to the nucleus [71–73]. The protein composition of FAs will change according to the size of the adhesion. In the smallest FAs, known as focal complexes, at 1 μm or less in length, talin, vinculin, paxillin, and integrins can be found and they are the point of initial contact between the ECM and cell-surface receptors. In mature focal adhesions (up to 5 μm in length), paxillin, vinculin, focal adhesion kinase (FAK), zyxin, and integrins are present. At the largest size, fibrillar, or supermature FA (>5 μm in length) greater amounts of these connective proteins and also tensin are found. All three types of adhesion are force sensitive. Forces from inside or outside of the cells are triggers for integrin activation and FA formation. Traditionally, studies focus on biochemical signals arising from changes in cytoskeletal assembly, especially actin cytoskeleton, as a result in changes in adhesion. Both FA- and FAK-directed activation of the G-proteins aid “cell sensing” of the surface. For example, direction of lamellipodium (from Rac activation) helps cell/surface exploration. Various factors determine cell movements, such as: chemical attractant and mechanical forces such as gravity, liquid flow, physical shape, ECM stiffness, and protein gradient. These are more attractive to integrin bonding, and formation of advancing lamellipodium will guide cells to these sites. Other activations arise from cell division-control protein 42 homolog (CDC42) activation, causing filopodial formation. These fine membrane protrusions are integrin-containing and “sweep” in front of the cells to locate binding sites for FA growth. The G-protein Rho is also involved in motility and sensing through its role in myosin activation and resultant actin contraction. RhoA inhibition shows significant decreases in cell motility [74]. In MSCs, it has been demonstrated that on nanotopography there is a shift from stem cell to differentiated osteoblast. This occurs through changes in adhesion and intracellular tension. Osteoblasts are large cells requiring larger adhesions to support the tensile cytoskeletal scaffolding [75]. Such effects have also been demonstrated in MSCs cultured on hydrogels [76] or forced to confine to morphologies using microcontact printing of fibronectin [77, 78].

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Looking to gel stiffness, it was seen with very soft gels that neural marker expression was promoted as the cells dissipated adhesive force into the compliant substrates, while on stiffer gels (similar stiffness to precalcified bone at 40 kPa), cells were allowed to retain intercellular tension, form large adhesions, and differentiate to osteoblasts [76]. A seminal study on MSC confinement used small adhesive arrays of fibronectin (1000 μm2) to prevent cell spreading, and hence development of intracellular tension, and lead to formation of adipose (fat) tissue, while when spreading was actively promoted on large, 10,000 μm2, fibronectin arrays, cells formed large adhesions, well-organized cytoskeleton, and differentiated to osteoblasts [77]. An elegant update on this study used fibronectin stars and flowers to demonstrate that even if the MSCs are the same size, features that promote adhesion (sharp corners of the stars) promote intracellular tension and osteogenesis, and features that reduce adhesion (soft ‘corners’ of the flower petals) reduce intracellular tension and differentiation [78]. The common feature of all the above studies is their presentation of Rho-A kinase (ROCK) as a key modulator of osteogenesis. This is sensible as, as has been described, Rho-A is a small G-protein involved in activation of actin/myosin contraction. Cells require this cytoskeletal contraction to spread onto substrates and to migrate. This contraction against focal adhesions allows formation of intracellular tension and motility. Osteoblasts are a large progeny of MSCs and thus are very well spread, therefore a highly contractile cytoskeleton is required to support this phenotype and this generates high intracellular tension. In addition, focal adhesion and the cytoskeleton are in a force– balance relationship, as highly contractile cytoskeleton requires large focal adhesions and focal adhesion requires force to gather integrins and hence to grow [79–83]. It is notable that osteoblasts have a high proportion of the aforementioned supermature adhesions [84, 85]. Pharmacological inhibition of ROCK prevents osteogenesis and promotes lowtension adipocyte-specific differentiation from MSCs (please note that adipocytes are round with few, small, adhesions). It is further noteworthy that the MSC phenotype itself appears to be of intermediate cytoskeletal tension, with the stem cells employing many, smaller adhesions; the cells are close to a fibroblastic phenotype, but with fewer adhesions and slightly less intracellular tension [86, 87], as was demonstrated using nanoscale topographies [86, 87]. The biochemical basis behind these phenotypical changes is not so well understood. Downstream of FAK and the G-proteins are major biochemical signaling hubs such as extracellular-signal related kinase (ERK1/2) and other mitogen activated protein kinase (MAPK) modulators such as p38 and c-jun n-terminal kinase (JNK), all of which have been shown to be important in nanoscale MSC interactions [86, 88, 89]. Such hubs have also been illustrated as central in other materials approaches such as microcontact printing [90] and changing substrate stiffness [76]. It is likely that the MSC phenotype is altered through regulation of transcription factors. For example, the osteogenic transcription factor, runt-related transcription factor 2 (RUNX2), requires phosphorylation to be active, while the adipogenic transcription factor, peroxisome proliferator-activated receptor gamma (PPARGγ), is inactivated by phosphorylation. As adhesion-related signaling increases, ERK activation also increases. At very low adhesion, ERK is largely downregulated and thus PPARGγ and RUNX2 are unphosphorylated permissive to adipogenesis, at medium adhesion, ERK is upregulated to a degree permissive for proliferation with little net adipogenesis or osteogenesis. However, if supermature adhesions are allowed to form, ERK is stimulated to the point of negative feedback and this slows proliferation and allows phosphorylation of the transcription factors, inactivating PPARGγ and activating RUNX2 to promote osteogenesis [85, 91, 92].

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Linking the Cytoskeleton to the Nucleus and Direct Mechanotransduction Maniotis et al. [93] reported that in reaction to tension, the intermediate filament cytoskeletone-oriented, the nucleus distorts, and the nucleoli rearranged along the applied axis. Thus, it was concluded that the nucleus is mechanically integrated within the physical entity of the cell via intermediate filaments and that active or passive cell extension can lead to passive nuclear deformation [72, 94]. Micromanipulation experiments have illustrated that transmission of force can occur from the peripheral cytoplasm into the nucleus [95], demonstrated the mechanical interconnection of chromosomes and nucleoli [96], and shown the importance of the nuclear lamina in force transmission [97, 98]. Topography has provided a useful tool in investigation of force transduction into the nucleus and has been used to indicate that redistribution of chromosomal territories (the space occupied by a given chromosome within the nucleus, reviewed in [99], or intraterritory loci, the location of particular genes) could affect the cellular transcriptional profile. Such movements could bring genes closer to choromosome factories (areas rich in RNA polymerase II) [100, 101], or away from/towards the periphery of the nucleus – an area that can promote downregulated transcription. Wang et al. [72] suggested a number of models by which tensile forces from FAs could modulate gene expression. It was proposed that the assembly or activity of transcription factor complexes could be affected directly or indirectly by tension-mediated alterations in the nucleoskeleton and that the telomeric ends of the chromosomes are attached to the lamina via matrix attachment regions (MARs). Also, the authors suggested that gene expression or mRNA transport could also be affected by tension-mediated changes in nuclear pores. Even at the nanoscale, topography provides a useful means to investigate mechanotransductive effects on the nucleus. Nanocolumn substrates were shown to induce repositioning of chromosome 3 (Ch 3) [102, 103] relative to planar controls. The Ch 3 centromeres were more closely apposed in cells cultured on the nanocolumns, probably as a consequence of the decreased cell spreading and nuclear area. On hexagonally arrayed nanopits, the intercentromeric distance was reduced for both chromosomes 3 and 11, and the cells were  markedly less spread than controls [103]. As the authors noted, the changes were consistent with the tensegrity model (see papers by Ingber et al. [72, 104, 105]). Tension on the cytoskeleton and nucleus should be reduced, which is likely to have lessened the force exerted on chromosomes. Furthermore, micro- and nanotopography have been used to illustrate movement of large chromosomes in fibroblasts [106] and MSCs [88], with the nuclear lamina indicated as central in the conveyance of mechanical signals to permit these movements. This is particularly interesting with MSCs, which, as has been described, differentiate in response to intracellular tension. The genome is surveyed with respect to gene positioning in relation to the telomeres (attached to lamins and hence mechanically sensitive) and the centromeres (not attached and thus not mechanically sensitive). It is seen that many osteogenic genes reside on larger chromosomes (which all these studies show to be more mechanically sensitive) at the telomeric regions (e.g., osteocalcin at 1q25-31, osteopontin at 4q22, osteonectin at 5q31 and alkaline phosphatase at 2q37). This indicates that the “osteogenic” genome of MSCs may be in a position to respond rapidly to changes in tension, as the telomeric ends may “unravel”, or at least become less dense and more euchromatic, faster in response to tension. This change in DNA density may be permissive to transcription factor and polymerase access, thus aiding phenotype selection.

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Direct Stimulation of Nanomechanotransduction “Nanokicking” To this point, we have described the nature of mechanotransduction and how it can be studied on the nanoscale using, for example, topography. What we have not yet covered is how to apply nanoscale mechanotransductive forces directly to MSCs. A recent report by us achieves just this, harnessing the reverse piezo effect to convert an applied voltage into nanoscale impulses, or “nanokicking.” Our report demonstrated that MSCs could be induced to form mature osteoblasts through nanoscale mechanical cues alone [61]. Adopting an open hypothesis for input frequency (the study used a set 15 nm vertical displacement and varied the frequency of input) lead to the use of a 1 kHz frequency to stimulate osteoblastogenesis (see Figure 12.2 for typical set-up). While this appears fast for the time-scale of many cell actions, Pierres et al. [107] provide an interesting perspective article on cell membrane undulations based on their original data [108] and the literature. They note that many familiar, conventional, membrane deformations occur at the tens of seconds time-scale, for example, filopodial probing. However, they next noted that many cell–membrane interactions occur much faster than this. For example, neutrophils completely ingest pathogens within seconds [109] and endothelial arrest of flowing leukocytes involves subsecond integrin activation [110]. The authors point out that subsecond membrane undulations may be key to these phenomena. Interestingly, erythrocyte membranes display high-frequency undulations (as high as 1 kHz) in the tens of nanometers range called flickering [111, 112] – this is very similar scale to the stimulation we supply the cells within this report. However, it is noted that such movements are much slower in nucleated cells, for example, lymphocytes and monocytes from the blood displaying 20–30 nm undulations at up to 30 Hz and fibroblasts (descendants of MSCs) displaying 1–4 nm displacements at up to 0.5 Hz [113–115]. Over a number of years there has been discussion of piezoelectric effects in bone. This is because while collagen is piezoelectric when dry, for the effect to be seen in hydrated bone [116], a kilohertz-range stimulus is required; that is, beyond that considered physiological [117]. Furthermore, it is known that electricity can be used to guide cells (galvanotaxis [118, 119]) and stimulate bone repair [120]. It is of further interest that the report demonstrated that this osteogenic response was ROCK dependent, in agreement with previous materials-based strategies for osteoinduction [76–78, 87, 121]. That ROCK inhibition blocked osteogenesis demonstrates that the stimulation is a

Petri dish Al disk Piezo actuator Electrical feed

Figure 12.2  Image of the typical set-up, with 60 mm diameter Petri dish and attached piezo actuator glued between the large aluminum block (below) to ensure upwards movement on expansion of the piezo, and the aluminum disk to provide low dispersion in the center to edge displacements. Reproduced from [61] with permission of the American Chemical Society.

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Figure 12.3  Network analysis, showing involvement of FA, FAK, ERK, actin, sonic hedgehog, and RhoA signaling in response to 1 kHz nanomechanostimulation: N = 3. Note that CP is the canonical pathway and the lines into the network indicate where these biochemical pathways are involved in the network. Reproduced from [61] with permission by the American Chemical Society.

real effect. The study also demonstrated that BMP2 was activated by 24 h and RUNX2 by 7 days via nanokicking. This is logical, as RUNX2 activity is mediated by BMP2 signaling [91, 122], and this would be followed by increased osteocalcin expression by day 14 according to the Stein and Liano steogenesis time lines [123]. Microarray-based pathway analysis (Figure 12.3) also implicated purine and nicotinamide metabolism (involved in energy demand) and sonic hedgehog signaling (involved is stem-cell differentiation) [124] as well as adhesion, FAK, ERK, and actin-mediated biochemical signaling (all known to be implicated in RUNX2 activation [125–129]) being central to nanomechanotransduction [130, 131].

Bone Bioreactors Now that we have demonstrated nanoscale mechanotransduction, what is the future direction for this technique? Use of osteoblasts and stem cells to make bone tissue in the laboratory (ex vivo) is possible [132, 133], but for clinical use (massive bone production), a special device

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to accelerate tissue growth and production is needed. Different research groups have developed different bioreactors [134]. They usually have been used to improve seeding efficiency, proliferation, and osteoblast differentiation [8]. In these culture chambers the cells will grow, proliferate, and differentiate to bone-forming cells, all in sterile conditions. These bioreactors mainly fall into mechanical (compressive, rotatory, and spinning) and perfusion (running a flow of fresh media over cells) categories in order to optimize the environment for osteoblast growth and differentiation [135, 136]. The efficacy and problems with each type are under investigation and it seems the perfusion systems show the best results for osteoblastic differentiation and mineralized matrix deposition [137]. Bone bioreactors have been divided into three main groups. 1. A spinner flask is the simplest type: cells on a three-dimensional scaffold are suspended in a media container and stirred by about 50 rpm. The cell viability, proliferation, and distribution compared to a static group are improved but the size of the produced tissue is limited because by increasing the flow for increasing the tissue perfusion, especially for deeper cells, generates a shear force, leading to necrotic damage in the surface cells [137]. 2. A rotating vessel (sometimes called the NASA bioreactor) is another type of rotational reactor that usually employs horizontal rotation of solid, bubble free, vessels. In this type of vessel the necrotic effect of shear stress is lessened. In one comparison between these two methods and control, osteoblastic activity was increased by spinner flask culture after 21 days due to superior media mixing [138, 139]. 3. The perfusion method is designed to mimic blood supply. In this method various types of flow with different speeds, frequencies, and continuity (unidirectional, oscillating, pulsating) can run the media over the growing cells and at the same time gas exchange is provided by another pomp system. Frequency of flow, flow rate, and shear stress are important variables for the perfusion system method [137, 140]. While it is premature perhaps to describe our nanoimpulse system as a bone bioreactor, it is certainly simple, efficient, and scalable. However, it is currently better defined as an osteoblast bioreactor as it is two-dimensional. Our future work, however, will focus on building this into a three-dimensional system for bone-tissue engineering.

Conclusions and Future Outlooks Stem cells show a wide range of reactions to their nanoenvironment. These mechanotransductory reactions are likely largely due to changes in FA formation and concomitant intracellular pathways activation. Nuclear responses and gene expressions could also be generated by these nanoscale stimuli. Conduction of stem-cell fate with induction of mechanical stimulation could introduce a new generation of bioreactors. To this end, we have introduced a new nanoscale method of MSC stimulation for targeted osteoblastogenesis. This does not rely on novel materials, complex chemistry, or electronic clean-room facilities. Rather, it is based on traditional cell-culture plastics with simple addition of piezo ceramics. Upscaling to bioreactors that can prime autologous MSCs to form osteoblasts without recourse to soluble factors can be easily envisaged. It could also be envisaged that in vitro experiments could be used to inform therapy (whole-body vibration) with the noted caveat that the modeling from cell to whole body is nontrivial and much research aimed at practical/theoretical scaling between nanoscale cell culture and the human body is required. Such techniques could be complementary to existing external stimuli for musculoskeletal regeneration such as extracorporeal shock wave treatment.

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Acknowledgments Nikukar is funded by the Iranian Ministry of Health and Medical Education. Reid is funded by grants from SUPA, STFC, RS, RSE, and SCI. Dalby is funded by grants from the BBSRC, EPSRC, MRC, and the AO Foundation.

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Chapter 13

Delivery of Molecules and Genes/Small Interfering RNA into Stem Cells by Nanoengineering Mohsen Ashjari1,2

Institute of Nanoscience and Nanotechnology, University of Kashan, Kashan, Iran Department of Stem Cells and Developmental Biology at Cell Science Research Center, Royan Institute for Stem Cell Biology and Technology, ACECR, Tehran, Iran

1  2 

Introduction Nanotechnology, the engineering and manufacture of efficient systems at molecular and atomic scales, has overshadowed many aspects of today’s medicine. This technology has revolutionized pharmaceutical, medical therapeutics, and diagnostics through the development of ingenious nanodevices [1]. Stem cells are unspecialized, clonogenic cells that, according to their potency, can differentiate into multiple types of differentiated cells from one or all three primitive embryonic germ layers (ectoderm, mesoderm, and endoderm) in response to specific signals. Stem cells possess two capacities to self-renew and make copies of themselves and differentiate into organs and tissues. Generally, stem cells differentiate into intended cellular tissue by induction of growth factors, genes, and proteins. The ability to accurately and efficiently differentiate stem cells into all lineages is the subject of intense research [2]. Recently, engineering has become an important strategy for the induction and regulation of differentiation of stem cells into specific cell types. Because stem cells can be grown long term in vitro, their genetic modification prior to transplantation provides a unique opportunity for correction of genetic defects [3]. Conventionally, cellular delivery strategies, such as electroporation and microinjection, nucleofection, retro-viral transduction, and cationic liposomes are well-known, however, their application in embryonic stem cells (ESC) is both a time and labor-intensive process. Nonviral carriers have emerged based on polymers or similar materials. These carriers in comparison with viral carriers exhibit low gene efficiency and cytotoxicity in numerous trials [4]. Nanotechnology-based intracellular delivery is a relatively recent concept in stem-cell research and development, and “nanoengineering” is the application of nanotechnology in stem-cell engineering. The concept of nanoengineered nonviral delivery into stem cells may be of value if biologically active agents can be incorporated onto nanocarriers. The goal of such nanocarriers is to enhance agents’ uptake by stem cells without the limitations found in viral delivery that include infection-related cell damage and immune response issues [5]. For example, the complexation of DNA with a nanoparticle avoids numerous viral delivery

Stem-Cell Nanoengineering, First Edition. Edited by Hossein Baharvand and Nasser Aghdami. © 2015 John Wiley & Sons, Inc. Published 2015 by John Wiley & Sons, Inc.

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problems and shows promise for gene delivery into the stem cell. Intracellular delivery of specific proteins/peptides may be used to influence signaling pathways and manipulate stem-cell fate. Efficient intracellular delivery occurs following the carrier moving to the cell surface, cellular uptake, endosomal escape, and carrier unpackaging [6]. Nanocarriers, due to their small size, can efficiently penetrate across the cell membrane barrier and increase the efficiency of intracellular delivery. These nanocarriers can be synthesized chemically and they modify the condensation and physicochemical state of the loaded gene/small interfe ring RNA (siRNA) to protect them against cytoplasmic damage. Functionality of the nanocarriers can be adjusted for controlled release of genes/siRNA. Factors that affect delivery efficiency of nanoparticles include cell type, cell-cycle stage, cell-culture conditions, cell density, and size of passaging nanoparticles, controlled intracellular release of bioactive agents, cytotoxicity, stability, storage, and shelf-life of nanocarriers [7]. To attain a successful outcome in stem cells, several factors in delivery systems should be taken into account. First, dependent on the target and type of bioactive agent, the size and type of nanocarrier can vary. From the standpoint of the nanocarrier characteristics, size, size distribution, surface charge, and the nature of the systems play critical roles for successful intracellular delivery [8].

Delivered Bioactive Agents into Stem Cells Molecules Application of small molecules to regulate stem cell behavior is particularly beneficial as they provide a high degree of temporal control over protein performance by either rapid activation or inhibition of single or multiple targets within a protein family. Retinoic acid (RA) is a good candidate in controlling stem-cell-derived neuronal differentiation [9], and has been used to tune neuralization and positional specification during mouse ESC differentiation. However, RA is rapidly metabolized by cells, has low water solubility in aqueous solutions, and requires a fine tuning of the concentration window to obtain results. These problems pose difficulties in the delivery of therapeutic doses. Moreover, the use of this bioactive small molecule in an in vivo setting for the differentiation of stem cells remains elusive [10]. Biological systems use a complex transportation network to deliver RA at the cell nucleus so that it can activate RA receptors. Initially, RA is taken by biological systems as retinol, which is removed from the blood and bound to cellular retinol-binding proteins in the cytoplasm. The retinol dehydrogenase enzymes metabolize retinol to retinal, which in turn is metabolized to RA by the retinaldehyde dehydrogenases. The RA is then bound to cytoplasmic RA-binding proteins and this complex finally enters the nucleus and binds to the RA receptors and the retinoid X receptors, which in turn heterodimerize and bind to a sequence of DNA known as the RA-response element. Targeted RA delivery is also of interest to manipulate stem cells. Increasing the specificity in the intracellular delivery of RA will reduce not only side-effects but also the necessary amount of the molecule and resultant costs. The chemical structure of RA is shown in Figure 13.1.

Gene/Small Interfering RNA Introduction of genes/siRNA to regulate specific genes involved in signaling pathways that control the cell phenotype can induce specific differentiation of stem cells into specific cell types. Noncoding RNAi molecules known as microRNA (miRNA) post-transcriptionally

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O OH

Figure 13.1  Chemical structure of retinoic acid (RA).

Plasmid DNA – –

+

– – –

Cationic polymer

+

+

+ +

Condensed complex

+ +

Figure 13.2  Plasmid DNA condensed with cationic polymers can be encapsulated into polymeric systems for gene delivery.

regulate gene expression to endogenously control a wide-range of biologic processes that include stem cell self-renewal, development, differentiation, growth, and metabolism. The siRNA is an exogenously delivered double stranded RNA that can function identically to known miRNAs. Effective delivery of siRNA may provide potent cues to direct stem-cell behaviors for regenerative medicine applications [11]. Most delivery strategies for siRNA agents use cationic organic materials to facilitate cell uptake and endosomal escape. Targeted delivery toward the ribosome or the endoplasmic reticulum rather than the cytoplasm may enhance translation inhibition elicited by interfering RNA [12]. DNA is negatively charged and can be condensed by cationic polymers to form a p ­ olyplex. Because condensed DNAs are compacted, they are less accessible to nuclease degradation in the cytoplasm and can cross the nuclear membrane at an increased efficiency [3]. Schematic representation of condensation of DNA is shown in Figure 13.2.

Proteins Intracellular delivery of proteins presents an exciting alternative to regulate signaling events. Intracellular signaling pathways play a significant role in maintaining the undifferentiated state and self-renewal of ESCs and progenitor cell populations. It has been shown that β-catenin plays a marked role in tuning cell growth and the balance between differentiation of neural stem cells (NSCs) and proliferation [13]. Inhibition of glycogen synthase kinase 3 beta (GSK-3b) by external stimuli may thus operate as a way to regulate the potential fate of stem-cell populations.

Nanoengineered Carriers for Intracellular Delivery into Stem Cells Nonviral carriers are nanoengineered with various biocompatible materials to form nanoparticulated devices (or nanoparticles) for intracellular delivery of bioactive agents with the intent to effectively overcome cellular barriers and improve stability and uptake of agents. Limitations with viral approaches that include small loading capacity, resistance to repeated infection, difficulties in production, quality control, and low safety can be potentially overcome by the application of a nonviral and nanoscale carrier approach.

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Control of Stem-Cell Fate by Engineering of Microenvironment

Nucleus

Lysosome

Small molecules

Proteins DNA siRNA

Endosome

A typical loaded nanoparticle

Figure 13.3  Scheme of nonviral delivery nanocarriers. Internalized nanoparticles escape the endosome and release their cargo.

Delivery of bioactive agents into stem cells can be a potent strategy to direct their differentiation into specific cell types, as seen in Figure  13.3. When the nanoparticles are formed the next key step in delivery is endocytosis. For efficient uptake it is important for nanocarriers to have an overall positive charge so that they will be electrostatically attracted to the negatively charged proteoglycan cell surface. The transfection efficiency of carriers is dependent on surface charge. When the surface charge becomes less positive, adsorptive endocytosis decreases, which results in reduced transfection efficiency. The size of the nanocarriers affects uptake and smaller particles (about 100 nm) have the most efficient uptake. Throughout uptake and internalization, nanocarriers are encased into endosomes and become part of the endosomal sorting pathway. As the next step, the bioactive agent must be released from the nanocarrier. Polycationic carriers have enhanced intracellular delivery due to their ability to directly enable endosomal escape through the “proton sponge” mechanism [4, 5]. Nanocarriers are formed by numerous biomaterials for intracellular delivery into stem cells [7]. These biomaterials consist of organic, inorganic, and organic–inorganic hybrid nanocomposites. Organic materials are classified into two categories: natural and synthetic polymers. The natural organic materials include chitosan (CS), hyaluronic acid (HA), and dextran, among others. Synthetic organic polymers allow a high level of design flexibility for preparation of nanocarriers. These polymers are divided into three groups. Bioreducible polycationic polymers show reduced cytotoxicity and controlled intracellular release of genes, leading to increased transfection efficiency. Examples of these polymers include branched/linear polyethylenimine (PEI) and branched poly(disulfide amine) (B-PDA) among others. Biodegradable polymers have the advantage of being eliminated after the bioactive agents are released, in the form of nontoxic degradation products such as the polyesters poly(lactideco-glycolide) (PLGA), poly (β–amino ester) (PBAE), polybutylcyanoacrylate (PBCA), and poly(ethylene glycol) (PEG). The final group comprises dendrimers or poly (amido amine)s (PAMAM). Polymeric nanocarriers are of interest due to their low cost, facilitated production, and controllable toxicity, based on parameters such as ionic charge, chemistry, and chain length. Inorganic carriers consist of silica (SiNP), calcium phosphate (CPNP) and magnetic (MNP) nanoparticles. The organic–inorganic hybrid carriers are combinations of polymers

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N

NH2

N H

N

N H

NH2 N

N

H2N

O

H N

N

N H

NH2

S

N H

HN

n

G0

O

H N

N H

PEI

G1

S

NH2

n

B-PDA

G3

G2 PAMAM

O O HO

HO

OH O OH

O

HO O

NH O

OH O n

O HO

O

NH

O

O OH

O N R

O n

Chitosan

O x O

H

H

O

O n

Y

PLGA

n

PBAE

O

O O

O

O

HA

OH

NH2

HO

O

PEG

O

H N

n

PBCA

Figure 13.4  Available biomaterial used as nonviral nanocarriers for delivery into stem cells.

and inorganic nanoparticles such as PEI–MNP or PEI–SiNP nanocarriers. The chemical structures of some biomaterials are shown in Figure 13.4. Tables 13.1 and 13.2 summarize various nanocarriers that have been nanoengineered for intracellular delivery into stem cells. In this chapter, recent advances in the delivery of bioactive agents by employment of nanoscale carriers are discussed along with common problems such as bioactive agent– nanoparticle conjugation, uptake efficiency, intracellular fate, and toxicity.

Organic Nanocarriers Polyethylenimine Some of the commonly used biological active agents are packaged with polycationic polymers to create nanoparticles that have the capability to effectively overcome extra- and intracellular barriers during agent transfer. Polyethylenimine is a polycationic polymer that can be easily taken up by cells through endocytosis. The polymer’s amine groups within the endosome cause swelling, increase osmotic pressure, and disrupt the endosome, leading to an efficient endosomal escape [10]. Polyethylenimine can coat a range of organic materials to form nanocarriers. The PEI coat on an organic polymeric core causes polyplexing of DNA, while the organic polymeric core reduces cytotoxicity and increases cellular uptake. Polyethylenimine conjugated with HA [14], PEG [15], or acetic anhydride [16] facilitate the controlled release of biologically active agents. The RA−PEI complex nanoparticle is a potentially powerful tool for controlling stemcell fate. Maia et al. [9] have reported neuronal differentiation of stem cells by delivery of RA-loaded nanoparticles. They prepared 200-nm-diameter nanoparticles by complex coacervation through the electrostatic interaction of a polycation (PEI) and a polyanion (dextran sulfate). These researchers investigated the effects of dissolution of nanoparticles at

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Table 13.1  Recently reported organic nanocarriers for bioactive agent delivery into stem cells Origin

Type

Bioactive agent

Cell type

Year

Reference

Synthetic

PEI

RA

mESC

2013

[10]

Cyclodextrinmodified dendrimer

siRNA

NSC

2013

[19]

Cystamine- PBAE

siRNA

MSC

2012

[30]

Branched PDA

siRNA

MSC

2012

[44]

Diblock copolymer

siRNA

MSC

2012

[11]

PLGA-PEI

DNA/siRNA

hMSC

2012

[39]

Acetylated-PEI

DNA

MSC

2008

[16]

PAMAM

DNA

MSC

2008

[18]

Peptide-functionalized dendrimer

DNA

MSC

2010

[20]

PAMAM dendrimer

DNA

MSC

2010

[21]

Peptide-functionalized dendrimer

DNA

MSC

2011

[22]

dPAMAM

DNA

rMSC

2009

[23]

PBAE

DNA

hMSC

2009

[28]

PBAE

DNA

hESC

2008

[29]

PBAE

DNA

MSC

2011

[31]

PLGA

DNA

hMSC

2011

[38]

PLGA-PEI

DNA

hCBMSC

2008

[36]

PLGA-PEI

DNA

hMSC

2011

[37]

PEG-graft-PEI

DNA

MSC

2011

[15]

PBCA

DNA

iPSC

2013

[46]

Dextran/PEI

RA

SVZNSC

2013

[9, 17]

CS/dendrimer

Dex

rBMSC

2009

[42]

CS/PEI

DNA

rMSC

2010

[40]

HA/Branched PEI

DNA

MSC

2011

[14]

HA/Branched PEI

DNA

hMSC

2008

[43]

CS

DNA

hMSC

2003

[41]

Natural

PEI, polyethylenimine; PBAE, poly(β–amino ester); PDA, poly(disulfide amine); PLGA, poly(lactide-co-glycolide); dPAMAM, partially degraded PAMAM dendrimer; PEG, polyethylene glycol; PBCA, polybutylcyanoacrylate; CS, chitosan; RA, retinoic acid; Dex, dexamethasone; mESC, murine embryonic stem cell; NSC, neural stem cell; hCBMSC, human cord blood-derived mesenchymal stem cell; rBMSC, rat bone-marrow stromal cell; iPSC, induced pluripotent stem cell; SVZNSC, subventricular zone neuronal stem cells.

different pHs as well as the effect of RA-nanoparticles on cell proliferation and functional neuronal differentiation. Their intent was to assess the effectiveness of this delivery system. In another study [17], the same group investigated the application of complex nanoparticles as a vehicle to induce the neuronal differentiation of subventricular zone NSCs. This intracellular delivery system activates nuclear RA receptors, decreases stemness, and increases proneugenic gene expression. Recently, Ku et al. [10] synthesized functional RA−PEI complex nanoparticles from branched PEI chains with carboxyl groups of RA surfaces by electrostatic interaction to

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Table 13.2  Recently reported inorganic and organic–inorganic hybrid nanocarriers for stem-cell fate Origin

Type

Bioactive agent

Cell type

Year

Reference

Inorganic

SiNP

Chimeric protein (GFP-FRATtide)

rNSC

2011

[13]

SiNP

siRNA

NSC

2013

[52]

MNP

DNA

NSC

2011

[49]

MNP

DNA

MSC

2011

[50]

CPNP

DNA

rDPSC

2008

[51]

CPNP

DNA

MSC

2011, 1012

[56, 57]

PEI-MNP

miRNA

hMSC

2013

[59]

PEI-MNP

DNA/Tat peptide

NSC

2010

[58]

PEI-MNP

DNA

hMSC

2013

[48]

PEI-SiNP

DNA

hMSC

2010

[60]

Organic

SiNP, silica nanoparticles; MNP, magnetic nanoparticles; CPNP, calcium phosphate nanoparticles; rNPC, rat neural stem cells; rDPSC, rat dental pulp stem cells; miRNA, microRNA.

form cationic RA−PEI complex nanoparticles for differentiation of ESC-derived neurons. The RA − PEI complex nanoparticles had a mean size of 70 nm and homogeneous circular shape morphology. They showed that pH enabled the control of amounts of RA delivered from the RA−PEI complex nanoparticles and also observed that these nanocarriers induced ESC-derived neuronal differentiation. Hosseinkhani et al. [16] acetylated PEI with acetic anhydride. This acetylated PEI had a more efficient enhanced gene delivery compared to unmodified PEI for mesenchymal stem cells (MSCs). They have mixed acetylated PEI solutions and plasmid DNA to cause homogeneous bone formation throughout the sponges. This approach can be of importance for siRNA delivery, which is recognized to have a short half-life. Polyamidoamine Dendrimers such as PAMAM are synthetic spherical macromolecules composed of a central core moiety from which multiple branches radiate. Due to the cationic property of dendrimers, they are a particularly interesting system for use in intracellular delivery, both in vivo and in vitro [18]. These molecules display a well-defined architecture and can be functionalized to improve gene expression and integration for stem-cell fate [19]. The use of cationic dendrimers as nanocarriers for gene delivery and their efficacy on MSCs with a view to applications in tissue engineering has been studied by Santos et al. [18]. They explained the efficacy of dendrimers in gene delivery according to the charge-based interactions between these molecules and DNA. These researchers genetically engineered MSCs to express human bone morphogenic protein 2 (hBMP-2) using PAMAM, different generations of dendrimers and plasmid DNA that encoded β-galactosidase for in vitro transfection of MSCs. The differentiation of the MSCs was studied in the absence of other osteoinductive factors. This was achieved through the analysis of established markers of the osteoblastic phenotype, which included alkaline phosphatase (ALP) activity, osteocalcin, and deposition of a calcified matrix. The PAMAM/hBMP-2 system strongly induced in vitro differentiation of MSCs to the osteoblast phenotype. Additionally, dendrimers can be functionalized with other molecules to enhance gene integration and expression, such as peptides that present a high binding affinity for MSCs

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Control of Stem-Cell Fate by Engineering of Microenvironment

[20]. This novel system has shown low cytotoxicity and transfection effectiveness superior to those of native dendrimers. By saturating cell receptors with the osteotrophic peptide prior to transfection, the transfection efficiency was observed to be similar to that of the native dendrimer, which demonstrated the receptor-mediated identity of the process. In order to improve gene delivery in MSCs, PAMAM dendrimers were functionalized with alkyl chains of different lengths at their periphery [21]. These systems have shown a remarkable capacity for mediating the internalization of pDNA with minimum cytotoxicity. In this delivery system the cationic nature of the dendrimer facilitates the binding of plasmid DNA, which confers its protection against serum nucleases. Indeed, the lipidic nature of external cores of dendrimer facilitates the interaction with biological membranes and assists with escape of the complexes to the cytosol. Evaluation of cellular uptake, Luc gene expression, enhanced green fluorescent protein (EGFP) expression, and cytotoxicity have shown a remarkable capacity of these nanocarriers for internalizing pDNA with very low levels of cytotoxicity, which correlates with the CH2 content present in the hydrophobic moiety. The same group employed a biomimetic approach with RGD nanocrystals based on ­dendrimers to deliver genes into MSCs [22]. They used the peptide-conjugated PAMAM and integrin targeting capabilities of the arginine-glycine-aspartate (RGD) motif to improve gene delivery. In these nanocarriers, dendrimers were used as scaffolds for RGD clustering and, by manipulating the number of peptides linked to each dendrimer, it was possible to evaluate the effect of RGD density on the gene-delivery process. There was enhanced transfection efficiency of the MSCs by means of the new vehicle compared to native dendrimers. It was found that gene expression depended on the electrostatic interaction established ­between the dendrimer moiety and the cell surface, in addition to the RGD density of the nanoclusters. Holladay et al. [23] studied a dendrimer and pDNA complex entrapped in a three-­ dimensional collagen matrix for gene delivery to rat MSCs. Their results showed a slow, sustained rate of transfection with a longer time of transgene expression. The authors ­postulated that primary cells, being more sensitive to matrix interactions, would benefit from the three-dimensional environment, which would act as a powerful adjuvant for gene delivery to these cells. Application of dendrimers as a nanocarrier to increase intracellular delivery of quantum dots (QDs) into stem cells for imaging purposes has been investigated [24]. In the in vivo monitoring of stem cells, due to the small amount of QD uptake in these cells, it is crucial to increase the cellular uptake of QDs to promote endosomal escape into the cytosol. A conjugation of polyamidoamine to QDs has been used to raise the efficiency of primary c­ ultured MSCs. In this study a dendrimer that could enhance cellular uptake to QDs through electrostatic interactions and a buffering capacity was used. The results showed that the fluorescence intensity of MSCs increased through conjugation of QDs with PAMAM. Dendrimers were verified for their capacity to deliver small molecules and siRNA. Shah et al. [19] have used a cyclodextrin-modified dendrimer (DexAM) for differentiation of NSCs. DexAM is a nanocarrier that solubilizes hydrophobic small molecules in physiological solutions and forms complexes with siRNA molecules. This makes it an attractive delivery system for programming stem-cell differentiation. Application of DexAM simultaneously releases hydrophobic small molecules and siRNA into NSCs with a significant enhancement of their neuronal differentiation. The researchers have applied them in inducing stem-cell differentiation, which allows simultaneous delivery of factors to direct stem-cell differentiation, ensuring long-term cell growth and survival. The prepared nanocarriers provide: (i) the capability to simultaneously deliver nucleic acid and small molecules; (ii) high transfection efficiency of siRNA; and (iii) minimal cytotoxicity.

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Poly(β-amino esters) One interesting type of cationic and hydrolytically degradable polymeric nanocarrier for intracellular delivery is PBAE, which was first developed by Lynn and Langer in 2000 [25]. Poly(β-amino ester)s are synthesized by the conjugate addition of amine monomers to diacrylates. The condensation of the gene inside PBAE nanocarriers can greatly increase the retention time and provide better protection against nuclease degradation. Poly(β-amino ester)s have the potential for structural diversity as well as the ability for ligand-specific uptake. They can efficiently condense DNA to form nanocarriers. These nanocarriers have the capability to buffer the endosome and facilitate endosomal escape, as well as having a high efficacy both in vitro and in vivo. In comparison with PEI, they possess reduced cytotoxicity and biodegradability, and the ability for triggered genes release within the stem cell [26, 27]. Yang et al. [28] developed PBAE nanoparticles that could deliver the vascular endothelial growth factor (VEGF) gene to hMSCs and hECS-derived cells to promote angiogenesis. Interestingly, their effort led to enhancement of human VEGF production, cell viability, and engraftment into tissue target-producing vessels. This delivery system showed that stem cells nanoengineered with PBAE could be therapeutic tools for vascularizing tissue constructs and treating ischemic disease. Green et al. [29] assembled PBAE with DNA to prepare submicron and positively charged polymeric nanoparticles for efficient gene delivery in hESCs. These nanocarriers exhibited low toxicity and did not unfavorably influence colony morphology or cause nonspecific differentiation of the hESCs. In an additional work [30], the same group have studied the capacity of cystamine-terminated PBAE for delivery of siRNA to hMSCs for osteogenic differentiation. In this delivery system the cystamine-terminated polymer is bound to siRNA prior to exposure to a reducing environment, which efficiently releases siRNA. Thus PBAE terminated by cystamine can form tight initial interactions with its cargo and then cause an efficient, environmentally triggered delivery in the cytoplasm. These researchers in another work [31] have used PBAE nanocarriers for intracellular delivery of genes to primary glioblastoma (GB) cells as well as GB tumor stem cells in vitro, with low nonspecific toxicity and transfection efficiencies. They developed DNA nanocarriers that remained relatively unchanged in normal serum and could also be stored for at least 3 months in a ready-to-use state with no detectable drop in efficacy, which showed their potential in practical or clinical settings. In this case the specificity and transfection for GB cells was higher than in healthy astrocytes and stem cells. Poly(lactide-co-glycolide) Poly(lactide-co-glycolide) is a biodegradable, biocompatible, and US Food and Drugs Administration approved biomaterial that has aroused considerable interest among researchers who seek to develop biodegradable nanocarriers [32]. The PLGA nanoparticles have widely been used for delivery of biologically active agents and have the capacity for cell  internalization [33]. Poly(lactide-co-glycolide) have the intrinsic capacity to escape ­endosomal degradation. The mechanism of this escape involves direct interaction of PLGA with the endosomal membrane owing to selective reversal of the surface charge of PLGA (from anionic to cationic) under acidic conditions [34]. The complexation of DNA with PLGA nanoparticles has allowed robust gene expression in stem cells. Poly(lactide-co-glycolide) can be modified with other polymers such as PEI. Polyplexing with PEI has been shown to enhance cellular uptake of DNA complexed to PLGA nanoparticles both in vitro and in vivo. Research has shown that the incorporation of PLGA nanoparticles that contain growth factor in embryoid bodies caused enhanced vascular differentiation of hESCs [35]. Incorporation of these nanoparticles minimally

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Control of Stem-Cell Fate by Engineering of Microenvironment

impacted cell viability and proliferation, yet positively impacted differentiation in this study. The internalization and perinuclear accumulation of PLGA nanoparticles within hESC ­colonies and aggregated hESCs forming embryoid bodies showed no adverse effects on cell viability or proliferation in cells that contained nanoparticles. Gwak et al. [36] developed PLGA particles as nanocarriers for gene delivery to human cord blood-derived MSCs. A comparison of cytotoxicity and long-term transgene expression between PLGA nanocarriers and PEI showed that PLGA were significantly less ­cytotoxic and had higher transgene expression in vitro for a longer duration (21 days) than PEI. The authors have stated that PLGA nanoparticles provided a higher potential as gene delivery carriers for use in gene therapy for diseases in which a long-term therapeutic gene expression regimen is necessary. Park et al. [37] investigated a PEI-modified PLGA nanoparticle to assess the ability of four genes (transcription factor) polyplexed with nanocarriers that were delivered intracellularly in hMSCs. For polyplexed genes with PEI–PLGA nanoparticles, the obtained transfection efficiency was approximately 80% and led to a significant increase in chondrogenesis in vitro. The cell-uptake ability of the gene-loaded nanocarriers was enhanced for both in vitro and in vivo culture systems, including hMSCs. The same group, in another study [38], used PLGA nanoparticles as gene carriers to mediate the transfer of the SOX9 gene in hMSCs. The nanocarriers that were complexed with high levels of SOX9 plasmid DNA allowed robust gene expression in hMSCs both in vitro and in vivo, and induced chondrogenesis. In an additional work [39], they reported inhibition of expression of unnecessary genes and enhanced expression of specific genes involved in hMSC differen­ tiation by the simultaneous application of siRNA and plasmid DNA that were incorporated into cationic PEI coated on PLGA nanoparticles as co-delivery factors. Use of nanocarriers has also allowed for simultaneous introduction of a DNA vector and siRNA, which enhances efficient differentiation. After transfection the percentage of GFPexpressing hMSCs decreased from 25.35 to 3.7% with GFP-DNA/PLGA or GFP-siRNA/ PLGA, whereas GFP-DNA/PLGA and scramble siRNA (MOCK)/PLGA had no adverse effect on GFP expression. Chitosan Chitosan is a nontoxic, biodegradable, natural polysaccharide consisting of repeating units of N-acetyl-glucosamine and glucosamine, the proportions of which determine the degree of deacetylation and the polymer’s properties of solubility, hydrophobicity, and the ability to interact with polyanions [7]. Chitosan can condense DNA and protect it against nuclease degradation. Furthermore, CS nanoparticles are stable during storage and their preparation does not require organic solvents, which minimizes possible damage to DNA during complexation. Thus, it is a good candidate as a nonviral gene delivery nanocarrier. Generally, a low molecular weight, highly deacetylated CS results in small sized nanoparticles with highly condensed DNA [40]. For the first time, Corsi et al. [41] synthesized a cationic CS-DNA plasmid complex to evaluate the potential for CS to develop a nonviral gene-delivery nanocarrier in a variety of cell types, including hMSCs. These CS-DNA nanoparticles were shown to have lower c­ ytotoxicity than lipoplexes and gene delivery efficiency was dependent on the cell type. Oliveira et al. [42] conducted research on the impact of combining dexamethasone (Dex) loaded carboxymethylchitosan/poly(amidoamine), or (Dex-loaded CMCht/PAMAM), nanoparticles and both hydroxyapatite and starch–polycaprolactone scaffolds (three-­dimensional system) on the in vitro expansion and osteogenic differentiation of rat bone-marrow stromal

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cells (rBMSCs). They demonstrated that rBMSCs seeded onto the surface of the scaffolds differentiated into osteoblasts when cultured in the presence of Dex-loaded CMCht/PAMAM nanoparticles. They confirmed that Dex-loaded CMCht/PAMAM nanoparticles combined with hydroxyapatite augmented osteogenesis by increasing ALP activity and mineralization of the extracellular matrix. The pre-incubation process confirmed by this study allowed the delivery of Dex inside the cells and directly influenced their cellular differentiation, which indicated its appropriateness for tissue engineering. Pimpha et al. [40] developed a nanocarrier for plasmid DNA by PEI-introduced CS nanoparticles for rat MSCs. The CS/PEI nanoparticles were prepared by the emulsion polymerization of a methyl methacrylate monomer in the presence of different concentrations of PEI mixed with CS. The introduction of PEI affected the surface charge, dispersing ­stability and buffering capacity of the nanoparticles from the viewpoint of the gene transfection carrier. Gene transfection and the prolonged time period of expression of CS/PEI nanoparticles for MSCs were achieved with greater efficiency than that of CS and ­lipofectamine. They have concluded that the combination of CS and PEI on the nanoparticles is promising as gene delivery nanocarriers for MSCs. Hyaluronic Acid Hyaluronan or HA has been used to modify the dispersing stability, surface charge, and buffering capacity of polymers such as PEI [43] to form nanoparticles for nonviral gene delivery. Nanoparticles made of HA and CS showed lower cytotoxicity and induced a higher rate of gene integration in NSCs and spinal cord slice tissue compared to those obtained with PEI. Mikos et al. [43] prepared branched PEI (bPEI) and HA as a nanocarrier for plasmid DNA in order to improve transfection of bPEI into hMSCs. The bPEI-HA nanocarrier formed a zwitterionic polymer capable of inter- and intramolecular interactions. The hMSCs transfected with smaller complexes showed a significant increase in transfection and bPEIHA performed significantly better than bPEI in terms of cell viability and maximum transfection efficiencies. Therefore, modifying bPEI by covalent conjugation with HA enhanced its performance as a gene-delivery nanocarrier in hMSCs. Also Mahor et al. [14] encapsulated plasmid DNA into HA biomaterials by using bPEI as a transfecting agent to produce a nonviral gene delivery nanocarrier for MSCs. The DNA-HA nanoparticles were formulated by ionic gelation followed by the crosslinking method with high encapsulation efficiency. Hyaluronic acid nanoparticles provided a surface for sustained release of DNA, thereby inducing transgene expression for a period of one month. The results of cellular localization studies illustrated that nanocarriers were rapidly internalized by the cells through nonspecific endocytosis. Consequently, long-term expression of the desired protein could be achieved with a smaller amount of DNA required. Poly(disulfide amine) Branched poly(disulfide amine) (B-PDA) is a bioreducible cationic polymer that contains disulfide bonds in its backbone which are capable of forming a nanocarrier for efficient intracellular delivery into the stem cell. This polymer with its flexible cationic amino branches can complex with a gene/siRNA and have a high proton-buffering capacity for efficient endosomal escape. Poly(disulfide amine) because of rapid degradation of disulfide bonds in the reductive environment facilitates release of siRNA in the cytoplasm. Also, it has a low toxicity and efficient body clearance owing to its efficient biodegradation [44].

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Previous research focused on enhancement of efficiency, production of growth factors, and survival of MSCs. For MSC spheroids, however, the therapeutic efficacy of this approach is restricted by the number of MSCs that can be included in the spheroids. When the size of the spheroid increased beyond a certain value, a large number of MSCs in the core underwent hypoxia and eventually died. Therefore, a strategy that enhanced their efficiency seemed essential. The strategy developed by Shim et al. [44] included increasing the survival rate of MSCs in a spheroid to inhibit their apoptosis activated by interactions between Fas and its ligand (FasL). They synthesized B-PDA nanocarriers for efficient intracellular delivery of Fas-silencing siRNA into hMSCs to improve the treatment of ischemic disease. The enlarged hMSC spheroids remarkably enhanced angiogenesis in mouse ischemic hind limbs. These researchers introduced an interdisciplinary approach of integrating sequence-specific RNAi, nonviral gene delivery, and spheroid-based stem cell delivery as a powerful therapeutic tool for the efficient treatment of ischemic disease. Diblock Copolymer Designing a new block copolymer that can be assembled into nanocarriers improves hydrophilicity, degradation rate, and crystallization, and shows tremendous potential for development in protein and gene delivery [45]. This copolymerization can increase loading, reduce the burst effect, prolong the in vivo residence time of bioactive agents, and prevent them from being engulfed by macrophages. Benoit et al. [11] have synthesized diblock copolymers for siRNA complexation, protection, and uptake in addition to pH-responsive blocks for endosomal escape. The designed copolymer consists of two blocks where the first block is composed of cationic dimethylaminoethyl methacrylate (DMAEMA), which complexes with negatively charged molecules (siRNA). The second block consists of relative amounts of DMAEMA/poly(propylacrylic acid) (PAA)/poly(butyl methacrylate) (BMA) essential for the endosomal escape of siRNA. This nanocarrier is cytocompatible and does not cause any negative effect on MSC survivability. The siRNA delivered via this copolymer does not alter MSC phenotype or MSC differentiation capacity. Thus, this polymer delivery system has been proven to be capable of siRNA delivery for controlling stem-cell behaviors for a variety of applications in regenerative medicine. Polyethylene Glycol Conjugation PEI with nonionic and hydrophilic polymers (such as polyethylene glycol (PEG)) minimizes toxicity. Polyethylene glycol improves the solubility of PEG-grafted PEI (PEG-g-PEI) complexes, minimizes aggregation, and reduces nonspecific interactions with proteins. This complex, PEG-g-PEI, holds promise in gene delivery due to ease of its preparation and potential targeting modification. Chen et al. [15] showed that a PEG–PEI graft copolymer had better gene delivery efficiency than cationic liposomes and did not affect the bionomics, proliferation, and differentiation potential of MSCs into adipocytes and osteoblasts (Figure 13.5). This copolymer c­ ompletely NH2 O H3C

O

O

11

O O

N H

H N m1

N

NH2 m2

Figure 13.5  Synthesized PEG–PEI copolymer by conjugating PEG onto polyethylenimine (PEI) [15].

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packaged the plasmid into stable nanocarriers. They indicated that PEG-PEI was a valid gene delivery agent with better transfection efficiency than cationic liposomes in MSCs. Polybutylcyanoacrylate Polybutylcyanoacrylate (PBCA) is also a nonviral nanocarrier for gene delivery due to its  biodegradability and nonimmunogenicity. Polybutylcyanoacrylate acquires negative charges. In order to prepare PBCA-DNA complexes, a cationic agent must be used to modify the surface of PBCA nanoparticles to enable the surface to acquire positive charges. Chung et al. [46] guided neuronal differentiation of induced pluripotent stem cells (iPSCs) with genetic regulation for regenerative medicine. They used PBCA nanocarriers to facilitate the intracellular delivery of plasmid DNA and mediate the transport of neurotrophin-3 (NT-3) into iPSCs. The PBCA nanoparticles could effectively package pDNA and increase the particle size after they were given a negative charge on the surface. Therefore, treatments with PBCA NP/NT-3 complexes could also enhance expressions of NT-3, TrkC, NH–H, NSE, and PSD95 by differentiating iPSCs.

Inorganic Nanocarriers Magnetic Nanoparticles Magnetic-based nanocarriers present many advantages for cell therapies, such as safety, delivery of bioactive agents (improvable by “magnetofection” approaches), magnetic cell targeting of magnetic nanoparticle-labeled cells to injury sites, and noninvasive imaging of magnetic nanoparticle-labeled transplant populations for cell tracking. Magnetic nanoparticles (MNPs) have gained interest as nanocarriers for gene delivery [47], while DNA–MNP complexes (called “Magnetoplex”) concentrate on the stem cells by applying an external magnetic field to increase the sedimentation of the complex [48]. Although MNPs have been used for nonviral gene delivery in hMSCs, their application for stem-cell transplantation therapies has received limited attention thus far. Pickard et al. [49] evaluated the potential of MNPs for gene transfer to neural precursor/stem cells using a neurosphere culture model system. They assessed repeat transfection (multifection) and repeat transfection plus applied magnetic field (magneto-multifection) methods to enhance transfection efficiency. They demonstrated for the first time that MNPs (neuromag particles with positive charges) can properly mediate gene delivery to neural precursor/stem cells (NPCs). Multifection approaches that markedly increased transfection with low toxicity showed no unfavorable influence on stem-cell proliferation/differentiation. The multifected NPCs that were used survived and differentiated in three-dimensional neural tissue arrays post-transplantation. Their findings showed that magnetic nanoparticle offered a simple, powerful alternative compared with viral vector systems used in neurobiology/neural transplantation research. Kim et al. [50] have used superparamagnetic iron oxide nanoparticles (SPION) to transfer genes into umbilical cord blood-derived mesenchymal stem cells (UCB-MSCs). This novel transfection method that uses SPION is safe and effective for UCB-MSCs and can be a tool for genetic optimization with a significant potential for cell tracing. Silica Nanoparticles Silica nanoparticles (SiNPs) can be functionalized to change their surface characteristics. The organically modified SiNPs show effective clearance in vivo without any sign of organ toxicity despite the fact that SiNPs are not biodegradable [51]. Silica nanoparticles have

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been shown to protect loaded DNA against denaturation and have potential for use as nonviral gene delivery carriers. Shah et al. [13] designed a strategy to influence signaling pathways and manipulate cell function, including stem-cell differentiation. They established an intracellular protein delivery through application of SiNP. They functionalized SiNP with n-octadecyltrimethoxysilane to adjust specifically targeted cell-signaling proteins. The protein (GFPFRATtide)-SiNP conjugates were delivered efficiently to the cytosol of human embryonic kidney cells and rat NSCs. This uptake impacted the Wnt signaling cascade, which lead to an elevation of β-catenin levels attributed to GSK-3b inhibition. Accumulation of β-catenin resulted in increased transcription of Wnt target genes, such as c-MYC, which guides the cell to actively proliferate and remain in an undifferentiated state. Therefore, in their study functional proteins operated in vitro intracellular delivery by means of a nanocarrier that acted as protein signals and regulated signaling pathways. Solanki et al. [52] successfully nanoengineered the delivery of siRNA for enhanced differentiation of NSCs by means of a self-assembled SiNP monolayer coated with extracellular matrix proteins. This nanocarrier improved cytotoxicity and eliminated undesirable side-effects of other carriers in the delivery of siRNA into stem cells. They have stated that while in conventional methods such as solution-mediated delivery or forwarded transfection, exogenous chemical materials are generally required to enhance cellular internalization of the siRNA (cationic lipids such as Lipofectamine™ 2000 and cationic polymers such as PEI), these exogenous materials are somehow cytotoxic and subsequently need to be removed after a certain incubation time. To overcome the abovementioned limitations they have proposed a delivery platform to transfect stem cells more efficiently. The effectiveness of this delivery platform was proven because stem cells only took up siRNAs, and not SiNP, on the surface of the platform. Therefore, this siRNA delivery may potentially help to overcome one of the critical barriers in stem-cellbased tissue engineering. Silver Nanoparticles Silver nanoparticles have been used in antibacterial materials and surfaces. Recently, the possibility of their use in wound dressing, surgical instruments, and bone-substitute biomaterials has been an interesting area of investigation. Silver nanoparticles have few potential cytotoxic and genotoxic effects in mammalian cells such as hMSCs [53]. Greulich et al. [54] conducted a study to evaluate the uptake and intracellular distribution of silver nanoparticles in hMSCs. They hypothesized that biomaterial silver nanoparticles might come into close contact with body tissues, including hMSCs. These researchers incubated hMSCs in the presence and absence of different concentrations of silver nanoparticles. They found that silver agglomerates mainly co-localized with late endosomal–lysosomal structures, but not in the cell nucleus, the endoplasmic reticulum, or Golgi complex. Based on previous studies and their experiment they noticed that the uptake of cells was size dependent. Calcium Phosphate Nanoparticles Calcium phosphate nanoparticles (CPNP) have shown potential as nonviral vectors for gene delivery. Calcium can form ionic complexes with phosphates on the nucleic acid backbone of plasmid DNA by the electrostatic interaction between the positively charged Ca2+ and the negatively charged nucleic acid. Yang et al. [55] induced bone morphogenetic protein 2 transfection in rat dental pulp stem cells (rDPSCs) using CPNP as gene nanocarriers. They also investigated the behavior of

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transfected cells when seeded on three-dimensional titanium fiber mesh scaffolds. Their results showed that nanoparticles could protect the DNA encapsulated inside from external DNase and release the loaded DNA in a low-acid environment. In vitro, nanoparticle transfection was shown to be effective and either accelerated or promoted the odontogenic differentiation of rDPSCs when cultured in three-dimesional scaffolds. Cao et al. [56] have developed a CPNP that encapsulates plasmid DNA (CP-pDNA) nanoparticles as a nonviral vector for gene delivery in MSCs. They have constructed CPNP incorporating plasmid transforming growth factor beta 1 (TGF-β1) and evaluated transfection efficiency, cell viability, and cytotoxicity of the CP-pDNA nanoparticles. The prepared CP-pDNA nanoparticles have exhibited significantly lower cytotoxicity than Lipofectamine™ 2000. The cellular uptake and transfection efficiency of the CP-pDNA nanoparticles into the MSCs were higher than needle-like CPNP and a standard calcium phosphate transfection kit. In another work [57], the same group prepared plasmid DNA/CPNP (pDNA-CP) by incorporating negative plasmid DNA-encoding TGF-β1 into CPNP in which they mixed fibronectin and loaded pDNA-CP into collagen/CS scaffolds to construct a three-dimensional nanoparticle gene delivery system (NGDS). They observed that the three-dimensional NGDS could successfully transfect MSCs and induce chondrogenic differentiation in vitro without Dex. An advantage of this delivery system was the sustained release with an elevated concentration for a relatively long period of time and high levels of transfection by TGF-β1 compared to MSCs transfected by the Lipofectamine™ 2000 method.

Organic–Inorganic Hybrid Nanocarriers Of note, SiNP and MNP can be hybridized with different polymers such as PEI with the possibility for intracellular delivery of bioactive agents into stem cells. Polymer–Magnetic Hybrid Magnetically assisted transfection was successfully considered for efficient and rapid delivery of gene/siRNA in different stem cells. For plasmid-based complexes, that transfection efficiency was enhanced by conjugation of PEI complexes to MNP even without the use of a magnetic field, as magnetic polyplexes provided a faster release of DNA into the cytosol compared with PEI polyplexes. Song et al. [58] prepared nanocarriers that consisted of PEI-coated MNPs, which were bonded with native transactivator of transcription (TAT) peptides and plasmid DNA encoding a luciferase reporter construct. The presence of the TAT peptide increased gene expression fourfold both in vitro (human NT2 NSCs) and in vivo (rat spinal cord injection). The magnetofection complexes in the cerebrospinal fluid responded to a moving magnetic field, shifting away from the injection site and mediating transgenic expression in a remote region. This type of combinatorial approach has implications for the development of TATmediated targeted gene therapies that are controllable in vivo. Schade et al. [59] reported a strategy for intracellular delivery of miRNA in hMSCs by MNPs. They have designed a delivery system (DNA/PEI/MNPs) beneficial for miR delivery, where miRs triggered their function in the cytosol close to the nucleus. This phenomenon is justifiable due to strong biotin–streptavidin connections between PEI and MNPs compared to DNAs where they need to pass the nuclear membrane. Polyplexes are effective vehicles to enter the nucleus. They have used bone-marrow-derived hMSCs with miRNA to enhance the therapeutic capacity of hMSCs in tissue engineering. The miR-335 was used, which is encoded in the second intron of the mesoderm-specific transcript (MEST) gene and is the

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regulating gene responsible for proliferation, differentiation, and migration in hMSCs. In addition, it was found to be upregulated during myogenic differentiation in vitro and induced during the regenerative phase after ischemia. In another work [48] the same group improved the transfection of hMSCs by an intracellular delivery of pDNA through a magnetic polyplex. They verified the possibility of substituting the pDNA/PEI delivery system by PEI-condensed pDNA incorporated with MNP. Indeed this new delivery system was fabricated and applied without application of a magnetic force considering the biotin–streptavidin interactions that could provide higher transfection effectiveness. The results of the study showed significantly more efficient capacity of PEI/DNA/MNPs to release pDNA than traditional pDNA/PEI alone. Polymer–Silica Hybrid An unsolved problem for the use of SiNP in nonviral gene delivery is the potential toxicity of these particles. In order to overcome this problem, coating of biomaterials onto the nanoparticle surfaces is an essential step. Unfortunately, the cytotoxicity of SiNP increases with increasing dose, exposure duration, and metabolic activity of the cell. Park et al. [60] prepared PEI/pDNA coated SiNP for nonviral gene delivery into stem cells, which stressed the ability of the system to facilitate high levels of gene expression. By polyplexing with PEI, there was enhanced ability of nanoparticles for cell-uptake in both in vitro and in vivo culture systems. The authors reported that 75% of hMSCs showed uptake for this nanocarrier. Although PEI has been previously identified as a potential candidate for gene delivery systems, PEI alone did not appear to sufficiently bind to specific genes. However, PEI complexed with nanoparticles seemed to have the necessary gene-binding capacity. Thus, PEI/pDNA was completely complexed by ionic binding with negatively charged nanoparticles. The result indicated that probably PEI alone did not have the capacity to deliver genes into stem cells since the stem cells proved unviable at high doses of PEI. This study showed that the PEI-mediated gene nanocarriers penetrated into the cell membrane and facilitated gene delivery.

Conclusions For delivery, numerous nanoengineered carriers, known as nanoparticles, with variable compositions are generally used as intracellular carriers to control stem-cell fate. The nanoengineered carriers are a promising technique to convert laboratory results into clinically viable applications with stem cells. The size, shape, charge, and surface chemistry of nanocarriers are the main factors that govern cellular uptake and the process of cellular delivery. Modified stem cells show superior characteristics of specific tissue differentiation, directional migration, and resistance to apoptosis. Despite remarkable enhancement, nanoparticulated delivery systems still suffer from low transfection efficiency. Several features are desirable in the development of delivery approaches of nanoparticles for therapeutic purposes – the ability to deliver sufficient amounts of nanoscale materials intracellular to mediate agent delivery of the desired function; the ability to deliver nanoparticles in a specific, controlled manner to only the targeted cell population; and elicitation of minimal cytotoxicity. Further research is needed for tailoring the size, content and surface properties as well as several problems regarding their fate, toxicity and safety must be addressed. Lastly, nanoengineered delivery offers new opportunities for transgenesis that should be investigated.

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Acknowledgments The author would like to thank the University of Kashan and Royan Institute for financial support. The author acknowledges Dr Baharvand for his helpful comments and promotion of this work.

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Part 5

Nanotissue Engineering – Biological Approach along with Differentiation

Chapter 14

Expansion of Stem Cells by Nanotissue Engineering Amir Mellati and Hu Zhang

School of Chemical Engineering, The University of Adelaide, Adelaide, South Australia, Australia

Introduction Tissue engineering is “an interdisciplinary field that applies the principles of engineering and the life sciences toward the development of biological substitutes that restore, maintain, or improve tissue function” [1]. There are three key components for regenerating functional tissues in vitro: cells, cell-supported matrices, and bioreactors for providing a cell expansion environment. Cell sources in tissue engineering may be tissue-specific cells or stem cells. These cells may also be autologous or allogeneic. Stem cells have been widely used in tissue engineering due to their ability for self-renewal in an undifferentiated state for a prolonged time and for multilineage differentiation after appropriate stimuli [2]. There are three broad types of stem cells: embryonic stem cells (ESCs), adult stem cells, and induced pluripotent stem cells (iPSCs). In order to be used for tissue engineering applications, large numbers of high-quality stem cells are needed. It is a necessary requirement to develop a culture system to produce pure populations of tissue-specific stem-cell numbers in vitro without the loss of stem-cell potential. Cell culture in two dimensions has been routinely and diligently undertaken in thousands of laboratories worldwide for the past four decades. In two-dimensional platforms, cells are typically exposed to a solid, rigid flat surface on the basal side and to liquid at the apical surface. Inhabiting such a two-dimensional rigid substrate requires a dramatic adaption for the surviving cells because they lack the extracellular matrix that is unique to each cell type,  and which may alter cell metabolism and reduce its functionality. Furthermore, two-dimensional systems cannot provide a complex and dynamic microenvironment for stem cells, thus leading to spurious findings to some extent by forcing cells to adjust to an artificial flat and rigid surface. The unique function of stem cells inside the human tissues might be understood by a specific, three-dimensional microenvironment. The microenvironment effective parameters consist of soluble factors, extracellular matrix, cell-to-cell interactions, and external mechanical and electrical forces. When stem cells are removed from their niche, their ­functionality, phenotype, and responses to environmental signals can often be affected [3].

Stem-Cell Nanoengineering, First Edition. Edited by Hossein Baharvand and Nasser Aghdami. © 2015 John Wiley & Sons, Inc. Published 2015 by John Wiley & Sons, Inc.

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For example, muscle stem cells grown on the tissue culture flask have been found to lose their regenerative potential [4]. This has led to the assumption that mimicking the stem-cell niche will assist stem-cell proliferation and controlled differentiation in vitro [3, 5–7]. Three-dimensional cell culture systems may create a biomimicking microenvironment for stem cells. Designing such a system requires a multidisciplinary approach and ­expertise. Advanced biomaterials have significantly contributed to three-dimensional cell culture systems in recent decades, and more unique and complex biomaterials have been proposed for improving stem-cell proliferation and controlled differentiation. Among them, nanostructured biomaterials are of particular interest because they have the advantageous feature of a high surface-to-volume ratio, and they mimic the physical and biological features of natural extracellular matrix (ECM) at the nanoscale. In this chapter, we will review nanostructured biomaterials developed for stem-cell-based tissue engineering. The scope of this review is focused on infiltration, adhesion, migration, and proliferation of stem cells within nanostructured biomaterials. Stem-cell differentiation into specific lineages due to nanoscale features is beyond the scope of this review. Design principles of nanostructured biomaterials will be discussed first, followed by the three ­different categories of these materials: nanofibers, nanocomposites, and nanostructured surfaces.

Design Principles of Nanostructured Biomaterials Except for the basic properties of biodegradability and biocompatibility, three-dimensional artificial matrices for facilitating stem-cell expansion and maintaining their stemness potential should be able to: promote cell adhesion and proliferation; provide appropriate mechanical, chemical, and biological features mimicking native ECM; facilitate efficient mass transfer within matrices for nutrients, gas, and metabolic wastes; and build an effective signal transduction for stem cells. Therefore, the matrices for tissue engineering will be able to match the properties of tissues in the macro-level, and also to recreate the nano features observed in the native tissues. Consequently, design criteria for three-dimensional artificial matrices must consider length scales that span macro-, micro-, and nanodimensions. Stem cells are highly sensitive to their surrounding niche and provide an answer to environmental cues at all length scales from macro- to nanometers. Macroscale structure is important to make the desired shape and size of the defect site, and to offer enough mechanical strength for tissue formation. Microscale level of scaffold design is usually related to tissue architectures. For example, oriented parallel fibers are beneficial for reconstructing peripheral nerves [8], while random nonwoven fibers may be more important for dermal replacement [9, 10]. Microstructural features are also essential for ensuring cell adhesion and migration and determining the overall mechanical properties [11]. Particular attention has been drawn to pore size, connectivity, and geometry of three-dimensional matrices [12]. Pore size should be relatively large because most of the adherent stem cells have a size ranging from 10 to 150 μm. A large-pore structure allows delivering a sufficient number of cells and cell migration, while the interconnected porosity offers efficient diffusion of nutrients and metabolic waste removal [13, 14]. Nanoscale elements of native cells and tissues have become more important for understanding stem-cell behaviors. Especially, nanofeatures of the ECM have a large impact on stem-cell adhesion, migration, proliferation, and controlled differentiation. Figure  14.1 summarizes the benefits of nanostructured matrices as a stem-cell microenvironment. The ECM consists of structural protein fibers (collagen and elastin), adhesive proteins (fibronectin and laminin), glycosaminoglycans (GAGs), and proteoglycans (PGs). These proteins

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Nanostructured matrice

Hierarchical organisation at nano-level

Mimicking ECM components’ dimensions

High surfaceto-volume ratio

Cell adhesion

Pore size

Cell migration

Enhanced proliferation rate

Nutrient delivery and waste removal

Controlled differentiation

Tissue acceptance after implantation

Minimum materials

Mechanical properties

Topographical properties

Figure 14.1  Advantages of nanostructured matrices for regulating stem-cell behavior.

have dimensions between 10 and several hundred nanometers [15]. The ECM also takes a variety of forms in different tissues and at different stages of development in the same tissue [16]. The nanoscale ECM plays a significant role in signal transduction. Stem cells respond to the ECM through membrane receptors that connect the matrix to the cytoskeleton. Integrin is one of the main receptors that can recognize arginine-glycine-aspartate (RGD) sequences of ECM proteins. When integrins bind to ECM proteins (such as fibronectin or vitronectin), the integrin ligation activates focal adhesion kinase (FAK) signaling. The activated FAK undergoes autophosphorylation and triggers a downstream signaling of extracellular signal regulated kinase (ERK)/mitogen activated protein kinase (MAPK) pathway to transfer mechanical information to the cell nucleus, where the cell DNA reacts to extracellular stimulus by producing changes such as proliferation and differentiation to cells. The Ras homolog gene family, member A (RhoA)/Rho associated coiled-coil protein kinase (ROCK) pathway can also be triggered by activated FAK which can subsequently activates the ERK/MAPK pathway to influence stem cells. A schematic description of this process is shown in Figure 14.2. The signaling pathways may be regulated from different ECM protein composition, density and distribution of ECM ligands. Through these pathways, cells detect and respond to the mechanical, chemical, and biological characteristics of their surrounding microenvironment [2]. A variety of strategies have been formulated for engineering ECM ligands, such as RGD, in artificial three-dimensional matrices through self-assembly of peptides and surface modifications. The ECM nanotopology also regulates stem-cell behavior, ranging from changes in cell adhesion, cell orientation, cell motility, surface antigen display, cytoskeletal condensation, activation of tyrosine kinases, and modulation of intracellular signaling pathways that regulate transcriptional activity and gene expression [17]. The nanotopology may include dimensional scale from nanometer to micrometer and types of topological forms such as ridges, steps, grooves, pillars and pits [17]. Structural ECM features, such as fibrils and pores, are often of a size compatible with cellular processes involved in migration, which may influence the strategy by which cells migrate through the ECM. This size ratio between cell (1–100 μm) and surrounding fibers (1–100 nm) enables the cell to be in direct contact with the fibers of the ECM thus creating a three-dimensional orientation [15]. As a result, the biophysical characteristics of ECM affect various cell functions, such as adhesion and migration [2]. Nanofibers, nanorods, nanotubes, and other nanotopologies have been created to mimic the ECM nanoscale features.

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Proliferation differentiation Nucleus

ERK/MAPK K FA P

Stem cell Integrins

RhoA/Rock

α β α β α β

Proteins e,g., fibronectin, vitronectin etc.

Nanomaterial scaffold

Figure 14.2  Schematic pathways involved in the interaction of stem cells and nanostructured matrices. When integrins on the cell membranes bind to nanostructured ECM components, cascades of signal transduction occur for translating physical contact with nanostructures into biological responses, such as cell morphological change, proliferation, and differentiation. Reprinted from [2] with permission from Elsevier.

Nature manages to make structures with the minimum amount of essential chemicals. The natural ECM includes less than 1% solid materials, yet they are mechanically strong and have various functionality. Nature regulates the mechanical characteristics of biological tissue by fine adjustments of its composition with an alteration of its nanoscale structure from molecular level up to macroscopic scale [18]. Therefore, investigators have tried to engineer artificial matrices that resemble the nanoscale features of the natural ECM and current literature strongly supports nanostructured matrices due to their greater cell attachment and phenotypic activity than macroscopic matrices [19]. The matrices with nanoscale features can be categorized into nanofibers, nanocomposites, and nanostructured surfaces.

Nanofibers Nanofibers, with diameters ranging from 1 to 1000 nm, are the most popular nanostructured biomaterials that have been widely used in tissue engineering, due to the similarity of diameter size scales between nanofiber structures and ECM fibers and large surface area, which is favorable for cell adhesion and bioactive factor encapsulation. There are three commonly used methods to produce nanofibers: electrospinning, self-assembly and phase separation [20], and the fiber composition, alignment, diameter, degradation, and mechanical characteristics can be controlled for different types of tissue regeneration. Fibers have been fabricated for stem-cell expansion using self-assembly peptides and a range of polymers such as poly(ε-caprolactone) (PCL), poly(l-lactic acid) (PLLA), poly( d, l-lactide-co-glycolide) (PLGA), and other synthetic or natural polymers and also their blends or copolymers (Table 14.1).

Poly(ε-Caprolactone) Poly(ε-caprolactone) is widely chosen as a Food and Drugs Administartion approved model polymer due to its low toxicity, low cost, and ease of fabrication. Disadvantages of unmodified PCL are slow degradation rates (weeks to months), weak mechanical properties, nonreactivity,

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Table 14.1  Nano-fibres for stem-cell expansion Category

Nanomaterial

Stem cell

Significance/result(s)

Reference(s)

Synthetic polymers

PCL

MSC

SEM images showed good cell attachment

[21]

MSC

Successful 7 days cell proliferation before adding differentiation medium

[19]

MSC

Surface modification led to improved cell adhesion and proliferation

[23]

CSC–MSC

The greater the scaffold thickness, the more the cell proliferation

[22]

ESC

ESCs successfully cultured

[24]

SSC

SSCs successfully cultured

[25]

NSC

NSCs successfully cultured

[26]

PCL nano/ microfiber

MSC

Multilayer of nano- (for higher ECM mimicry) and micro- (for better cell migration) fibers

[27]

PCL nano-fiber/ PCL strands

MSC

Combining good architecture control of strands with good cell attachment of nanofibers

[29]

Poly(lactic acid)

NSC

Cell adhesion is near TCPS

[31, 32]

MSC

Microfiber constructs: cells were aggregated within the larger interfiber space Nano-fiber constructs: cells spread across multiple nanofibers

[36]

TSC

Effect of fiber orientation on cell morphlogy and orientation Increased cell proliferation

[37]

MSC

Nano-fibers increased proliferation rather than microfibers

[44]

MSC

Good proliferation and morphology Retaining differentiation abilities after 14 days culture (prior to differentiation)

[43]

ESC

ESCs successfully cultured

[39, 40]

NSC

High cell attachment on nanofiber surfaces

[45]

PLGA/collagen

HSC

Rapid and rich cell attachment

[47]

Polyurethane

MSC

Enhanced attachment and proliferation

[48]

Polyamide

ESC

Enhanced attachment and proliferation

[49]

PES

HSC

Surface-aminated nanofibers enhance adhesion and expansion

[50]

PES

HSC

Aminated chain length is important factor

[51]

Self-assembled RAD16 peptide and its motifs

ESC

ESCs successfully cultured

[62]

NSC

NSCs successfully cultured

[64]

hASC

hASCs successfully cultured

[63]

Self-assembled multi domain peptide

MSC

Promoted cell adhesion, migration and expansion

[65]

Self-assembled peptideamphiphiles

NSC

NSCs successfully cultured

[66]

MSC

MSCs successfully cultured

[67–69]

PLGA

(Continued )

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Table 14.1  (Continued ) Category

Nanomaterial

Stem cell

Significance/result(s)

Reference(s)

Natural polymers

Collagen

MSC

MSCs successfully cultured

[52]

Silk/PEO

MSC

PEO was added to improve processability; Improved adhesion and proliferation

[53, 54]

Other modifications

PCLnano-fiber/ PLLA nano-fiber

ESC–MSC

Cells infiltrated into the scaffold rather than migration along the surface; Enhanced proliferation

[60]

P(LLA-CL)

SMC

Enhanced proliferation

[55]

Starch/PCL

MSC

Example of blending with natural materials; combination of micro- and nanofibers Better morphology and cell growth

[61]

P(EOT-BT)

MSC

Microfibers with nanopores are successful for cell adhesion and proliferation

[59]

MSC, mesenchymal stem cell; PCL, poly(ε-caprolactone); CSC, carcinoma stem cell; ESC, embryonic stem cell; SSC, somatic stem cell; NSC, neural stem cell; TCPS, tissue culture polystyrene; TSC, tendon stem cell; PLGA, poly(d, l-lactide-co-glycolide); HSC, hematopoietic stem cell, PES, polyethersulfone; hASC, human adipose stem cell; PEO, poly(ethylene oxide); P(LLA-CL), poly (l-lactic-co-ε-caprolactone); SMC, smooth muscle cell; P(EOT-BT), poly(ethylene oxide terephthalate)–poly(butylene terephthalate).

and hydrophobicity. Intensive attempts have been made at fabricating PCL fibers for tissue engineering based on mesenchymal stem cells (MSCs) [19, 21–23], ESCs [24], somatic stem cells (SSCs) [25] and neural stem cells (NSCs) [26]. In 2003, Yushimoto et al. explored PCL nanofiber matrices for expanding MSCs. Penetration of cells and abundant ECM were observed in the cell-fiber constructs after 1 week. Scanning electron microscopy (SEM) images showed that the surfaces of the constructs were covered with several layers of cells by the fourth week [21]. Ruckh et al. further quantified cell growth along with live cell imaging. After a short-term (7 days) culture of MSCs in PCL nanofibers, live cell fluorescence staining and MTT assay showed significantly higher proliferation of MSCs on nanofibers than two-dimensional control surfaces. The SEM analysis also supported the fluorescence microscopy results that the MSCs preferentially adhere, spread, and colonize on nanofiber matrices compared to two-dimensional surfaces [19]. The physical parameters of PCL nanofibers, such as diameter of the fibers and morphology of the fiber surface, can influence cell attachment and growth. This has been confirmed by a study using mouse ESCs (P19) and mouse MSCs. Matrices with a thickness of 0.6 mm were found to provide a better substrate for cell proliferation rather than scaffolds with a thickness of 0.1 mm, possibly due to more dimensional stability [22]. To demonstrate that nanofiber surface modification affects stem-cell behavior, the surface of the PCL nanofibers was modified by He+ irradiation, which led to a slight smooth surface and different nanoscale surface chemical structures. The results showed that early attachment, further proliferation as well as osteoblastic markers, were higher for MSC on He+ irradiated PCL [23]. One of the drawbacks of nanofiber matrices is their small pore size, which results in poor cell infiltration and migration. To capitalize on the properties of microfibers (i.e., pores large enough for cell migration) and nanofibers (i.e., physical mimicking of native ECM), multilayered matrices can be fabricated to increase the pore size for cell migration. Mesenchymal stem cells were attached well on both single and bi-layered matrices but were more spread

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251

when nanofibers were present. However, increasing the thickness of the nanofiber layer reduced the infiltration of cells into the matrices [27]. The pore size of a fibrous structure can be controlled with melt-plotted strands from computer-aided design (CAD) and computer-aided manufacturing (CAM) technologies. In this method, the melted polymer is plotted with a 250-mm dispensing needle tip, laid down layer by layer. The resulting fabricated matrices have smooth strand surfaces and large pore size between the strands. These characteristics limit the initial cell adhesion. To overcome this disadvantage, micro-/nanofiber electrospun with PCL were layered between microsized PCL strands [28]. The cell attachment was further improved by two natural biomaterials (small intestinal submucosa (SIS) and silk fibroin). Bone-marrow-derived rat MSCs revealed an incredible increase in initial cell attachment and cell expansion on the three-dimensional hierarchical PCL fibrous matrices modified with two natural biomaterials [29].

Poly(l-Lactic Acid) Poly(l-lactic acid) is a biodegradable, biocompatible polymer and it has better thermal processibility than other biopolymers such as poly(ethylene glycol) (PEG) and PCL. However, unmodified PLLA has limited applications in tissue engineering due to its poor toughness, very slow degradation (more than 3 years), relative hydrophobicity, which results in low cell affinity, and lack of reactive side-chain groups [30]. One modification for PLLA is to fabricate PLLA nanofibers using a liquid–liquid phase separation method [31]. This method can create nanofibrous matrices with controlled and carefully designed macroporous architecture. However, the nanofiber diameter is not adjustable in this method. An electrospinning method has been exploited to produce PLLA nanofibers with variable diameters. A liquid–surface separation method may produce nanofibrous matrices with soft surface, while electrospinning methods may result in nanofibers with increased surface roughness. Results have shown that the NSC attachment was better on a surface fabricated using the electrospinning method [32, 33], since the greater the roughness, the better the cell adhesion [34, 35]. However, increasing the roughness will also lead to an increase in the hydrophobicity, which impels the nutrients from pores. Micro- and nanosized PLLA fibrous matrices have also been fabricated in order to study the effect of architectural characteristics on cell spreading, migration, and proliferation. The microfibrous meshes with a large pore size enhanced cell aggregation while small-pore nanofiber structures presented a spread, spindle-shape morphology. Cell attachment may be higher on the nanofiber scaffolds because the fibers were highly packed in matrices. By increasing the fiber diameter size, cells were aggregated within the larger interfiber distance/ pore space rather than spread across multiple fibers [36]. Poly(l-lactic acid) nanofibers have also been fabricated to have either aligned or randomly oriented structures. Human tendon stem/progenitor cells (hTSPCs) were seeded onto both nanofibers. They adhered very well and the cell number was increased about threefold in 14 days for both nanofibers. However, histological staining and confocal images (Figure 14.3) showed that hTSPCs were spindle-shaped and well orientated on the aligned nanofibers [37], demonstrating that nanofibrous-aligned structure can influence stem-cell morphology and orientation.

Poly(d, l-Lactide-co-Glycolide) Poly(d, l-lactide-co-glycolide) is a biocompatible and biodegradable linear copolymer that can be prepared at different ratios between its constituent monomers, lactic and glycolic acid. Poly(d, l-lactide-co-glycolide) offers superior control on its degradation by varying the

(A)

(C) Random (6w)

H&E

Aligned (6w)

(D)

(E)

(F)

Mawsson

(B)

Figure 14.3  Effects of nanofiber alignment on human tendon stem/progenitor cells (hTSPCs) orientation and morphology. Well-oriented stem cells in an aligned nanofiber matrix at week 6 is shown by (A) H&E (hematoxylin and eosin) staining and (B) Masson (Masson trichrome) staining; random orientation of cells on a nonaligned (random) nanofiber matrix is shown by (C) H&E staining and (D) Massion staining; (E) stretched CFDA-stained hTSPCs on aligned nanofibers; and (F) spreading-like morphology of hTSPCs on the randomly oriented matrix. Scale bars: 20 μm (A–D) and 50 μm (E and F). Reprinted from [37] with permission from Elsevier. (See insert for color representation of the figure.)

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ratio between its monomers [38]. Poly(d, l-lactide-co-glycolide) nanofibrous matrices have been used to culture ESCs [39, 40] and MSCs [41–43]. Poly(d, l-lactide-co-glycolide) nanofibers promote stem-cell attachment and growth ­compared to two-dimensional culture systems. Mesenchymal stem cells were seeded onto PLGA nanofibers and supplemented with normal medium (without any differentiating ­supplements) for 14 days. Live/dead assays showed stem cells remained viable after 2 weeks. Progressive cell numbers in a DNA quantification assay revealed the ability of PLGA nanofibers to accommodate stem-cell proliferation [41, 42]. The SEM and confocal images ­suggested that MSCs were attached to nanofibers but they have the same elongated shape after 14 days as that in two-dimensional culture. These stem cells retained their ability to differentiate into chondrogenic or osteogenic lineage after 7 and 14 days, as indicated by histological staining [43]. Hybrid nano-micro-fibrous PLGA matrices were compared to knitted microfibrous PLGA. Hybrid nano-micro-fibrous matrices were prepared by electrospinning PLGA nanofibers onto the surfaces of the knitted matrices. The MSC seeding efficacy was slightly higher for knitted matrices and cell attachment was comparable. However, cell proliferation was faster for hybrid matrices, as cell population increased by 92% between days 2 and 7, while it was 21% for knitted matrices [44]. More morphological structures of PLGA matrices have been examined for cell attachment, including nanofibers, microfibers, aligned microfibers and PLGA films, with more C-17.2 NSCs being attached on the surface of PLGA nanofibrous matrices [45]. Poly(d, l-lactide-co-glycolide) nanofibrous matrices have also been explored for culturing bone-marrow-derived hematopoietic stem cells (BM-HSCs), which are conventionally cultured in suspension using special spinner flasks or stirred bioreactors because of their nonadherent property. Therefore, due to the lack of close cell–cell and cell–matrix interaction, this culture system could not maintain cell localization to the specific environment [46]. An electrospinning technique was employed for fabricating nanofiber matrices with PLGA blended with collagen type I. The matrices were further coated with E-selectin, a critical adhesive biomolecule. The BM-HSC capture efficiency significantly increased from 23.40 to 67.41% within 30 min and from 29.44 to 70.19% within 60 min of incubation at room temperature after blended nanofiber matrices were coated with E-selectin [47].

Other Synthetic Polymers Other synthetic polymers have also been explored for stem-cell expansion. Polyurethane (PU) nanofibers have been fabricated and integrated into the microfluidic chips to mimic vascularized tissues embedded in ECM nanofibers, due to their strong mechanical properties. The MSCs attached and expanded on PU nanofibers. Cell adhesion and proliferation was further enhanced by acrylic acid grafting to PU nanofibers to decrease the PU surface hydrophobicity [48]. However, nonbiodegradability is a major obstacle to the use of PU in tissue engineering. Three-dimensional nanofibrillar organization of polyamide nanofibers resembles the ECM/basement membrane. Proliferation and self-renewal of mouse ESCs on this nanofibrillar surface were greatly enhanced in comparison with tissue-culture surfaces without nanofibers. In addition, stem cells cultured on the three-dimensional nanofibrillar surface maintained their differentiation ability in the presence of differentiating factors [49]. Surface-aminated polyethersulfone (PES) nanofibers were found to enhance the expansion rate of hematopoietic stem/progenitor cells (HSPCs) from human umbilical cord blood. The HSPCs are multipotent cells that can proliferate and differentiate into all blood cell types 1 and 2. However, achievable HSPCs from umbilical cords are very low because of the

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small volume of blood and it restrains direct transplantation of HSPCs to patients. Surfaceaminated polyethersulfone nanofibers have been demonstrated to be one approach to expand HSPCs. Scanning electron microscopy images revealed that cell colonies were formed on nanofibers [50] and the chain length of the grafted amines have an impact on the proliferation rate of HPSCs [51].

Natural Polymers Natural polymers contain essential components of natural ECM and they have been fabricated into nanofibers for culturing stem cells. For example, collagen is found in abundance in natural ECMs. Type I collagen nanofibers by electrospining technology have been ­prepared for examining the morphology, growth, adhesion, cell motility, and osteogenic differentiation of human bone-marrow-derived MSCs. The MSCs grown on 500–1000 nm nanofibers showed significantly higher cell viability than on a two-dimensional surface [52]. Silk is another popular natural polymer to synthesize nanofibers. Silk nanofibrous mats with fibroin diameter 700 ± 50 nm were found to support extensive MSC proliferation and matrix coverage [53, 54].

Copolymers/Blends Great efforts have been made to modify polymeric nanofibers using copolymer electrospinning or blending with other polymeric materials in order to improve processability of polymers for nanofiber manufacturing, in order to resemble the natural ECM as much as possible, and to promote stem-cell interactions with matrices. Aligned poly(l-lactic-co-ε-caprolactone) [P(LLA-CL)] co-polymer nanofibers were electrospun for growth of human coronary artery smooth muscle cells (SMCs). Cell adhesion to the copolymer nanofibers was quite similar to a two-dimensional surface, while SMCs’ proliferation rate on nanofibrous matrices was twice as fast as on a two-dimensional surface in 7 days [55]. Poly(ethylene oxide terephthalate)–poly(butylene terephthalate) (PEOT/PBT) nanofibers were also explored for culturing stem cells due to their adjustable surface energies [56]. Higher surface energy (hydrophilic) material leads to a higher cell attachment with a spindle-like shape whereas lower surface energy (hydrophobic) material results in lower cell attachment and a rounded morphology [57, 58]. Recently, Moroni et al. found that nanoporous PEOT/PBT microfibers promoted MSC spread, attachment, and proliferation, while smooth microfibers without nanopores led to rounded aggregated cells [59]. Blending nanofibers with different polymer supports can form a three-dimensional network and help cell migration. An aligned nanofibrous mesh essentially behaves as a two-dimensional sheet on which cells can only migrate along the surface, rather than a three-dimensional matrix in which cells are capable of infiltrating. To overcome this problem, a novel three-dimensional unwoven macroporous nanofibrous (MNF) matrix was manufactured from PLLA and PCL (w/w 9:1) using an electrospinning-based yarn assembly technique. Human MSCs derived from ESCs were seeded onto the MNF matrix and a much higher cell proliferation were observed [60]. Blending with natural polymers is another attractive strategy to mimic natural ECM. Stem-cell responses to the blending matrices have been studied extensively through hybrid nano- and microfibrous matrices produced by blending starch and PCL (30/70 wt%). Microfibers were impregnated, as much as possible, with electrospun nanofibers. Bonemarrow MSCs growing on the hybrid matrices presented a different morphology, being able to bridge between microfibers. Cells along the nanofibers showed a much-stretched morphology. When cells stretch themselves, the receptors are also stretched and activated,

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which leads to gene expressions different from unstretched cells. Increasing metabolic activity and proliferation rates were seen for cells on hybrid matrices. When the diameter of the fibers is lower than the cells, they can adhere well around the fibers and organize themselves. In addition, filling large spaces of the microfiber meshes with nanofibers will result in higher cell-seeding efficiency because more cells could be retained inside the structure [61].

Self-assembled peptides Self-assembly is to fabricate nanofibers through weak noncovalent interactions from small molecules, proteins, peptides, and nucleic acids [15]. Several peptides such as short fibrillizing peptides, β-hairpins, peptide-amphiphiles, and peptide derivatives self-assemble to form networks of β-sheet-rich nanofibers, which further merge to build supramolecular hydrogel architectures for tissue engineering application. For example, the peptide RADA16-I (AcN-RADARADARADARADA-CONH2), which is an alternating 16-residue peptide with basic arginine, hydrophobic alanine, and aspartic acid, can undergo spontaneous assembly into well-ordered interwoven nanofibers in water and rapidly form hydrogels with ~10 nm fiber diameter, 5–200 nm pore size and > 99% water content under physiological conditions, which is similar to the structure of natural ECM. This mild cross-linking chemistry allows yielding viable encapsulated cells for three-dimensional culture. This self-assembled peptide and its several different functional motifs with other short peptide sequences have been used as a well-defined microenvironment for stem cells. Mouse ESCs encapsulated in RAD16 showed undifferentiated stem-cell maintenance [62]. Functioning peptide mixtures RAD/SKP (Ac-(RADA)4GGSKPPGTSSCONH2), RAD/FHR (Ac-(RADA)4GGFHRRIKA-CONH2) and RAD/PRGD (Ac-(RADA) 4GPRGDSGYRGDS-CONH2) onto three-dimensional matrices have been demonstrated to control human adipose stem cell (hASC) behaviors in vitro [63]. Modified RADA16 with bone marrow homing peptides (BMHPs) has also been successfully applied to NSCs [64]. Multidomain peptides (MDPs) are a type of amphiphilic self-assembling peptides with a modular ABA block design in which the amphiphilic B block drives self-assembly while the side A blocks, which are electrostatically charged, control the conditions of assembly procedure. Galler et al. synthesized a range of multidomain peptides for fabricating nanofibers. The peptides were modified with enzymatic cleavage and a supplement of cell adhesion motifs (RGD). Combination of these items led to an increase in MSC viability and proliferation and encouraged cell migration within the matrix [65]. Peptide-amphiphiles (PAs) are oligo-peptides including a hydrophobic N-terminal alkyl tail, a β-sheet-forming segment in the middle, and a hydrophilic C-terminal functional segment for increasing cell adhesion [20]. These molecules generally self-assemble into highaspect-ratio rods/cylinders with a hydrophobic core and a hydrophilic region on the exterior of the fiber for cell interaction. The exterior region can be tailored to provoke desired chemical and biological responses, such as cell adhesive ligands and cell-mediated degradable sites to control cellular behaviors [66]. Mesenchymal stem cells [67–69] and NSCs [66] have been incorporated into the PA nanofibers for proliferation and differentiation.

Nanocomposites Although nanofibers have gained wide application to support attachment and proliferation of stem cells, a macroporous structure and weak mechanical properties are key limiting factors for specific tissue regeneration. The nanofibers can be blended with nanoparticles,

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Table 14.2  Nanocomposites and surface modified nanostructures for stem-cell expansion Category

Nanomaterial

Stem Cell

Significance/Result (s)

Reference(s)

Nano-composites

HAp nanorods/ PVP nanofiber composite

MSC

Nano-rods in nanofibers Cells were attached

[70]

PCL/PLLA/nHAp nanofibers composite

SSC

Nanoparticles in nanofibers Not cytotoxic

[71]

PHBV/nHAp nanofiber composite

MSC

Nanoparticles in nanofibers Proliferation lower than PHBV nanofibers

[72]

PLGA/Collagen/ nHAp nanofiber composite

MSC

Nanoparticles in nanofibers Good adhesion and spreading

[73]

PCL/nHAp/TCP nanofiber composite

MSC

Nanoparticles in nanofibers Cell proliferation is inversely proportional to nanofiber roughness

[74]

PEG/silica nanoparticles

MSC

Nanoparticles in gel Cells are alive and good proliferation

[76]

Multiwalled carbon nanotubes in PLLA

MSC

Nanotube in nanofiber Cells are alive, attached and proliferation is higher Makes conductive fibers

[75]

PGA nanofiber sheet/collagen

CSC

Nanosheet in sponge Enhanced cell attachment and proliferation

[77]

Titanium oxide nanotube

MSC

Nanotubes on surface Promote cell adhesion and proliferation without differentiation

[88]

(PS-b-P2VP)/ (PS-b-P4VP) nanopatterned surface

MSC

Nanopatterned surface Effect of nanotopography

[78]

Nanolayer of PNIPAAm

_

Nanoscale layer on surface Optimum cell adhesion and detachment depends on thickness of layer

[90]

Surface-modified nano-structures

HAp, Hydroxyapatite; PVP, poly (vinyl pyrolidone); MSC, mesenchymal stem cell; PCL, poly(ε-caprolactone); PLLA, poly(l-lactic acid); nHAp, nano-hydroxyapatite; SSC, somatic stem cell; PHBV, poly(3-hydroxybutyrateco-3-hydroxyvalerate); PLGApoly(d, l-lactide-co-glycolide); TCP, β-tricalcium phosphate; PEG, poly(ethylene glycol); PGA, poly(glycolic acid); CSC, carcinoma stem cell; PS-b-P2VP)/(PS-b-P4VP), poly(styrene)-blockpoly(2-vinylpyrindine) diblock copolymer/poly(styrene)-block-poly(4-vinylpyrindine) diblock copolymer; PNIPAAm, poly(N-isopropylacrylamide).

nanorods, nanotubes, and microfibers to form nanocomposites. Examples of these materials are described here and have been summarized in Table  14.2. The nanocomposites in the biological tissues provide a guideline for fabricating the nanocomposite matrices for tissue engineering. For example, bone tissue is composed of rigid hydroxyapatite (HAp) nanocrystals (tens of nanometers in length and width, 2–3 nm in depth) precipitated into collagen fibers (50–70 nm in diameter). The HAp crystals are one of the main constituents of bone tissues, and they provide compressive strength. To manufacture nanocomposites including HAp and nanofibers, Chen et al. prepared HAp nanorods with an average diameter of ~7 nm

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and length of ~27 nm and then the nanorods were blended with poly(vinyl pyrolidone) (PVP) to form composite nanofibers by electrospinning. The MSCs were well attached to the HAp fabric substrates after culture for 24 h [70]. The same approach has been used to electrospin aligned nanofibrous PCL/PLLA/nHAp (nanohydroxyapatite) composites, and human unrestricted somatic stem cells (USSCs) were seeded into the nanocomposites. Results showed that stem cells were viable in the matrices [71]. However, higher HAp content in the poly(3-hydroxybutyrate-co-3-hydroxyvalerate) (PHBV)/HAp nanocomposites decreased the MSC proliferation rate [72], while a blend of PLGA, collagen, and HAp nanoparticles was electrospun for fabricating nanofibers, which supported MSC adhesion and spreading [73]. Composites consisting of nanoparticles of 20% Hap and 80% β-tricalcium phosphate (TCP) and PCL were fabricated to demonstrate cell proliferation was inversely proportional to the nanofiber roughness [74]. Multiwalled carbon nanotubes (MWNTs) have also been explored for their potential to form nanocomposites. The MWNTs were encapsulated into PLLA nanofibers to provide three specific enhancements to fibrous tissue matrices: modified fiber size, electrical conductivity, and increased mechanical strength. Adipose-derived human MSCs were integrated into the PLA nanofibers with 1 wt% MWNT and were viable after day 14. The proliferation rate of the hMSCs increased drastically by day 14 and MWNTs in the nanocomposites promoted MSC proliferation [75]. As well as nanofibrous nanocomposites, nanocomposites can also be made from incorporation of nanoparticles or nanofibers into microporous hydrogels or macroporous sponges. A thixotropic polyethylene glycol (PEG)–silica gel was prepared by combining multi-arm PEG with hydrolyzed tetraethoxysilane (TEOS). The viscosity of thixotropic nanocomposites decreases under stress and returns to its original situation after stress removal. Good nutrient and gases delivery through the matrix improved proliferation and viability of MSCs over 3 weeks [76]. Electronspun nanofibers can also serve as fillers in the macroporous structure to form a new type of nanocomposite. Poly(glycolic acid) (PGA) nanofibers were formed as a sheet and added as a layer within the collagen sponge. The sheets noticeably improved the compressive strength of the collagen sponge. More cardiac stem cells (CSCs) were adhered to the collagen sponge with PGA nanofibers than with the sponge alone. Nanofibers also promoted cell ­proliferation [77].

Surface-Modified Nanostructures Introducing nanofeatured elements such as nanotubes or ultrathin layers on the surfaces or modifying their topography using nanopatterning techniques are other applications of nanotechnology in stem cell three-dimensional culture (Table 14.2). Nanopatterned surfaces with different topographies can be achieved through synthesis methods or introduction of nanostructures to the surfaces. For example, di-block copolymers of polystyrene and poly-2-vinylpyrindine or poly-4-vinylpyrindine formed dot-like (6 nm) or worm-like (3 nm) surface nanotopography, respectively, via controlled microphase separation. The worm-like surfaces supported greater human mesenchymal progenitor cell proliferation. More elongated cells and thicker ECM deposits were found on the worm-like surfaces [78]. Nanotopography of the surfaces can be changed by coating nanomaterials onto the ­surfaces. Nanotubes on the matrix surface improved cellular tracking, sensing of microenvironments, delivering of transfection agents along with scaffold enhancement [79]. Carbon nanotubes provided the needed structural reinforcement for tissue scaffolding [80–83] and

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mimicked some ECM fibers [84]. Nanotubes were placed in an array to simulate neural ­networks [85] and high electrical conductivity of the carbon nanotubes was found to be beneficial for directing cell growth [86]. Carbon nanotubes can also be functionalized to release bioactive factors. It has been shown that such factors, for example glucose oxidase, can be attached to nanotubes without losing the enzymatic activity [87]. Titanium oxide nanotubes have also been demonstrated to affect human MSC behavior. A dramatic change in MSC responses was found when they were exposed to various ranges of nanotube dimensions. Small (~30 nm diameter) nanotubes enhanced attachment without noticeable differentiation, while larger (~70–100 nm diameter) nanotubes stimulated a dramatic stem-cell elongation (~10 fold increase), which generated cytoskeletal stress and resulted in differentiation of MSCs into osteoblast-like cells [88]. Surface properties such as wettability can be tuned through surface modification. Introducing a thin layer of thermoresponsive polymers, poly(N-isopropylacrylamide) (PNIPAAm), imparts tunable hydrophobic/hydrophilic characteristics to the surface, which could be interchanged by temperature [89]. The thickness of the layer was found to play an important role in cell adhesion. The optimum thickness of the polymer on the surface was determined to be 15 nm. Beyond this thickness, the hydrophobic environments in its vicinity no longer promoted the dehydration and shrinkage of the PNIPAAm chains and were not suitable for cell adhesion [90, 91].

Conclusions Nanoscale artificial design of the stem-cell microenvironment is key to control stem-cell fate. Nanofibers from natural or synthetic materials, nanocomposites, and surface-modified nanostructures have been created to mimic the natural ECM nanoscale features, including presentation of ECM ligands, nanotopography, and cell–ECM interactions. Although these nanoscale features have been demonstrated to improve cell attachment and proliferation, how stem cells respond to the environmental cues has not been clearly understood. The challenge for biomaterial scientists will be to create more refining and subtle structures for mimicking nanoscale ECM configurations, and for stem-cell biologists to employ these nanostructures to explore cellular responses. Currently, nanostructures prepared by material scientists are confined to only cell attachment and proliferation, while there is lack of biological contributions for cellular-signal pathways and gene-expression profiles. Crossdisciplinary collaboration will elucidate the mechanism of the stem-cell microenvironment, and this will further provide greater insight for creating more functional artificial matrices for tissue-engineering applications. Most efforts in nanotissue engineering have focused on controlling stem-cell differentiation into desired tissues and very few reports have contributed to stem-cell self-renewal, especially in three-dimensional artificial matrices. Hydrogels and nanogels have been demonstrated as an excellent platform for stem-cell culture and expansion [92, 93]. However, nanoscale features in the ­hydrogels need to be further studied, and new composites, such as nanoparticles– hydrogels, nanofibers–hydrogels, and nanogels–hydrogels, need to be explored for stem-cell proliferation and directed differentiation. In addition, fine-scale assays should be developed, especially for highly hydrated microenvironment, to examine the exact interactions of stem cells and three-dimensional nanostructures in situ. Techniques for both probing cellular biological responses (gene expression and protein synthesis) and characterizing nanoscale features of artificial matrices are key limiting factors for revolutionizing nanotissue engineering. More efforts will be required for accelerating the development of these new techniques.

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Chapter 15

Nanotissue Engineering of Neural Cells Sasan Jalili-Firoozinezhad1,2, Fahimeh Mirakhori1,3, and Hossein Baharvand1

Department of Stem Cells and Developmental Biology at Cell Science Research Center, Royan Institute for Stem Cell Biology and Technology, ACECR, Tehran, Iran 2  Departments of Surgery and of Biomedicine, University Hospital Basel, University of Basel, Basel, Switzerland 3  School of Biology, College of Science, University of Tehran, Tehran, Iran 1 

Introduction Stem-cell niches form highly complex, dynamic microenvironments with numerous factors involved, in which cells must constantly interact and reply back in response to the state of the tissue. These specialized regions provide physical, structural, and architectural cues in the form of a specialized basal lamina and various extracellular matrix (ECM) components, which sequester growth factors and other bioactive molecules to modulate signaling-molecule availability [1]. Moreover, the presence of a different range of supportive cells (e.g., astrocytes, ependymal cells, and endothelial cells) and differentiated cells (e.g., different types of mature neurons) within stem-cell niches add further complexity. They act to integrate signals from within the niche environment and provide proper responses to modulate resident stem-cell behavior through a dynamic feedback loop [2]. Supporting and differentiated cells can also further modify the niches by their paracrine secretion, including growth/trophic factors, mitogens, morphogens, and differentiation cues [3–7]. Additionally, different conformations and densities of the matrix, matrix stiffness, and the location of cells within these niches can affect stem-cell behavior [8–11]. There has been much progress in the attempt to confirm the existence of neural stemcell (NSC) niches in regions of the adult central nervous system (CNS), such as the ­subventricular zone [2] and dentate gyrus [12]. However, individual components that form these constantly changing niches, specific cell–microenvironment interactions and the mechanisms by which these all interact together to modulate precise control of the NSC fate are still largely unknown. Therefore, for recapitulating a complex synthetic NSC niche environment, a much more comprehensive understanding of the exact nature of neuronal niches, including spatial organization and dynamic modulation of cells, soluble factors, and the matrix, is required [13]. However, recreating even an elegantly simple NSC niche is invaluable for in vitro NSC self–renewal, assessment for differentiation, and its potential for regenerative medicine. In this chapter we provide an extensive overview on ongoing efforts in nanotechnology related neural-tissue engineering. In the first section, different aspects of biological

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factors important for neural repair will be reviewed. In the second section, the effect of physico-mechanical cues will be summarized in terms of micro-/nanotopography, mechanics, and piezoelectricity of natural and synthetic substrates. We believe that this extensive overview of studies that examine the relation of cells to their physical and chemical microenvironment will identify some key bioengineering milestones necessary for successful nerve regeneration.

The Role of ECM Components/Engineering ECM Signals into Biomaterials The interplay between CNS cells and their surrounding ECM influences most aspects of ­nervous system development and function such as adhesion, survival, migration, proliferation, and differentiation of cells by providing a physical and biochemical cellular microenvironment [14–17]. The CNS microenvironment is primarily composed of a complex blend of glycoproteins, proteoglycans (such as collagens, laminins, and fibronectins), glycosaminoglycan (GAG) hydrated gel, other link proteins, and a stack of signaling molecules, growth factors, and morphogens [14, 18, 19]. This framework is dynamically created, secreted, transported, and depleted by CNS cells (e.g., glia cells), hence acellular allografts can be used as ideal ­scaffolds [19]. However, variations in biochemical and mechanical properties following the extraction process should be taken into consideration in order to prevent immunogenic responses and incomplete repair in foreign hosts, allograft-limited sources, imperfections in the chemical processes for removing epitopes/globular proteins and immune stimulating factors [20]. Potentially, a scaffold can be supplemented with ECM in the hope of generating neural-like constructs and providing structural stability. By doing so, desirable bioactivity that can induce a neural stem/progenitor’s fate toward lineages of interest is applied. Thus, three-dimensional ECM-based synthetic or natural implants with robust biomechanical properties and biocompatibility can be promising alternatives to allografts and autografts [20]. Natural scaffolds composed of collagen, Matrigel, laminin, or fibronectin have been fabricated to enhance neuron survival and neurite outgrowth in response to signals [20]. Interaction of fibronectin and laminin as the most important CNS basement-membrane components with specific cell-surface molecules (integrin receptors) initiates a cascade of signal transduction that lead to varied cellular responses [21]. The concentration and type of these components can affect neuronal cell behavior. For example, embryonic motor neurons (MNs) require laminin 2/4/8 to improve their neurite outgrowth. Additionally, implants made of natural materials have been shown to exert neuroprotective effects on an injured CNS and provide a favorable bridge for the regenerating axons [22, 23]. The CNS native environment is mainly composed of GAGs. Some GAGs such as chondroitin sulfate (CSPG) and heparin sulfate (HSPG) inhibit axonal outgrowth at the injury site [24, 25]. However, HSPGs such as syndecan-1 and glypican-1 participate in neural survival and regeneration via controlling the IGF-I interaction with their binding proteins on the neural surface [25, 26]. In addition to the above-mentioned materials, other ECM proteins such as growth-factorreduced Matrigel (gfrMG) have been shown to support neural cell survival, migration, and neurite outgrowth both in vitro and in vivo. This can promote proliferation of grafted cells and increase the number of tyrosine hydroxylase (TH)-positive dopaminergic neurons. The gfrMG increases the number of TH-positive cells by suppressing the inhibitory effects of the host-brain microenvironment on neural differentiation [27]. Thus, scaffolds based on natural materials that depend on the desired microenvironmental cues can be fine-tuned to properly mimic the particular neuronal-niche’s biophysical and

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biochemical traits. However, the precise role of natural matrices in this regard remains an oddity since they may induce immunological and inflammatory responses due to the presence of undefined components. On the contrary, synthetic biomaterials possess advantages in manufacturability and lower lot-to-lot variability, which permit us to more easily and precisely adjust key parameters of substrates such as architecture, stiffness, and electrical properties. Biofunctional cues can be added to the synthetic scaffolds to overcome their poor biocompatibility and lack of recognition signals. The role for this type of scaffold will be further discussed later in the ‘Biophysical’ section.

Role of Growth Factors, Signals, and Bioactive Molecules Protein Incorporation for Improving Cell Function Spatio-temporal cues such as growth factors, mitogens, neurotrophic factors, and morphogenes are biomolecules that are actively involved in the regulation of neural development, survival and function, and progenitor differentiation [28, 29]. In order to promote cell function, artificial biomaterials can be modified with several types of these bioactive molecules. Presentation of bioactive molecules to biological substrates also allows for the study of cellular internalization dynamics and roles of specific signal-transducing ligands in neural cell-fate decisions. Through introducing the bioactive factors to biomaterials, besides the enhancement of attachment, proliferation, and differentiation of transplanted cell, endogenous NSCs can be induced to proliferate and differentiate at the legion sites [20]. Such being the case, developing bioactive scaffolds with proper chemical properties can increase the value of neural tissue engineering in terms of regenerative medicine. The roles of neural master growth factors in this matter will be discussed.

Growth Factors During development, growth factors, or soluble-secreted polypeptides, can use their signal transduction ability to instruct cell behavior by binding to specific cell-surface receptors and trigger changes in the cytoskeleton, metabolism, gene expression, and protein synthesis. These changes result in specific cellular responses [30]. In order to properly affect cellular target receptors by diffusion, the soluble growth factors must be present for long periods of time. As the half-life of growth factors is short, the efficacy of delivery can be boosted through their incorporation within the bioengineered three-dimensional scaffolds [31]. However, undesired levels of delivered growth factor can lead to numerous side-effects due to overextension of neural projections beyond their specific targets, which are implicated in progressive neuronal death [32]. Adding growth factors into the scaffolds allows for control of their multiple functions, enables the desired delivery, and simultaneously presents multiple cues to make them highly favorable for neural-tissue regeneration. Protection against proteolysis in local protein-delivery systems can be achieved by their incorporation into materials such as heparin and increase the duration of their activity [33]. Immobilized bioactive molecules have enhanced biological activities compared to their soluble forms and can assist neuronal cell survival, proliferation, differentiation, and fate modification in vitro and in vivo [34]. For example, membrane proteins such as notch ligands or delta that are not activated by soluble ligands, can be immobilized for improved activities in vitro [35]. The most commonly used growth factors for neural cells are nerve growth factor (NGF), brain-derived neurotrophic factor (BDNF), glial-derived neurotrophic factor (GDNF), ­ciliary neurotrophic factor (CTNF), insulin-like growth factors (IGFs), epidermal growth factor (EGF), and fibroblast growth factors (FGFs).

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During CNS development, the FGF family has a number of varied roles that include ­ euronal survival, axonal growth, synapse formation, and nerve regeneration [29, 36, 37]. The n combination of EGF/FGF2 is usually employed for maintenance of neural or progenitor cells. The presence of EGF in culture media can ensure NSC proliferation and differentiation potential [38, 39]. The NGF, the first growth factor identified in 1951, is necessary for neurogenesis [40], controls neuronal differentiation and survival, directs axonal outgrowth during development, modulates neural plasticity in the mature CNS, and induces neuroprotective effects under pathological conditions by providing a trophic support to endangered host neurons [41]. The BDNF can promote neural survival by its neurotrophic effects by activation of ­tyrosine kinase receptors TrkB on axons/dendrites [42, 43] and the ERK signaling pathway [44]. Besides supporting neural survival in vitro, BDNF has also been shown to rescue MNs from cell death in vivo. The BDNF can induce proliferative effects on NSCs and stimulate the formation of differentiated cell types in different proportions [45]. Local co-delivery of GDNF and BDNF has been shown to significantly increase the number of MNs on chronic and acute axotomy, compared to BDNF alone [46, 47]. Incorporation of GDNF as microspheres in a poly(caprolactone) (PCL) conduit has been shown to significantly increase tissue integration.

Morphogenes During embryogenesis, morphogenes are the signaling molecules whose local concentrations affect how various specialized cell types within a tissue will develop with relation to their location in the body [48]. For example, retinoic acid (RA) and sonic hedgehog (Shh) are two distinct morphogenes that play a role in neural-tissue development and promote stem-cell differentiation into a neural lineage in vitro [49–51]. Additionally, RA in combination with FGFs and Shh control the onset of neurogenesis and contribute to dorso-ventral patterning [52]. During mammalian neurogenesis, neural precursor cells within the ventral neural tube come in contact with signaling molecules such as Shh to specify MNs [53]. The notochord underneath the neural plate and the floor plate at the ventral midline of the neural tube both produce Shh [54]. The endogenous neural precursor cell population has been shown to increase as a result of the neuroprotective effects of delivered Shh at the spinal cord injury site and additionally improve neuronal function and axonal guidance [55–57]. Co-administration of RA and Shh can create a favorable environment for motor neuron differentiation of stem cells [58, 59].

Small Chemical Molecules Compounds that modulate intercellular signaling mechanisms of the developmental pathways have the potential to be mediators of stem-cell self-renewal promotion and their differentiation to specific cell types. These compounds can also serve as the cell-fate regulators or the reprogramming factor of somatic cells. Small molecular-mass chemical entities or small molecules (SMs) are powerful tools in this regard as they selectively interact with biological macromolecules in a reversible and dose-dependent manner [60, 61]. Several of these SMs have been applied toward the regulation of neural cell fates in vitro and in vivo. For instance, Shh has been replaced with a smooth agonist small molecule (SAG) in order to generate MNs from mouse embryonic stem cells (ESCs) and to expand neuronal precursor cells in spinal cord injury animal models [62, 63]. In principle, SMs that mimic key fragments and functions of the mentioned macromolecules can incorporate into the scaffolds and embark on any of these events to substitute

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growth factors and morphogenes. Therefore, application of small chemical molecules that target signaling pathways such as Shh, Wnt, FGF, or notch can be useful in neural-tissue engineering.

Biosignaling Molecules (ECM, Growth Factors, and SMs) Presentation Biosignaling molecules can be physically adsorbed and/or deposited onto the biomaterial matrices through weak forces such as van der Waals forces, hydrogen bonding, or electrostatic interaction. For example, laminin, polylysine, and collagen have been used to modify poly(lactide-coglycolide) (PLGA) films [64], plasma-treated PLGA films and chitosan films [65], and poly(l-lactic acid) (PLA) nanofibers [66]. Despite its technical simplicity, the poor stability of biomolecule layer limits its usage in neural-tissue engineering. Blending ECM reagents with biomaterials is a simple alternative approach to direct their deposition that results in more stable and uniform distribution. For example, neural adhesion has been significantly improved by the use of a blend of chitosan and 3% weight polylysine [67]. Electrostatic attachment is based on electrostatic interactions between ECM molecules and biomaterials. It is similar to physical deposition and blending, and includes layer-by-layer (LbL) assembly and electrochemical polymerization. For instance, LbL films comprised of hyaluronic acid (HA)/collagen have been shown to promote cortical neuron adhesion on a nonpermissive substrate, such as a glass coverslip [68]. The advantages of these films include versatility and applicability to any charged substrate. The introduction of multiple linkages, chemical or photocoupling can improve the films’ stability. However, these methods are relatively complicated compared to former techniques and factors such as pH, polyelectrolyte loading, and ionic strength of the polyelectrolytes can affect composite stability. Two main strategies for growth-factor presentation in tissue engineering exist to improve its integration to biomaterials and prevent rapid diffusion, chemical immobilization, and physical encapsulation. Chemical immobilization to the biomaterials includes chemical cross-linking or affinity-based interaction of the growth factor to the polymer substrate and/or cells chosen for tethering proteins to solid substrates. Physical encapsulation includes the encapsulation, diffusion and pre-programmed sustained release of growth factor from a degrading substrate or dissolving delivery systems into the surrounding tissue microenvironment (Figure 15.1) [31]. For example, encapsulated FGF2 slowly diffuse from biodegradable acidic gelatin hydrogels in vivo, whereas it does not release from the gelatin in vitro [69]. Incorporation of biodegradable PLGA microspheres within a porous scaffold can assist with slowing the release of FGF-2 [70]. The immobilizations of a wide range of growth factors and other biomolecules to different natural and synthetic solid substrates can be performed by means of covalent or noncovalent chemical approaches. Noncovalent attachment includes direct hydrophobic interactions with excipient molecules through secondary interactions between growth factors and biomaterials or indirectly via intermediate proteins/biomolecules [71]. In this way, chemically or physically coated biopolymeric gels have been used as ECM-mimicking materials (e.g., heparin, fibronectin, laminin, collagen, and gelatin) [72–74]. Small oligopeptides mimicking key fragments such as RGD [71, 75] and a variety of synthetic hydrogels can provide specific biological binding sites that can tether growth factors, morphogens or their small molecule agonists into the area of interest. Several other techniques have been developed to overcome loose binding of ECM biochemical cues and/or growth factors to scaffolds. In this regard, covalent attachments that permit molecular escape via diffusion and provide more stable delivery systems are of great interest. These attachments include the use of heterofunctional cross-linkers such as thiol, amine, carboxylate, and hydroxyl to conjugate ECM components or growth factors to

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(A)

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Inactivated Diffusion of cell growth ? factor

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? Tissue regeneration

? ?

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Figure 15.1  Schematic illustration of two main strategies for growth factor presentation using synthetic ECMs, (a) Physical encapsulation. (b) Chemical immobilization [31].

biomaterials. In some cases they require pretreatment for surface functionalization by treatment with plasma or ionizing radiation graft copolymerization [76, 77]. 1-Ethyl-3-(3dimethylaminopropyl)carbodiimide (EDC) cross-linking is one of the most popular methods for covalent modification of neural biomaterials [66, 78].

Effects of Physico-Mechanical Cues Rationale for Micro- and Nanotopography Researchers have generated micro- and nanopatterning on biological substrates with the intent to appropriately mimic the complex structure of the ECM and introduce spatiotemporal controlling parameters to positively affect neural cell behavior. These substrates play a pivotal role in development of new therapies [79]. As with modulation of cellular function via intricate topographical characteristics in vivo, the application of synthetic topographical cues, cell polarization, proliferation, alignment, adhesion, and differentiation in vitro will be promoted to hopefully result in better nerve regeneration [80, 81]. Micro-/nanoscale structures of the human body are the leading hallmarks that help researchers to design and fabricate biomimetic materials. Figure  15.2 presents nanotopographical features of four types of human tissues [82].

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– Brain (aligned neurite connecting cerebral cortex and ventricular surface) – Skeletal muscle (aligned muscle fibers) – Heart (aligned myocardial muscle fibers) – Skin (organized ECM matrix fibers in dermis)

– Vesssel (nanofibrous matrices in basal lamina) – Bone (brick-and-mortar like structure in cortical bone) – Ligament (aligned and hierarchical collagen fiber bundles) – Tendon (aligned and hierarchical collagen fiber bundles)

Figure 15.2  Different nanotopographical forms in various human tissues [82].

To study the effect of different topographies and mimic the anisotropic structures of neural cells, techniques such as electrospinning and nanolithography have been used to design a number of micro-/nanostructures in the form of fibers or gratings [82, 83]. The behavior of nerve cells in the context of differentiation, neurite outgrowth, attachment, migration, and alignment has been investigated on these micro-/nanostructures. Microgrooved substrates guide the development, differentiation, and neurite extension of human NSCs related to the microchannel width. Small patterns not only induce proper neurite alignment but also induce longer neurite length, whereas larger micropatterns have been shown to hinder alignment [84]. Another study has researched the alignment and elongation of human pluripotent stem cells (hPSCs) along the micron-sized grating axis. While the cells exhibited random morphology on the unpatterned substrate, filopodia aligned greatly with the introduction of 2 μm gratings and neural markers highly expressed on this surface (Figure 15.3) [85]. Previously, it has been shown that by using 350 nm ridge/groove topographic surfaces, human ESCs (hESCs) could be differentiated to neurons without the introduction of chemical factors [86]. Recently, the effect of polydimethylsiloxane (PDMS) nanograting substrates with various widths on growth and differentiation of human-induced pluripotent stem cells (hiPSCs) showed that cellular contact-guidance enhanced considerably on the smaller pitch. The expression of neural markers significantly upregulated on grating with a width of 350 nm compared to the flat surface [87].

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(A)

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Figure 15.3  hPSC elongates along the axis of the 2-mm gratings. (A) The 2-mm gratings. (B) Elongated morphology is observed on 2-mm gratings. (C) Filopodia of hPSC on 2-mm gratings extend along the axis [85].

In terms of stem-cell engineering prospects, nanofibers support stem-cell expansion and neural differentiation, meanwhile they direct alignment of neural cells and enhance neurite outgrowth along aligned fibers [88]. The majority of nerve-tissue engineering studies have focused on investigating neural proliferation and differentiation, and neurite outgrowth of numerous cell types on aligned fibrillar substrates of various materials and sizes [89]. It has been proven in many investigations that compared to tissue culture polystyrene (TCPS), nanofiber topography provides more three-dimensional topographical cues that mimic the neural niche and induces morphological and functional changes for various cell types, including mesenchymal stem cells (MSCs). For example, PCL aligned nanofibers significantly upregulate MSC expression of neural markers (Tuj-1, MAP2, GalC and RIP) compared to two-dimensional substrates and induce MSCs to elongate along the axes of aligned fibers. Alongside the alignment of nanofibers, improving the hydrophilicity and mechanical properties of these constructs through blending with natural polymers enhances the proliferation, adhesion and orientation of Schwann cells (SCs) around the aligned fibers [90]. Aligned PCL

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nanofiber meshes are less receptive to adhesion and oligodendrocyte survival compared to random fibers, thus a higher fraction of adult NSCs (ANSCs) exhibit markers of neuronal differentiation on aligned fibers. Generally, surface alignment rather than fibrous topography alone is the key parameter that favors the yield of neuronal progenitors [91]. Other than fiber alignment, various studies have shown that fiber diameter affects cell adhesion, migration, proliferation, and differentiation. When compared to 930 nm (38%) and 260 nm (21%) random fiber substrates, 480 nm (47%) fibers have been shown to induce significantly higher neuronal differentiation of ANSCs [91]. Similarly, in a study, polyamide nanofibrous architecture with a mean fiber diameter of 400 nm was able to greatly induce differentiation of hESCs to neural progenitors (NPs) within 12–18 days and mature MNs within 36 days [92]. With the intent to determine the proper fiber diameter for committed differentiation of NSCs on laminin-coated polyethersulfonate (PES) electrospun nanofibers, three fiber meshes with mean diameters of 283 nm, 749 nm, and 1452 nm were fabricated. While NSCs showed a 40% promotion in oligodendrocyte differentiation, the 749 nm and 1452 nm substrates increased neuronal differentiation compared to TCPS. The 749 nm fiber mesh alone yielded the highest fraction of neuronal differentiation among the samples. In conjunction with decreased fiber diameter, cell proliferation, migration, and spreading increased considerably [93]. To investigate the simultaneous impact of fiber diameter and alignment, neurite extension and migration of SCs from dorsal root ganglia (DRG) explants were studied on different diameters of aligned poly(l-lactic acid) (PLLA) fibers. Neurite length on the small-diameter (293 nm) fibers was shorter than those on intermediate (759 nm) and large (1325 nm) diameter meshes (42% and 36%, respectively). Based on the neurofilament-stained image of DRG, it was evident that intermediate fibers had the longest neurites compared to other groups (Figure  15.4) [94]. While aligned small-size fibers (280 nm) induced NSC differentiation towards the neural lineage [82], 600 nm and 1.6 μm fibers were most effective in promoting SC differentiation compared to random fibers, which favored differentiation of these cells into oligodendrocytes [93, 95]. .

(A)

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Figure 15.4  Neurofilament-stained image of DRG on (A) large, (B) intermediate, and (C) small diameter fibers. Scale bar = 100 μm [94]. (See insert for color representation of the figure.)

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Substrate Mechanical Properties Biomechanical cues can be transferred to cells through micro-/nanoscale features of the ­substrates. Since neurons and glial cells reactively respond to mechanical signals, it is of utmost importance to gain adequate information regarding the mechanics of the nervous tissue microenvironment and its cellular constituents [96]. Numerous cell types in the n ­ ervous system can adapt their morphological properties, mechanical properties, and biological function based on their microenvironment stiffness. For instance, due to differences in stiffness between white and gray matter, neuronal and glial cells show different mechanical behavior [96]. Studies investigating the co-culture of neurons and glial cells have shown that neurons grow mainly on top of glial cells, which provide stiffness similar to brain tissue [97–100]. As a nonlinear viscoelastic tissue, the CNS does not raise internal stresses and it is also not exposed to mechanical forces. Therefore, the spinal cord and brain are devoid of fibrillar stiff ECM; they are mainly surrounded by soft hyaluronan and glycosaminoglycans, and exhibit a low elastic modulus [96]. By taking this into consideration, polymeric scaffolds fabricated from synthetic or natural-based biomaterials can be designed to consist of low mechanical properties that resemble native brain tissue [101]. Decellularized CNS ECM that is used as a soft biological scaffold has been shown to retain native biochemical and topographical features of in vivo conditions, and stimulate proliferation, migration, and differentiation of the PC12 cell line in vitro [102]. Neural tissues have stiffness values less than 100 kPa (native brain tissue modulus  2 weeks of monitoring, the impedance of noninduced cells leveled off as the plate reached confluency, while osteogenesis and adipogenesis, which were induced by commercially available differentiation mediums, were characterized, respectively, by a steady increase and gradual fall in Z*. More detailed statistical analysis indicated that the impedance of osteo-induced cells was significantly higher than that of the adipo-induced cells as early as 12 h post induction in a 4–16 kHz frequency interval and continued for the full 2-week differentiation period. Moreover, using a threeparameter model developed by Giaever and Keese [14], it was reported that this difference in impedance is due to the fact that: (i) osteo-induced cells established intercellular junctions over the course of 48 h after induction as opposed to ADSCs undergoing adipogenesis; and (ii) the cell-membrane capacitance of osteo-induced cells was significantly lower than that of ADSCs undergoing adipogenesis. Similarly, Reitinger et al. adopted a radiofrequency identification-based sensing platform to characterize the proliferation and differentiation of human bone-marrow-derived multipotent stem cells (bmMSCs) over periods from several days to weeks [15]. In particular, ECIS was achieved by fabricating electrodes comprised of two comb-shaped interdigitated gold electrodes onto glass substrates (placed in six-well culture plates) (Figure  19.1). Continuous measurements could then be collected using a sinusoidal voltage of 35 mV at a frequency of 10 kHz. In agreement with Bagnaninchi and co-workers, Reitinger et al. observed that impedance increased significantly as bmMSCs adhered and spread. In addition, upon differentiation into osteogenic or adipogenic lineages there was a steady rise (Figure  19.2A and B) and gradual fall in impedance, respectively (Figure  19.2C and D). Importantly, the authors emphasized that the differentiation could be validated in the bmMSC culture within a few days and monitored continuously for weeks at a time. In contrast, conventional protocols are end-point assays that require several weeks of cultivation before they can be evaluated. Finally, Öz and co-workers used an 8W10E+ ECIS Cultureware assay (Applied Biophysics), which contained forty 250-μm gold electrodes per well, and could be measured on an ECIS Model 1600 (Applied Biophysics) at 45 kHz for the screening of cancer drug therapies [16]. Specifically, the purpose of this work was to monitor the differentiation of cancer stem cells

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Figure 19.2  Sensor osteogenic and adipogenic differentiation [15]. (A) The MSCs undergoing osteoblastogenesis were monitored 7 days post-induction. Compiled impedance signals derived from eight parallel measurements of differentiating or untreated controls are shown. (B) Sensor osteogenic differentiation was verified by histological staining with Alizarin red S, 18 days post-induction. (C) Adipogenic differentiation was assessed as described in (A). Compiled impedance signals derived from nine parallel measurements of differentiating or untreated controls are shown. (D) Lipid vesicles were visualized by means of Oil Red O, 12 days post-induction. Reproduced from Reitinger et al., 2012 with permission from Elsevier. (See insert for color representation of the figure.)

after induction by different drug treatments (e.g., decitabine (DAC) and cytarabine (araC)). This work is particularly significant owing to the fact that the differentiation of cancer stem cells, which are a subpopulation of tumor cells that exhibit stem-like properties such as self-renewal and differentiation potential, by drug treatment, represents a promising approach for cancer therapy [17]. To this end, the authors monitored the differentiation of the embryonic cancer-stem-cell line, NTERA 2 D1 (NT2), upon exposure to DAC [18], araC [19], and retinoic acid [20], which have previously been shown to induce cancer-stem-cell differentiation by causing DNA-damage, cellular stress, and the proteolytic depletion of stem-cell factors, in order to monitor and better understand the mechanisms that regulate the forced exit of cancer stem cells from malignant pluripotency. As early as 24 h after treatment, it was reported that treatment with DAC, araC, or retinoic acid could trigger specific, concentration-dependent increases in the dielectric resistant of NT2 cultures. Furthermore, in terms of its mechanism of action, it appears that the impedance profiles of

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drug-induced NT2 cell differentiation correlated well with the depletion of a key stem-cell self-renewal factor, OCT4. Overall, the key advantage of ECIS over conventional assays is that differentiation can be monitored such that its onset can be identified when other phenotypic changes of differentiation-specific marker-gene expression patterns are not yet apparent. Furthermore, by ­calculating the slope/time ratios for each condition, the degree of induced differentiation may also be obtained. As such, it is clear that ECIS represents a powerful technique for the ­high-throughput screening of stem cells, which can be applied to nanomaterials and can be performed noninvasively, in real-time, and label-free.

Microfluidic Flow Cytometry Microfluidic flow cytometry (FC) is a technique that is also based on electrical impedance. In the case of FC, single cells are continuously flowed between electrode pairs that are ­energized using AC excitation signals at one or multiple frequencies [21] (Figure  19.3). Owing to the fact that the area enclosed by electrode pairs approaches the diameter of single cells, minute electrophysiological variations of individual cells can be detected and then correlated to the impedance measurements obtained using this technique [22]. Unlike ECIS, which is also based on impedance-based measurements, FC utilizes microfluidic technology [23]. This renders the system highly compact by minimizing the size, weight, and power requirements as well as the need for user intervention, resulting in a highly autonomous system. In particular, microfluidics generally can be described as a set of microchannels that are either etched or molded into a material (e.g., glass, silicon, or polymer such as PDMS). These microchannels, which form the microfluidic chip, are connected in a specific organization to achieve a desired function, which can include mixing, pumping, cell sorting, or for chemical reactions. Finally, the microchannels are (A) Flow profile

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Figure 19.3  Microfluidic impedance flow cytometer [23]. The cell passes over electrodes and the impedance is measured. Information can be obtained about cell size, membrane, and cytoplasm properties. Reproduced from Cheung et al., 2010 with permission from Wiley.

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connected to the outside by input and outputs that pierce through the chip. Specifically, the FC technique exploits the microfabricated electrodes in these microchannel/microchambers to sense minute variations in impedance that are caused by morphological or electrophysiological changes within cells. To this end, Song and co-workers recently presented a microfluidic ECIS-based flow cytometer that could be used to identify the differentiation states of single stem cells [24]. Specifically, they utilized a novel dual microprobe design, which not only allowed for high-throughput screening, but also allowed for ease of ECIS measurements owing to the electrode configuration used. For their demonstration, Song et al. characterized the differentiation state of a mouse embryonic carcinoma cell line, P19. In particular, these cells were chosen because they are originally derived from an embryonic tetracarcinoma in mice and are known to readily differentiate into neuronal cells in the presence of retinoic acid [25]. First, to demonstrate that their microfluidic device could be used to measure impedance signals they performed three experiments where 20 μm polystyrene beads, undifferentiated P19 cells, or a mixed sample were flowed through the device with a measurement frequency of 50 kHz. In the case of the beads alone, it was observed that there was a negative and positive spike when the bead passed through the bottom micropore and the top micropore, respectively. Similar spikes were observed in the undifferentiated P19-cell case with smaller amplitude. In the case of the mixed sample, two populations were observed through impedance measurements: a larger spike which corresponded to the polystyrene, and a smaller spike corresponding to undifferentiated P19 cells. When trying to identify undifferentiated P19 cells from differentiated P19 cells, the size and electrical behavior of these populations were comparable when observed in the resistance-dominant domain. However, using differences in the membrane capacitance [26], which could be observed at 1 MHz, even when a mixture of undifferentiated and differentiated P19 cells was introduced, two distinct peaks were observed corresponding to the different populations. Altogether, FC represents a powerful and advantageous technique when compared to  conventional end-point protocols to quantify proliferation and/or differentiation. Specifically, not only is FC able to provide noninvasive real-time monitoring of stem cell self-renewal and differentiation, but it is also able to do this in a label-free and single-cell fashion.

Electrochemical Methods Electrochemical sensors have been used for a wide variety of applications owing to their many advantages, including their high sensitivity and selectivity as well as their fast and noninvasive nature [27]. In particular, a number of cell properties can be measured using electrochemical techniques, including volume, concentration, electrical, and morphological parameters, which are all important when monitoring biomass, sterilization control, quantitative evaluation of drug effects (e.g., on cancer), and, more recently, the self-renewal and differentiation of stem cells in a high-throughput, noninvasive, and label-free method on various surfaces including nanomaterials. In this section, we will cover electrochemical cyclic voltammetry-based sensors. Cyclic voltammetry (CV) is a type of potentiodynamic electrochemical measurement and is the most widely used technique to acquire qualitative and quantitative information about electrochemical reactions and has recently been applied to the measurement of reactions in living cells [28]. In particular, to perform CV, a voltage is swept between two values (from V1 to V2) at a fixed rate and when the voltage reaches V2 the scan is reversed and swept back to V1. The result is a current/voltage plot where, during the forward

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sweep (from V1 to V2), the reactant is converted to product and as a result a current begins to flow and eventually reaches a peak before dropping, in accordance with the Nerst Equation. When the scan is reversed, the curve simply moves back through the equilibrium position gradually converting product back to reactant. Using this mechanism, investigators have developed screening methods that utilize electrochemically active molecules to monitor the concentration of stem cells in order to gain information about stem-cell self-renewal or stem-cell differentiation. The advantages of electrochemical detection systems lie in their ability to miniaturize the whole detecting sensor instrument while allowing for facile monitoring and analysis of cell signals, which can then be correlated to stem-cell differentiation. To this end, Yea et al. recently developed an electrochemical cell-based sensor to ­monitor the differentiation of mouse embryonic stem cells (ESCs) by utilizing 1-naphtyl phosphate (1-NP) [29] (Figure 19.4a). 1-Naphtyl phosphate is a phosphate containing a double benzene ring that is known to be dephosphorylated into 1-naphthol by reacting with alkaline phosphatase (AP), an embryonic stem cell marker [30]. The product, 1-naphthol, has unique electrochemical properties from 1-NP and, as such, by monitoring 1-naphthol concentration the differentiation state of ESCs can be determined. Specifically, the electrochemical cell consisted of a two-chamber slide with a working gold electrode, Ag/AgCl reference electrode, and a platinum counter electrode. After introducing ESCs and allowing for cell attachment, CV experiments were performed across a 0.6 V to −0.2 V potential window with 0.1 V s−1 scan rate. After confirming that the electrochemical signals of 1-NP could be detected using their electrochemical device (Figure 19.4b and c), Yea and co-workers cultured undifferentiated mouse ESCs on the electrode. Upon its introduction, 1-NP was dephosphorylated to 1-naphthol by AP on undifferentiated mouse ESCs, resulting in a decrease of the reduction peak current. Moreover, the reduction peak decreased further in a linear fashion as the number of cells increased. In this way, owing to the fact that differentiated cells do not exhibit AP, the reduction peak correlates with the differentiation state of the mouse ESCs, resulting in a tool with which to rapidly screen the differentiation of stem cells. In summary, though the electrochemical detection of stem-cell differentiation is still in its infancy, it represents a promising method that holds a number of advantages over conventional assays, including easy, rapid, and noninvasive screening.

Raman Scattering-Based Methods Raman spectroscopy is a well-known “fingerprinting technology” that can be used to analyze the chemical structure of target materials as well as to quantify specific chemical bonds. In particular, there are two major categories of interactions that can induce a Raman shift. The first is known as a “Stokes shift,” which normally occurs when the final vibration state of the molecule has higher energy than its initial state, while the second shift is the anti-Stokes shift, which occurs when the initial state of the molecule is more energetic than its final state [31]. The energy transfer that occurs during the interaction between photons and molecules is very specific for almost all types of chemical bonding, resulting in sharp characteristic Raman peaks in the spectra. Hence, each peak that appears in the Raman spectra can be correlated to the molecular structure of the target material, thereby providing very specific information regarding the structural characteristics of the molecule of interest. For stem-cell applications, Raman spectroscopy can also be a valuable tool, as it can act as a simple and efficient technique to rapidly analyze the state of stem cells.

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Figure 19.4  Electrochemical detection of ESC differentiation [29]. (a) Schematic representation of the electrochemical detection system used to monitor the differentiation of mouse ESCs. (b) Cyclic voltammetry at different concentrations of 1-naphthyl phosphate (1-NP; from 0.2 mm to 1 mm) and a scan rate of 100 mV s−1. (c) The reduction peak current increased linearly with 1-NP concentration. Reproduced from Yea et al., 2013 with permission from Elsevier.

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For instance, Turner et al. reported a novel technique for the in situ analysis and ­monitoring of live embryonic stem cells that is based on Raman microscopy [32]. In particular, a coherent anti-Stoke Raman spectroscopic method (CARS) was utilized to obtain biomolecular spectroscopy due to the fact that the signals generated by coherent excitation and signal emission are remarkably stronger than that of s­ tandard, spontaneous Raman spectroscopy (Figure 19.5a). To this end, the authors focused on genetic materials such as DNA, RNA, and its complexation with ribosomal proteins, which correlated with Raman peaks at 788 cm−1 (OP–O stretch, DNA) and 811 cm−1 (O–P–O stretch, RNA) (Figure 19.5b and c). By analyzing the CARS mapping images that were obtained and the mean CARS intensities (A) Lamp OPA

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Figure 19.5  Coherent anti-Stoke Raman spectroscopy (CARS) analysis of living ESCs [32]. (a) Instrumentation set-up and optical path. For all image data, the pump and Stokes beam pulse energies were both ~1 nJ pulse−1 at the sample. The CARS images of undifferentiated mouse ESCs are shown at (b) 788 (O–P–O stretch, DNA) and (c) 811 cm−1 (O–P–O stretch, RNA) in picosecond mode (20 cm−1 resolution). Reproduced from Konorov et al., 2007 with permission from ACS.

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over a Raman shift range of 675–950 cm−1, the authors determined that the CARS intensities at 811 cm−1, which is assigned to the RNA backbone, is significantly lower for differentiated stem cells, while intensities over the range of 730–770 cm−1 were higher for this population. As such, Turner and co-workers were able to conclude that the Raman spectra obtained from undifferentiated and differentiated stem cells are distinct and characteristic of their differentiation state and can be used as an efficient indicator for the label-free m ­ onitoring of stem-cell differentiation based on Raman spectroscopy. In a similar research effort, the same group utilized normal Raman spectroscopy instead of CARS for the analysis of ESCs [33]. In this study, the authors detected normal Raman signals from ­undifferentiated and differentiated ESCs and generated a library of Raman intensities for each peak. Finally, the Raman intensities of nucleic acids (DNA–RNA composites) at 784 cm−1 were divided by protein-related bands (e.g., tryptophan), which exist at 757cm−1, in  order to discriminate undifferentiated stem cells from differentiated stem cells. Remarkably, the distribution of the Raman peaks for the two protein/­nucleic-acid intensity ratios (757/784 cm−1 and 853/784 cm−1) were found to be separated clearly ­between the two cell populations. This indicates that normal Raman spectroscopy is also effective for the identification of undifferentiated and differentiated stem cells. More recently, surface-enhanced Raman spectroscopy (SERS) has been applied extensively in live cell research to overcome the weak intensities that are characteristic of normal Raman techniques, often considered a critical disadvantage of Raman-based monitoring tools. Surface-enhanced Raman spectroscopy is normally generated from ‘hot spots” that exist in the nanogap present between noble metal structures (gold, silver, copper, etc.) and gives 10–15-fold higher signal intensities than normal Raman peaks [34]. Hence, SERS can be used as an efficient tool for the identification of stem-cell differentiation. Recently, Tamiya and co-workers reported a novel SERS-based tool that is capable of monitoring mouse ESCs, including undifferentiated single cells, embryoid bodies (EBs), and terminally differentiated cardiomyocytes [35] (Figure 19.6). To this end, cells were treated with gold nanoparticles of different diameters (40, 60, and 100 nm) and subjected to SERS studies. Interestingly, the SERS spectra of undifferentiated stem cells showed strong Raman signals at the characteristic DNA and RNA peaks (O–P–O stretch DNA: 787 cm−1 and O–P–O stretch RNA: 813 cm−1), while EBs and differentiated cardiomyocytes showed high SERS intensity for protein peaks (amide I, amide II, and amide III) and mitochondria (1604 cm−1). According to the authors’ report, the increase in protein components in differentiated ­cardiomyocytes is mainly caused by the increase in cellular activity related to protein translation, post-translation, and cell signaling, which is markedly different from undifferentiated single ESCs. Hence, it is obvious that SERS-based techniques are also very powerful for the noninvasive and label-free monitoring of stem cell differentiation, which can overcome the disadvantages of current stem-cell-related characterization techniques.

Microscopy-Based Methods In conventional immunocytochemical and flow cytometrical methods, cell-surface markers are detected using fluorescently labeled antibodies. This method represents a noninvasive method with which to determine stem cell self-renewal and differentiation. However, these methods detect only the end-point of the antibody–antigen reaction as opposed to the real-time binding reaction kinetics (e.g., equilibrium dissociation constant Kd), which can be  used to reveal both the presentation and population of the surface antigens from one  differentiation stage to another. Moreover, not only does the actual process of

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labeling antibodies with extrinsic fluorescent agents alter the strength and specificity of the protein–ligand interactions [36], but techniques such as flow cytometry can also negatively impact cell viability and alter the differentiation potential of the sorted stem cells [37]. As such, significant effort has been invested in developing microscopy-based methods with which to achieve the real-time and label-free screening of stem cell self-renewal and differentiation. To this end, Lo and co-workers have demonstrated that oblique-incidence reflectivity difference (OI-RD) microscopy is an effective method for the label-free, real-time detection of cell-surface markers and applied this technique to analyze the presence of stage-specific embryonic antigen 1 (SSEA1), a well-established stem-cell marker, on mouse ESCs and mouse induced pluripotent stem cells (iPSCs) [38]. Specifically, Lo et al. immobilized mouse stem cells and non-stem cell controls (A19 fibroblasts and HEK293T cells) onto a glass surface as a type of “cell microarray.” Afterwards, the cell microarray was reacted with unlabeled SSEA1 antibodies and the status of SSEA1 antibody binding on the cell surface was monitored with OI-RD microscopy (Figure 19.7). Oblique-incidence reflectivity difference microscopy utilizes a p-polarized He–Ne laser beam at λ = 633 nm that passes through a photoelastic modular so that the output beam is polarization modulated at Ω = 50 kHz [39]. This polarized laser beam, with adjustable phase, is reflected off the glass slide surface containing cells, and when the surface is exposed to a probe solution, the binding event that occurs between the antigen and its substrate results in a magnitude and phase shift of the electromagnetic field that is associated with the reflected light [40]. Using this method, binding curves were obtained for individual pixels and used to determine the Kd. By obtaining binding curves for undifferentiated mESCs and miPSCs as well as differentiated mESCs and mIPSCs (Figure 19.8), it was determined that the Kd is lower for differentiated cells when compared to undifferentiated cells. As higher Kd would correspond with lower binding affinities, and considering the fact that the number of SSEA1 surface markers should be higher on undifferentiated cells than differentiated cells, the observation that SSEA1 antibody shows higher binding affinity with differentiated cells is surprising. However, the authors explain that this can be caused by differences in the presentation of the SSEA1 surface marker at different stages of differentiation. In particular, given that the

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Figure 19.7  Sketch of an oblique-incidence reflectivity difference (OI-RD) scanning optical microscope consisting of illumination and detection optics and a sample cartridge that holds a functionalized glass slide and a fluidic inlet/outlet assembly for each of eight chambers [38]. By scanning a focused optical beam along the y axis and moving the sample-holding stage along the x axis, the scanner detects real-time changes on the microarray as a result of reaction or other processing by measuring the amplitude and phase changes of the reflected beam. PEM, photoelatic modulator; PS, phase shifter; FTL, f-theta lens; OBJ, objective; A, analyzer; PD, photodiode. Reproduced from Lo et al., 2011 with permission from BioTechniques.

 19  High-Throughput Screening of Stem Cell Self-Renewal

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surface markers may be at low densities on stem cells during differentiation, it is possible that binding of the antibody is independent of the density and the variation in binding affinity constants can be attributed to the variation in the presentation of the surface markers [41].

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In another example, Stringari et al. utilized fluorescence lifetime microscopy (FLIM) to image and discriminate undifferentiated human ESCs from differentiated progenies by monitoring their metabolic activity [42]. In particular, FLIM is an imaging technique that produces an image based on the differences in the exponential decay rate of fluorescence and can be used to measure the time-decay characteristics of the cell and tissue microenvironment, thereby allowing for the molecular localization and identification of intrinsic fluorophores or endogenous proteins [43]. By using cluster analysis of the phasor distribution of FLIM images [44], this technique provides a powerful technique with which to obtain a unique signature that can be used to monitor the metabolism of intrinsic fluorophores, like nicotinamide adenine dinucleotide (NADH) [45] and lipiddroplet-associated-granules (LDAG), which were found to correlate with the state of stem-cell differentiation [46]. Specifically, by monitoring NADH and LDAGs, ESCs that were induced to differentiate using BMP4 or retinoic acid, phasor FLIM could discriminate between undifferentiated ESCs from differentiating ESCs (Figure 19.9a–d). Moreover, this correlated well with measurements of the ESC marker OCT4, where undifferentiated ESCs have a high expression of OCT4, while differentiating ESCs have a characteristically low expression of OCT4 (Figure  19.9e–g). In addition, it was reported that the FLIM phasor distribution, which is characteristic of undifferentiated ESCs, was dominated by LDAG while differentiating ESCs treated with BMP4 were shifted toward the central region of the phasor plot, representing an increase in NADH. In a similar study, Stringari and co-workers also demonstrated that NADH metabolism could be used to monitor stem-cell differentiation of Caenorhabditis elegans germ line cells [44]. Overall, there are a number of microscopy-based screening methods that are now available, which can be utilized for the noninvasive, label-free, and real-time monitoring of stem cell self-renewal and differentiation.

Conclusions Given the tremendous potential that stem cells hold in the clinic as well as the increasing interest in using nanomaterials [47], there is an urgent need for high-throughput methods to screen stem cell self-renewal and differentiation in a simple, noninvasive, real-time, and label-free fashion. To this end, recent efforts have been placed in developing and adapting techniques from other fields, including electrical cell–substrate impedance, microfluidic flow cytometry, electrochemical, Raman, and microscopy-based methods. Each technique has its own individual advantages; for example, microfluidic flow ­cytometry allows for the high-throughput screening of individual stem cells whereas Raman and microscopy-based techniques can be used to monitor the unique chemical structure and metabolism of stem cells, respectively. However, when compared to conventional techniques used for the characterization of stem cells, including RT-PCR, Northern and Western blotting, immunofluorescence, and flow cytometry, the methods described in this chapter hold a number of general advantages. First, these methods are simple to perform. Second, these methods can provide real-time monitoring of stem cell self-renewal and differentiation, whereas conventional methods tend to be end-point assays that can give only a snapshot of what is occurring in the stem cells. Third, and most importantly, the described methods are noninvasive and, as such, stem cells can be used even after being screened. Despite the fact that the field of high-throughput screening of stem cell self-renewal and differentiation is still in its infancy, advances are rapidly being made in this field and we are moving ever closer to having the ability to screen the

 19  High-Throughput Screening of Stem Cell Self-Renewal

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Figure 19.9  Label-free identification of hESCs [42]. (a) A fluorescence lifetime microscopy (FLIM) phasor histogram of the FLIM image excited at 760 nm from one H9 hESC colony co-cultured with mouse embryonic fibroblasts (MEFs). The color scale (from blue to purple) corresponds to the 64 levels of the contours that indicate the percent occurrence in the phasor histogram of the pixels of the image. Different clusters within the phasor distribution correspond to bright lipid droplet-associated granules within hESCs (red), the hESCs themselves (green) and the MEF feeders (blue). (b)  A transmission image and (c) a two-photon fluorescence intensity image of an undifferentiated hESC colony grown on MEF feeders. (d) Phasor color map: pixels of different colors correspond to the color of the cluster in the phasor plot A (arrow indicates a MEF). (e) Expression of the pluripotency marker OCT4 in the same colony as in (b) after cell fixation and immunostaining. (f) 4′,6-­Diamidino-2phenylindole (DAPI) staining. (g) Merge of DAPI and OCT4 staining. Arrow indicates a MEF with a nucleus that does not express OCT4. Reproduced from Stringari et al., 2012 with permission from SPIE. (See insert for color representation of the figure.)

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status of individual stem cells at a high-throughput, which would greatly facilitate the translation of stem-cell-based therapies to the clinic.

Acknowledgments We would like to acknowledge generous support by the NIH Director’s Innovator Award [(1DP20D006462-01), K.B. Lee] and the N.J. Commission on Spinal Cord grant [(09-3085SCR-E-0), K.-B. Lee]. P.T. Yin would like to acknowledge the NIH Biotechnology Training Grant for their support. We are also grateful to the KBLEE group members for their useful comments on the manuscript.

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Part 6

Nanotechnology in Stem-Cell Imaging

Chapter 20

Nanotechnology for Cellular Imaging Miroslaw Janowski1–4, P. Walczak1,2,5, and J.W.M. Bulte1,2,6–8

Russell H. Morgan Department of Radiology and Radiological Science, Division of MR Research, The Johns Hopkins University School of Medicine, Baltimore, MD, USA 2  Cellular Imaging Section and Vascular Biology Program, Institute for Cell Engineering, The Johns Hopkins University School of Medicine, Baltimore, MD, USA 3  Department of NeuroRepair, Mossakowski Medical Research Centre, Polish Academy of Sciences, Warsaw, Poland 4  Department of Neurosurgery, Mossakowski Medical Research Centre, Polish Academy of Sciences, Warsaw, Poland 5  Department of Radiology, Faculty of Medical Sciences, University of Warmia and Mazury, Olsztyn, Poland 6  Department of Oncology, The Johns Hopkins University School of Medicine, Baltimore, MD, USA 7  Department of Biomedical Engineering, The Johns Hopkins University School of Medicine, Baltimore, MD, USA 8  Department of Chemical and Biomolecular Engineering, The Johns Hopkins University School of Medicine, Baltimore, MD, USA 1 

Introduction There is much enthusiasm and interest in the application of stem-cell therapy in practically every field of medicine. However, despite the promising preclinical results, stem-cell therapy has proven to be difficult to translate into routine clinical practice. The major problem is in the interpretation of the outcomes of stem-cell therapy, which is due to the lack of accurate information about the fate of the cells in vivo. In parallel, for the development of new pharmaceuticals, an assessment of drug kinetics has been the gold standard [1]. The outcomes of surgery, in turn, are typically confirmed by post-operative imaging. These methods allow for a conclusive assessment of pharmacological or surgical treatment efficacy. Due to its recent emergence, such principles have not yet been established for stem-cell therapy. However, the value of neuroimaging for the monitoring of transplanted cells has been recently emphasized [2] as a strategy for more rational stem-cell therapy, but, to date, only a small number of clinical studies have followed this recommendation [3]. This is attributable to the lack of robust, flexible, and proven techniques for the monitoring of transplanted cells. The advances in nanotechnology and material science have been remarkable in recent years, with new agents and effective methods for labeling of stem cells. As a result of these advances, stem-cell therapy for internal diseases of the liver, heart, kidney, or pancreas can be assessed more objectively; however, the evaluation of the clinical course of neurological disorders is much more difficult. While a meta-analysis of preclinical studies has shown positive results of stem-cell treatment for neurological diseases [4], the mechanisms mediating cell-dependent effects have yet to be fully elucidated, which precludes inferences about the injection dose and timing, cell type, etc. [5]. It is anticipated that cellular imaging will play an essential role in defining the mechanisms governing stem-cell therapeutics, which is crucial for further augmentation of stem-cell treatment [6]. Stem-Cell Nanoengineering, First Edition. Edited by Hossein Baharvand and Nasser Aghdami. © 2015 John Wiley & Sons, Inc. Published 2015 by John Wiley & Sons, Inc.

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The Need for Cellular Imaging in a Clinical Setting There are various imaging modalities used routinely in clinical practice and the selection of a modality depends on the exact investigative inquiry, accessibility, and cost-effectiveness. Other aspects that define the appropriateness of an imaging technique are sensitivity, specificity, resolution, and radiation exposure. Imaging plays an essential and consistently increasing role in medicine and there are finely honed algorithms that are routinely employed for the assessment of the progression/recession of many disorders. Therefore, it is of paramount importance to ensure that procedures required for stem-cell tracking do not interfere with the current imaging algorithms used for disease assessment. The existing stem-cell imaging methods using iron oxide nanoparticles were shown to obstruct routine imaging, as they deplete the signal on magnetic resonance imaging (MRI) scans. Therefore, the choice of a stem-cell imaging method should be determined by the requirement for a certain structural or functional imaging method mandatory for correct assessment of the principal disorder. This concept can be referred to as an “imaging window,” which should be tailored to each patient, taking into consideration patient’s co-morbidities.

The Ultimate Goals of Stem-Cell Imaging Stem-cell imaging is performed to achieve several distinctive goals with different levels of complexity. First, the most basic task is the monitoring of cell distribution. Despite the relative simplicity of such monitoring, it provides highly valuable information on the initial placement of transplanted cells, whether the cells reached the desired target, or even whether misinjection is a health threat. This task requires high-resolution imaging technology, but it does not probe the actual biological status of grafted cells. Preferably, any stem-cell transplantation should be accompanied by monitoring of cell distribution. Stem-cell tracking is being applied in a variety of disorders and with many different transplantation routes [7]. There is an abundance of nanoparticle formulations suitable for cell labeling to accomplish this task. The task of evaluating cell survival is more complex. It has been successfully addressed in small animals by the application of bioluminescence imaging, based on efficient reporter genes such as firefly luciferase. Although this technique is invaluable for preclinical studies in small animals, it cannot be successfully translated to human studies. The highly desirable task of monitoring cell survival with nanotechnology techniques is in its infancy. Only very recently were arginine-containing liposomes used to monitor pH changes with MRI as a surrogate marker for transplanted cell death [8]. In general, it mandates specific characteristics of the nanomaterial, with a change in properties depending on cell viability or associated changes of the microenvironment, and, most importantly, that change should be easily detectable by noninvasive, clinically applicable imaging. The most complex task is to evaluate not only cell survival, but also cell function. This would require the imaging probe to detect the metabolic/gene expression changes during the process of cell differentiation, and specifically, in the transplanted cells. That task has not yet been achieved, but further advances in nanotechnology could potentially address this issue and equip scientists with an invaluable tool for studying stem-cell biology. The evaluation of dopaminergic neuron function in grafts deposited in patients with Parkinson’s disease was feasible using positron emission tomography (PET) technology; however, this method cannot discriminate transplanted cells from endogenous counterparts.

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Nanotechnology for Stem-Cell Imaging The fields of stem cells and cell-based therapy have benefitted tremendously from ­nanotechnology, which has provided many resolutions for the problems associated with stem-cell imaging. Nanotechnology delivers a diversity of choices for straightforward stem-cell tagging, and does not preclude the use of additional methods, including reportergene approaches. Stem-cell imaging methods preferably use intracellular labeling, which may produce contrast for one imaging modality (unimodal type) or for several modalities (multimodal type).

Iron Oxide Nanoparticles Iron oxide nanoparticles are characterized by a strong negative (hypointense) signal on MRI. The excellent detectability of iron oxide was initially observed as artifacts related to the use of mascara [9–11]. The coating of iron oxide nanoparticles with polymers facilitated biocompatibility and protection against clumping [12]. There is an assortment of iron oxide nanoparticles for cell tagging in various sizes, ranging from micron-sized [13] to nanometersized particles [14]. Iron oxide particles are the most often used agents for stem-cell imaging. Initially, iron oxide nanoparticles were used for ex vivo tagging of peripheral blood mononuclear cells [15], and then this technology was successfully applied to neural stem cells (NSCs) [16–19]. It was found that the size of iron oxide nanoparticles governs the effectiveness of their uptake by cells, with a higher tagging efficacy by nanoparticles ~70 nm diameter (SPIO) as compared for ultrasmall iron oxide nanoparticles (USPIO) [20]. One of the SPIO formulations (Feridex®) has been approved by the Food and Drug Administration (FDA) for clinical use as a contrast agent. This spurred a preclinical research on labeling efficiency with the eventual establishment of methods for tagging of nonphagocytic cells, including stem cells, and the eventual application of Feridex® for clinical stem-cell transplantations [21–23]. Unfortunately, the company ceased production of Feridex® in 2008 for economic reasons, which left the field without a clinical grade formulation of SPIO. While another iron oxide nanoparticle-based drug (Feraheme®) has been recently cleared by the FDA for treatment of iron deficiency anemia its application to cell tagging is more challenging due to the USPIO. However, the efficient tagging of cells by Feraheme® has been recently demonstrated [24, 25]. Further advances through nanocomplex self-assembling, by a combination of Feraheme®, heparin, and protamine, resulted in larger, more easily internalized complexes of ~100 nm in size [26]. It should be emphasized that all these agents were approved by the FDA for a different application; thus, they would have to be used off-label for clinical cellular imaging. The contrast enhancement of iron oxide nanoparticles is exceptionally strong, and has enabled single-cell visualization [27, 28]. However, the persistence of SPIO and contrast within the tissue despite cell death complicates the interpretation of imaging results [29–31]. Consequently, iron oxide nanoparticles are excellent for short-term monitoring of cell distribution, however, the persistent presence of SPIO in the tissue precludes its utility for the long-term imaging of stem-cell fate. Due to the current limitations of iron oxide nanoparticle formulations, there is continued effort to advance their properties by surface modifications, through the application of new coatings characterized by superior labeling and safety profiles. Poly(N,N-dimethylacrylamide)coated maghemite [32], aminosilane coating [33], amine surface modification [34], carboxymethyl chitosan modification [35], 1-hydroxyethylidene-1.1-bisphosphonic acid (HEDP) coating [36], nontoxic protein transduction domain (PTD) conjugation [37], higher density carboxyl groups modification [38], and D-mannose modification [39] have been used for this reason.

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The iron oxide contrast in MRI depends on the size and cellular clustering of nanoparticles. The SPIOs are typically used as negative T2 contrast, but it has been shown that USPIOs also produce a more desirable positive T1 contrast [40]. Scaling up production of the extremely small-sized iron oxide nanoparticles (ESION) and providing positive T1 contrast may enhance the application of iron oxide for cellular imaging [41]. Another application for iron oxide nanoparticles that takes advantage of their strong magnetic characteristics is directing the movement of labeled cells after transplantation. An external magnetic field applied focally can guide cells to the target organ/structure, facilitating stem-cell trafficking to the desired sites [42–44].

Gadolinium Nanoparticles Gadolinium (Gd) chelates are widely used in the clinical arena, due to their strong positive T1 contrast mechanism, circumventing the problems associated with contrast agents that produce signal voids, such as obscured images, after the administration of SPIO [45]. Unfortunately, those gadolinium chelates do not yield a positive T1 contrast following uptake by the cells [46]. The wider application of gadolinium for cell tagging maybe hindered by problems with the production of water-soluble solutions with satisfactory stability [47]. One solution to overcome these obstacles has been to integrate the metal-based nanoparticles or polymer-based scaffolds with gadolinium [48, 49], but the gadolinium chelates still produce a relatively low signal per molecule. In this context, production of gadolinium oxide nanoparticles has shown to be successful in the tracking of hematopoietic cells [50]. This occurred in concert with the commercial production of dextran-coated gadolinium oxide nanoparticles – for example, GadoCELLTrack (BioPAL) – which should generate even more interest in this method [51]. Auxiliary work to further improve the coatings of gadolinium oxide nanoparticles is progressing, with a choice of variations, including shells made of polyethylene glycol (PEG) [52] and diethylene glycol (DEG) [53], silica [54], and albumins [55]. An alternative method consists of capturing gadolinium atoms in cages made of carbon nanotubes [56] or fullerenes [57]. However, the principal advantage of gadolinium oxide is believed to be the clearance following the death of labeled cells. It has been shown that free Dex-DOTA-Gd3+ can clear from the grafted area, in contrast to iron oxide nanoparticles [58]. However, this also has the disadvantage of releasing free Gd3+ ions that are known to be very toxic. In this respect, it may be difficult for Gd-based cell imaging to enter the clinical arena.

Manganese Oxide Nanoparticles The use of manganese and gadolinium as both being positive T1 contrast agents has been recognized for some time [59]. However, the neurotoxicity of manganese precluded its initial use in clinical applications [60–62], and gadolinium became the agent of choice for use in patients [63]. Further investigations have shown that the toxic effect of manganese is highly specific to neurons, without affecting any other cell type [64]. The development of methods that facilitate the manufacture of manganese oxide nanoparticles, with subsequent coating by a biocompatible PEG-phospholipid, has rejuvenated the interest in manganese, especially in the context of stem-cell labeling and imaging [65]. While positive T1 contrast is highly desirable, this formulation of manganese oxide nanoparticles proved to be of inferior sensitivity for stem-cell imaging compared to iron oxide-based contrast agents [66]. A recent advancement in the manufacture of a hollow structure of manganese oxide nanoparticles, coated with mesoporous silica, yielded increased T1 signal through better entrance of water molecules to the magnetic core [67].

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Fluorine Nanoparticles The major advantage of fluorine is the near-absence of this element in the human body, thus providing contrast with no tissue background. The additional advantage of 19 F is the ability to acquire both anatomical 1H and 19 F “hot spot” scans, which then can be automatically superimposed [68] and the fluorine signal quantified [69]. Since the signal-to-noise ratio (SNR) of fluorine is almost the same as that of proton images, the contrast agents based on fluorine necessitate a high load of 19 F nuclei, comparable to the presence of 1H in tissues [70]. Luckily, a method to achieve this high concentration has been known for years and consists of a compound synthesis by the exchange of 1H to 19 F nuclei [71]. Moreover, the biocompatibility of fluorocarbons was tested in a clinical setting far before the advent of MRI, and these compounds were employed as X-ray contrast agents [72], or blood substitutes [73]. Thus, they are ideally suitable to be used as MRI contrast agents. For example, perfluoropolyether nanoparticles have already been employed for the in vivo imaging of dendritic cells [74], beta islets [75], and neural stem cells [76, 77]. Despite promising results, the application of perfluorocarbons for “hot spot” MRI is cumbersome due to the requirement for labeling cells with a high load of fluorine to reach an acceptable SNR [78], or the large amount of cells to be transplanted for visualization in vivo [79].

PARACEST Nanoparticles There is continuous search for noninvasive and clinically applicable novel contrast mechanisms, especially for MRI. A newer method that has recently gained a wider use is chemical exchange saturation transfer (CEST) MRI, termed as such because it depends on the exchange of protons between compounds or their specific groups and bulk water as an effect of saturation by an off-resonance pulse sequence. The same phenomenon has also been observed for the rare earth metals, which are characterized by the paramagnetic shift in the frequency of resonance (PARACEST). The benefit of CEST and PARACEST contrast agents is that this contrast is completely switchable, as the signal is present only as an effect of the application of a frequency-specific saturation pulse, and is otherwise absent; thus, there is no interference with regular MR imaging, such T1-weighted, T2-weighted, or diffusion imaging. While proteins are relatively insensitive in providing CEST contrast, the PARACEST nanoparticles have been shown to reveal a stronger signal. The superior PARACEST signal has been achieved by imaging of europium and ytterbium chelates. These chelates were then introduced to dendrimer nanoparticles to produce an in vivo applicable contrast agent [80]. Since the contrast is dependent on pulse frequency, this enables the visualization of different cell populations by the application of PARACEST nanoparticles [81]. However, given the issues mentioned above, it is unlikely that PARACEST agents will enter the clinic any time soon. The CEST agents, on the other hand, are readily clinically translatable as they consist of naturally occurring carbohydrates and proteins, without the presence of metal ions.

Gold and Tantalum Nanoparticles Heavy metals are characterized by a very high X-ray absorption rate, far better compared to iodinated compounds, making them attractive cell-tracking agents that could be potentially detectable by computed tomography (CT). Gold nanoparticles have been evaluated as a contrast agent over recent years [82, 83]. An attractive feature of gold is that it is bioinert,

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and its biocompatibility has long been established in dentistry [84, 85]. The low toxicity of gold nanoparticles for labeled cells has been confirmed in vitro [86]. That was followed by effective imaging with micro-CT of cells tagged with gold nanoparticles [87]. Despite the favorable characteristics of gold nanoparticles, the cost of gold is still prohibitive for widespread application. Thus, there is a search for an inexpensive replacement for gold. Tantalum oxide nanoparticles have been considered as an alternative attractive candidate, since they are also characterized by a high X-ray absorption, biocompatibility, bioinertia, and low toxicity [88, 89], all of which comes at a much lower cost.

Polymer Dots While quantum dots have been used with increased frequency for cell labeling, a new organic alternative has arisen, in the form of π-conjugated polymer-based semiconductor nanoparticles, characterized by high fluorescence [90]. There are many advantages of these nanoparticles, such as a great emission rate, minimal “blinking,” and outstanding photostability. The diversity of emitted photon wavelengths, depending on polymer type, opens up the possibility of multicolor imaging. The progress in the assembly of near-infrared π-conjugated polymers heralds a wide utility in stem-cell imaging in vivo [91]. The synthesis of inorganic–organic/nanocrystals–polymer hybrids with these exceptional characteristics will enable a better fit for specific experimental conditions [92].

Nanodiamonds Fluorescent nanodiamonds are another group of optical labels. They are highly photostable, chemically nonreactive, biocompatible, and, most importantly, can emit photons of far-red bandwidth, which makes them advantageous for in vivo stem-cell imaging [93]. The atomic composition of nanodiamonds enables enhancement by fluorescence resonance energy transfer (FRET), but at the cost of the introduction of another label into the cells [94, 95]. Nanodiamonds have been shown to be relatively safe for the cells [96–98], and are characterized by a low exocytosis rate [99]; however, a slight increase in DNA repair proteins in nanodiamond-labeled embryonic stem cells has been reported [100].

Upconverting Luminescence Nanoparticles Nanoparticles characterized by near-infrared fluorescence have been shown to be suitable for in vivo stem-cell tracking; however, the tissue autofluorescence interferes with the process of detection of specific fluorescence, thus decreasing sensitivity. In fluorescence imaging, the absorption of light is followed by the emission of photons of longer wavelength, which must be detected against a usually high autofluorescence signal. The phenomenon of upconverting luminescence (UCL) may address this constraint. However, there are specific substances characterized by the emission of shorter wavelengths, called the anti-Stokes process, which is practically never encountered in living organisms; thus, any signal for UCL materials is devoid of background [101]. The anti-Stokes emission has been recognized for some time, but only after suitable techniques for the manufacture of UCL nanoparticles for use in vivo were developed did this field develop further. In addition, near-infrared upconversion observed in rare-earth nanophosphors resulted in the production of photoluminescent probes suitable for in vivo imaging in small animals [102]. Proof-of-principle experiments performed on tissue

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phantoms revealed the potential for stem-cell imaging by NaYF4 nanocrystals doped with Tm3+, with up to a 3-mm detection depth [103]. That was followed by a successful modification based on replacing yttrium with lutetium, which lead to a dramatic intensification of the signal by NaLuF4-based upconversion nanophosphors, reaching a sensitivity applicable to imaging in large animals [104].

Nanobubbles Echogenic microbubles are routinely used contrast agents in ultrasonography [105, 106]. The advantage of a background-free signal was achieved by the detection of the second harmonic wave [107]. However, the second harmonic wave depends on the bubble size; thus, their diameter must fit the needs of clinical ultrasonography. While the micrometer-size bubbles are produced for clinical purposes and they are easily available on the market, their large size is not compatible with stem cell labeling. The appropriate formulation of nanobubbles is very important, as the toxicity of some nanobubble types has been reported [108] and clinical application should be approached cautiously. Fortunately, there is continuous research and further advances in the detection of nanobubbles with ultrasound microscopy [109], as well as the development of nanobubbles with sufficient stability and safety [110, 111], indicating an important role for ultrasonography in stem-cell imaging.

Multimodal Nanoparticles Homogeneous Nanoparticles There are a few nanostructures that can be detected by multiple imaging modalities, making them desirable in specific experimental conditions. Europium doping of mesoporous gadolinium nanoparticles produced not only a MRI signal, but also the emission phosphorescence, which enables optical imaging. The applicability of cellular imaging with this probe was confirmed on HeLa cells [112]. The exchange of europium to lanthanum within gadolinium oxide results in an addition of upconversion luminescence [113]. The excellent production of a MR signal that is well-known for iron oxide is even exceeded by metals with higher magnetic moments, such as cobalt. However, the suboptimal stability of cobalt nanoparticles precluded their use in medical applications [114]. A recently manufactured alloy made of hollow cobalt and platinum (CoPt) surmounted the obstacle of stability while not influencing the strength of the MRI signal, and adding platinum-based X-ray visibility [115]. The nanoparticles made of this CoPT alloy were shown to be a viable option for stem-cell labeling [116]. The embedding of CoPt within the dendrimer matrix resulted in nanoparticles of novel architecture [117]. The major advantage of radionuclides is an extreme sensitivity with almost imperceptible amounts of label sufficient for signal acquisition. The short life-span of radionuclides, however, is a major obstacle [118]. Fortunately, nanotechnology has provided a tool with which to incorporate radionuclides into existing types of nanoparticles. Due to the rapid decay of radioactivity, which prohibits storage of the radionuclides, the production of such multimodal nanoparticles must be rapid and simple to accommodate at the application site just before use. This challenge has been addressed by the fast and straightforward microwavebased production of dextran-coated 64Cu-doped iron oxide [119]. Perfluorooctylbromide is an excellent compound for nanoparticle synthesis and multimodality imaging. The presence of fluorine is detected by MRI, while bromide is visualized by

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X-ray [75]. Another fluorine compound, a perfluoropropane, has been successfully used for nanobubble production, which, in addition to MRI, is also visible by ultrasound imaging [120].

Hybrid Nanoparticles Not many materials are visible by more than one modality. Thus, there is a trend to produce hybrid nanoparticles, which are usually manufactured in a multistep process. Typically, the particles are composed of a core with a substance detectable by a specific modality, surrounded by another substance specific to a different modality. The coating of an iron oxide core by a gold shell is an example of a typical hybrid nanoparticle, in this case providing detection by both MRI and CT [121]. A more complex architecture can be based on a perfluorooctylbromide core for detection by fluorine MRI and CT, conjugated with europium chelate, and producing a PARACEST MRI signal [122]. The coating of gadolinium nanoparticles with fluorescein [123] or rhodamine [124] allows for both optical and magnetic resonance imaging. The incorporation of near-infrared optical tags into magnetic nanoparticles is potentially interesting for clinical translation [125]. The addition of Cy5.5 to iron oxide nanoparticles is one such example of this technique [126, 127]. Gadolinium, Cy5.5, and/or dozens of other tags can be attached to the dendrimer core [128]. Even triple-modality nanoparticles are available with one example: an X-ray visible core covered by optical and paramagnetic labels [129].

Summary Interest in the field of stem-cell therapy is steadily growing, in particular for disease conditions where no effective treatment exists. The emergence of nanotechnology offers an array of techniques for stem-cell imaging. In general, the selection of imaging modality and stemcell label should not interfere with the imaging of the patient’s primary disorder, or better still, should not interfere with the imaging of any co-morbidities. There are a variety of nanotechnologies available for stem-cell labeling, ranging from unimodal nanoparticles to the highly specialized multimodal approaches. The selection of the best imaging modality and contrast agent should be based on a careful assessment of individual requirements.

Acknowledgments The authors thank Mary McAllister for editorial assistance.

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Part 7

Nanotissue Engineering and Clinical Applications

Chapter 21

Advancing Translational Nanotechnology to Clinical Application Michelle Griffin1, Shima Salmasi1, Naghmeh Naderi1, Peter E. Butler1,2, and Alexander M. Seifalian1,2

UCL Centre for Nanotechnology and Regenerative Medicine, Division of Surgery and Interventional Science, University College London, London, UK 2  Department of Plastic and Reconstructive Surgery, Royal Free London NHS Foundation Trust Hospital, London, UK 1 

Introduction The Clinical Need for Organ Tissue Engineering Patients’ tissue or organs may fail due to disease, injury, or aging and require treatment to allow for either repair or replacement. Treatment often involves harvesting tissue from another part of the body (autograft) or an organ from another individual (transplant or allograft). Although both therapeutic treatments will provide life-saving treatment there are limitations to these approaches. Autografts are limited by their donor site morbidity, overall tissue availability, and capacity for long-term survival. Unfortunately, organ transplantation is severely limited by donor shortage, harsh consequences of immune suppression medication, and difficulty monitoring rejection leading to chronic rejection. Artificial joint replacements or manmade synthetic vessels are also limited by their failure to mimic the native organs complex function, potential of noncompatibility, and failure for long-term survival [2]. The need to replace organs has sparked huge research interest into tissue engineering as a replacement modality for organ transplantation. The goal of tissue engineering is to rebuild or replace damaged or lost tissues [3]. The two essential components of tissue engineering are cells and biomaterials [4]. The introduction of cells into the nonfunctional tissue or organ is for the regeneration of new tissue or the local production of growth factors or  hormones to promote tissue growth or angiogenesis. Biomaterials, commonly called ­scaffolds, can be implanted with or without cells [4]. The scaffold is used to provide the structure and substrate for cell attachment and consequential tissue growth, allowing for organ regeneration. Scaffolds may also be loaded with growth factors or biophysical stimuli to direct the in vivo cell population or transplanted cells towards desired cell l­ineages on the scaffold, leading to specifically desired tissue formation [4]. We will now provide an overview of the cell choice and scaffolds that have been used for organ engineering.

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Stem Cells for Organ Engineering When considering how to create cell-based tissue engineering techniques it is vital to understand which cell type will be appropriate [5]. The ideal cell type would be one that can be easily isolated, maintained in culture, allows for safe implantation, and has the ability to form the functional cell type that is required [5]. The current drawback of applying ­cell-based techniques to organ replacement is the difficulty in harvesting and expanding enough cells for tissue transplantation [5]. Differentiated cells have several disadvantages, including inability to be maintained in culture for long periods to gain enough cells for transplantation [6]. They have also been shown to lose their capacity to form tissue when expanded, being unable to support tissue repair and regeneration [7]. Hence, stem cells have proven to be a key choice for designing tissue-engineering organs due to their unique properties. Stem cells are undifferentiated cells that have self-renewal and multidifferentiation properties. Stem cells can be divided into two types depending on their differentiation potential: ­pluripotent and multipotent stem cells. Pluripotent cells are human embryonic stem cells (hESCs) and induced pluripotent stem cells (iPSC) [8]. Their pluripotency allows them to self-renew indefinitely and differentiate into any of the three germ layers including e­ ndoderm, ectoderm, and mesoderm [8]. Human ESCs are isolated from the inner cell mass of blastocysts [9]. The mechanism by which these cells allow for self-renewal is still not understood but certain genes including Oct3/4 and Sox2 have been implicated [10]. Embryonic stem cells have been shown to differentiate in vitro into several cell lineages, including neurons, (cardio) myocytes, skin, blood, endothelial, cartilage, and pancreatic cells [11–16]. Despite their ability to differentiate into many specialized cell types, their progress for clinical application is limited due to ethical implications of the isolation techniques, their capacity to evoke immune responses, and possibility to cause teratomas [17]. Multipotent stem cells exist in the differentiated tissue and renew themselves for the lifetime of the organism and differentiate into all the cell types in the lineage they originate from [18]. Adult stem cells have become the primary target of many tissue-engineering strategies due to their multidifferentiation potential and suitable availability [18]. The most commonly used sources of adult stem cells are those of mesenchymal stem cells (MSCs) [18]. They are easily isolated from several adult tissue sources, including blood, adipose tissue, bone, dermis, umbilical cord, and blood [18–23]. Several studies have shown that MSCs have the potential to differentiate into several lineages, including osteogenic, chrondrogenic, and adipogenic, and with certain cues in neurogenic and hepatogenic lineages [24, 25]. With a potential easier route to clinical application, the majority of tissue-engineering efforts are heading towards finding a suitable scaffold for adult MSCs.

Scaffold for Organ Regeneration Scaffold materials are three-dimensional structures, which will guide the organization and growth of cells. Therefore, the selection of good quality scaffolds is vital for the success of tissue engineering. Numerous scaffolds, both naturally occurring and synthetically manufactured have been used to replace and restore tissues and organs in the body. Nearly all human cells reside in a tissue-specific three-dimensional matrix called an extracellular matrix (ECM) [26]. The ECM provides the anchor for the support of the cells as well as directing cell fate and function through cell–matrix interactions. The ECM consists of a complex network of nanoscale protein fibers forming local microenvironments, for cellular communication, oxygen and nutrient transport, and removal of waste material [26]. In addition the basement membrane consists of pores, ridges, and fibers that are nanometers

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in size [27]. In tissue engineering, biomaterials replace the native ECM found in the body to allow cells to create a highly ordered assembly of tissue formation. Hence, ideally, the artificial matrix should support cell growth and maintenance by providing the appropriate mechanical, chemical, and biological characteristics of the native ECM [28]. On this basis three-dimensional nanomaterials appear to be the most suitable scaffold to influence stemcell behavior for tissue engineering.

Nanomaterials for Organ Regeneration An appropriate three-dimensional scaffold should facilitate normal cellular organization and behavior, define and maintain the desired tissue volume, and at the same time promote host integration and implant vascularization [29]. Ultimately, the scaffold should undergo nontoxic degradation as it is replaced by the healthy host tissue. Hence, a biodegradable scaffold is ideally used in tissue engineering. Geometric variables of the scaffold such as porosity, pore size, and pore morphology are also important. A porous scaffold provides a large surface area for neovascularization, nutrient and waste exchange, cell migration, and matrix deposition [29]. Understanding of the cell-biomaterial interface is important in the choice of scaffold due to a direct impact on cell adhesion, an important step in the survival of anchorage-dependent cells [29]. Both cell proliferation and differentiation can be ­controlled by functionalization of the biomaterial surface. Multiple studies have shown that the interactions between cells and biomaterials occur at the nanoscale [30–32]. With the advent of nanotechnology, nanoparticles are being merged to synthetic scaffolds to develop nanostructured biomaterials and enhance interaction of proteins that control cell adhesion and, thus, tissue formation. Nanocomposites can be defined as multiphase solid materials where one of the phases has a dimension of less than 100 nm [33]. Nanomaterials have been developed as promising cell-carrier scaffolds due to their ability to mimic the nanoscale properties of the ECM [28]. Nanofibrous composite scaffolds are found to decrease ­immunogenicity, improve the capacity for cell interaction [34], and to harbor increased ­concentrations of fibronectin and vitronectin; the adsorption proteins that reduce apoptosis of transplanted cells [35]. For example collagen is a self-assembled triple helical bundle of nanofibers of 300 nm in length and 1.5 nm in diameter. Scaffolds manufactured from nanofibers, nanotubes, and nanoparticles have all shown to be useful in tissue engineering of organs. Nanofibers are the most highly documented nanoscaffolds, including polylactic acid (PLA) and polycaprolactone (PCL). They have emerged as an important nanomaterial due to their high surface area and highly interconnected pores [26]. These properties would allow ­efficient cell nutrient and metabolic waste between the scaffold and environment [26]. In this chapter, we will now explore the use of nanomaterials as scaffolds for tissue-engineering strategies, which have reached clinical application or are undergoing current clinical trials.

Clinical Application of Nanopolymers Nanoparticles The use of nanomaterials in medicine provides unique freedom to modify essential p ­ roperties such as solubility, diffusivity, drug half-life and release properties, and immunogenicity [36]. In the past few decades, a number of nanoparticle-based diagnostic and therapeutic agents have been developed for the treatment of cancer, asthma, endocrine and neurological ­conditions, pain, infections, and for tissue-engineering applications [36, 37].

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These nanoscale agents may provide more effective and convenient routes of administration, lower toxicity, allow targeted delivery and controlled release, and lead to reduced healthcare costs [37]. Nanoparticle imaging contrast agents have also been shown to have improved sensitivity and specificity [37]. Nanoparticle-based therapeutic agents have already been introduced in the world of m­edicine, and numerous products are currently under clinical investigation. Among these products, liposomal drugs and polymer–drug conjugates are two dominant classes, accounting for more than 80% of the total amount (see Table 21.1). Nanofibers Nanofibers can be assembled to form porous scaffolds that aid in the process of tissue ­engineering. These can be synthesized artificially or naturally. They can be prepared by ­electrospinning, phase separation, or self-assembly [38]. Electrospinning is a simple and cost-­ effective method for producing nanofibers that involves applying a high voltage to extruded polymer solution [38]. Drugs and growth factors can also be encapsulated in the form of nanoparticles, as an embedded layer, or incorporated within the polymer solution itself [39–42]. The phase-separation technique is based on thermodynamic demixing of a homogeneous polymer–solvent solution into a polymer-rich and a polymer-poor phase, usually by either exposure of the solution to another immiscible solvent or cooling the solution to a point below the bimodal solubility curve [43]. One limitation of nanofibrous materials generated using the phase-separation technique is the lack of interconnected pores, which are critical for cell seeding, vascularization, and tissue organization [43]. Phase-separation techniques are often used in combination with other scaffold fabrication techniques, such as porogen leaching, to overcome this problem. The combined technique provides better control over the porous architecture of the nanofibers [44]. Nanofibers stimulate cell colonization, successfully mimic the natural ECM because of their large surface area, and aid in efficient exchange of nutrients and metabolic waste ­between the scaffold and its environment [45]. In a recent study, a spatial distribution of the different cells with a three-dimensional scaffold culture system resulted in better structural organization as compared with two-dimensional culture systems [45]. In addition, nanofibers can be tailored to have controlled drug release properties. Nanobandage  Wound injury, due to burns and trauma is a serious health condition which if left untreated can lead to major infection and eventually death of the patient [46]. With around 10 million people suffering from major burn and chronic wounds worldwide, researchers have investigated various natural and synthetic materials such as collagen, gelatin, chitin, poly(lactic acid), poly(urethane) and poly (ethyleneimine), to develop gauze and bandages for a fast and effective treatment of such injuries [46, 47]. In most cases of major injuries the primary focus is to stop heavy bleeding of the site and the current commercially available options of bandages such as fibrin dressings and glue, either have short shelf-life or bring about adverse immune response at the site of the injury. Furthermore, they are not suitable for applications on many parts of the body such as the neck. In an effort to develop a product to rapidly seal and promote healing of major wounds and injuries, specifically those caused on battlefields, researchers at Massachusetts Institute of Technology (MIT), USA, designed thrombin-coated nanobandages [48, 49]. These were prepared using thrombin, a natural clotting agent, in solution together with tannic acid sprayed on gauge sponges using a nanosprayer. The use of nanospray allowed for very high effective area of absorption of the sponge as well as the area of action of thrombin [48]. Conventionally, gelatin sponges are used in hospitals to halt bleeding. However, such

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 21  Advancing Translational Nanotechnology to Clinical Application

Table 21.1  Examples of nanomaterials that are clinically approved or under current clinical trial [1] Nanomaterials

Trade name

Composition

Usage

Current status

Liposome-PEG doxorubicin

HIV-related Kaposi’s sarcoma, metastatic breast and ovarian cancer

FDA approved

Abelcet

Liposomal amphotericin B

Fungal infections

FDA approved

AmBisome

Liposomal amphotericin B

Fungal and protozoal infections

FDA approved

DepoCyt

Liposomal cytarabine

Malignant lymphamatous FDA approved meningitis

DaunoXome

Liposomal daunorubicin

HIV-related Kaposi’s sarcoma

FDA approved

Epaxal

Liposomal IRIV vaccine

Hepatitis A

FDA approved

Myocet

Liposomal doxorubicin

Combination therapy with cyclophosphamide in metastatic breast cancer

Clinical trial

Inflexal V

Liposomal IRIV vaccine

Influenza

FDA approved FDA approved

Nanoparticle – liposome Doxil/Caelyx

Nanoparticle – other

Nanofiber

Polymeric platforms

DepoDur

Liposomal morphine

Post-surgical analgesia

Visudyne

Liposomal verteporfin

FDA approved Age-related macular degeneration, pathologic myopia, ocular histoplasmosis

Estrasorb

Micellular estradiol

Menopausal treatment

FDA approved

Feridex

Iron oxide + dextran

Cell tracking (MRI contrast)

FDA approved

Resovist

Iron oxide + carboxy-dextran

Liver-specific contrast agent for MRI of focal liver lesion

FDA approved

FDA approved

Gd-DTPA

Gadolinium + DTPA

Drug delivery

Abraxane

Albumin + paclitaxel

Metastatic breast cancer FDA approved

Aurimune

Gold +TNF + PEG + therapeutic payload

Being tested for a variety of solid tumours

Clinical trial

NanOss

Calcium phosphate + gelatin

Bone regeneration

FDA approved

Nanobandage Synthetic polymer + thrombin

Bimolecular delivery; ECM mimics

Clinical trial

DermaFuse

Bioactive glass

Wound healing; tissue repair

Clinical trial

PuraMatrix

Self-assembled peptide

Dental bone void filler and as a hemostat

Clinical trial

Copaxone

l-Glutamic acid, l-alanine, l-lysine, and l-tyrosine copolymer

Multiple sclerosis

FDA approved

Genexol-PM

Methoxy-PEG-poly( d, Metastatic breast cancer Clinical trial l-lactide) taxol (continued)

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Nanotissue Engineering and Clinical Applications

Table 21.1  (Continued) Nanomaterials

Other

Trade name

Composition

Usage

Current status

Adagen

PEG-adenosine deaminase

Severe combined immunodeficiency disease associated with ADA deficiency

FDA approved

Macugen

PEG-anti-VEGF aptamer

Macular degeneration

FDA approved

Pegasys

PEG-α-interferon 2a

Hepatitis B and C

FDA approved

Neulasta

PEG-GCSF

Neutropenia associated with chemotherapy

FDA approved

Somavert

PEG-HGF

Acromegaly

FDA approved

Oncaspar

PEG-asparaginase

Acute lymphoblastic leukaemia

FDA approved

Renagel

Poly(allylamine hydrochloride)

End-stage renal disease

FDA approved

Emend

Nanocrystalline aprepitant

Antiemetic

FDA approved

Tricor

Nanocrystalline fenofibrate

Antihyperlipidemic

FDA approved

Rapamune

Nanocrystalline sirolimus

Immunosuppressant

FDA approved

DTPA; diethylenetriamine penta-acetic acid,; HGF; hepatocyte growth factor, PEG; Polyethylene glycol, GSCF; Granulocyte-colony stimulating factor; TNF; Tumour Necrosis Factor, VEGF; Vascular endothelial growth factor

­ andages are not suitable for application on all parts of the body and in the case of battleb fields, their portability is limited. In contrast, precoated gelatin sponges with layers of thrombin/tannic acid offer a highly stable and portable treatment system. In vivo animal studies carried at on such nanobandages showed blood clotting within 60 s, a dramatic achievement compared to the normal gelatin-dipped bandages, which take 150 s on average to present the same effect [48]. After proving the potential of nanobandages, researchers at MIT developed a next generation of nanobandages, which in addition to thrombin and tannic acid contain antibiotic (acid-vancomycin) to avoid microbial infection. Other than rapid healing and hemorrhage control, the nanobandages potentially offer great advantages towards effective and rapid sealing of internal injuries that are too large for primary closure [50]. Adherence of the nanobandage to the moist surface of the internal tissue of the body, whilst maintaining its flexibility, biocompatibility, and degradability is a prerequisite for it to serve its purpose [50]. To this end, the nanobandage surface is patterned, an inspiration drawn from the nanoscale hills and valleys observed in gecko lizards [50, 51], using a micropatterning technology and accommodated within a thin layer of gecko-like dry adhesives to achieve secure attachment of the bandage to the site of the injury. A key advantage of these nanobandages, both for instant healing and rapid sealing of internal injuries, is that the fibers of the sponge can be coated with thrombin at the nanoscale, therefore, increasing the amount of hemostatic agents that will be delivered to the wound, leading to a faster and more effective healing process [50, 51]. Doxycycline poly(ε-caprolactone) Nanofibers  Controlled-drug delivery is another potential avenue of application of nanomaterials in humans that is becoming more prevalent. Nanofibers in particular have been studied to develop a system in which drugs, such

 21  Advancing Translational Nanotechnology to Clinical Application

(A)

HV

371

(B)

WD

mag

10 µm

det HFW

15.00 kV 9.7 mm 5000 × LFD 29.8 μm

Doxy

NST

BHU

Figure 21.1  Poly(ε-caprolactone) nanofibers containing doxycycline were successfully electronspun and evaluated clinically for peridontal diease. (A) Scanning electron micrograph to show the surface mor­ phology of nanofiber. (B) Clinical photograph showing the insertion of the nanofiber into the periodontal pocket. Reproduced from [52] with permission by the Journal of Clinical and Diagnostic Research.

as antibiotics, can be delivered in a noninvasive, precise, timed, and more importantly, targeted manner to the desired location within the body [52]. Advantages of such local drug-delivery systems include: better accessibility, low enzymatic activity, and painless administration [52]. An example of such a system was reported recently whereby the effectiveness of doxycycline nanofibers, in conjunction with nonsurgical periodontal therapy or deep cleaning, were evaluated in the treatment of chronic periodontitis [52]. Doxycycline is one of the most common broad-spectrum antibiotics that is effective in managing most periodontal infections, owing to its low minimum inhibitory concentration (MIC), its antibacterial protein synthesis and anti-inflammatory properties [53]. In addition, doxycycline can bind to hard tissue of a periodontal pocket and act as a drug reservoir. Doxycycline incorporated with biodegradable polymers such as poly(ε-caprolactone) (PCL) nanofibers have been investigated for the treatment of periodontal infections in which the nanofibers have been inserted directly into the base of the infected periodontal pocket to fill its depth and curves (Figure  21.1) [53]. It was shown that such nanofibers were able to ­maintain drug concentrations while achieving a sustained release of doxycycline for a minimum of 19 days with low burst release. Furthermore, in all cases, nanofibers maintained their smoothness and flexibility and did not shrink during the treatment period [53]. These properties, together with the ease of implantation of the nanofibers, their cost, and time effectiveness, can prove ideal in the treatment of infected periodontal diseases. POSS-PCU Nanocomposite Material  The materials currently used in soft tissue regeneration, which include collagen, hyaluronic acid, silicon, and other filler materials, have s­ everal disadvantages such as high cost, immunogenicity, and the risk of transmitting infectious diseases. This lack of progress has inspired a fresh perspective and provoked further investigation and development in this field of tissue engineering. Since 2005, our laboratory have  developed biodegradable and nonbiodegradable nanocomposite scaffolds based on ­incorporating a polyhedral oligomeric silesquioxane (POSS) nanocage into polyurethanes for organ regeneration [54, 55]. Polyurethanes are adaptable copolymers that have been used for biomedical devices for several decades [56]. The nonbiodegradable nanocomposite polymer POSS-PCU consists of POSS combined with poly(carbonate-urea) urethane, a synthetic polymer already used in medical implants and devices [55]. The POSS molecule is a nanoparticle, which has dimensions in the order of

372

Nanotissue Engineering and Clinical Applications

6 nm [57, 58]. The POSS modification of the PCU polymer enhances several characteristics of the polymer, including its mechanical, thermodynamic stability, and cytocompatibility [59]. The POSS is derived from a class of compounds closely related to silicones and has the chemical composition of a hybrid, intermediate between that of silica and silicone. The chemical diversity of POSS technology is broad and POSS molecules are cytocompatible and suitable for synthesis of nanocomposite materials [54, 60]. They possess high tensile and elastomeric properties, good biocompatibility, and are easily fabricated and modified. The contrasting segmental characteristics of the soft amorphous phase and hard crystalline phase of polyurethanes provide the foundation for its versatility [61]. The hard segment provides this material with advantageous mechanical properties and the soft segment allows for elastomeric properties. The side chains of this copolymer may be modified to enhance particular properties for biomedical applications [62–64]. Numerous studies have shown that POSS-PCU scaffolds are antithrombogenic, biocompatible [58], nontoxic [54, 60], and do not cause inflammatory reactions to the surrounding host tissue [55]. Several cell lineages have been shown to attach to and grow on POSS-PCU, including fibroblasts, adipose stem cells, epithelial cells, endothelial cells, and chondrocytes, illustrating the cytocompatiblity and nontoxic properties of the biomaterial [55, 60, 65, 66]. Furthermore POSS-PCU scaffolds implanted in a sheep model illustrated no signs of ­degradation and inflammation over 36 months [55]. The POSS-PCU scaffold has already been taken to clinic use to replace tissues and organs, being implemented as the world’s first synthetic trachea in 2011 (Figure 21.2) in a 36-year-old O

(A)

NH

O

(C)

O i-Bu O Si Si i-Bu O O Si O Si O i-BuO O Si O i-Bu Si O O i-Bu O Si Si O i-Bu i-Bu

Urea hard segment

HN

O

Urethane group

O

O

i-Bu

Si O O Si O

O i-Bu Si O Si O i-Bu

n

O O

O O

O Si

O

O

O

O

O

Si O i-Bu O

O

NH

NH

O O

O

Si

O

NH

ata

O

O

NH

O

O

i-Bu

NH

O

POSS Nanocage

(B)

NH

(D)

(E)

Si O

i-Bu

i-Bu POSS Nanocage

Figure 21.2  Translational applications of POSS-PCU nanocomposite polymer already used for humans. (A) Schematic drawing of the poly-oligomeric silsesquioxane (POSS) nanocage. (B) The chemical structure of POSS–PCU with the polycarbonate back bone with the POSS particle. (C) The POSSPCU manufactured for a lower limb bypass graft. (D) The POSS-PCU lacrimal duct that has been successfully used in a patient. (E) The world’s first synthetic trachea, which was implanted to replace the trachea of a man with carcinoma of the trachea. Reproduced from [58] with permission by the Journal of Tissue Engineering and Regenerative Medicine. (See insert for color representation of the figure.)

 21  Advancing Translational Nanotechnology to Clinical Application

(A)

373

(B)

Figure 21.3  The next generation of artificial heart valves. (A) Tri-leaflet heart-valve designed, by University College London (UCL) with enhanced geometry for improved hemodynamic design. (B) A valve prototype manufactured using POSS-PCU and Dacron suture ring. Reproduced from [68] with permission by the International Journal of Nanomedicine.

man who presented with recurrent primary cancer of the distal trachea and main bronchi, previously treated with debulking surgery and radiation. The nanocomposite POSS-PCU implant was tailor-made for the patient, seeded with autologous bone-marrow mononuclear cells and placed in a bioreactor for 36 h before implantation [67]. This is to our knowledge the only case that has used stem cells on a nanocomposite polymer for clinical application. One of the most promising areas of application of POSS nanomaterials is the development of cardiovascular implants [68]. Synthetic heart valves based on POSS-PCU represent an attractive alternative to the existing devices as they resemble the superior durability of mechanical valves and enhanced hemodynamic properties of bioprosthetic valves at the same time [68]. The superior mechanical and chemical properties of POSS-PCU make this material suitable for developing the next generation of artificial heart valves (Figure 21.3). Currently, the POSS-PCU nanocomposite is under investigation in our centre for developing a novel aortic valve suitable for transcatheter aortic valve implantation [68]. Its design strategy is aimed at achieving high hydrodynamic performances while reducing stress levels [68]. In this design three polymeric leaflets (POSS-PCU nanocomposite) are attached to a self-expandable nitinol wire stent, which creates a high expanded/collapsed diameter ratio. It also improves anchoring by creating sufficient radial and axial forces. What makes this design even more interesting is that these heart valves are fully retrievable and repositionable, two properties that can play a major role in determining the success of the operation. Tissue-engineering of vascular bypass grafts is another area of research in which the potential application of POSS-PCU is being investigated (Figure  21.2). To this end, ­progenitor cells, extracted from peripheral blood of adult healthy patients, were seeded on a porous biofunctionalized POSS-PCU nanocomposite under static conditions and endotheliazation was evaluated [69]. The results were promising and demonstrated that they have the ability to promote rapid endotheliazation from progenitor cells [69]. Further studies on POSS-PCU bypass grafts revealed that they match the viscoelastic properties of the human arteries, a crucial aspect for the success of the implant in humans. So far the in vivo implantation of the POSS-PCU nanocomposite small diameter bypass grafts generated favorable outcomes and the human clinical trial of these is due to proceed

374

Nanotissue Engineering and Clinical Applications

in the near future. However, in a single case, a POSS-PCU bypass graft has already been used as a femoral bypass graft in a patient and he is currently being monitored to evaluate the outcome of the bypass. Lastly, we have utilized POSS-PCU for patient use as a lacrimal duct conduit (LDC) (Figure 21.2). Patients suffering from extensive tearing, blurry vision, and infections have an obstructed lacrimal drainage system [70]. The obstruction can be at different levels of the drainage apparatus [70]. When it is at the canalicular level, the standard treatment of stenting or reopening the obstruction fails [70]. Therefore the placement of a bypass is often the only treatment option [70]. This is done by the insertion of a polyurethane tube to allow tears to flow into the nose [70]. However, this damages normal tissue and function, including mucous blockage, and the tube may become dislocated causing a source of complications [70]. Owing to the viscoelastic and biocompatibility properties of POSS-PCU a small diameter conduit was made to replace the canaliculus and overcome lacrimal duct obstruction [70].

The Ethics of Nanotechnology in Regenerative Medicine We have illustrated with various nanomaterials the tremendous potential that nanotechnology could contribute to the improvement in health care. Nanotechnology deals with particles that are dominated by forces of physics and chemistry that are not present in traditional materials [71]. This unique and novel behavior can be exploited and manipulated by scientists for tissue engineering. However, understanding of this full potential will not be accomplished until the social and ethical issues are delineated. In general, as nanotechnology is a relatively new technology there are still ethical concerns about consumer and occupational safety and health. The key concern will be ensuring that medical nanodevices are classified appropriately for clinical use [72]. A new “drug” must demonstrate safety and efficacy using a multistep approval process before regulatory approval, often taking years to complete [72]. However, the premarket approval for a medical “device” is much less rigorous and for “cosmetic” use the FDA does not have a premarket approval system. Hence, labeling the nanodevice as a “drug” or “device” or “cosmetic” will prove to be vital to ensuring patient safety [72]. As nanotechnology will be used for many things other than medicine, there is concern that there will be an effect on the environment and nature. Further research is required to fully understand the impact of the accumulation of nanoparticles in plants and microorganisms to fully appreciate the extent on the environment [73]. It is important to remember that whether toxic hazardous materials will be released into the environment during the manufacture of the use of nanoproducts is still not known [73]. Lastly the main ethical debate is where the use of nanotechnology will be utilized to alter our human function rather than only restoring human function from disease and injury [74]. Currently there is a spectrum of beliefs, at one end people will argue to embrace new ­technology to advance our health but others will feel we should recognize the dangers and brake further technological development [74, 75].

Translating Science to Product Nanotechnology will improve the quality of life of patients, extend life expectancies, and reduce overall health costs. However, the clinical translation of regenerative medicine requires more than scientific discovery.

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375

Despite huge advancements into delivering nanomedicine strategies for improving healthcare, further research is required to fill the gap in the knowledge regarding the safety of nanotechnology [76, 77]. Complete research needs to be conducted to adequately assess the life cycle of the engineered nanomaterials on the impact human health and environment [77]. After satisfactory completion, communication with stakeholders needs to be ­undertaken to outline the risks and benefits of nanotechnology [77]. Epidemiologic research is a powerful healthcare tool to determine distribution of diseases in populations to manage healthcare problems. Social behavior and environmental regulation impact on the health outcomes [78]. Understanding and analyzing how nanomedicine could impact on public health using epidemiological studies could better predict the effect of nanomedicine enhancing health outcomes and for whom this technology would benefit [76]. As nanomedicine draws closer to becoming the first line of treatment in healthcare, understanding health outcomes and a long term cost benefit analysis is vital to its successful implementation [76]. At first nanomedicine will be more expensive than traditional methods and this challenge will need to be overcome with sufficient data to demonstrate that the application of nanomedicine is beneficial [76]. It will be important to ensure public perceptions of risks and benefits of nanomedicine are adequate to ensure correct medical decision-making by patients. For example, noncompliance with vaccines already highlights how public perception can be vital to medical care [79]. Stakeholders must be clear and transparent with regards to the capability of nanotechnology when describing to the public and policy boards [76]. To ensure implementation of emerging technologies into healthcare systems it will be important to train healthcare professionals in areas of nanotechnology, strengthening the bond between industry, government, and academia [76]. Applications in certain medical specialties will be essential, for example radiology, infectious disease, and oncology, to advance research to commercialization [76]. Finally, it will be important for regenerative medicine to develop clinical therapies that will revolutionize healthcare outcomes, but at the same time it is important not to overstate, or be unrealistic, about what is possible to deliver to the patient. We must be careful not to be too overenthusiastic and advocate only what is reasonable for today in order to ensure that we maximize the benefits of nanotechnology.

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[30]  Guo X, JE Gough, P Xiao, J Liu and Z Shen (2007). Fabrication of nanostructured hydroxyapatite and analysis of human osteoblastic cellular response. Journal of Biomedical Materials Research A 82: 1022–1032. [31]  Anselme K (2000). Osteoblast adhesion on biomaterials. Biomaterials 21: 667–681. [32]  Anselme K, M Bigerelle, B Noel, E Dufresne, D Judas, A Iost and P Hardouin (2000). Qualitative and quantitative study of human osteoblast adhesion on materials with various surface ­roughnesses. Journal of Biomedical Materials Research 49: 155–166. [33]  Lock J and H Liu (2011). Nanomaterials enhance osteogenic differentiation of human mesenchymal stem cells similar to a short peptide of BMP-7. International Journal of Nanomedicine 6: 2769–2777. [34]  Sanders JE, CE Stiles and CL Hayes (2000). Tissue response to single-polymer fibers of varying diameters: evaluation of fibrous encapsulation and macrophage density. Journal of Biomedical Materials Research 52: 231–237. [35]  Wei G and PX Ma (2004). Structure and properties of nano-hydroxyapatite/polymer composite scaffolds for bone tissue engineering. Biomaterials 25: 4749–4757. [36]  Brannon-Peppas L and JO Blanchette (2004). Nanoparticle and targeted systems for cancer therapy. Advanced Drug Delivery Reviews 56: 1649–1659. [37]  Kawasaki ES and A Player (2005). Nanotechnology, nanomedicine, and the development of new, effective therapies for cancer. Nanomedicine 1: 101–109. [38]  Jang JH, O Castano and HW Kim (2009). Electrospun materials as potential platforms for bone tissue engineering. Advanced Drug Delivery Reviews 61: 1065–1083. [39]  Yoo HS, TG Kim and TG Park (2009). Surface-functionalized electrospun nanofibers for tissue engineering and drug delivery. Advanced Drug Delivery Reviews 61: 1033–1042. [40]  Kim HW, HH Lee and JC Knowles (2006). Electrospinning biomedical nanocomposite fibers of hydroxyapatite/poly(lactic acid) for bone regeneration. Journal of Biomedical Materials Research A 79: 643–649. [41]  Song JH, HE Kim and HW Kim (2008). Electrospun fibrous web of collagen-apatite precipitated nanocomposite for bone regeneration. Journal of Materials Science and Materials Medicine 19: 2925–2932. [42]  McCullen SD, S Ramaswamy, LI Clarke and RE Gorga (2009). Nanofibrous composites for  tissue engineering applications. Wiley Interdisciplinary Reviews Nanomedicine and Nanobiotechnology 1: 369–390. [43]  Nam YS and TG Park (1999). Porous biodegradable polymeric scaffolds prepared by thermally induced phase separation. Journal of Biomedical Materials Research 47: 8–17. [44]  Wei G and PX Ma (2008). Nanostructured biomaterials for regeneration. Advances in Functional Materials 18: 3566–3582. [45]  Di Maggio N, E Piccinini, M Jaworski, A Trumpp, DJ Wendt and I Martin (2011). Toward ­modeling the bone marrow niche using scaffold-based 3D culture systems. Biomaterials 32: 321–329. [46]  Kumar, VK Lakshmanan, M Raj, R Biswas, T Hiroshi, SV Nair and R Jayakumar (2013). Evaluation of wound healing potential of beta-chitin hydrogel/nano zinc oxide composite ­bandage. Pharmaceutical Research 30: 523–537. [47]  Albertini B, M Di Sabatino, N Calonghi, L Rodriguez and N Passerini (2013). Novel ­multifunctional platforms for potential treatment of cutaneous wounds: development and in vitro characterization. International Journal of Pharmacology 440: 238–249. [48]  Trafton A (2012). New way to stop the bleeding; nanoscale biological coating developed at MIT could prevent battlefield deaths. News Office, Massachusetts Institute of Technology. [49]  Mahdavi A, L Ferreira, C Sundback, JW Nichol, EP Chan, DJ Carter, CJ Bettinger, S Patanavanich, L Chignozha, E Ben-Joseph, et al. (2008). A biodegradable and biocompatible gecko-inspired tissue adhesive. Proceedings of the National Academy of Sciences of the USA 105: 2307–2312. [50]  Dougherty E (2008). MIT creates gecko-inspired bandage. News Office, Massachusetts Institute of Technology. [51]  Yanik MF (2009). Towards gecko-feet-inspired bandages. Trends in Biotechnology 27: 1–2.

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[52]  Chaturvedi TP, R Srivastava, AK Srivastava, V Gupta and PK Verma (2013). Doxycycline poly e-caprolactone nanofibers in patients with chronic periodontitis – a clinical evaluation. Journal of Clinical and Diagnostics Research 7: 2339–2342. [53]  Javali MA and KL Vandana (2012). A comparative evaluation of atrigel delivery system (10% doxycycline hyclate) Atridox with scaling and root planing and combination therapy in treatment of periodontitis: A clinical study. Journal of the Indian Society of Periodontology 16: 43–48. [54]  Kannan RY, HJ Salacinski, KM Sales, PE Butler and AM Seifalian (2006). The endothelialization of polyhedral oligomeric silsesquioxane nanocomposites: an in vitro study. Cell Biochemistry and Biophysics 45: 129–136. [55]  Kannan RY, HJ Salacinski, JE Ghanavi, A Narula, M Odlyha, H Peirovi, PE Butler and AM Seifalian (2007). Silsesquioxane nanocomposites as tissue implants. Plastic and Reconstruction Surgery 119: 1653–1662. [56]  Santerre JP, K Woodhouse, G Laroche and RS Labow (2005). Understanding the biodegradation of polyurethanes: from classical implants to tissue engineering materials. Biomaterials 26: 7457–7470. [57]  Vara DS, HJ Salacinski, RY Kannan, L Bordenave, G Hamilton and AM Seifalian (2005). Cardiovascular tissue engineering: state of the art. Pathologie Biologie (Paris) 53: 599–612. [58]  Oseni AO, PE Butler and AM Seifalian (2013). The application of POSS nanostructures in cartilage tissue engineering: the chondrocyte response to nanoscale geometry. Journal of Tissue Engineering and Regenerative Medicine. doi: 10.1002/term.1693. [59]  Rahmani B, S Tzamtzis, H Ghanbari, G Burriesci and AM Seifalian (2012). Manufacturing and hydrodynamic assessment of a novel aortic valve made of a new nanocomposite polymer. Journal of Biomechanics 45: 1205–1211. [60]  Punshon G, DS Vara, KM Sales, AG Kidane, HJ Salacinski and AM Seifalian (2005). Interactions between endothelial cells and a poly(carbonate-silsesquioxane-bridge-urea)urethane. Biomaterials 26: 6271–6279. [61]  Kannan RY, HJ Salacinski, M Odlyha, PE Butler and AM Seifalian (2006). The degradative resistance of polyhedral oligomeric silsesquioxane nanocore integrated polyurethanes: an in vitro study. Biomaterials 27: 1971–1979. [62]  Sudaryanto, T Nishino, S Asaoka and K Nakamae (2001). Incorporation of methyl groups into hard segments of segmented polyurethane: microphase separation and adhesive properties. International Journal of Adhesion and Adhesives 21: 71–75. [63]  Chuang FS, WC Tsen and YC Shu (2004). The effect of different siloxane chain-extenders on the thermal degradation and stability of segmented polyurethanes. Polymer Degradation and Stability 84: 69–77. [64]  Ebert M, B Ward, J Anderson, R McVenes and K Stokes (2005). In vivo biostability of polyether polyurethanes with polyethylene oxide surface-modifying end groups; resistance to biologic oxidation and stress cracking. Journal of Biomedical Materials Research A 75: 175–184. [65]  Nayyer L, M Birchall, AM Seifalian and G Jell (2014). Design and development of nanocomposite scaffolds for auricular reconstruction. Nanomedicine 10: 235–246. [66]  Guasti L, B Vagaska, NW Bulstrode, AM Seifalian and P Ferretti (2013). Chondrogenic differentiation of adipose tissue-derived stem cells within nanocaged POSS-PCU scaffolds: A new tool for nanomedicine. Nanomedicine 10: 279–289. [67]  Jungebluth P, E Alici, S Baiguera, K Le Blanc, P Blomberg, B Bozoky, C Crowley, O Einarsson, KH Grinnemo, T Gudbjartsson, et al. (2011). Tracheobronchial transplantation with a s­ tem-cell-seeded bioartificial nanocomposite: a proof-of-concept study. Lancet 378: 1997–2004. [68]  Ghanbari H, A de Mel and AM Seifalian (2011). Cardiovascular application of polyhedral ­oligomeric silsesquioxane nanomaterials: a glimpse into prospective horizons. International Journal of Nanomedicine 6: 775–786. [69]  de Mel A, G Punshon, B Ramesh, S Sarkar, A Darbyshire, G Hamilton and AM Seifalian (2009). In situ endothelialization potential of a biofunctionalised nanocomposite biomaterialbased small diameter bypass graft. Biomedical Materials and Engineering 19: 317–331. [70]  Chaloupka K, M Motwani and AM Seifalian (2011). Development of a new lacrimal drainage conduit using POSS nanocomposite. Biotechnology and Applied Biochemistry 58: 363–370.

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[71]  Drexler KE (1987). Engines of Creation, the Coming Age of Nanotechnology. Anchor Press/ DoubleDay: New York. [72]  Stander L and L Theodore (2011). Environmental implications of nanotechnology – an update. International Journal of Environmental Research and Public Health 8: 470–479. [73]  Jotterand F (2008). Emerging Conceptual, Ethical, and Policy Issues in Bionanotechnology. Springer: Berlin;183–192. [74]  Glenn LM (2003). A legal perspective on humanity, personhood, and species boundaries. American Journal of Bioethics 3: 27–28. [75]  Glenn LM, Boyce JS (2008). Nano-Technology: Considering the Complex Ethical, Legal, and Societal Issues with Parameters of Human Performance. Springer: Netherlands. [76]  Pautler M and S Brenner (2010). Nanomedicine: promises and challenges for the future of public health. International Journal of Nanomedicine 5: 803–809. [77]  Som C, M Berges, Q Chaudhry, M Dusinska, TF Fernandes, SI Olsen and B Nowack (2010). The importance of life cycle concepts for the development of safe nanoproducts. Toxicology 269: 160–169. [78]  Satterfield T, M Kandlikar, CE Beaudrie, J Conti and B Herr Harthorn (2009). Anticipating the perceived risk of nanotechnologies. Nature Nanotechnology 4: 752–758. [79]  May T (2005). Public communication, risk perception, and the viability of preventive vaccination against communicable diseases. Bioethics 19: 407–421.

Chapter 22

Stem-Cell Nanoengineering from Bench to Bed Omid Mashinchian1, Shahin Bonakdar2, Shahriar Sharifi3, and Morteza Mahmoudi4,5,6

Department of Medical Nanotechnology, School of Advanced Technologies in Medicine, Tehran University of Medical Sciences, Tehran, Iran 2  National Cell Bank, Pasteur Institute of Iran, Tehran, Iran 3  Department of Biomedical Engineering, University Medical Center Groningen, Groningen, The Netherlands 4  Department of Nanotechnology, Faculty of Pharmacy, Tehran University of Medical Sciences, Tehran, Iran 5  Nanotechnology Research Center, Faculty of Pharmacy, Tehran University of Medical Sciences, Tehran, Iran 6  Division of Cardiovascular Medicine, Stanford University School of Medicine, Stanford, CA, USA 1 

Introduction During recent decades, numerous therapeutic approaches and constructs have been p ­ roposed for replacement, repair, and regeneration of damaged organs and tissues. These constructs can be created from the merged autologous or xenogeneic cells with natural or synthetic matrix materials, or even pharmaceutical agents, for either in vivo or ex vivo implantation. Additionally, these materials have the capability of providing either a structural, mechanical, and or metabolic function like natural tissues [1, 2]. The objective of this field is to mimic specific features of target organ/tissue in order to motivate cellular differentiation and organize into functional tissue assembly [3, 4]. Various developed constructs have been utilized in both in vitro and in vivo environments. Among the broad range of constructs, including autologous cells for cartilage healing or regeneration, tissue-engineered ligament, bonegraft substitutes, manufactured constructs for regeneration of the cardiovascular system such as myocardium, valves, and vessels, spinal-cord repairing and nerve regeneration, and functional restoration of metabolic organs such as the pancreas and liver [4], just skin and musculoskeletal substitutes have been approved for use in the clinical phase in the United States by the Food and Drug Administration (FDA) [5]. In contrast, other predetermined applications are still under preclinical studies or regulatory evaluation [6]. The second-generation of engineered tissues could be achieved by multidisciplinary approaches, including stem-cell biology, materials science, mechanical engineering, bioengineering, chemistry, and computer-assisted modeling/simulation. Advancement of this ­technology provides promising approaches to establish a service industry (e.g., libraries for customized scaffold-matrix materials or cell/stem-cell banks). The National Institutes of Health (NIH) referred to these engineered-tissue approaches as “regenerative medicine,” which is the use of cells, biomolecules, and other materials to restore the functional architecture of an individual’s diseased or deficient tissue/organ [7]. Regenerative medicine

Stem-Cell Nanoengineering, First Edition. Edited by Hossein Baharvand and Nasser Aghdami. © 2015 John Wiley & Sons, Inc. Published 2015 by John Wiley & Sons, Inc.

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embraces the life, physical and engineering sciences to improve failed biological functions due to congenital abnormalities, disease, and aging. For successful translation of research in regenerative medicine into products for the clinical application and marketplace, the integration of sciences with several key factors, such as market opportunity, public/private funding, regulatory approval, cost reimbursement, and public perception, is an indispensable requirement [5]. The FDA has developed proper strategies for the regulatory oversight of tissue-based products since 1990 that cover most of the engineered products. Some of these strategies, such as ethical concerns, are potential barriers to the further commercial development of regenerative medicine.

Legislative Authority The responsibility for management of the artificial organs/tissues and their commercialization inside the US Federal Government has been distributed among several regulatory agencies, such as the Health Resources Services Administration (HRSA). These agencies manage the organ transplantation and bone marrow donor program. The FDA as a sciencebased regulatory agency in the US Public Health Service (PHS) regulates the remaining products [5]. The PHS regulates the legislative authority for premarket approval and ­post-market surveillance [8]. The FDA is responsible for the evaluation of tissue-engineered products in gaining approval for the marketplace, and takes corrective action to eliminate products from the marketplace when they are unsafe, misbranded, or corrupted (Figure 22.1). The FDA assures the safety of foods, cosmetics, and also the effectiveness of human and veterinary pharmacological agents together with biological/medical devices. These agencies are composed of different centers and offices: Center for Biologics Evaluation and Research (CBER), Center for Devices and Radiological Health (CDRH), Office of Regulatory Affairs (ORA), Office of Orphan Products (OOP), and Office of Combination Products (OCP) [9].

Medical device

OK

Compliance with good manufacturing practices? Yes

Registration and listing

Low risk and exempt from intense premarket evaluation?

No

Similar to previously approved, legally marketed “predicate” device? Yes Premarket approval application mandated by FDA for this type of device?

Figure 22.1  The pathway for medical device development from initiation stage, to product launch, to the premarket evaluation stage.

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The FDA determines the approval procedures based on the product’s classification, which is categorized as a drug, biological drug, and combination product (device) for human medical devices. The combination product is a mixture of either drug/biological entity or drug/ device (e.g., tissues or cells combined with a scaffold for wound healing application and drugs encapsulated in a delivery device). It is very important for sponsors to comprehend the main requirements for the FDA’s approval process and then develop their product plan. This plan should comprise the correct and meaningful studies at preclinical and clinical phases in order to validate the safety and effectiveness of a given product. The sponsor should be provide evidence for a product’s safety and effectiveness, and then regulatory assessment is conducted on a case-by-case basis. One of the important considerations for tissue engineering products is the assessment of the preclinical information, such as evolution of toxicity and immunogenicity for local/ systemic responses [10, 11]. Moreover, there are no detailed and exact regulatory mechanisms to provide an outline for introducing tissue engineering into clinical setting in the European Union (EU).

The FDA Regulations: Human Tissue for Transplantation The FDA created regulatory outlines for human tissues to prevent the introduction, transmission, or expansion of communicable diseases. The agency’s methodologies for regulation of cellular/tissue-based products are considered in order to avoid use of contaminated tissues with potential to spread infectious diseases such as AIDS and hepatitis. The manufacturers of this type of product are demanded to register their company establishments and then list their products with the Center for Biologics Evaluation and Research (CBER) at the FDA [12]. Based on the FDA regulation, every specific unit of the tissue-engineered product must be designed, produced, and assembled in a way that can substitute all the biological functions of a particular damaged and injured organ. In 1994, the FDA established a Tissue Engineering Working Group (TEWG) to recognize and report the emerging scientific and regulatory issues of tissue-engineered medical products (TEMPs). The participating members of this group are from FDA centers and offices, the Center for Veterinary Medicine (CVM), and the Center for Food Safety and Applied Nutrition (CFSAN). This group is responsible for advancing regulatory consistency for TEMPs through intercenter communication and cooperation [13, 14]. For example, Biobrane (Bertek Pharmaceuticals) and Integra (Integra Life Sciences Corp) were the first biologically based wound dressing products to be approved by the FDA as regulated devices (synthetic materials and animal tissue products) [15]. The FDA is currently reviewing their regulatory approach for tissueengineering products in order to address a wider scope of engineered products and to introduce complete requirements intended for preventing the transmission of communicable disease (www.fda.gov/cber/tiss.htm).

Ethical Issues The research subfield of regenerative medicine with the largest number of ethical issues is that involved in the study of human embryonic stem cells (hESCs). The primary concern raised in relation to hESC research is the need for donors to consent. Additionally, derivation of embryonic germ cells from fetal tissue is associated to ethical concerns about abortion. Currently, there are extensive varieties of national regulations for study on early embryos and no legislation has been approved in several member states of the USA. In the

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EU, utilization of cell nuclear replacement was permitted by the Swedish Research Council in 2001, and then The Netherlands and the UK accepted a regulation that permits research on surplus embryos and cell nuclear replacement [13]. Finally, most countries allow surplus frozen embryos from in vitro fertilization clinics to be used for stem-cell research with the permission of the donors [16]. In the United States, there is no specific regulation in some states to control the hESC research in private research facilities, but some have banned hESC research. Privately funded projects are subjected to state regulation, but federal funded projects are allowed only if hESCs are derived from stem-cell lines [13]. Use of xenogeneic cells for therapy is another area related to regenerative medicine which raises ethical discussion.

Public Information Websites Related to Tissue-Engineered Products Targeted search results from specific databases holding information about tissue engineering and regenerative medicine, such as PubMed, in addition to general databases such as Google and Google Scholar, can pinpoint various applications of nanomedicine and related products. The data gathered from journal articles, websites of consumers and clinics, commercial websites, patents, and manufacturer documents, can reveal information about the number of registered clinical trials for tissue-engineered products. ClinicalTrials.gov is the main US database for web-based clinical trial registries that is maintained by the National Library of Medicine (NLM) at the NIH. Moreover, there are numerous supplementary sources, such as the Australian New Zealand Clinical Trials Registry (ANZCTR) (www.anzctr.org.au), the Chinese Clinical Trial Registry (ChiCTR) (www.chictr.org) which is supported by Chinese Evidence-Based Medicine Center, the European Union (EU) Clinical Trials Register website (www.clinicaltrialsregister.eu), the WHO International Clinical Trials Registry Platform (WHO ICTRP) (http://www.who.int/ictrp/), Current Controlled Trials (www.controlledtrials.com), International Federation of Pharmaceutical Manufacturers & Associations (www.ifpma.org) and Stroke Trials (www.strokecenter.org/trials/), that can be used for obtaining data about tissue-engineered products.

Nanoengineering in Clinical Application Potential applications of adult or embryonic stem cells have evoked a great amount of attention in regenerative medicine. Stem cells offer hope for a wide range of diseases or deficiencies such as type I diabetes mellitus, parkinson, liver destruction, or congestive heart failure [17, 18]. Therefore, investigations have been widely designed for appropriate regulation of stem-cell function [19]. Numerous physically or chemically inductive signals, such as growth factors, have recently been suggested to control stem-cell behavior or direct the differentiation process in vitro and in vivo (Table 22.1) [20, 21]. Metal ions, inorganic substances, and organic molecules with different quantities in culture can also tailor the stem-cell response. These molecules can be utilized within the structure or delivered to the substrate for retaining or reprogramming cell fate (Table 22.2) [22]. The surface properties of the natural tissues, such as topography or energy, can be well mimicked by nanodimensional materials (less than 100 nm) [23]. It is widely believed that cells are naturally surrounded by a nanodimensional environment and respond to nanometric alteration [23, 24]. On the other hand, precise intercellular bioactivity can be achieved by using nanostructures in macromolecular delivery, which is essential for commercialization of cellular-based products [25]. Advances in nanotechnology have lead to the development

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Table 22.1  Examples of FDA approved growth factors that can be delivered by nanocarriers Commercial Name

Company

Disease and treatment

FDA approval status

Insulin like growth factor

Increlex

Tercica

Severe primary IGF-1 deficiency

Approved

Insulin like growth factor

Iplex

Insmed

–

Approved in 2005 and with drawn in 2007

PDGF-BB (becaplermin)

REGRANEX

Healthpoint Biotherapeutics

Neuropathic diabetic ulcers

–

Bone graft consists of a genetically engineered human protein (rhBMP-2)

InFUSE™

Medtronic Sofamor Danek

Lumbar tapered fusion device

Approved

BMP-7

OP-1

Stryker

Treatment of long-bone nonunion fractures

–

Platelet-derived growth factor

GEM 21S®

Osteohealth

Periodontally related bone defects

Approved in 2008

Name

of efficient targeting of therapeutic agents with controllable concentration at the appropriate time and site [26, 27]. Growth factors are soluble polypeptides that initiate intercellular signals to regulate cell functions such as proliferation or differentiation. Due to the short biological half-life and nonspecific distribution, these molecules can be delivered to the site of injuries via nanoparticles as molecular carriers to enhance the efficacy and retention effect [28]. In order to achieve the specific and accurate identification of target sites for appropriate delivery, ­surface modification of carriers is suggested. However, several limitations including insufficient purification, costly preparation, and risk of adverse immune reaction have restricted the clinical investigations on biological recognition molecules. For this reason, some researchers have probed the physical variables (such as size or geometry) instead of chemical modifications to improve the degree of specific targeting [29]. The nanosized carriers are categorized from different points of view, such as material type, structure or application. Some important examples of nano delivery systems are highlighted in Figure 22.2. The characteristics of nanostructures are critical for regulating cytotoxicity or cell function. For example, investigation on inorganic structures with different dimensions showed that regulatory procedures require extra attention when the size falls below 30 nm [30]. Figure 22.3 shows the developmental stages of nanomedicine from beginning to commercialization. Current application of nanotechnology in medicine is ­categorized according to diagnostics or therapy. However, it has been predicted that “nanomachines” with the capability of combined therapy and diagnostics (theranostics) in human beings will emerge in future [31].

Nanocarriers Liposomes A liposome is an artificially constructed spherical vesicle (tiny bubble) composed of a phospholipid bilayer, which can enhance the pharmacokinetics and biodistributions of drugs or  growth factors [32, 33]. Tanaka et al. reported that injection of magnetic liposomal

Endorem DOTAP: Cholesterol-fus1

Acute myeloid leukemia

Sarcoma

Stable angina, heart failure and atherosclerosis

Multiple sclerosis

–

–

Crohn’s disease

Multiple myeloma and plasma cell neoplasm neurotoxicity

Acute myeloid leukemia

Multiple myeloma

Myocardial infarction

Allogeneic hematopoietic progenitor cell

Chemotherapy-resistant sarcoma stem cell

Allogeneic stem cells

Mesenchymal stem cells

Healthy volunteer/cell for tracking

Transfer a gene (fus1) into cancer cells, using the drug DOTAP: Cholesterol-fus1

Adipose-derived stem cells for treatment of complex perianal fistulas

Hematopoietic stem cell and peripheral blood stem cell

Hematopoietic stem cell and peripheral blood stem cell

Autologous stem-cell

bone marrow stem cell

Intracoronary cardiac stem cells injection

Intracoronary transfer of bonemarrow cells augment left ventricular function recovery of the heart

–

–

–

Coronary artery disease and congestive heart failure

Myocardial infarction

–

–

–

G-CSF granulocyte-colony stimulating factor

Bortezomib and dexamethasone

–

Phase 2

Phase 1

Phase 4

Phase 2

Phases 2 and 3

Phase 2

–

Bortezomib, dexamethasone, melphalan, thalidomide Cyclophosphamide, busulfan, sargramostim

Phases 1 and 2

Fibrin glue

Phase 1

–

Cx601

–

Superparamagnetic particles of iron oxide

Phases 1 and 2

Superparamagnetic iron oxide particle (Feridex)

Tagging MSCs with cell tracking drug for detection by MRI

Phases 1 and 2

Transplantation of iron-bearing nanoparticles

–

Phases 1 and 2

Phase 2

Phase 2

Stage

Liposome

Liposome

Liposome

Carrier

Doxorubicin

Amphotericin B

Amphotericin B

Fungus diseases

Allogeneic hematopoietic stem cell

Drug

Disease

Cell/procedure

Table 22.2  Stem-cell clinical trials with the potential of nanocarriers for delivery of molecules (data updated from clinicaltrials.gov)

NCT00264316

NCT00474461

NCT00126100

NCT00458822

NCT006305650

NCT00792142

NCT01372969

NCT00059605

NCT00972946

NCT00781872

NCT01270139

NCT00949325

NCT00391014

NCT00148148

Number

 22  Stem-Cell Nanoengineering from Bench to Bed

PEGylated Liposome 1990

Liposome and drug delivery in 1960s

First liposomebased siRNA delivery in 2001 Long circulating Controlledrelease PLGA-PEG nanoparticle 1994 microchip in 1999

First published dendrimer in 1978

387

First targeted Dendrimersome delivery of first described siRNA in 2010 entered clinical trials in 2008

Drug delivery First clinical study of Co-delivery of First targeted by microneedle a targeted polymer- drugs and siRNA liposome in Polymersome drug conjugate 2002 to treat multi-drug 1998 1980 in 1999 resistance in 2008 First polymerFirst controlled drug conjugate release polymer approved by the “PRINT” technique system for FDA in 1990 used to encapsulate macromolecules Doxil approved bioactive agents 2005 in 1976 by FDA in 1995

Figure 22.2  Examples of delivery mechanisms that serve as important milestones throughout the history of drug delivery systems. Reprinted with permission from ref. [27] © 2010 American Chemical Society. Nanomedicine technology development pipeline Years to commercialization 7–20 years

1–7 years

Nanomedicine applications in development

Nanomedicine clinical investigations

Basic nanoscience Application development research Theory

Preclinical animal study

Clinical study

Near-term impact Now Commercial nanomedicine products

Commericial products

Use

Figure 22.3  General steps of nanomedicine development. Reprinted with permission from ref. [30]. © 2013 Elsevier.

t­ransforming growth factor (TGF) beta 1 into articular cartilage defect accelerated the cartilage repair [34]. Transduction of plasmid containing vascular endothelial growth factor genes was also examined [35]. In  another study, transportation of nerve growth factor through blood brain barrier was promoted by liposomal encapsulation [36].

Polymeric Nanoparticles Nanoparticles based on biocompatible and biodegradable polymers have been widely investigated as therapeutic carriers [37]. Polymers can be designed and modified physically or chemically to enhance the efficiency of targeting. Size, shape, charge, stiffness, or

388

Nanotissue Engineering and Clinical Applications

s­ urface groups may be engineered to increase the cellular uptake or tissue accumulation [38, 39]. The size of particles is critical for the effectiveness of many therapeutic agents during delivery to the brain due to the blood brain barrier. Research has shown that particles with a diameter of less than 64 nm can move though the brain, while larger diameter (114 nm) particles densely coated with polyethylene glycol can diffuse within the brain [40]. Polymeric nanoparticles are able to be functionalized with biological molecules that accumulate in tumors while bypassing healthy cells [41]. Farokhzad et al. investigated clinically nanoparticles with targeting molecules that recognize an antigen expressed on the surface of most prostate tumor cells [42]. Although, much research has been conducted on the concept of targeted delivery, few formulations reached clinical trials due to the potential risks of conjugation challenges, such as finding the appropriate ligand receptor. In addition, some cancer cells, including cancer stem cells, do not show any upregulation in specific receptors [43]. Tumor cells can be specifically identified by monoclonal antibodies and lysed by tumorkilling substances without harming normal cells. For example, FDA approved Cetuximab for use in squamous cell carcinoma and colorectal carcinoma [44]. Conjugation of peptide sequences, such as arginine-glycine-aspartate (RGD), to the nanostructures has been proposed previously. For example, it was reported that a poly(d, l-lactide-co-glycolic) (PLGA) nanoparticle conjugated with RGD can be used to deliver therapeutic agents at inflammatory sites expressing the upregulated intercellular cell-adhesion molecule-1 [45]. In another study, RGD conjugated to the poly(amidoamine) dendrimers was utilized for gene delivery into mesenchymal stem cells (MSCs) [46]. These oligopeptides, as well as extracellular matrix (ECM) molecules can directly regulate stem-cell functions through the interaction between integrins on the cell surface and ligands on the ECM molecules [47]. Other small molecules like retinoic acid or fluoxetin can be utilized for gene expression modulation [48, 49].

Ceramic Nanostructures Targeted drug delivery by ceramic nanoparticles has drawn special attention during recent years. The porous structure of ceramic materials at the nanoscale size, stability of their size at body temperature or pH, and also nonswelling behavior in aqueous media are the main characteristics of these structures [50]. In addition, biocompatibility, alteration in their surface-charge density, and functionality by changing their stoichiometry, make them suitable for both drug and growth factor delivery [51]. Moreover, surface functionalization of ceramic-based particles can be easily utilized for conjugation of biomolecules to target specific ligands [52]. Prasad et al. synthesized silica-based nanoparticles with different functional groups, such as hydroxyl, thiol, amine, and carboxyl [53]. Entrapment of a water-insoluble anticancer drug in silica [54, 55], ibuprofen release from alumina [56], valproic acid delivery to the brain by titania [57], and insulin delivery by a calcium phosphate dehydrate core [58], are examples of investigated ceramic-based structures for controlled release. Hui et al. [59] evaluated the ability of silica, hydroxyapatite, and zirconia nanoparticles as gene delivery vehicles. Adair et al. [60, 61] developed colloidally stable calcium phosphate nanoparticles for hydrophobic antineoplastic and fluorescence agent delivery. In other research [62], doxorubicin was loaded on PEGylated silica nanoparticles. Titania nanotubes were also suggested for delivery of different drugs such as albumin, sirolimus, or paclitaxel [63]. The release rate of drugs can be controlled by alteration of nanotube geometry, including diameter, length, and wall thickness [64].

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Nanoparticle and Tissue Regeneration The mineral phase of natural bone consists of a calcium phosphate material widely known as hydroxyapatite (HA, Ca10(PO4)6 OH2) [65]. A variety of inorganic compositions have been commercialized for biomedical application, including bioactive calcium-phosphate-derived materials (such as HA or tricalcium phosphate (TCP, Ca3 (PO4)2)), bioactive bioglass– ceramic, or bio-inert materials (e.g. alumina, zirconia ,or pyrolytic carbon). Coating of metallic prosthesis with the aim of enhancement in abrasion resistance, blood compatibility, therapeutic delivery systems, or bone-tissue integrations are the common application for bioceramics [66]. In addition, bioactive bioceramics, alone or in combination with polymers, can be utilized as bone regenerative materials. Nanocomposites of gelatin–HA [67], collagen–HA [68], poly(lactide-glycolide)–HA [69], polyurethane–fluorohydroxyapatite ­ [70], polycaprolactone–bioglass® 45S5 [71], and carbon nanotubes–bioglass [72], are a few examples of nanostructures in “regenerative medicine.” However, it is well established that for defects larger than a critical size, cell–scaffold construct together are required to be approved in clinical setting [73]. On the other hand, incorporation of osteogenic factors (such as TGF-β2, bone morphogenic protein 2 (BMP2), or insulin-like growth factor (IGF)) into the scaffold structure may be considered as a promising therapy for regenerating bone tissue in critical defects [74]. Some researchers have introduced the process of immobilization of organic molecules on ceramic structures, for example, Pompe et al. described a protocol for immobilization of growth factors on glass surfaces by means of a thin layer of poly(octadecene-alt-maleic anhydride) [75].

Nanostructure Role in Clinical Tissue Regeneration The intrinsic characteristic of nanostructures increases the chance of noncovalent interactions with macromolecules due to the large contact area [76]. The same as the nanoparticles, nanostructures such as nanofibers can be designed for delivery of bioactive molecules [77]. It has been shown that the profile of protein adsorption to fibrous and nonfibrous structures with the same chemistry is different [78]. In addition to the importance of the chemical interactions, our findings also confirmed that the protein adsorption on a nanoweb physically depends on the fiber alignment, size distribution, bead formation, and porosity (unpublished data). Incorporation of nerve growth factor in ε-caprolactone and ethyl ethylene phosphate [79], basic fibroblast growth factor in collagen coated with perlecan [80], and plasmid DNA into poly(lactide) –poly(ethylene glycol) nano-fibers [81], are examples of biofunctional tissue-engineered fibrous structures. Moreover, biologically inductive molecules such as drugs, enzymes, or cytokines can be immobilized on the surface of nanofibers through chemical (e.g. covalent bonding) reactions [82]. It has been suggested that nanofibrous structures possess the inherent potential for differentiation of stem cells to specific cell lineages. These fibers can induce differentiation signals due to their similarity to the nanofibrous structures of natural ECM [83]. Differentiation of MSCs cultured on silk fibers to ligament fibroblasts [84], PCL fibers ­containing retinoic acid to neural cells [85], and also myogenesis [86] have been reported. Conductive and inductive chemical or physical cues introduced by ECM are essential for altering stem-cell phenotypes [87]. According to the aforementioned properties, electrospun fibers have drawn much attention for commercialization of biomedical applications. However, several drawbacks, such as protein denaturation (by using ultrahigh voltage and severe solvents) or disproportional fiber diameters (related to the natural ECM), have confined their usage [88, 89].

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Nanotopographies The stem-cell niche is well known as a nano-/microenvironment with inductive factors that controls the cell function [36, 90, 91]. The size and shape of the specialized cells and tissues are considered as determinant factors for cell function [92]. Since micro- or nanoscale variations in topographical shape and geometry (ridge, groove, and pillar) can be sensed by the cells, cell attachment, proliferation, and differentiation are affected by these variations (Figure 22.4) [93]. The cells can be embedded in vertical structures formed by columns, pits, or pillars and their function changes with these configurations. Based on this hypothesis, specified topographies have been designed to mimic the structure of the natural ECM [94]. Nanostructured surfaces were created from polycaprolactone using electron beam lithography to maintains stem-cell function over eight weeks [95]. It was shown that nanofeatures on the surface of the bone-tissue engineered scaffolds play an inductive role in osteogenicity of MSCs in the absence of osteogenic supplements [96]. The migration speed of fibroblasts was increased when cultured on a pillar form structure compared to a flat surface [97]. Moreover, nanopillars can increase the rate of transfection efficiency and endocytosis of drugs and genes [98]. It seems that in future, specific topographies for each cell lineage will be applied on tissue culture plates to control the proliferation or differentiation of stem cells. In spite of much research on cell morphology, regulation in two-dimensional and cellular alignment in three-dimensional engineered structures remain a challenge [99]. Khademhosseini et al. proposed methacrylated gelatin hydrogels as a three-dimensional environment to control the morphology of fibroblasts, myoblasts, endothelial cells, and cardiac stem cells [99].

Ridge width

Height

Top width Bottom width

Figure 22.4  Schematic cross-sectional image showing different topographical parameters.

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Smart Substrates Overall control of stem-cell activity, such as proliferation and differentiation, are affected by a varied array of environmental factors (e.g., nanofeatures on the substrates). Specifically, numerous investigations in this field have demonstrated the impact of the ECM on stem-cell fate by physical interactions, ECM geometry or topography, ECM mechanical properties, and cell geometry at the nanoscale [100, 101]. Very recently, Mahmoudi et al. [100] produced smart nanoenvironments using cell-imprinted substrates based on various matured cell types as templates. Stem cells seeded on these cell-imprinted substrates were driven to adopt the specific shape and molecular characteristics of the cell types that had been used as the template for imprinting the cell substrate. These results suggest that the dynamic plasma membrane of stem cells is capable of adopting the imprinted shape of the matured cells in artificial substrates. The specifically induced plasma membrane shapes, which are fingerprinted according to the respective mature cell types used as the template, can control the selective activation of genes of the printed matured cells, followed by autoactivation of specific complex cell signaling and metabolomic pathways. This new strategy could pave the way for a reliable, efficient, and cheap way of controlling stem-cell differentiation and, ­consequently, change the future of commercial substrates.

Conclusions Employment of rapid advances in the nanoengineering field of regenerative medicine has resulted in the evolution of a new multidisciplinary research interest dealing with the control of stem-cell fate by nanostructures and controlled macromolecular delivery by nanoplatforms. Although products based on regenerative medicine, and specially those including stem-cell therapies, show excellent promise for many types of treatments, there still exist safety, scientific, manufacturing and ethical challenges. Many countries began establishing regulatory and guidance documentation for tissue engineering research and products, as well as for stem-cellbased therapies. This review examined the different regulatory processes involved in stem-cell nanoengineered research and products for clinical applications, and highlighted the guidelines that must be followed to obtain certification from regulatory agencies such as the FDA. Control of stem-cell fate using nanostructures as well as nanoparticulate biomolecule delivery will likely be necessary for most advanced cell and tissue therapies. Nevertheless, combinations of nanotechnology and stem-cell research for development of new regenerative medicine products will most likely increase the risk associated with the therapy, rendering the product to be regulated under a higher risk category, resulting in delayed commercialization.

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Index

Note: Page references in italics refer to Figures; those in bold refer to Tables acellular collagen matrices  54 α-actin 40 actin microfilaments  207 Activin/Nodal signaling pathways  28 acute myocardial ischemia (AMI)  13 adipocyte-derived mesenchymal stem cells (AD-MSCs) 7–8 adipocytes 7 adipogenic differentiation  92 adipogenic transcription factor  210 adipose-derived stem cells (ADSCs) definition 7–8,49 differentiation 328 adult stem cells (ASCs)  3–17, 49 aging of  13–14 plasticity 6–7 agarose scaffold  306 agenesis 55 aging of ASCs  13–14 alamar blue dye  174 alcoholic liver fibrosis  13 ALDH 197 alginate 53 nanofibrous matrices  103 scaffold 306 alkanethiolates 91 alkoxysilanes 150 allografting  299, 317 aluminum nanomaterials  76 aluminum oxide  76 amide scattering peaks  146 amine surface modification  349 aminosilane coating  349 amniocentesis 51 3-aminopropyl-trimethoxysilane (APTMS)-functionalized ITO  117

amniotic fluid and placental stem cells (AFPSCs) 51 amyotrophic lateral sclerosis (ALS)  13 Annexin V  173 anodic alumina oxide (AAO) nanoporous mask  114 anodization 99 anthropogenic namomaterials  70–2 antimony oxide  76 anti-Stokes shift  333 apoptosis (programmed cell death)  172–3 arginine-glycine-aspartate (RGD)-coupled silk films and scaffolds  305 asc-MYC 236 atomic-force microscopy (AFM)  72, 172 nanografting 100, 101–2 atomic-layer-deposition devices  72 atresia 55 Australian New Zealand Clinical Trials Registry (ANZCTR)  384 autografting  299, 317 AXIN2 195 5-azacytidine (AZA)  14 basic fibroblast growth factor (bFGF)  57, 148 beta-cells 53 Biobrane 383 biocides directive  80 bioelectrospraying 135–6 biomimetic nanofibrous scaffolds  123–4 effect on stem-cell behavior  130–1 biochemical 130 biomechanical 131 biophysical 130

Stem-Cell Nanoengineering, First Edition. Edited by Hossein Baharvand and Nasser Aghdami. © 2015 John Wiley & Sons, Inc. Published 2015 by John Wiley & Sons, Inc.

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398

Index

biomimetic nanofibrous scaffolds (cont’d) regulation of electrospun nanofiber  131–5 architecture designing  133–5 biofunctionalization of electrospun scaffolds 131–3 bulk biofunctionalization  131–3, 132 fiber alignment  133, 134 fiber diameter  133 polymer selection  131 pore size and porosity135 surface biofunctionalization  133 bioelectrospraying and cell-electrospinning 135–6 assessing cellular viability  135–6 possible applications  136 bionanotechnology 205–6 bioprinting 89 2,3-bis-(2-methoxy-4-nitro-5-sulfophenyl)2H-tetrazolium-5-carboxanilide (XTT) 173, 174 bisacrylamide cross-linkers  90 bismuth oxide  76 BIX 14 bone, nanotissue engineering of  304–6 bone bioreactors  213–14 perfusion method  214 rotating vessel  214 spinner flask  214 bone-marrow-derived stem cells (BMSCs)  7, 10, 12 bone ECM  301 bone morphogenetic proteins (BMP)  148, 195 BMP-2  150–1, 213 covalent conjugation to silk film  305 in PLEOF NPs  149–50 signaling 213 BMP4  28, 92 bone-marrow stromal cells  6, 7 bone-marrow transplantation  15 boron nitrides (BN)  78 Brachyury 30 brain-derived neurotrophic factor (BDNF)  148, 267–8 branched poly(disulfide amine) (B-PDA)  226, 233–4 branched/linear polyethylenimine (PEI)  226 bPEI-HA nanocarrier  233 Buckminsterfullerene 74 cadherins 208 cadmium selenide (CdSe) quantum dot  173 cadmium sulfide quantum dots  78 cadmium telluride (CdTe) quantum dot  145–6 calcium phosphate nanoparticles (CPNP)  145, 152, 226–7, 236–7

calponin-1 40 carbohydrate-based scaffolds  306 carbon fibers  306, 308 carbon nanomaterials  73–4 carbon nanotubes (CNTs)  73, 74, 75 patterns 105 carboxymethyl chitosan modification  349 cardiac stem cells (CSCs)  286 cardiomyopathy 207 cardiovascular tissue engineering  285–93 advantages/disadvantages of approaches  286 cardiac tissue engineering  287–8 cell therapies in cardiovascular diseases  285–7 nanoscaffolds and  291–3 nanotechnology and  288–93 polymeric nanofiber scaffolds  289–91, 290 vascular tissue engineering  288 catecholconjugated poly(l-aspartic acid)-b-poly(l-phenylalanine) 150 β-catenin  151, 190 cationic liposomes  223 cationized polysaccharide (CPEPS)  154 CAVEOLIN-1 (siCAV)  153 CD105  6, 206 CD11 12 CD11b  6, 206 CD14  6, 12, 206 CD19  6, 2–6 CD34  6, 9, 12, 206 CD45  6, 12, 206 CD47 175 CD49f 27 CD73  6, 206 CD79a 6 CD79α 206 CD90  6, 206 cell sensing  209 cellular imaging, nanotechnology for  347–54 fluorine nanoparticles  351 gadolinium nanoparticles  350 goals of  348 gold and tantalum nanoparticles  351–2 homogeneous nanoparticles  353–4 hybrid nanoparticles  354 iron oxide nanoparticles  349–50 manganese oxide nanoparticles  350 multimodal nanoparticles  353–4 nanobubbles 353 nanodiamonds 352 need for, in clinical setting  348 PARACEST nanoparticles  351 polymer dots  352 upconverting luminescence nanoparticles  352–3 Center for Biologics Evaluation and Research (CBER)  382, 383

Index

Center for Devices and Radiological Health (CDRH) 382 ceramic nanostructures 388 cerium oxide  76 Cetuximab 388 characteristics of human stem cells  5 chemical exchange saturation transfer (CEST) MRI  351 chemical immobilization  269, 270 Chinese Clinical Trial Registry (ChiCTR)  384 Chinese Evidence-Based Medicine Center  384 chitosan (CS)  226, 291 films, plasma-treated  269 nanocarrier 232–3 nanofiber scaffolds  291, 306 nanofibrous matrices  103 chondrocyte technology  53–4 chondrocytes, definition  7 chondroitin sulfate (CSPG)  266, 307 chorionic villous sampling  51 chromium oxide  76 chronic wounds  316–17 CHST6 197 cidofovir (Cido)  108 ciliary neurotrophic factor (CTNF)  267–8 c-jun n-terminal kinase (JNK)  210 c-kit 9 classification of adult stem cells  7–8 of stem cells  187 clinical applications  384–6, 386 clinical tissue regeneration, nanostructure role in  389 ClinicalTrials.gov  384 c-Myc  25, 27, 28, 50 coherent anti-Stoke Raman spectroscopic method (CARS)  335–7, 336 collagen 266 collagen gel/membrane scaffold  396 collagen/electrospun poly(glycolic acid) (PGA) fibers  291 gels 91 colony forming units (CFUs)  11 colony-forming unit fibroblasts (CFU-F)  303 colorectal carcinoma  388 Comet (single-cell gel electrophoresis) assay  176, 177 computed tomography  172 computer-aided design (CAD)  251 computer-aided manufacturing (CAM)  251 copper oxide  76 Cosmetics Directive  80 COX-2 176 crystal violet dye  173, 174 CS/PEI nanoparticles  233

399

Current Controlled Trials  384 cyclic voltammetry (CV)  114, 332–3 cyclodextrin-modified dendrimer (DexAM)  230 α-cypermethrin 114 cysteine-aspartic proteases (caspases)  173 cytochalasin D  194 Dacron 306 databases 384 DCN 195 decellularization of tissues  287 dendrimers  79, 178, 226 dendritic polymers  79 dental pulp stem cells  7 deoxynucleotidyl transferase dUTP (deoxyuridine triphosphate) nick-end labeling method see TUNEL DexAM 230 dexamethasone (Dex) loaded carboxymethylchitosan/ poly(amidoamine)/(Dex-loaded CMCht/ PAMAM) nanoparticles  232, 233 Dex-GMA 150 dextran 226 dextran sulphate nanofilms  292 diabetes mellitus  53, 384 4ʹ,6-diamidino-2-phenylindole (DAPI)  172, 173 diamond nanoparticles  176 diblock copolymer nanocarrier  234 dichlorodihydrofluorescein diacetate (DCHFDA) for iron oxide nanoparticles  176 differential pulse voltammetry (DPV)  116–17 2′,7′-difluorescein diacetate (DFDA) for cerium oxide nanoparticles  176 3-[4, 5-dimethyl thiazol-2-y1]-2, 5-diphenyl tetrazolium bromide (MTT)  114, 173, 174 dimethylaminoethyl methacrylate (DMAEMA) 234 2,4-dinitrophenyl (DNP) functional groups  100 dip-pen nanolithography (DPN)  99, 100 disadvantages of human stem cells  5 D-mannose modification  349 DNA hybridization  27 DNA polymerase  173 DNA–MNP complexes (“Magnetoplex”)  235 Dolly the sheep  50 doxycycline nanofibers  371 doxycycline poly(ε-caprolactone nanofibers  370–1 drawing 125, 125 drug-delivery systems  72 Duchene muscular dystrophy (DMD)  53 EB-differentiated cardiomyocytes  146 electric cell-substrate impedance sensing system (ECIS Zθ) 328

400

Index

electrical cell-substrate impedance (ECIS) 328–31 electrochemical methods  332–3 electron beam lithography  99, 191 electroporation 223 electrospinning  26, 127–8, 128, 129, 135–6, 271, 290, 368 electrospraying 127–8, 128, 129 embryoid bodies (EBs)  30, 38, 146 embryonic antigen  1 (SSEA1),  338, 339 embryonic germ cells (EGC)  25 embryonic stem cells (ESCs)  25, 27, 49–50, 146 endocytosis 226 endothelial progenitor cells (EPCs)  6, 7 endothelial stem cells  7 Eomes 30 ependymal cells  8 epidemiological studies  375 epidermal growth factor (EGF)  38, 148, 267–8 ethical concerns  6, 383–4 ethics of nanotechnology in regenerative medicine 374 European Union (EU) Clinical Trials Register website  384 europium oxide  76 extracellular matrix (ECM)  188–9 2D surfaces vs cells cultured in a native 3D environment  91 chemical features  37–8 composition  35, 37, 246–7 from varying tissues  37 interactions of stem cells and  35–42, 36 nanotopography of  39,91 perspective 41–2 stiffness 39–41 topographical features  38–9, 40 extracellular signal regulated kinases (ERK)  196, 210 extracellular signal-regulated kinase/mitogen activated protein kinase (ERK/MAPK) signalling pathway  191, 247 extremely small-sized iron oxide nanoparticles (ESION)  350 FDA (US)  382, 383 approved growth factors  385 Center for Food Safety and Applied Nutrition (CFSAN).  383 Center for Veterinary Medicine (CVM) Tissue Engineering Working Group (TEWG) 383 Feraheme 349 Feridex® 349 fibrin clot  316 fibrin glue scaffold  306

fibroblast growth factors (FGFs)  267–8 fibroblast growth factor-2 (FGF-2)  38 signaling pathways  28 fibronectin 290 fibronectin patterns  91 fibronectin scaffolds  266 fibronectin stars and flowers  210 fibronectin-coated poly(l-lysine) nanofilm  292 flickering 212 flow cytometry  173, 340 fluorescence lifetime microscopy (FLIM) 340, 341 fluorescence resonance energy transfer (FRET) 352 fluorine nanoparticles  351 focal adhesion kinase (FAK)  209 signaling 247 focal adhesions (FAs)  91, 197 focal adhesion (FA) complex  206 force isotropy  198, 198 formamidopyrimidine glycosylase (FPG)  176 Formazan dyes  173, 174 FRAT (frequently rearranged in advanced T-cell lymphomas) protein  151 fullerenes  73, 74–5 Fullerene C60 (Buckminsterfullerene)  74 fumed silica  77 G-protein-gated inwardly rectifying potassium (GIRK) channels  51 GadoCELLTrack (BioPAL)  350 gadolinium nanoparticles  350 gadolinium oxide  76 GalC 272 gallium oxide  76 Gap43 (axonal marker)  109 GATA6 195 gelatin scaffolds  292 Gene Klf4  29 glial fibrillary acidic protein (GFAP)  8 glial-derived neurotrophic factor (GDNF)  148–9, 267–8 glucose oxidation scattering peaks  146 glycidyl methacrylated dextran (Dex-GMA) 150 glycogen synthase kinase  3 (GSK3)  28 glycogen synthase kinase  3 beta (GSK-3β), inhibition of  225 glycoprotein CD133  8 glypican-1 266 gold chloride  75 gold nanoclusters  75 gold nanoparticles  145, 351–2 applications 171 as optical scattering agents  146

Index

for detection and imaging of stem cells  147 toxicity to stem cells  145–6 Goosecoid 30 graft-versus-host disease (GVHD)  13, 15 granulocyte/macrophage colony stimulating factors (GM-CSF)  176 graphene  73, 74, 74, 75, 145 hydrogenated 75 methods for the mass-production of  331 graphite  73, 74 growth-factor reduced Matrigel (gfrMG)  266 H-2 K  9 β-hairpins 255 HAuCl4 micelles  154 Health Resources Services Administration (HRSA) 382 heart valves, artificial  373, 373 hemangioblast 7 hematopoiesis, definition  7 hematopoietic stem cells (HSCs)  6, 7, 9–10 cell-surface markers  9, 9 characteristics 6 differentiation of  9 long-term 19 short-term 10 heparin sulfate (HSPG)  266 hepatocyte growth factor  148 hepatocyte-like cells  6 higher density carboxyl groups modification  349 history of stem cell research  4 HLA-DR  6, 206 human adult GSCs (haGSCs)  30–1 human embryonic stem cells (hESCs)  49–50 human leukocyte antigen G5  12 human mesenchymal stem cells (hMSCs)  88 human multipotent germ-line stem cells (hmGSCs) 27 hyaline (articular) cartilage, nanotissue engineering of  306–7 hyaluronic acid (hyaluronan (HA)  226 in bone engineering  304 in embryonic development  38 HA/gelatin scaffold  305 hydrogels 88 nanoparticles 233 scaffold 306 hybrid nanomicrofibrous PLGA matrices  253 Hydrogels 274 hydrogenated graphene  75 hydroxyapatite scaffolds  232 hydroxybutyl chitosan (HBC) electrospun nanofibrous meshes  196 1-hydroxyethylidene-1.1-bisphosphonic acid (HEDP) coating  349

401

5-hydroxymethyl-cytosine (5hmC)  14 hypospadias 54 hypoxanthine guanine phosphoribosyltransferase (HPRT) mutation assay  177 imaging window  348 immunofluorescence 340 immunoglobulins 208 in situ nick-translation (ISNT) method  173 indium oxide  76 indium tin oxide (ITO) microelectrode arrays 102, 103 inductively coupled plasma atomic emission spectroscopy (ICP-AES)  176 Inkjet-based bioprinting of cells andbiomaterials 57, 58 inorganic nanocarriers  235–7 inorganic nanotubes  78 iNOS 176 insulin 53 insulin-like growth factors (IGFs)  267–8 Integra 383 integrins  197, 208–9 interferon gamma  176 interleukins IL-1 176 IL-1β 17 6 IL-4 176 IL-6  12, 176 intermediate filaments  207 International Federation of Pharmaceutical Manufacturers & Associations  384 intestinal stem cells  7 iron nanoparticles  76 iron oxide nanoparticles  76, 349–50 KERA 197 kidney creation, by tissue engineerinng  56 kidney transplantation  47 KLF4 (Kruppel-like factor  4)  28 Klf4  25, 27, 29, 50 L-glutamate 51 lab-on-a-chip devices  38, 105 lactate dehydrogenase (LDH)  172 laminin  207, 266 laminin-derived peptides  92 laminopathies 207 laser-guided direct writing (LGDW)  57 latrunculin B  194 Lefty1 (left-right determination factor  1)  29 legislative authority  382 leukapheresis 31 leukemia 7 leukemia inhibitory factor (LIF)  28, 38, 92

402

Index

LGTIPG peptide  92 ligaments, nanotissue engineering of  308–9 Lin lineage marker  9 Lin−/low 9 Lin28  28, 51 LIN28A 14 linkers of nucleoskeleton and cytoskeleton complexes (LINC)  207 lipid droplet-associated-granules (LDAG)  340 Lipofectamine™ 2000  236 liposomes 385–7 liver-derived stem cells  7 luminescence nanoparticles  352–3 lymphoma 7 macromolecules 79 macular degeneration  13 Mafa 53 magnesium nanoparticles  76 magnesium oxide  76 magnetic activated cell sorting (MACS)  27 magnetic ferrous oxide NPs  145 magnetic nanoparticles (MNP)  26–7, 153, 226–7, 235–6 magnetic resonance imaging (MRI)  172 magnetofection approaches  235 “Magnetoplex” 235 malathion 114 mammary stem cells  7 manganese oxide nanoparticles  76, 350 Map2 (late stage neuronal marker)  109 MAP2 272 mass spectrometry (MS) technique  136 Matrigel  38, 91, 266 matrix attachment regions (MARs)  211 mechanical shear stress  90 mesenchymal stem cells (MSCs)  6, 7, 10–13, 31, 35, 49 expression of profile surface markers  12 sources of  10–11, 11 identification of  11–12 immunosuppressive properties of  12–13 nanoengineering, effects of  205–14 bone bioreactors  213–14 cell structure 207 cells and mechanical forces  207–8 cytoskeleton 207 focal adhesion complexes  208–9, 209 mechanotransduction  208, 211 nanokicking 212, 213 plasticity 303 mesenchyme 7 mesoderm-specific transcript (MEST) gene  237 metal anodization  101 metal nanmaterials  75–6

metal oxide nanomaterials  75–6 methacrylated biotin-PEG  155 methacrylated PEG-b-polylactide (PLA)-b-PEG 155 5-methyl cytosine (5mC)  14 methyl 4-hydroxybenzoate (M4H) 108 microarray-based pathway analysis  213, 213 microcontact printing  92, 99 microelectro-mechanical systems (MEMS)  72, 89 microfilaments 207 microfluidic alginate hydrogel  92 microfluidic device  38 microfluidic flow cytometry (FC)  331–2 microfluidics 92 microinjection 223 microRNA (miRNA)  224–5 microscopy-based methods  336–40 microstamps 89 microstensils 89 microtubules 207 MIP-2 176 mitogen-activated protein kinase (MAPK)  28, 210 Mixl1 30 molecules, delivery by nanoengineering  224–5 molybdenum 77 molybdenum oxide  76 Mooney–Rivlin model  88 MoS2 78 multidomain peptides (MDPs)  255 multimodal magnetic core-shells (MCNPs)  147 multiwalled carbon nanotube (MWNT)  75, 257 toxicity of  176 muscle, nanotissue engineering of  307–8 muscular dystrophy  207 musculoskeletal cells, nanotissue engineering of  299–308 bone ECM  301 cell-nanomaterial interaction  303–4 challenges 308–9 for bone  304–6 for hyaline (articular) cartilage  306–7 for ligaments and tendons  308–9 for muscle  307–8 normal, nanostructures in  301, 302 stem cells for  301–3, 302 Myc 28 myocytes 7 nanobandage 368–70 nanobelts 77 nanobubbles 353 nanocarriers  225–6, 226, 385–91 inorganic 235–7 nanoparticle and tissue regeneration  389

Index

organic 227–35 organic–inorganic hybrid  237–8 nanocolumns 211 nanocomposites 255–7, 256 nanodiamonds 352 nanodot, two-dimensional  105 nanoelectronic memory tools  72 nanofabrication 88 nanofiber fabrication techniques 124–9, 125 nanofibrous scaffold structures 26 nanofibrous structures 195–7, 196 nanoflowers 77 nanofluidics  88, 92 Nanog  14, 28, 29, 50, 51 nanografting 100 nanograting surfaces  193–5, 193, 194 nanogrooved (nanograting, nanostripes) surfaces 193–5, 193, 194 nano-iron 76 nano-islands  189, 190 nanokicking 212, 213 nanolithography  72, 99, 271 nanomachines 385 nanomagnesium 76 nanomaterials 69–80 classification of  72–3 consumer goods  70, 71 design principles  246–8 economic impact  70 environmental and health effects of  79–80 general characteristics  70–2 size limit  70 uses in nanotechnology  72 nanomechanics 88 nanomedicine 205 nanonickel 76 nanoparticles 143–55 for detecting stem cells  146–8 for gene delivery to stem cells  151–4, 152 for growth factor delivery to stem cells 148–51, 149 for patterning stem cells  154–5, 154 immobilization in scaffold  144 toxicity to stem cells  145–6 nanopatterned surfaces of cell chips  99–118 applications 106–18 3D gold nanoparticle (GNP) cell chip encapsulated in graphene oxide 108–9, 111–13 graphic oxide-nanoparticle hybrid structure 109–13, 111–13 RGD-containing polypeptides  113–18, 114–18 technology 100–6

403

nanophosphors 353 nanopillar arrays  105 nanopillars  39, 189–90 nanopits  39, 189, 211 engineered, effects of  191 nanoporous surfaces  190–3, 191 cell responses on pore-size gradients  192 nanoprotrusion surfaces  189–90 reduced cell adhesion  189–90 nanoribbons 189 nanorods 77 three-dimensional 105 nanoscaled PCL with polyvinyl alcohol (PVA) 307 nanoscience 69 nanosensors 72 nanosilver  75, 76 nanostripes surfaces  193–5, 193, 194 nanostructure scaffold  102–5 nanostructured biomaterials  88 nanotemplating 99 nanotetrapods 77 nanotissue engineering, stem-cell expansion by  245–58 advantages of nanostructured matrices for  247 design principles of nanostructured biomaterials 246–8 nanocomposites 255–7, 256 nanofibers for248–55, 249–50, 252 copolymers/blends 254–5 natural polymers  254 self-assembled peptides  255 synthetic 248–54 surface-modified nanostructures 257–8 nanotitanium dioxide  77 nanotools 72 nanotopographies 390, 390 of ECM  39,91 nanotubes 39 nanowires 77 nanozinc oxide  77 1-naphthyl phosphate (1-NP)  333 NASA bioreactor  214 National Nanotechnology Initiative (NNI)  80 natural nanomaterials  70, 254 NaYF4 nanocrystals  353 near-infrared spectroscopy  177 neocartilage formation  53 nerve growth factor (NGF)  30, 148, 267–8 nestin 8 NeuN 8 neural cells, nanotissue engineering of  265–77 effects of physico-mechanical cues  270–6 rationale for micro- and nanotopography 270–3, 271

404

Index

neural cells, nanotissue engineering of (cont'd) role of piezoelectric (SZ) substrates  275–6 substrate mechanical properties  274–5 role of ECM components/engineering ECM signals into biomaterials  266–7 role of growth factors, signals, and bioactive molecules 267–70 biosignaling molecules (ECM, growth factors, and SMs) presentation  269–70 growth factors  267–8 morphogenes 268 protein incorporation for improving cell function 267 small chemical molecules  268–9 neural-crest stem cells  8 neural stem cells (NSCs)  8 neural tissue engineering  274 neuroblasts, precursor cells and astrocytes  8 NeuroD1 195 neuron-specific enolase, and calbindin  8 neurophases 8 neurotrophin 3  30 neutral red dye  173, 174 Ngn3 53 nickel nanoparticles  76 nickel oxide  76 nicotinamide adenine dinucleotide (NADH)  340 niobium oxide  76 NMAN 5040  176 nomenclature 6 Northern blotting  340 Notch 89 NP–protein–surfactant ternary systems  148 nuclear cloning  50 nucleofection 223 oblique-incidence reflectivity difference (OI-RD) microscopy  338–9, 338 OCT 14 Oct3 50 Oct4 (Octamer-4)  27, 28, 29, 50, 51, 113, 115 Oct4 25 n-octadecyltrimethoxysilane 236 Office of Combination Products (OCP)  382 Office of Orphan Products (OOP)  382 Office of Regulatory Affairs (ORA)  382 olfactory stem cells  7 oligonucleotides (ODN) NPs  154 organ tissue engineering, clinical need for  365–6 clinical application of nanopolymers  367–74 nanofibers 368–74 nanomaterials for organ regeneration  367 nanoparticles 367–8, 369–70 scaffold for organ regeneration  366–7 stem cells for organ engineering  366

organ transplantation, history of  47 organic nanocarriers  227–35 organic–inorganic hybrid nanocarriers  237–8 osteoblasts  7, 209–10 osteocalcin 195 osteogenesis imperfecta  15 osteogenic differentiation  92 osteogenic transcription factor  210 oxygen–phosphorous–oxygen (OPO) scattering peaks  146 p38 210 p53 29 PARACEST nanoparticles  351 Parkinson’s disease  348, 384 paxillin 209 PDK4 197 Pdx1 53 PEGylation of NPs  145, 175 PEI–MNP nanocarriers  227 PEI–SiNP nanocarriers  227 peptide amphiphiles (PAs)  255, 305 peptide derivatives  255 perfluorocarbon (PFC)  57 perfluorooctylbromide 353–4 perfluoropropane 354 perfusion decellularization technology  57 peroxisome proliferator-activated receptor gamma (PPARGγ) 210 phagocytosis 175 phase separation  26, 125, 126, 290, 368 phosphoinositide  3-kinase (PI3K) signaling pathway 29 physical encapsulation  269, 270 piezoelectric effects in bone  212 Plant Protection Products Directive  80 platelet-derived growth factor (PDGF)  38, 148 platelet-rich plasma (PRP) scaffold  306 platinum nanoparticles  76, 145 pluripotent markers  115 pluripotent stem cells (PSCs)  25–31, 49–51 application of nanotechnology to  29–30 differentiation of PSCs  29–30 proliferation and self-renewal of PPSC  29 from germ cells of the testis  27–8 induced (iPSCs)  27, 30 regulation of  28–9 reprogramming and generation of  50–1 schematic of generation  26 poly (β–amino ester) (PBAE) nanocarrier  226, 231 poly(caprolacton fumarat) polymers for bone engineering  304 poly(caprolactondiacrylate polymers for bone engineering  305

Index

poly(caprolactone) (PCL) nanofibers  248–51, 371 in bone engineering  304 nanofibrous meshes  195 poly(N,N-dimethylacrylamide)-coated maghemite 349 poly(disulfide amine) nanocarrier  233–4 branched poly(disulfide amine) (B-PDA) 233–4 poly(ethylene argininylaspartate diglyceride) (PEAD) 150 poly(ethylene glycol) (PEG)  226 poly(ethylene oxide terephthalate)–poly(butylene terephthalate) (PEOT/PBT) nanofibers 254 poly(glycolic acid) (PGA)  52, 257, 306, 322 in bone engineering  304 PGA–collagen cell-seeded scaffolds6 55 poly(3-hydroxybutyrate-co-3-hydroxyhexanoate) fibrous meshes  196 poly(lactic acid) (PLA)  52, 306, 322 in bone engineering  304 nanofibers  248, 251, 269, 290, 291, 293 nanofibrous mesh  197 PLLA-co-poly-(3-caprolactone) [PLCL] 291 poly(l-lactic acid-co-e-caprolactone) (PLLA-CL) fibers  292 co-polymer nanofibers  254 poly(d, l-lactide-co-glycolic) (PLGA)  226, 322, 388 biomaterials 292 in bone engineering  304 films 269 hybrid nanomicrofibrous matrices  253 nanocarrier 231–2 nanofibers  248, 251–3 NPs  52, 148, 149, 306 plasma-treated 269 scaffold 308 poly(lactide ethylene oxide fumarate) (PLEOF) macromers  149–50, 151 poly(d, l-lactideco-glycolide)/gelatin (PLGA/Gel) composite  291 poly(1,8-octanediol-co-citrate) (POC)  291 poly(propylene fumarate) (PPF) in bone engineering  304 polyacrylamide gels  90 polyamide nanofibers  253 polyamido amine (PAMAM)  226 dendrimers  152, 229, 230 nanocarrier 229–30 PAMAM/hBMP-2 system  229–30 polyaniline (PANi)  307 polybutylcyanoacrylate (PBCA)  226 nanocarrier 235 polycaprolactone 367

405

polycaprolactone-collagen nanofibrous substrate scaffold  29 polycaprolactone-gelatin nanofibrous substrate scaffold  29 polydimethylsiloxane (PDMS)  30, 89, 90, 91, 109, 113 nanograting substrates  271 pillars 39 polyethersulfone (PES) nanofibers  253–4 polyethylene glycol (PEG)  175 hydrogel arrays  291 microsphere scaffolds  291 nanocarrier 234–5 polyethylenimine (PEI) nannocarrier  226, 227–9 polyhedral oligomeric silsesquioxane (POSS) nanocages  293 polylactic acid (PLA)  367 poly-l-lysine (PLL) nanodots  114 polymer dots  352 polymer phase separation  100–1 polymer-based microwells  38 polymer-demixing 189 polymeric nanoparticles  387–8 polymer-magnetic hybrid nanocarrier  237–8 polymer-silica hybrid nanocarrier  238 polypyrrole (PP) nanowires  101, 102 polystyrene-block-poly(2-vinylpyridine) (PS-b-P2VP) 154 polystyrene-block-polyethylene oxide (PS-b-PEO) 155 polyurethane (PU) nanofibers  253 polyvinyl alcohol (PVA)  307 polyvinylidene fluoride (PVDF) films,  276 polyvinylidene fluoridetrifluoroethylene (PVDF-TrFE) 276 positron emission tomography (PET)  172, 348 POSS-PCU nanocomposite material  371–4, 372 PPAR-gamma (peroxisome proliferator-activated receptor gamma) signalling pathway  196 pre-adipocytes 7 prevascularization 57 propidium iodide (PI)  172, 173 prostaglandin E2  12 protein-based scaffolds  306 protein transduction domain (PTD) conjugation 349 proteins, delivery by nanoengineering  225 proton sponge mechanism  226 PTGDS 197 Public Health Service (PHS) (US)  382 Public Information Websites Related to Tissue-Engineered Products  384 public perception  375

406

Index

quantitative polymerase chain reaction (qPCR) 177 quantum dots (QD)  78–9, 230 nanostructures 26 Rac signaling pathways  29 Rad51 29 Raman scattering-based methods  333–6 Raman signals  108, 110 Raman spectroscopy (fingerprinting technology)  177, 333 regenerative medicine  48–59, 205–6, 381–2 basis of  48 biomaterials in  51–2, 52 cell therapies  53–4 cell types used in  49–51 adult stem cells  49 amniotic fluid and placental stem cells (AFPSCs) 51 embryonic stem cells  49–50 pluripotent stem cells  49–51 reprogramming and generation of induced pluripotent state cells (PSCs)  50–1 stem cells from somatic cell nuclear transfer  50 50–1 challenges and future directions  56–7 definition 48 in vivo: components and behavior of cells on  51–2 overview  48 stem cells in  14–15, 16 tissue therapies  54–6 Registration, Evaluation, Authorization and Restriction of Chemicals (REACH)  80 regulation 382 reproductive cloning  50 retinal pigment epithelial cells  6 retinaldehyde dehydrogenases.  224 retinoic acid (RA)  224, 268 delivery by nanoengineering  224, 225 RA−PEI complex nanoparticles  227, 228–9 structure  225 retinol dehydrogenase  224 retro-viral transduction  223 Rex1 29 RGD-containing polypeptides  113–18, 114–18 RGD neural cell chip, three-dimensional 105–6, 105 RGD peptide conjugated SAMs  92 rhenium oxide  76 Rho-A 210 Rho-A kinase (ROCK)  210, 212–13 ROCK inhibition  212 RhoA/ROCK-dependent actomyosin contractility mechanism.  92

RIP 272 RNIAEIIKDI laminin peptide  92 ROCK pathway  247 RT-PCR 340 runt-related transcription factor  2 (RUNX2)  195, 210, 213 safety of nanotechnology  375 satellite cells  307 scaffold-free tissue engineering  287 scaffolds, surface modification of  300 scanning electron microscopy (SEM)  172 scanning tunneling microscopes (STM)  72 screening of stem cell self-renewal and differentiation on nanomaterials  327–42 electrical cell-substrate impedance (ECIS) 328–31 electrochemical methods  332–3 microfluidic flow cytometry (FC)  331–2 microscopy-based methods  336–40 Raman scattering-based methods  333–6 sebaceous duct cells  6 selectins 208 selenide quantum dots  78 self-assembled monolayer (SAM)  91, 92 self-assembly 26, 125, 127, 290–1 self-powered nanosystems  77 severe combined immunodeficiency (SCID) 27 short fibrillizing peptides  255 signal transducer and activator of transcription 3 (STAT3) 28 silica nanoparticles (SiNP)  145, 226–7, 236 silica-coated CdTe quantum dot  145–6 silicon dioxide  77 silicon oxide  76 silicone 47 silver nanoparticles  76, 145 single-walled carbon nanotube (SWCNT)  75 toxicity of  176 skeletal stem cells  6 skin cells, nanotissue engineering of  315–23 adult stem cells for skin regeneration  317–18 mesenchymal stem cells  317 adipose-derived stem cells  318 epithelial sem cells – bulge cells  318 bioactive nanofibers  322–3 electrospinning 319–20, 320 engineered ECMs  319 epithelial cells  6 extracellular matrix  318 nanotopography 319 natural polymers  320–2 chitosan 321 collagen 320–1

Index

hyaluronic acid  321 silk 321–2 skin and skin loss  315–17 healing  316, 323 procurement and grafting  317 structure  316 synthetic polymer blends  322 Smad pathways  89 SMAD1 28 small interfering RNA (siRNA), delivery by nanoengineering 224–5, 225 smart substrates  391 somatic cell nuclear transfer  25, 27, 50–1 limitations of  50 Sonic hedgehog (Shh)  89, 268 sources of human stem cells  5 SOX2  14, 51 Sox2  25, 27, 28, 29, 50, 113, 115 SOX9 (siSOX9)  153 spermatogonial stem cells (SSCs)  25, 54 squamous cell carcinoma  388 starch–polycaprolactone scaffolds  232 Stella1 29 stem-cell differentiation, ECM and  40 stem cell factor  38 stem-cell nanoengineering  87–93 delivery through scaffolds  123, 124 nanoengineering approaches to study stem-cell fate  92–3 nanoscale approaches to elucidate stem-cell and ECM Interaction  90–1 need for  87–8 physical and biological responses  88 to study stem-cell homing  90–1 to study stem-cell niche  89–90 stem-cell niches  187, 265 stenosis 55 stiffness, in stem cells  35, 36 Stokes shift  333 stress urinary incontinence (SUI)  53 stroke 13 Stroke Trials  384 stromal cells see adult stem cells (ASCs) stromal vesicular fraction  7 superparamagnetic iron oxide nanoparticles (SPION)  146, 235 surface nanotopographies, stem-cell responses to  187–98 mechanisms governing  197–8, 198 nanofibrous structures 195–7, 196 nanogrooved (nanograting, nanostripes) surfaces 193–5, 193, 194 nanoporous surfaces  190–3, 191 cell responses on pore-size gradients  192

407

nanoprotrusion surfaces  189–90 reduced cell adhesion  189–90 surface-enhanced Raman spectroscopy (SERS)  117, 146, 147, 156, 337, 337 Synapsin (late stage neuronal marker)  109 syndecan-1 266 tantalum nanoparticles  351–2 tantalum oxide  76 Taylor cone  127 Tbx3 29 TCF3 195 TCPS 196 Teflon  47, 306 telomerase 10 template synthesis  125–6, 125 tendons, nanotissue engineering of  308–9 ten-eleven translocation (TET) proteins  14 tensegrity model  211 teratoma  15, 26, 50 teratoma formation assays  27 testicular stem cells  7 tetrafluoroethylene (Teflon)  47, 306 therapeutic cloning  50 tin oxide  76 tissue engineering (TE)  47, 143 definition 245 tissue-engineered medical products (TEMPs) 383 titanium dioxide  77 titanium oxide  76 TNF-α 176 toxicology of nanobiomaterials  171–9 blood compatibility of nanomaterials  178–9 carcinogenicity and reproductive toxicity 176–8 colorimetric assays for semi-quantitative analysis of cytotoxicity  174 cytotoxicity assessment  172 genotoxicity of nanomaterials  175 genotoxicity tests  177, 178 immunotoxicity of nanomaterials  175–6 in vitro cell toxicity/viability assays  174 time-scale representation for toxicity assessment. 174 traction-force microscopy  88 trans-differentiation 6 transforming growth factor-beta (TGF-β)  12, 148 transmission electron microscopy (TEM)  172 tricalcium phosphate (TCP) in bone engineering 304 trypan blue dye  172 tubulin 207 Tuj-1  109, 272

408

Index

TUNEL 173 tungsten 77 tungsten oxide  76 ultrasmall iron oxide nanoparticles (USPIO) 349 ultrasonography 172 umbilical cord blood-derived mesenchymal stem cells (UCB-MSCs).  235 unorthodox differentiation  6 urethral stricture disease  54 vanadium oxide  76 vascular endothelial growth factor (VEGF)  53, 57, 148, 150 VEGF-165 gradient  292 vesicoureteral reflux (VUR)  53–4

vinculin 209 vitronectin 290 Water Framework Directive  80 Western blotting  340 WHO International Clinical Trials Registry Platform  384 Wingless(Wnt)  89, 195 WS2 78 yttrium oxide  76 Z* 328 zero valence iron  76 zinc oxide  76, 77 zirconium oxide  76 zyxin 209

Natural killer (NK) cell

Neutrophil

Bone

T lymphocytes

Lymphoid progenitor cell

Basophil

Eosinophil B lymphocyte

Hematopoietic stem cell Multipotential stem cell

Monocyte/macrophage Myeloid progenitor cell Platelets

Red blood cells Bone (or cartilage) Stromal cell

Bone matrix

Stromal stem cell

Hematopoietic supportive stroma

Osteoblast Lining cell

Blood vessel

Osteocyte Pre-osteoblast

Pericyte

Skeletal muscle stem cell?

Osteoclast

Marrow a dipocyte

Hematopoietic stem cell

Hepatocyte stem cell?

Adipocyte

Figure 1.1  Differentiation of hematopoietic and stromal cells.

Cell-matrix Adhesion

Growth factors Autocrine

Cell-cell Adhesion

Paracrine

Receptors

Other growth factors

Stem cell

Nucleus Niche cell

ECM

Figure 3.1  The interaction between stem cell and its niche through autocrine and paracrine growth factors, ECM molecules, and direct cell contact with the ECM and another cell.

Stem-Cell Nanoengineering, First Edition. Edited by Hossein Baharvand and Nasser Aghdami. © 2015 John Wiley & Sons, Inc. Published 2015 by John Wiley & Sons, Inc.

(B)

(A)

EMS

SEI 4.0 kV ×50.000 100 mm WD9.0 mm

(D)

(C)

Figure 3.3  Natural ECMs of varying tissues. (A) 4’,6-Diamidino-2-phenylindole (DAPI) nuclei staining shower arrangement of cells in ECM of neural tubes [63]. (B) Scanning electron microscopy (SEM) image of an osteoblast cell in ECM [64]. (C) Staining reveals arrangement of neural stem cells in the subventricular zone around a blood vessel [65]. (D) Aligned fibroblasts and myocytes in cardiac tissue [66].

(A)

(B)

25 μm

Figure 3.4  Recent achievements in stem-cell engineering using nanoscale-based approaches. (A)  Myoblast differentiation on electrospun fibers of polycaprolactone and polyaniline of varying diameters (ranging from 0.25 to 2 μm). Differentiation and morphology were found to be highly dependent on the alignment of the fibers [63]. (B) Neuronal differentiation on PDMS scaffolds, including flat and nanopillar topographies. Higher pillar heights (400 nm) increased and accelerated neuronal differentiation. Immunostaining for Tuj1, Nestin and Hoescht on 50, 120, 220, and 400 nm nanopillars (left to right, top to bottom) [36].

(A)

Glial differentiation

Neuron formation

1500

1650

0.0 900

1050 1200 1350

Raman shift/cm–1

2.0 1.6 1.2 0.8 0.4

1650

900

Raman shift/cm–1

3.2

1.6 0.8 900

Raman shift/cm–1

(H)

1 –

1.6

0.8 * 0.0

te A te B te D te C stra stra stra Sub Sub Sub

stra

Sub

Substrate D

Substrate C

1.6

*

1050 1200 1350 1500 1650

Raman shift/cm–1

†

2.4

Substrate D

2.4

1050 1200 1350 1500 1650

(G) Difference of I1656cm /a.u. ×103

β-lll tubulin / DAPI

(F)

1500

Substrate C

Intensity/a.u.×103

0.1

(E) 1656 cm–1

0.2

2.4

1470 cm–1

(D) Substrate B

Differentiated mNSC D15

D14

1656 cm–1

0.3

Undifferentiated mNSC

I1656cm–1/I1470cm–1

1050 1200 1350

Dispersed neuronal cells

Intensity/a.u.×103

0.0

Intensity/a.u.×103

1656 cm–1

Intensity/a.u.×103

(C)

900

Cell seeding

D7

0.1

GO GNP

1656 cm–1

Neurosphere

D0

Substrate A

GO GNP

+RA (10–6M)

Mouse neural stem cell (mNSC)

0.2

Low Raman signal Saturated

1470 cm–1

+RA (10–6M) Differentiation

(B)

Strong Raman signal (π–π Stacking) Polyunsaturated

*

1.2

0.8

0.4

ted

ia rent

iffe Und

d d ted iate tiate entia rent iffer d n Diffe U

ren Diffe

Figure 7.10  Raman signals from undifferentiated and differentiated mouse neural stem cells (NSCs). (a) Schematic diagram representing the method to detect the differentiated and differentiated state of NSCs using three-dimensional graphene oxide encapsulated Au nanostructures. Raman spectra of undifferentiated (black) or differentiated (blue) NSCs on (b) substrate A, (c) substrate B, (d) substrate C and (e) substrate D. (f) Confocal fluorescence images of differentiated mNSCs on substrate D showing the successful differentiation of NSCs to neuronal cells. (g) Intensity difference of Raman peak at 1656 cm−1 (C = C bond) achieved with undifferentiated NSCs subtracted from differentiated cells. (h) Relative values of the Raman intensity at 1656 cm−1 divided by the intensity at 1470 cm−1. All the Raman spectra of NSCs were subtracted by the Raman spectra of the same substrates without cells to eliminate the background signals. Results are medians of Raman signals obtained from ten different spots (*p