Stem Cells and Tissue Repair : Methods and Protocols [2nd ed.] 9781071606544, 9781071606551

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Stem Cells and Tissue Repair : Methods and Protocols [2nd ed.]
 9781071606544, 9781071606551

Table of contents :
Front Matter ....Pages i-xi
Culturing and Manipulating Mouse Embryonic Stem Cells (Chrissa Kioussi)....Pages 1-9
Generation of an Induced Pluripotent Stem Cell Line with the Constitutive EGFP Reporter (Kiel T. Butterfield, Patrick S. McGrath, Chann Makara Han, Igor Kogut, Ganna Bilousova)....Pages 11-21
Engraftable Induced Pluripotent Stem Cell-Derived Neural Precursors for Brain Repair (Ourania Zygogianni, Georgia Kouroupi, Era Taoufik, Rebecca Matsas)....Pages 23-39
In Vitro Direct Reprogramming of Mouse and Human Astrocytes to Induced Neurons (Katerina Aravantinou-Fatorou, Dimitra Thomaidou)....Pages 41-61
Preparation of Bioscaffolds Delivering Stem Cells for Neural Regeneration (Li Yao, Ashley DeBrot)....Pages 63-70
Improved Isolation of Human Vascular Wall–Resident Mesenchymal Stem Cells (Diana Klein)....Pages 71-81
In Vitro Generation of Vascular Wall–Typical Mesenchymal Stem Cells (VW-MSC) from Murine Induced Pluripotent Stem Cells Through VW-MSC–Specific Gene Transfer (Jennifer Steens, Hannes Klump, Diana Klein)....Pages 83-97
Analysis of ToothInnervation in Microfluidic Coculture Devices (Pierfrancesco Pagella, Thimios A. Mitsiadis)....Pages 99-106
Peripheral Nerve Regeneration in a Novel Rat Model of Dysphagia (Kiyoshi Sakai, Takeshi Tsuruta, Junna Watanabe, Yukiko Sugimura, Kohei Sakaguchi, Wataru Katagiri et al.)....Pages 107-113
Healing of Full-Thickness Murine Skin Wounds Containing Nanofibers Using Splints for Efficient Reepithelialization and to Avoid Contracture (Nilika Bhattacharya, Arup K. Indra, Gitali Ganguli-Indra)....Pages 115-123
Enrichment and Characterization of Human and Murine Pulmonary Mesenchymal Progenitor Cells (MPC) (Megan Summers, Karen Helm, Susan M. Majka)....Pages 125-140
Isolation and Culture of Quiescent Skeletal Muscle Satellite Cells (Francisco Hernández-Torres, Lara Rodríguez-Outeiriño, Amelia Aránega)....Pages 141-150
Isolation, Culture, Cryopreservation, and Identification of Bovine, Murine, and Human Spermatogonial Stem Cells (Pedro M. Aponte)....Pages 151-164
Long-Term Ex Vivo Expansion of Murine Spermatogonial Stem Cells in a Simple Serum-Free Medium (Hiroshi Kubota, Kazue Kakiuchi)....Pages 165-182
Generating Kidney Organoids from Human Pluripotent Stem Cells Using Defined Conditions (Sara E. Howden, Melissa H. Little)....Pages 183-192
Painting the Pancreas in Three Dimensions: Whole-Mount Immunofluorescence Method (Maricela Maldonado, Jeffrey D. Serrill, Hung-Ping Shih)....Pages 193-200
Induction of Osteoblasts by Direct Reprogramming of Mouse Fibroblasts (Hui Zhu, Joy Y. Wu)....Pages 201-212
Back Matter ....Pages 213-214

Citation preview

Methods in Molecular Biology 2155

Chrissa Kioussi Editor

Stem Cells and Tissue Repair Methods and Protocols Second Edition



METHODS

IN

MOLECULAR BIOLOGY

Series Editor John M. Walker School of Life and Medical Sciences University of Hertfordshire Hatfield, Hertfordshire, UK

For further volumes: http://www.springer.com/series/7651



For over 35 years, biological scientists have come to rely on the research protocols and methodologies in the critically acclaimed Methods in Molecular Biology series. The series was the first to introduce the step-by-step protocols approach that has become the standard in all biomedical protocol publishing. Each protocol is provided in readily-reproducible step-bystep fashion, opening with an introductory overview, a list of the materials and reagents needed to complete the experiment, and followed by a detailed procedure that is supported with a helpful notes section offering tips and tricks of the trade as well as troubleshooting advice. These hallmark features were introduced by series editor Dr. John Walker and constitute the key ingredient in each and every volume of the Methods in Molecular Biology series. Tested and trusted, comprehensive and reliable, all protocols from the series are indexed in PubMed.



Stem Cells and Tissue Repair Methods and Protocols Second Edition

Edited by

Chrissa Kioussi Department of Pharmaceutical Sciences, College of Pharmacy, Oregon State University, Corvallis, OR, USA



Editor Chrissa Kioussi Department of Pharmaceutical Sciences College of Pharmacy Oregon State University Corvallis, OR, USA

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-0654-4 ISBN 978-1-0716-0655-1 (eBook) https://doi.org/10.1007/978-1-0716-0655-1 © Springer Science+Business Media, LLC, part of Springer Nature 2020 This work is subject to copyright. All rights are reserved by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, expressed or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. This Humana imprint is published by the registered company Springer Science+Business Media, LLC part of Springer Nature. The registered company address is: 1 New York Plaza, New York, NY 10004, U.S.A.



Preface Humans have long dreamed of replacing aging, ailing, and failing body parts, with the hope of cure, or even immortality. The myth of the titan Prometheus tells us that ancient Greeks were aware of the liver’s strong potential for self-repair. After stealing fire from the Olympian gods to aid mankind, Prometheus was chained to a rock for the eagle Ethos to feed on his liver daily, and for it to grow back at night. The word ηπαρ (hepar) means “to repair oneself.” In a more spectacular way, the regenerating heads of the Hydra, of Hercules lore, may reflect poetic extrapolations of Speman-type double-headed animals, regenerating lizard tails or re-growing amphibian limbs. Certainly, the idea of regeneration had been observed in nature. Thousands of years later, Angelico depicts a black-skinned limb grafted onto an injured white-skinned soldier in the renaissance painting “Healing of Justinian by St. Cosmas and St. Damien,” and Goethe wrote about Faust’s dream of “engineering” life, all at a time when the ancient Greek word root “hepar” was being incorporated into new words associated with the liver (hepatic, hepatocyte, heparin), as its regenerative capacity was being rediscovered by modern science. The dream of regenerative tissues is very much alive and may be realized before we know it. We have discovered how to recognize and grow stem cells, and we have developed the ability to alter them with targeted genetic manipulations in animals. Humans seem to lack a built-in regenerative capacity for whole appendages observed in reptiles and amphibians. However, all vertebrates appear to repair and regenerate injured tissue using resident stem cells in tissue niches. Deriving such stem cells, from the induced pluripotent stem cells generated from patient biopsies, remains a central challenge. Modifying and implanting such stem cells appropriately presents an opportunity for technical innovation, as does the derivation of entire organs in vitro prior to implantation. It is our hope that by sharing concise experimental details the academic community can continue to contribute effectively to this field, despite the growing cost. Regenerative medicine is the process of creating living, functional tissues to repair or replace those lost due to age, disease, injury, or congenital defects. Stem cells, smart implants, and 3D printed organs are no longer just futuristic dreams but imminent realities to be crafted by science and technology. The purpose of this book is to provide a current compendium of stem cells and regenerative techniques for beginners and experienced scientists alike. Tissue can be repaired by expansion and reprogramming of (1) embryonic stem cells (Chapter 1), (2) induced pluripotent stem cells (Chapters 2, 3, and 7), or (3) mesenchymal stem cells (Chapters 6 and 11). Induced pluripotent stem cells can be driven into neuronal (Chapters 3, 4, and 9) or vascular (Chapter 7) differentiation, while mesenchymal stem cells can be used to derive skeletal muscle (Chapter 12), osteoblasts (Chapter 17), and spermatogonial cells (Chapters 13 and 14). A novel technique for monitoring the development of sub-organ microenvironments in the developing pancreas (Chapter 16) may help expand multi-cell type approaches become more relevant. Such 3D organ formation from multiple cell types is underway for tooth (Chapters 5 and 8) and kidney (Chapter 15), as are nanofibers for skin wound healing (Chapter 10). The chapters will provide state-of-the-art method descriptions and the references therein will provide a suitable starting point for exploring the vast literature that has already developed for potential future regenerative medical techniques.

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Current clinical trials on skin grafts, leukemias, and cardiac valves are underway, while many other important challenges, such as restoring proper insulin-producing cells to a defective pancreas or regenerating the motor neurons of a paraplegic, remain. The liver is no longer the only organ known to regenerate. Aging baby boomers have increased the demand for tissues and organs, with approximately half a million Americans benefiting from transplants annually. The development of alternative sources of replacement organs, generated from the patient’s own cells, will be a game changer in medicine. The scientific community needs to effectively foresee and ethically manage the social implications of these emerging technologies, or risk the eagle Ethos picking at their liver for an eternity. Corvallis, OR, USA

Chrissa Kioussi



Contents Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Contributors. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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1 Culturing and Manipulating Mouse Embryonic Stem Cells . . . . . . . . . . . . . . . . . . Chrissa Kioussi 2 Generation of an Induced Pluripotent Stem Cell Line with the Constitutive EGFP Reporter . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Kiel T. Butterfield, Patrick S. McGrath, Chann Makara Han, Igor Kogut, and Ganna Bilousova 3 Engraftable Induced Pluripotent Stem Cell-Derived Neural Precursors for Brain Repair. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ourania Zygogianni, Georgia Kouroupi, Era Taoufik, and Rebecca Matsas 4 In Vitro Direct Reprogramming of Mouse and Human Astrocytes to Induced Neurons. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Katerina Aravantinou-Fatorou and Dimitra Thomaidou 5 Preparation of Bioscaffolds Delivering Stem Cells for Neural Regeneration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Li Yao and Ashley DeBrot 6 Improved Isolation of Human Vascular Wall–Resident Mesenchymal Stem Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Diana Klein 7 In Vitro Generation of Vascular Wall–Typical Mesenchymal Stem Cells (VW-MSC) from Murine Induced Pluripotent Stem Cells Through VW-MSC–Specific Gene Transfer. . . . . . . . . . . . . . . . . . . . . . . . . . . . Jennifer Steens, Hannes Klump, and Diana Klein 8 Analysis of Tooth Innervation in Microfluidic Coculture Devices . . . . . . . . . . . . . Pierfrancesco Pagella and Thimios A. Mitsiadis 9 Peripheral Nerve Regeneration in a Novel Rat Model of Dysphagia . . . . . . . . . . . Kiyoshi Sakai, Takeshi Tsuruta, Junna Watanabe, Yukiko Sugimura, Kohei Sakaguchi, Wataru Katagiri, and Hideharu Hibi 10 Healing of Full-Thickness Murine Skin Wounds Containing Nanofibers Using Splints for Efficient Reepithelialization and to Avoid Contracture. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Nilika Bhattacharya, Arup K. Indra, and Gitali Ganguli-Indra 11 Enrichment and Characterization of Human and Murine Pulmonary Mesenchymal Progenitor Cells (MPC) . . . . . . . . . . . . . . . . . . . . . . . . . . Megan Summers, Karen Helm, and Susan M. Majka 12 Isolation and Culture of Quiescent Skeletal Muscle Satellite Cells. . . . . . . . . . . . . ˜ o, Francisco Herna´ndez-Torres, Lara Rodrı´guez-Outeirin and Amelia Ara´nega

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Contents

Isolation, Culture, Cryopreservation, and Identification of Bovine, Murine, and Human Spermatogonial Stem Cells . . . . . . . . . . . . . . . . . . . . Pedro M. Aponte Long-Term Ex Vivo Expansion of Murine Spermatogonial Stem Cells in a Simple Serum-Free Medium. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Hiroshi Kubota and Kazue Kakiuchi Generating Kidney Organoids from Human Pluripotent Stem Cells Using Defined Conditions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sara E. Howden and Melissa H. Little Painting the Pancreas in Three Dimensions: Whole-Mount Immunofluorescence Method . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Maricela Maldonado, Jeffrey D. Serrill, and Hung-Ping Shih Induction of Osteoblasts by Direct Reprogramming of Mouse Fibroblasts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Hui Zhu and Joy Y. Wu

Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Contributors PEDRO M. APONTE • Colegio de Ciencias Biologicas y Ambientales, Universidad San Francisco de Quito (USFQ), Quito, Ecuador; Colegio de Ciencias de la Salud, Escuela de Medicina Veterinaria, Universidad San Francisco de Quito (USFQ), Quito, Ecuador; Instituto de Investigaciones en Biomedicina “One-health”, Universidad San Francisco de Quito (USFQ), Quito, Ecuador AMELIA ARA´NEGA • Faculty of Experimental Sciences, Department of Experimental Biology, University of Jae´n, Jae´n, Spain; Fundacion Medina, Granada, Spain KATERINA ARAVANTINOU-FATOROU • Neural Stem Cells and Neuroimaging Group, Department of Neurobiology, Hellenic Pasteur Institute, Athens, Greece NILIKA BHATTACHARYA • Department of Pharmaceutical Sciences, College of Pharmacy, Oregon State University , Corvallis, OR, USA GANNA BILOUSOVA • Department of Dermatology, University of Colorado School of Medicine, Aurora, CO, USA; Charles C. Gates Center for Regenerative Medicine, University of Colorado School of Medicine, Aurora, CO, USA KIEL T. BUTTERFIELD • Department of Dermatology, University of Colorado School of Medicine, Aurora, CO, USA; Charles C. Gates Center for Regenerative Medicine, University of Colorado School of Medicine, Aurora, CO, USA ASHLEY DEBROT • Department of Biological Sciences, Wichita State University, Wichita, KS, USA GITALI GANGULI-INDRA • Department of Pharmaceutical Sciences, College of Pharmacy, Oregon State University , Corvallis, OR, USA; Knight Cancer Institute, Portland, OR, USA CHANN MAKARA HAN • Department of Dermatology, University of Colorado School of Medicine, Aurora, CO, USA; Charles C. Gates Center for Regenerative Medicine, University of Colorado School of Medicine, Aurora, CO, USA KAREN HELM • Division of Pulmonary and Critical Care Medicine, Department of Medicine, National Jewish Health, Denver, CO, USA; University of Colorado Cancer Center, Aurora, CO, USA FRANCISCO HERNA´NDEZ-TORRES • Faculty of Experimental Sciences, Department of Experimental Biology, University of Jae´n, Jae´n, Spain; Fundacion Medina, Granada, Spain HIDEHARU HIBI • Department of Oral and Maxillofacial Surgery, Nagoya University Graduate School of Medicine, Nagoya, Japan SARA E. HOWDEN • Murdoch Children’s Research Institute, Parkville, VIC, Australia; Department of Paediatrics, The University of Melbourne, Melbourne, VIC, Australia ARUP K. INDRA • Department of Pharmaceutical Sciences, College of Pharmacy, Oregon State University, Corvallis, OR, USA; Knight Cancer Institute, Portland, OR, USA; Department of Biochemistry and Biophysics, Oregon State University , Corvallis, OR, USA; Linus Pauling Science Center, Oregon State University , Corvallis, OR, USA; Departments of Dermatology, Oregon Health and Science University , Portland, OR, USA KAZUE KAKIUCHI • Laboratory of Cell and Molecular Biology, Department of Animal Science, School of Veterinary Medicine, Kitasato University, Towada, Aomori, Japan

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WATARU KATAGIRI • Division of Reconstructive Surgery for Oral and Maxillofacial Region, Niigata University Graduate School of Medical and Dental Sciences, Niigata, Japan CHRISSA KIOUSSI • Department of Pharmaceutical Sciences, College of Pharmacy, Oregon State University, Corvallis, OR, USA DIANA KLEIN • Institute for Cell Biology (Cancer Research), Medical Faculty, University Hospital Essen, University of Duisburg-Essen, Essen, Germany HANNES KLUMP • Institute for Transfusion Medicine, Medical Faculty, University of Duisburg-Essen, Essen, Germany IGOR KOGUT • Department of Dermatology, University of Colorado School of Medicine, Aurora, CO, USA; Charles C. Gates Center for Regenerative Medicine, University of Colorado School of Medicine, Aurora, CO, USA GEORGIA KOUROUPI • Laboratory of Cellular and Molecular Neurobiology—Stem Cells, Department of Neurobiology, Hellenic Pasteur Institute, Athens, Greece HIROSHI KUBOTA • Laboratory of Cell and Molecular Biology, Department of Animal Science, School of Veterinary Medicine, Kitasato University, Towada, Aomori, Japan MELISSA H. LITTLE • Murdoch Children’s Research Institute, Parkville, VIC, Australia; Department of Paediatrics, The University of Melbourne, Melbourne, VIC, Australia SUSAN M. MAJKA • Division of Pulmonary and Critical Care Medicine, Department of Medicine, National Jewish Health, Denver, CO, USA; University of Colorado Cancer Center, Aurora, CO, USA MARICELA MALDONADO • Department of Translational Research and Cellular Therapeutics, Diabetes and Metabolic Research Institute, Beckman Research Institute, Duarte, CA, USA REBECCA MATSAS • Laboratory of Cellular and Molecular Neurobiology—Stem Cells, Department of Neurobiology, Hellenic Pasteur Institute, Athens, Greece PATRICK S. MCGRATH • Department of Dermatology, University of Colorado School of Medicine, Aurora, CO, USA; Charles C. Gates Center for Regenerative Medicine, University of Colorado School of Medicine, Aurora, CO, USA THIMIOS A. MITSIADIS • Unit of Orofacial Development and Regeneration, Institute of Oral Biology, University of Zurich, Zurich, Switzerland PIERFRANCESCO PAGELLA • Unit of Orofacial Development and Regeneration, Institute of Oral Biology, University of Zurich, Zurich, Switzerland LARA RODRI´GUEZ-OUTEIRIN˜O • Faculty of Experimental Sciences, Department of Experimental Biology, University of Jae´n, Jae´n, Spain; Fundacion Medina, Granada, Spain KOHEI SAKAGUCHI • Department of Oral and Maxillofacial Surgery, Nagoya University Graduate School of Medicine, Nagoya, Japan KIYOSHI SAKAI • Department of Oral and Maxillofacial Surgery, Nagoya University Graduate School of Medicine, Nagoya, Japan JEFFREY D. SERRILL • Department of Translational Research and Cellular Therapeutics, Diabetes and Metabolic Research Institute, Beckman Research Institute, Duarte, CA, USA HUNG-PING SHIH • Department of Translational Research and Cellular Therapeutics, Diabetes and Metabolic Research Institute, Beckman Research Institute, Duarte, CA, USA JENNIFER STEENS • Institute for Cell Biology (Cancer Research), Medical Faculty, University of Duisburg-Essen, Essen, Germany YUKIKO SUGIMURA • Department of Oral and Maxillofacial Surgery, Nagoya University Graduate School of Medicine, Nagoya, Japan



Contributors

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MEGAN SUMMERS • Division of Pulmonary and Critical Care Medicine, Department of Medicine, National Jewish Health, Denver, CO, USA; University of Colorado Cancer Center, Aurora, CO, USA ERA TAOUFIK • Laboratory of Cellular and Molecular Neurobiology—Stem Cells, Department of Neurobiology, Hellenic Pasteur Institute, Athens, Greece DIMITRA THOMAIDOU • Neural Stem Cells and Neuroimaging Group, Department of Neurobiology, Hellenic Pasteur Institute, Athens, Greece TAKESHI TSURUTA • Department of Oral and Maxillofacial Surgery, Nagoya University Graduate School of Medicine, Nagoya, Japan JUNNA WATANABE • Department of Oral and Maxillofacial Surgery, Nagoya University Graduate School of Medicine, Nagoya, Japan JOY Y. WU • Division of Endocrinology, Stanford University School of Medicine, Stanford, CA, USA LI YAO • Department of Biological Sciences, Wichita State University, Wichita, KS, USA HUI ZHU • Division of Endocrinology, Stanford University School of Medicine, Stanford, CA, USA OURANIA ZYGOGIANNI • Laboratory of Cellular and Molecular Neurobiology—Stem Cells, Department of Neurobiology, Hellenic Pasteur Institute, Athens, Greece

Chapter 1 Culturing and Manipulating Mouse Embryonic Stem Cells Chrissa Kioussi Abstract Mouse embryonic stem cells (mESC) have the ability to self-renew due to their rapid proliferation and high telomerase activity while maintaining their pluripotency. Depending on the environment, mESC can differentiate into a broad range of cell types. These characteristics have established mESC as a tool for modeling human disease, genetic engineering, lineage specificity, stem cell-based therapies, and tissue regeneration. Here we describe a protocol for mESC expansion and differentiation. Key words Mouse embryonic stem cells, Mouse embryoid bodies, Differentiation, Mouse embryonic fibroblasts

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Introduction Mouse embryonic stem cells (mESC) were first derived in 1981 by culturing early embryos for generation of teratocarcinoma stem cells [1, 2]. Their ability to transmit to germline and generate healthy genetically normal offspring would allow them to be used routinely in genetic manipulations a decade later. mESC are karyotypically normal pluripotent cells which derive from the inner cell mass of blastocyst-stage embryos with the ability to differentiate into any adult somatic lineages in vivo and in vitro [1, 2]. They can self-renew symmetrically and maintain their karyotype and function indefinitely [3]. The pluripotent nature of mouse ES cells was demonstrated by their ability to contribute to germline and adult tissues after injection into host morula or blastocyst stage embryos [4]. mESC have also the remarkable ability to differentiate into specific lineages when in culture [5] which makes them a unique system for establishing in vitro models for early mammalian development, cell replacement therapy and tissue regeneration, with applications in drug discovery. mESC can differentiate into cells derived from all three germ layers, mesoderm, ectoderm, and mesoderm, under appropriate conditions. Three differentiation strategies are typically used to

Chrissa Kioussi (ed.), Stem Cells and Tissue Repair: Methods and Protocols, Methods in Molecular Biology, vol. 2155, https://doi.org/10.1007/978-1-0716-0655-1_1, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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differentiate mESC, (1) mESC can aggregate and form threedimensional colonies, the embryoid bodies (EBs) [6], (2) cultured directly on stromal cells [7], and (3) cultured as a layer on extracellular matrix proteins [8]. mESC have initially been established and maintained in their undifferentiated state in coculture with mouse embryonic fibroblasts (MEFs). The leukemia inhibitory factor (LIF), a feederderived molecule, promotes the proliferating and nondifferentiating state of mESC and can replace the MEF feeder layer in the presence of fetal bovine serum (FBS) [9]. Addition of the bone morphogenetic factor (BMP) 4 in the presence of LIF is capable to replace the serum requirement. Several transcription factors, including Oct3/4 [10, 11], Sox2 [12], and Nanog [13, 14], maintain the pluripotency of early embryos and mESC. Several genes that are frequently present in high levels in tumors are also contributing to the long-term maintenance and proliferation of mESC. mESC maintain their ground state in the presence under a dual inhibition of a defined medium (2i) in which the Erk1/2 signaling pathways is totally inhibited and the glycogen synthase kinase if partially inhibited [15–17]. In this chapter we provide protocols for maintenance and expansion of mESC currently used for transgenic and gene knockout studies for analysis of gene function. The in vitro differentiation protocols of mESC are used to further understand the molecular mechanisms involved in organ development.

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Materials

2.1 Tissue Culture Equipment

1. Biosafety cabinet. 2. Water-jacketed CO2 incubator, 37  C, 5% CO2 with HEPA filtration and 95% humidity. 3. Vacuum pump. 4. Water bath. 5. Inverted microscope. 6. Microcentrifuge. 7. Electroporator. 8. Microcentrifuge tubes, 1.5 mL. 9. Conical test tubes, 15 mL. 10. Serological pipettes 5, 10, 25 mL. 11. Automated pipettes, multichannel pipettes, various sizes. 12. Tissue culture treated plastic plates 15, 10, 6, 3.5 cm, 4-, 24-, 96-well. 13. Bacteria grade plastic plates, 10, 6, 3.5 cm. 14. Cryovials.

Mouse Embryonic Stem Cells Cultures

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15. Dissection tools: microdissecting scissors, forceps. 16. 0.22 μM filter units. 17. Pasteur pipets, borosilicate 900 unplugged. 18. Manual hemocytometer. 19. Glass coverslips 24  60 mm. 20. Electroporation cuvette, 0.4 cm. 2.2 Tissue Culture Reagents and Supplies

1. Mouse embryos at postpartum day 13.5. 2. MEF Medium: DMEM high-glucose medium supplemented with 15% fetal bovine serum (FBS), nucleosides, 2 mM glutamine, 0.1 g/mL penicillin–streptomycin, 0.002% ß-mercaptoethanol. 3. FBS, fetal bovine serum (see Note 1). 4. Leukemia inhibitory factor (LIF), 500–1000 U/mL. 5. mESC medium: MEF medium with LIF. 6. Trypsin 0.25% (w/v)–EDTA 0.04% (w/v). 7. Phosphate-buffered saline (PBS): 137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, and 1.76 mM KH2PO4 in ddH2O, pH 7.4 (with HCl). 8. Mitomycin C (10 μg/mL) (see Note 2). 9. Gelatin (0.1% in ddH2O, autoclaved). 10. EmbryoMax® Electroporation Buffer. 11. Neomycin (G418; 250 μg/mL). 12. Hygromycin B (50 μg/mL). 13. All-trans retinoic acid 102 M dissolved in dimethyl sulfoxide. Split into 100 mL aliquots and stored in the dark at 80  C. 14. Laminin (20 μg/mL) (see Note 3). 15. Poly-L-ornithine hydrochloride (1.5 μg/mL) (see Note 4). 16. 30 ,30 ,5-Triiodo-L-thyronine (T3) (20 ng mL) (see Note 5). 17. Human recombinant insulin (500 μL/mL).

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Methods

3.1 Preparation of Mouse Embryonic Fibroblasts

1. Aseptically dissect mouse embryos at day 13.5–15.5. Usually two pregnant ICR mice (about 12 embryos per litter) (see Note 6). 2. Remove yolk sack, head, and organs. Isolate limbs with some parts of the body by using forceps, wash with PBS, and place into a 15 mL conical tube containing 3 mL 0.25% trypsin– EDTA. Incubate for 20 min. Invert gently every 5 min (see Note 7).

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3. Stop the reaction by adding 3 mL FBS. Centrifuge cells at 300  g for 5 min. 4. Remove supernatant and resuspend pellet in 1 mL of MEF Medium. Pipette up and down to mix (see Note 8). 5. Count cells and dilute them to a final density of 2  105/mL (see Note 9). 6. Incubate cells O/N at 37  C, 5% CO2, 95% humidity. 7. The next day remove medium from plates and wash twice with PBS. 8. Dissociate cells by adding 1 mL of trypsin–EDTA and incubate at 37  C, 5% CO2, 95% humidity for 4 min. 9. Stop Trypsin activity by adding 2 mL of medium and transfer to a 15 mL conical test tube. 10. Centrifuge at 300  g for 5 min. 11. Remove supernatant and resuspend pellet in 7 mL of medium by gentle pipetting. 12. Place 1 mL in 6 and 3.5 cm gelatinized plates (see Note 10). 3.2 Grow and Passage mESC

1. Thaw a vial of mESC (see Note 11) by quickly warming it in a 37  C water bath. 2. Aseptically transfer cells into a 15 mL conical tube, add 2 mL of media, and centrifuge for 5 min at 300  g. 3. Remove supernatant. Resuspend in 4 mL of media and plate on a 3.5 cm dish with MEFs. 4. Change the medium the next day. When the culture reaches 70% confluence, aspirate the medium, rinse the dish with PBS. 5. Add 0.3 mL trypsin–EDTA and incubate at 37  C, 5% CO2, 95% humidity for 5 min. Check under the microscope for detached clumps. 6. Stop reaction by adding 2 mL of mESC Medium. Transfer to a 15 mL conical tube, centrifuge for 5 min at 300  g. Aspirate supernatant and resuspend in fresh mESC Medium. 7. Count cells and plate them on MEF feeder dishes, 106 cells/ 6 cm dish or 2  106 cells/10 mm dish. 8. Change the medium daily and keep track of the passaging number. 9. Passage mESC every 2–3 days for maintenance of freezing. Check mESC in microspore daily before and after changing the medium (Fig. 1).

Mouse Embryonic Stem Cells Cultures

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Fig. 1 Phenotypes of mESC. mESC cultured (a) onto mEFs in the presence of LIF (b) in the presence of LIF, (c) in the presence of DMSO, (d) in the presence of RA and (e) with medium only. All panels represent cultured mESC for 5 days with basic mESC culture media 3.3

Freeze ES Cells

1. Prepare fresh freezing medium, by adding 10% FCS and 10% DMSO in mESC medium (see Note 12). 2. Remove the medium from dish, wash cells with PBS, and add trypsinized cells as indicated in subheading 3.2. 3. Resuspend mESC in an appropriate volume a cold freezing medium to reach a concentration of 5  106 cells/mL/vial. 4. Place vials in a cryo-container at 80  C freezer for 24 h. Transfer vials to liquid nitrogen container for long-term storage.

3.4 Electroporate mESC

1. Three days before electroporation is to be performed, prepare 8  10 cm plates with MEF cells (as described in Subheading 3.1). 2. The morning that electroporation is to be performed feed mESC fresh medium. 3. After 2–3 h harvest mESC (as described in Subheading 3.1) and determine the cell count. 1  107 mESC is the minimum number of cells required for electroporation. Freeze any excess of mESC (as described in Subheading 3.2). 4. Centrifuge mESC required for electroporation at 300  g for 10 min, aspirate and discard the medium. 5. Resuspend mESC in 600 μL of Electroporation Buffer. 6. Add 25–40 μg of linearized DNA plasmid (purified) in 30 μL Electroporation Buffer into mESC. Mix well and leave for 5 min at room temperature. 7. Place mESC in a 0.4 cm electroporation cuvette. Electroporate at 500 μFD, 0.24 kV. The time constant produced should be around 7 ms. Place the cuvette on ice for 10 min. 8. Transfer electroporated mESC to 80 mL of mESC Medium. 9. Mix gently. Plate mESC, 10 mL per feeder plate. 10. Incubate for 24 h at 37  C, 5% CO2, 95% humidity prior to antibiotic selection.

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3.5 Select and Pick mESC Colonies

1. Transformant mESC are usually selected for resistance to neomycin (G418) or hygromycin B. 2. After 48 h, cell death should be apparent. Change the media daily. mESC-resistant colonies should reach 800 cells in approximately 7 days following electroporation. 3. The day prior to picking mESC colonies prepare an appropriate number of 24-well plates with and without MEFs. 4. Colonies selected for picking should be spaced well enough apart to ensure no contamination from surrounding colonies with defined borders. Set pipette to 15 μL, scrape the colony with the pipette tip to dislodge the colony, aspirate and transfer the colony to an empty well in a 96-well plate (see Note 13). 5. Use a new tip for each colony. Usually 20–30 mESC colonies can be picked from one 10 cm plate. 6. Add 5 μL of trypsin to each well and incubate at 37  C, 5% CO2, 95% air for 2 min. Replace MEF media in the 24-well plates with 500 μL of mESC medium. 7. Disperse mESC colony in the 96-well plate by using a multichannel pipette to break up each colony. Transfer the suspension to the 24-well plate containing 500 μL of mESC medium. 8. Remove 250 μL of mESC and transfer to a new 24-well plate without MEFs, add 250 μL ES medium in both plates with and without MEFs. The plate without MEFs will be used for genotyping, while the one with MEFs will be stored for further use and mESC expansion. 9. Change the media daily with medium supplemented with neomycin or hygromycin (50 μg/mL). 10. New colonies should be evident within a few days. If colonies are too close disperse them by breaking the colonies using a pipette tip and spread the cells. Each well should be covered with colonies in 7–10 days.

3.6 Differentiate mESC into Embryoid Bodies (EB)

1. Resuspend mESC in MEF media (without LIF). 2. Plate mESC onto bacterial grade dishes, 5  105 cells per 10 cm dish (see Note 14) (Day 0). 3. Two days later change media (Day 2). Tilt plate and aspirate very carefully the media. Cells are detached and easy to be aspirated. 4. Change media every day as in Day 2. As mESC differentiate, they form bigger clumps and settle at the bottom. 5. Keep mESC for up to 2 weeks. At Day 15, EBs develop cavities that are filled with fluid and these structures are now called cystic EBs.

Mouse Embryonic Stem Cells Cultures

3.7 NeuronalInduced mESC

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1. For induction of neurogenesis, EBs were collected (as described in Subheading 3.6) and incubated in 10 mL MEF medium in the presence of 2 μL of 500 μL RA. 2. Change the medium daily for a total of 4 days. 3. Collect EBs in a 15 mL conical tube, let them settle at the bottom, aspirate the medium carefully. 4. Plate 10–20 EBs onto a laminin/poly-L-ornithine–coated glass inside a chamber of a 4-well plate. 5. Change the MEF medium in the presence of 2 μL of 500 μL RA every 2 days for a total of 8 days. Neuronal like cells will appear around Day 4 (see Note 15).

3.8 AdipocyteInduced ES Cells

1. For induction of adipogenesis, EBs were collected (as described in Subheading 3.6) and incubated overnight in 10 mL MEF medium in the presence of 2 μL of 500 μL RA. 2. Change media daily for a total of 3 days. 3. Collect EBs in a 15 mL conical tube, let them settle at the bottom, aspirate media carefully. 4. Plate 10–20 EBs onto a gelatin-coated glass inside a chamber of a 4-well plate. 5. Change MEF medium in the presence of 20 ng/mL T3 and 500 μL/mL insulin, every 2 days for a total of 8 days. Adipocyte-like cells will appear around Day 4 (see Note 16).

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Notes 1. Serum needs to be tested for its effect in mESC differentiation. 2. If MEFs will be used as feeders for mESC using media with antibiotic selection such as G418, then mice that carry G418resistant gene need to be used. 3. Recombinant laminin is stored at 80  C. Thaw vial before use and dilute in PBS. Used freshly diluted laminin all times. Apply 300 μL laminin on each glass coverslip, incubate at 37  C for at 2 h. 4. Prepare 1.5-mg/mL (100) aliquots of poly-L-ornithine hydrochloride by diluting in H2O. Store at 20  C. 5. To prepare 20 μg/mL stock solution add 1 mL 1 N NaOH per in 1 mg 3,30 ,5-triiodo-L-thyronine; gently swirl to dissolve and add 49 mL DMEM medium. Stock solutions are stored at 20  C. Working solutions are stable for 30 days at 4  C. 6. Store as solution 1 mg/mL in PBS for 1 week at 4  C protected from light. Wear gloves for handling.

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7. The medium should be cloudy, if not incubate for another 5–10 min. 8. Supernatant will be gooey and difficult to remove with a pipette. A cutoff pipette tip is suggested for better aspiration. 9. One 15 cm confluent plate can generate five 10 cm or ten 6 cm or twenty 3.5 cm plates. 10. If MEFs are used for feeders, then they need to be plated onto gelatinized dishes. Gelatinized dishes can be prepared the day before by adding 1, 2, and 4 mL 0.1% gelatin in 3.5, 6, and 10 cm dishes, respectively, for at least 3 h. Aspirate gelatin and let dishes dry inside the biosafety cabinet. Dried gelatinized dishes can be stored for up to 2 weeks. 11. Different strains of mESC are commercially available. 12. The final concentration of FBS of the freezing medium should be 30%. 13. Before start picking mESC colonies, ensure that you are wearing gloves and face mask. All surfaces including the microscope should be wiped with ethanol prior to use. 14. Cell density influences mESC differentiation. Several cell concentrations can be used for pilot studies. 15. Cells can be fixed and immunolabeled with anti-ß III-tubulin to observe the neuronal like state. 16. Cells can be fixed and stained with Oil Red to confirm the lipidcontaining adipocytes. References 1. Evans MJ, Kaufman MH (1981) Establishment in culture of pluripotential cells from mouse embryos. Nature 292(5819):154–156 2. Martin GR (1981) Isolation of a pluripotent cell line from early mouse embryos cultured in medium conditioned by teratocarcinoma stem cells. Proc Natl Acad Sci U S A 78 (12):7634–7638 3. Smith AG (2003) Embryo-derived stem cells: of mice and men. Annu Rev Cell Dev Biol 17 (1):435–462. https://doi.org/10.1146/ annurev.cellbio.17.1.435 4. Bradley A, Evans M, Kaufman MH, Robertson E (1984) Formation of germ-line chimaeras from embryo-derived teratocarcinoma cell lines. Nature 309(5965):255–256 5. Keller GM (1995) In vitro differentiation of embryonic stem cells. Curr Opin Cell Biol 7 (6):862–869 6. Doetschman TC, Eistetter H, Katz M, Schmidt W, Kemler R (1985) The in vitro

development of blastocyst-derived embryonic stem cell lines: formation of visceral yolk sac, blood islands and myocardium. J Embryol Exp Morphol 87(1):27–45 7. Nakano T, Kodama H, Honjo T (1994) Generation of lymphohematopoietic cells from embryonic stem cells in culture. Science 265 (5175):1098–1101 8. Nishikawa SI, Nishikawa S, Hirashima M, Matsuyoshi N, Kodama H (1998) Progressive lineage analysis by cell sorting and culture identifies FLK1+VE-cadherin+ cells at a diverging point of endothelial and hemopoietic lineages. Development 125(9):1747–1757 9. Williams RL, Hilton DJ, Pease S, Willson TA, Stewart CL, Gearing DP et al (1988) Myeloid leukaemia inhibitory factor maintains the developmental potential of embryonic stem cells. Nature 336(6200):684–687 10. Nichols J, Zevnik B, Anastassiadis K, Niwa H, Klewe-Nebenius D, Chambers I et al (1998) Formation of pluripotent stem cells in the

Mouse Embryonic Stem Cells Cultures mammalian embryo depends on the POU transcription factor Oct4. Cell 95(3):379–391 11. Niwa H, Miyazaki J-I, Smith AG (2000) Quantitative expression of Oct-3/4 defines differentiation, dedifferentiation or self-renewal of ES cells. Nat Genet 24(4):372–376 12. Avilion AA, Nicolis SK, Pevny LH, Perez L, Vivian N, Lovell-Badge R (2003) Multipotent cell lineages in early mouse development depend on SOX2 function. Genes Dev 17 (1):126–140 13. Chambers I, Colby D, Robertson M, Nichols J, Lee S, Tweedie S et al (2003) Functional expression cloning of nanog, a pluripotency sustaining factor in embryonic stem cells. Cell 113(5):643–655 14. Mitsui K, Tokuzawa Y, Itoh H, Segawa K, Murakami M, Takahashi K et al (2003) The

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homeoprotein nanog is required for maintenance of pluripotency in mouse epiblast and ES cells. Cell 113(5):631–642 15. Wray J, Kalkan T, Gomez-Lopez S, Eckardt D, Cook A, Kemler R et al (2011) Inhibition of glycogen synthase kinase-3 alleviates Tcf3 repression of the pluripotency network and increases embryonic stem cell resistance to differentiation. Nat Cell Biol 13(7):838–845 16. Ying Q-L, Wray J, Nichols J, Batlle-Morera L, Doble B, Woodgett J et al (2008) The ground state of embryonic stem cell self-renewal. Nature 453(7194):519–523 17. Mulas C, Kalkan T, von Meyenn F, Leitch HG, Nichols J, Smith A (2019) Defined conditions for propagation and manipulation of mouse embryonic stem cells. Development 146(6): dev173146

Chapter 2 Generation of an Induced Pluripotent Stem Cell Line with the Constitutive EGFP Reporter Kiel T. Butterfield, Patrick S. McGrath, Chann Makara Han, Igor Kogut, and Ganna Bilousova Abstract The discovery of induced pluripotent stem cell (iPSC) technology has provided a versatile platform for basic science research and regenerative medicine. With the rise of clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated protein 9 (Cas9) systems and the ease at which they can be utilized for gene editing, creating genetically modified iPSCs has never been more advantageous for studying both organism development and potential clinical applications. However, to better understand the behavior and true therapeutic potential of iPSCs and iPSC-derived cells, a tool for labeling and monitoring these cells in vitro and in vivo is needed. Here, we describe a protocol that provides a straightforward method for introducing a stable, highly expressed fluorescent protein into iPSCs using the CRISPR/Cas9 system and a standardized donor vector. The approach involves the integration of the EGFP transgene into the transcriptionally active adeno-associated virus integration site 1 (AAVS1) locus through homology directed repair. The knockin of this transgene results in the generation of iPSC lines with constitutive expression of the EGFP protein that also persists in differentiated iPSCs. These EGFPlabeled iPSC lines are ideal for assessing iPSC differentiation in vitro and evaluating the distribution of iPSC-derived cells in vivo after transplantation into model animals. Key words Induced pluripotent stem cells, iPSCs, Nucleofection, CRISPR/Cas9, Fluorescent protein

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Introduction The technology that allows for the generation of induced pluripotent stem cells (iPSCs) from somatic cells provides an ideal platform for disease modeling, drug discovery and therapy development. iPSCs have the potential to model the diversity of genotypes within the human population, as these cells can be generated from both normal and disease-associated somatic human cells [1]. When combined with gene editing using programmable nucleases, such as clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated protein 9 (Cas9), iPSC technology

Chrissa Kioussi (ed.), Stem Cells and Tissue Repair: Methods and Protocols, Methods in Molecular Biology, vol. 2155, https://doi.org/10.1007/978-1-0716-0655-1_2, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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becomes even more valuable for studying gene function and for lineage tracing analyses [2, 3]. The ability to correct mutations or introduce de novo mutations into otherwise healthy iPSCs further broadens iPSC applications in modeling human diseases and developing novel cell therapeutics [2]. To better understand the behavior and true therapeutic potential of iPSCs, the functionality and distribution of iPSC-derived cells need to be assessed following the transplantation of these cells into model organisms. This can be achieved by the generation of iPSC lines with constitutive expression of a fluorescent protein that also persists in differentiated iPSCs. In this chapter, we present a protocol that generates iPSCs with the constitutive enhanced green fluorescent protein (EGFP) reporter using the CRISPR/Cas9 system to enable the visualization of these cells and their derivatives with the use of fluorescence microscopy. Using CRISPR/Cas9, the generation of fluorescently labeled iPSCs has become a relatively efficient and straightforward process. The CRISPR/Cas9 system is composed of two main components: the Cas9 endonuclease that can introduce a DNA double stranded break (DSB) at a specific genomic location and a guide RNA (gRNA) that directs Cas9 to a predetermined genomic locus [4]. The gRNA consists of a short variable 20-nucleotide sequence that is complementary to the DNA sequence being targeted and a constant region with several stem loop structures that serve as scaffolding for Cas 9 binding. The gRNA recognizes the complementary sequence on the genomic DNA upstream of the three base protospacer adjacent motif (PAM), preferably 50 -NGG-30 (where “N” can be any nucleotide base followed by two guanine nucleobases), and directs the endonuclease Cas9 to introduce a sitespecific DSB. In the presence of a DNA donor template that contains a sequence of interest flanked by regions homologous to those adjacent to the Cas9 cleavage site, the DSB can trigger homology directed repair (HDR) to incorporate the exogenous DNA sequence, such as the EGFP reporter, into the predetermined genomic locus of iPSCs [5, 6]. An important criterion for the successful integration of a fluorescent reporter into iPSCs is the selection of an appropriate integration site. The adeno-associated virus site 1 (AAVS1) locus has long been used a safe harbor for transgenic integration and has been shown to be transcriptionally active in multiple organs in humans, similar to the ROSA26 locus in mice [7, 8]. Incorporation of the EGFP transgene into the AAVS1 locus allows for robust and stable expression in a variety of cell types and ensures continuous and stable expression in iPSCs and iPSCderived cells following differentiation and transplantation into animal models.

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Fig. 1 The targeting donor construct for insertion of the EGFP reporter into the AAVS1 locus of human iPSCs. The donor DNA plasmid used to target the AAVS1 locus contains 50 and 30 homologous sequences of approximately 800 bp flanking the Cas9-induced cleavage site together with sequences encoding EGFP under the control of CAGGS promoter and puromycin resistance [11]. SA splice acceptor

There are several approaches to deliver the CRISPR/Cas9 complex into the cells of interest: (1) DNA plasmids encoding both the Cas9 protein and the gRNA; (2) mRNA encoding Cas9 alongside a separate gRNA; and (3) the Cas9 protein preassembled with the gRNA into a ribonucleoprotein (RNP) complex [4]. Although the CRISPR/Cas9 complex was originally delivered via plasmid DNA, the advances in the generation of mRNA, particularly modified mRNA (mod-mRNA), which is characterized by lower cellular immunogenicity than unmodified mRNA [9, 10], and RNP complexes [4] allow for a more efficient and controllable way to introduce functional Cas9 into cells. In addition, both mod-mRNA and RNP complexes are readily available through commercial sources. The provided protocol for the introduction of the EGFP reporter into the AAVS1 locus of human iPSCs relies on using mod-mRNA encoding the Cas9 nuclease delivered together with plasmid expressing the AAVS1-specific gRNA. A DNA plasmid with homology arms targeting the AAVS1 locus is used as a donor template for HDR to introduce the constitutive EGFP transgene. Additionally, the targeting plasmid employs a gene-trap strategy to convey puromycin resistance for positive selection of transfected cells (Fig. 1). This donor plasmid has been previously used to knockin EGFP into AAVS1 using zinc-finger nucleases [11]. The AAVS1-specific gRNA, which has been previously used for modification of the AAVS1 locus together with Cas9 [12], is expressed from the human U6 polymerase III promoter of the DNA plasmid delivered together with Cas9 mod-mRNA and the donor template. Nucleofection is used to deliver the plasmids and Cas9 mod-mRNA. Nucleofection is followed by selection with puromycin to eliminate nontargeted iPSCs, since any cell that does not fully incorporate the donor construct into the AAVS-1 locus will not survive puromycin treatment. Following selection with puromycin and outgrowth, iPSC colonies with uniform EGFP expression can be manually picked and expanded for downstream applications (see Fig. 2a for the schematic of the protocol). Although the protocol describes the generation of iPSCs with the constitutive EGFP reporter, a similar procedure can be used for the introduction of any other marker into the AAVS1 locus of human iPSCs.

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Fig. 2 The protocol for the generation of iPSCs with constitutively expressed EGFP. (a) Schematic representation of the procedures described in the protocol. (b) Healthy iPSCs before nucleofection. (c) A colony of iPSCs following incubation with Y-27632 on day 1 of the protocol. (d, e) Clusters of iPSCs 24 h post-nucleofection, expressing EGFP. (f, g) A colony of iPSCs with mosaic expression of EGFP on day 13 of the protocol. (h, i) A colony of iPSCs uniformly expressing EGFP on day 13 of the protocol. Scale bar, 200 μm

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Materials

2.1 Targeting the AAVS1 Locus by Nucleofection

1. Matrigel human Embryonic Stem Cell (hESC)-Qualified Matrix, LDEV-free (Corning). Matrigel solidifies rapidly at room temperature (RT). Therefore, it is recommended to aliquot each new batch of the matrix upon arrival following the manufacturer’s instructions. Use prechilled pipette tips,

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racks and tubes while working with the reagent. Store at 80  C. 2. Dulbecco’s Modified Eagle Medium/Nutrient Mixture F-12 (DMEM/F12). 3. Tissue culture (TC)-treated 6-well plates. 4. mTeSR1 basal medium supplemented with mTeSR1 5 supplement according to the manufacturer’s instructions (mTeSR1 complete medium). 5. TrypLE Select Enzyme (1), no phenol red. 6. 10 mM Rock inhibitor diluted in DMSO (Y-27632 2HCl). 7. Human Stem Cell Nucleofector Kit 2 (Lonza). 8. AAV-CAGGS-EGFP [11], available from Addgene (Plasmid #22212, a gift from Rudolf Jaenisch). Prepare a minimum of 500 ng/μL in Qiagen EB Buffer, store at 20  C (see Note 1). 9. gRNA_AAVS1-T2 [12], available from Addgene (Plasmid #41818, a gift from George Church). Prepare a minimum of 500 ng/μL in Qiagen EB Buffer, store at 20  C (see Note 1). 10. Mod-mRNA Cas9 is available commercially or can be synthesized in vitro as previously described [10]. Mod-mRNA should be diluted/resuspended in Nuclease-Free Water at no less than 500 ng/μL. 2.2 Chemical Selection for Targeted Cells

1. TC-treated 6 well plate.

2.3 iPSC Colony Picking and Clonal Expansion

1. mTeSR1 complete medium.

2. Puromycin dihydrochloride (10 mg/mL). Add 100 μL of 10 mg/mL stock solution of puromycin to 900 μL of mTeSR1 complete medium to make a 1 mg/mL working solution (see Note 2).

2. TC-treated 6-well plates. 3. 1 DPBS. 4. 0.5 mM EDTA in DPBS. Add 500 μL of 0.5 M EDTA to 500 mL of DPBS to create a working solution of 0.5 mM EDTA in DPBS and filter-sterilize using a 0.22 μm vacuum filtration system.

2.4

Equipment

1. Biological safety cabinet. 2. 37  C water bath or bead bath. 3. 37  C/5% CO2/5% O2 humidified tissue culture incubator (see Note 3). 4. Nucleofector 2b Device (Lonza). 5. Hemocytometer. 6. Centrifuge.

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Methods Work under RNase-free conditions and use aseptic techniques when possible. Perform all cell culture-related manipulations in a biological safety cabinet using aseptic techniques. Follow institutional biosafety standards for work with human cells. Once the nucleofection procedure is initiated, daily maintenance is required for approximately 15–17 days. Be sure to plan accordingly.

3.1 Targeting the AAVS1 Locus by Nucleofection

1. Prepare 1 well of a 6-well plate of iPSCs. Confirm that iPSCs are healthy with minimal differentiation (Fig. 2b). iPSC colonies should be at approximately 75–85% confluency before the nucleofection procedure is performed. If confluency is not adequate, delay the protocol until the cells have reached the required confluency. (see Note 4). 2. Coat 1 well of a 6-well plate with hESC-qualified Matrigel following the manufacturer’s instructions (see Note 5). Seal plates with parafilm and incubate for 1 h at RT to polymerize inside the biological safety cabinet. Set aside. 3. Prepare an 8 mL aliquot of mTeSR1 complete medium in a 15 mL conical tube and supplement it with 10 μM Y-27632. Invert the tube 3–4 times to mix. Place the mTeSR1 aliquot supplemented with Y-27632 in a bead bath to warm to 37  C. 4. Aspirate the spent mTeSR1 complete medium from the iPSCs to be nucleofected. Replace with 2 mL of mTeSR1 complete medium with Y-27632 prepared in step 3. Return the cells to the low O2 incubator for 2 h. The exposure to Y-27632 may slightly change the morphology of iPSC colonies (see Fig. 2c). 5. After incubation, aspirate mTeSR1 complete medium with Y-27632 from the cells and add 1 mL of DPBS to rinse. Gently rock the plate to ensure complete coverage of the cells, then aspirate DPBS. Repeat the rinse with DPBS. 6. Aspirate DPBS and add 1 mL of RT TrypLE to the well. Gently rock the plate to ensure complete coverage of the cells and return the plate to the low O2 incubator. Incubate for 3 min. 7. Remove the plate from the incubator and firmly but gently tap the side of the plate to dislodge cells. Check the cells under the microscope. If 90% of the cells are detached and floating, proceed. If the majority of cells are still attached, incubate for another 3 min. Continue to check cells every 3 min until 90% of the cells are detached (see Note 6). 8. Quickly rinse/collect the detached cells using 3 mL of mTeSR complete medium supplemented with Y-27632 from step 3 to neutralize TrypLE. Transfer the iPSC suspension into a 15 mL conical tube.

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9. Invert the cell suspension five times to fully mix and then count the cells using a hemocytometer (see Note 7). 10. Transfer 1  106 iPSCs into a new 15 mL conical. Centrifuge at 200  g for 3 min at RT. 11. During centrifugation, prepare complete nucleofection solution. For each reaction, add 81.8 μL of Nucleofector Solution to 18.2 μL of Supplement 1 in a clean 1.5 mL Eppendorf tube (4.5:1 ratio). Pipet up and down five times to mix thoroughly, taking care to avoid bubbles (see Note 8). 12. Aspirate medium from the pelleted cells. Resuspend the pelleted cells in 100 μL of complete nucleofection solution. 13. Add 2 μg of Cas9 mod-mRNA, 2 μg of AAV-CAGGS-EGFP plasmid, and 1 μg of gRNA_AAVS1-T2 plasmid to the resuspended cells from step 12. The total volume of mod-mRNA and plasmids should not exceed 10 μL. 14. Gently mix the complete nucleofection cell suspension by pipetting up and down, taking care to avoid bubbles. Transfer the cell suspension to the nucleofection cuvette. Pipet the solution along the side to avoid bubbles (see Note 9). 15. Place the nucleofection cuvette into the nucleofection chamber and execute program B-016 on the Nucleofector. Once the nucleofection is complete, add 500 μL of mTeSR1 complete medium supplemented with Y-27632 directly into the cuvette. 16. Aspirate the Matrigel solution from the well prepared in step 2, do not allow the surface of the well to dry. Add 2 mL of prewarmed mTeSR1 complete medium supplemented with Y-27632 to the well. 17. Use the provided dropper to resuspend cells in the cuvette and gently transfer the entire transfection mix from the cuvette into the prepared well. 18. Place the plated cells into a tissue culture incubator with O2 set to 5% (low-O2). Once the plate is set down, disperse the cells by alternating between an up/down then left/right motion. Repeat the motions two more times. Do not swirl the plate to mix. Incubate the cells overnight. 19. The following day (day 2), place 2.5 mL of mTeSR1 complete medium into a bead bath to warm to 37  C. Do not supplement with Y-27632. Remove the plate with nucleofected cells from the low O2 incubator and aspirate the spent medium. Significant cell death is expected within the 24 h following nucleofection. Add 2 mL of warm mTeSR1 complete medium and assess the expression of EGFP in cells using a fluorescence microscope. Cells should appear as small clusters with elongated and spindly morphology (Fig. 2d, e).

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20. Replace medium with mTESR1 complete medium 48 h post transfection (day 3), as cells continue to recover from nucleofection. 3.2 Chemical Selection for Targeted Cells

1. On day 4, transfer 6.5 mL of mTeSR1 complete medium to a new 15 mL conical tube and add 1.63 μL of a 1 mg/mL working solution of puromycin to achieve 250 ng/mL final concentration of puromycin. Close the tube and invert five times to thoroughly mix. Place the tube in a bead bath to warm to 37  C. 2. Remove the plate with nucleofected cells from the incubator and aspirate the spent medium. Add 2 mL of mTeSR supplemented with puromycin from step 1 and return the plate to the incubator. Closely monitor iPSCs for GFP expression and colony formation. 3. Replace mTeSR1 complete medium supplemented with puromycin daily for the next 2 days (days 5 and 6), for a total of 72 h of selection. Since correct targeting is a rare event, approximately 1–20 colonies should survive selection. 4. On day 7, replace the spent selection medium to fresh mTeSR1without puromycin. Continue changing medium every day. Check cells for EGFP expression and colony formation daily. Some colonies may show mosaic expression of EGFP (Fig. 2f, g) while many should uniformly express the marker (Fig. 2h, i). Colonies will be large enough for picking and passaging approximately 2 weeks following nucleofection.

3.3 iPSC Colony Picking and Clonal Expansion

1. Use a fluorescent microscope to mark iPSC colonies that uniformly express EGFP for subsequent picking and expansion. The colonies should be large and clearly formed. Plan to pick as many colonies as possible - up to 12. 2. Prewarm 30 mL of mTeSR1 complete medium to 37  C. 3. Coat up to 12 wells of a 6-well plate with hESC-qualified Matrigel, depending on the number of observed colonies (see Subheading 3.1, step 2 for the coating procedure). Seal plates with parafilm and incubate for 1 h at RT. 4. Aspirate the Matrigel solution from the coated plates and add 2 mL of mTeSR1 complete medium per well. Do not supplement with Y-27632. Do not allow the surface of the wells to dry. Set the plates aside. 5. Aspirate spent mTeSR1 complete medium from wells with targeted cells. Rinse once with 1 mL of 0.5 mM EDTA and aspirate (see Note 10). 6. Add 1 mL of 0.5 mM EDTA per well. Incubate for 4 min at 37  C.

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7. Gently remove the plate from the incubator and place it in the biosafety cabinet (see Note 11). 8. Carefully aspirate EDTA from the well. Very gently add 3 mL of prewarmed mTeSR1 complete medium, taking care not to dislodge iPSC colonies. 9. Move the plate to an inverted or dissecting microscope to better visualize colonies. Prepare a sterile 1 mL pipette. Fully depress the plunger. Then, use the pipette tip and gently scrape a colony while slowly drawing liquid into the tip to collect the colony. Draw as little medium as possible while picking the iPSC colony. 10. To transfer the colony, pipet up and down 3–4 times in a single previously prepared well from step 4. Repeat steps 9 and 10 until as many as 12 colonies have been picked and transferred into individual wells (see Note 12). 11. Move plated cells back into the low O2 incubator. To ensure even cell distribution, shake each plate back and forth and side to side. Do not swirl. Replace mTeSR1 complete medium daily.

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Notes 1. Bacterial stocks are grown and plasmid DNA is isolated using the Qiagen Midi Prep kit following the manufacturer’s instruction. 2. The efficiency of puromycin may decline with repeated freezing and thawing. Prepare a 1 mg/mL working stock solution of puromycin immediately before use. 3. This protocol has been optimized for iPSCs cultured in a Tri-Gas incubator set for 5% oxygen (low O2). The procedure should work with similar efficiency for ESCs or iPSCs cultured under normoxic conditions (i.e., 20% oxygen). 4. iPSCs are cultured in mTeSR1 complete medium in a 6-well plate, following standard human cell culture procedures. Rock inhibitor is not used for routine passaging. One well of a 6-well plate yields approximately 1–2  106 cells when grown to a confluency of 75–85%. If differentiation is present, manually pick and remove the differentiated areas, passage cells, and nucleofect when optimal confluency is reached. If the confluence of cells is higher than 85%, split cells at a lower density and proceed with the protocol when cells have reached ideal confluency. 5. The dilution of Corning Matrigel is calculated for each lot based upon the protein concentration. Appropriately sized aliquots should be prepared according to the dilution factor provided by the manufacturer in the certificate of analysis.

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6. Do not leave TrypLE on the cells for more than 12 min, as this will greatly reduce cell viability in downstream applications. If cells are not detached within 12 min, a cell scraper can be used to mechanically dislodge iPSCs. 7. iPSCs can remain clustered together following the treatment with TrypLE. When counting cells using a hemocytometer, estimate the number of cells in the cluster. 8. Human iPSCs are very sensitive to changes in environmental conditions. Therefore, proceed with the nucleofection steps as fast as possible. 9. The presence of bubbles will negatively affect the nucleofection process by interfering with the ability of the solution to acquire a consistent electric charge. 10. To avoid cytotoxicity, EDTA must be diluted to a working stock of 0.5 mM in DPBS before using it to detach cells (see Subheading 2.3, item 4). 11. At this point, cells may be very loosely adhered and easily dislodged. 12. Do not combine colonies into the same well; each picked colony should receive its own well.

Acknowledgments We are grateful for funding support from the National Institutes of Health (P30 AR057212 and R21 AR074642). We also thank the Gates Frontiers Fund. References 1. Kim C (2014) Disease modeling and cell based therapy with iPSC: future therapeutic option with fast and safe application. Blood Res 49 (1):7–14 2. Kehler J, Greco M, Martino V, Pachiappan M, Yokoe H, Chen A, Yang M, Auerbach J, Jessee J, Gotte M, Milanesi L, Albertini A, Bellipanni G, Zucchi I, Reinbold RA, Giordano A (2017) RNA-generated and geneedited induced pluripotent stem cells for disease modeling and therapy. J Cell Physiol 232 (6):1262–1269 3. Roberts B, Haupt A, Tucker A, Grancharova T, Arakaki J, Fuqua MA, Nelson A, Hookway C, Ludmann SA, Mueller IA, Yang R, Horwitz R, Rafelski SM, Gunawardane RN (2017) Systematic gene tagging using CRISPR/Cas9 in human stem cells to illuminate cell organization. Mol Biol Cell 28(21):2854–2874

4. Lino CA, Harper JC, Carney JP, Timlin JA (2018) Delivering CRISPR: a review of the challenges and approaches. Drug Deliv 25 (1):1234–1257 5. Ahn JH, Chu HS, Kim TW, Oh IS, Choi CY, Hahn GH, Park CG, Kim DM (2005) Cell-free synthesis of recombinant proteins from PCR-amplified genes at a comparable productivity to that of plasmid-based reactions. Biochem Biophys Res Commun 338 (3):1346–1352 6. Ratner HK, Sampson TR, Weiss DS (2016) Overview of CRISPR-Cas9 biology. Cold Spring Harb Protoc 2016(12). https://doi. org/10.1101/pdb.top088849 7. Bak RO, Porteus MH (2017) CRISPRmediated integration of large gene cassettes using AAV donor vectors. Cell Rep 20 (3):750–756

Generation of Induced Pluripotent Stem Cells with the EGFP Reporter 8. Sadelain M, Papapetrou EP, Bushman FD (2011) Safe harbours for the integration of new DNA in the human genome. Nat Rev Cancer 12(1):51–58 9. Youn H, Chung JK (2015) Modified mRNA as an alternative to plasmid DNA (pDNA) for transcript replacement and vaccination therapy. Expert Opin Biol Ther 15(9):1337–1348 10. Vaidyanathan S, Azizian KT, Haque A, Henderson JM, Hendel A, Shore S, Antony JS, Hogrefe RI, Kormann MSD, Porteus MH, McCaffrey AP (2018) Uridine depletion and chemical modification increase Cas9 mRNA activity and reduce immunogenicity without

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HPLC purification. Mol Ther Nucleic Acids 12:530–542 11. Hockemeyer D, Soldner F, Beard C, Gao Q, Mitalipova M, DeKelver RC, Katibah GE, Amora R, Boydston EA, Zeitler B, Meng X, Miller JC, Zhang L, Rebar EJ, Gregory PD, Urnov FD, Jaenisch R (2009) Efficient targeting of expressed and silent genes in human ESCs and iPSCs using zinc-finger nucleases. Nat Biotechnol 27(9):851–857 12. Mali P, Yang L, Esvelt KM, Aach J, Guell M, DiCarlo JE, Norville JE, Church GM (2013) RNA-guided human genome engineering via Cas9. Science 339(6121):823–826

Chapter 3 Engraftable Induced Pluripotent Stem Cell-Derived Neural Precursors for Brain Repair Ourania Zygogianni, Georgia Kouroupi, Era Taoufik, and Rebecca Matsas Abstract Stem cell transplantation has attracted great interest for treatment of neurodegenerative diseases to provide neuroprotection, repair the lesioned neuronal network and restore functionality. Parkinson’s disease (PD), in particular, has been a preferred target because motor disability that constitutes a core pathology of the disease is associated with local loss of dopaminergic neurons in a specific brain area, the substantia nigra pars compacta. These cells project to the striatum where they deliver the neurotransmitter dopamine that is involved in control of many aspects of motor behavior. Therefore, cell transplantation approaches in PD aim to replenish dopamine deficiency in the striatum. A major challenge in developing cell therapy approaches is the ability to generate large numbers of transplantable cells in a reliable and reproducible manner. In recent years the technological breakthrough of induced pluripotent stem cells (iPSCs) has demonstrated that this is possible at a preclinical level, accelerating clinical translation. A second important issue is to efficiently differentiate iPSCs into dopaminergic neuronal progenitors with restricted proliferation potential in order to avoid cellular overgrowth in vivo and minimize the risk of tumorigenesis. Here we describe an effective protocol that includes human iPSC differentiation to the dopaminergic lineage and enrichment in neuronal precursor cells expressing the polysialylated form of the neural cell adhesion molecule PSA-NCAM, through magnetically activated cell sorting. The resulting cells are transplanted and shown to survive, differentiate, and integrate within a striatal lesion model generated by unilateral 6-hydroxydopamine administration in mice of the NOD/SCID strain that supports xenografts. Key words Induced pluripotent stem cells (iPSC), Neurodegeneration, Parkinson’s disease

1

Introduction The mammalian central nervous system (CNS) has limited capacity for self-repair after injury or disease. The ability to overcome this barrier would transform the lives of patients with debilitating and incurable age-associated neurodegenerative diseases. This is an important task given that the incidence of these diseases is rising worldwide due to the continuously increasing life expectancy [1]. Neurodegenerative diseases lead to progressive loss of neuronal cells, and are often manifested with overlapping clinical symptoms [2]. Recent progress in regenerative biology and medicine has

Chrissa Kioussi (ed.), Stem Cells and Tissue Repair: Methods and Protocols, Methods in Molecular Biology, vol. 2155, https://doi.org/10.1007/978-1-0716-0655-1_3, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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opened up new possibilities to foster CNS regeneration and restore function in the damaged or diseased CNS. A variety of approaches have been tried over the past years that include: cell rescue using neurotrophic factors; cell-based approaches involving transplantation of exogenously derived and in vitro-cultured cells; strategies comprising stimulation of resident neural precursors to differentiate and replace the cells that are lost; methods aiming at direct reprograming of endogenous brain cells to desired phenotypes [3]. At a preclinical level, such approaches have been developed and tested for CNS restoration in various animal models of injury or disease. Here we focus on the generation of appropriate engraftable cell populations from human induced pluripotent stem cells and discuss their transplantation in immunocompromised mice bearing a brain lesion induced by administration of the neurotoxin 6-hydroxydopamine (6-OHDA). 1.1 Human Pluripotent Stem CellDerived Neuronal Precursors for Transplantation

A major challenge in developing any cell-based approach is the ability to generate large numbers of transplantable cells in a reliable and reproducible manner. Recent developments using human embryonic stem cells (hESCs) or induced pluripotent stem cells (iPSCs) have demonstrated that this is possible at a preclinical level opening new perspectives for translation to the clinic. Parkinson’s disease (PD) has long been a preferred target for cell transplantation, rapidly advancing toward clinical studies [4]. This is because the movement impairment that characterizes PD relates primarily to local and relatively selective degeneration of neurons of a particular brain region, the substantia nigra pars compacta. These neurons produce the neurotransmitter dopamine and deliver it to the striatum, where it plays a crucial role in control of motor activity [5]. Therefore treatment of the motor features in PD involves restoration of dopamine activity in the striatum [6]. Lessons learned from early fetal graft transplants in PD patients [7, 8] indicated that ventral midbrain allografts show long-term efficacy and survival, release dopamine, and improve selective PD symptoms [9]. At the same time a number of limitations became apparent, including ethical concerns and practical issues associated with inadequate fetal tissue supply and inability for standardization toward clinical use, indicating the necessity for alternative transplantable cellular sources. In recent years the pioneering technology of hESCs and iPSCs has opened up new prospects for understanding and treating human neurodegenerative diseases. It has prompted, on one hand, the creation of in vitro patient-derived models of disease [10] and on the other it has raised hopes for cell replacement therapies [4]. Moreover, it has helped in resolving tissue accessibility and reproducibility whilst bypassing ethical dilemmas. Nevertheless, the acquisition of the desirable iPSC-derived dopaminergic subpopulation that could survive and differentiate in vivo without

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evidence of tumor formation has been a challenge. The last few years though, the use of nonautologous human iPSC-derived midbrain dopamine cells in rodent and nonhuman primates has suggested the feasibility of the approach [11–15]. An important issue in this respect is to acquire in vitro adequate cells for transplantation but also to generate cells with restricted proliferation potential so as to avoid cellular overgrowth in vivo. This may be achieved by implementing optimized directed differentiation protocols yielding the desired precursor cell types in combination with cellular enrichment procedures to remove unwanted cells. Here we describe a method for generation of such engraftable dopaminergic neuronal precursors derived from iPSCs further enriched by selection for cells expressing the polysialylated form of the neural cell adhesion molecule (PSA-NCAM) ([16]; Fig. 1a). In this way we obtained an enriched population of neuronal cells differentiated to the dopaminergic lineage, consisting of progenitors and early neurons with restricted proliferative capacity, appropriate for transplantation studies [17–20]. The resulting cells were transplanted and tested for survival, differentiation, and integration in a striatal lesion model obtained by unilateral 6-OHDA administration in immunocompromised NOD/SCID mice that supports xenograft survival [21] (Fig. 1b–e). Indeed previous studies have shown that lesions increase the survival and integration of transplanted cells within the host brain [22]. The 6-OHDA lesion model mimics the loss of dopaminergic neurons in PD and has been widely used for assessing PD treatments [23]. Nevertheless, in our case it was used as a means for sustaining and assessing graft survival and differentiation rather than for therapeutic studies [16]. 1.2 The 6-OHDA Lesion Model

When injected into the striatum, 6-OHDA is taken up by the nerve endings of dopaminergic neurons through the dopamine transporter and is retrogradely transported to and accumulates in their cell bodies residing in the midbrain [24]. 6-OHDA is then readily oxidized leading to the generation of reactive oxygen species that ultimately cause neurodegeneration by oxidative cytotoxicity [25]. Therefore this model has been used to model PD, initially in rats where it is best characterized [26] and more recently in mice [27]. Apart from the striatum, the substantia nigra pars compacta or the median forebrain bundle have been targeted to destroy the nigrostriatal dopaminergic pathway [28, 29]. Lesions in the latter areas usually result in more extensive degeneration and more severe motor phenotypes [30] than striatal lesions [31]. To bypass the blood–brain barrier, 6-OHDA needs to be injected stereotactically into the brain. The extent of the induced lesion depends not only on the region injected but also on the

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Fig. 1 Generation of PSA-NCAM enriched human iPSC-derived dopaminergic neuronal precursors and intrastriatal engraftment in a 6-OHDA lesion model. (a) Schematic representation of the dopaminergic differentiation and MACS sorting protocol. Upon floor plate induction, over 40% of the cells express FOXA2, at 11 days in vitro (DIV), while upon further differentiation during 11–28 DIV cells express the dopaminergic marker Nurr1. After MACS sorting (28 DIV), approximately 80% of replated cells express PSA-NCAM at 30 DIV, the day of transplantation, as shown in the photomicrographs. Scale bar, 40 μm. (b) Schematic representation of the time course of the in vivo experiment. (c) Drawing illustrating the two 6-OHDA injection sites within the striatum (asterisks), which are also the sites of cell transplantation. (d) Immunohistochemistry for tyrosine hydroxylase (TH) with HRP detection in coronal sections, showing loss of dopaminergic projections in the striatum, at the ipsilateral side, 2 weeks after 6-OHDA injection, by comparison to the intact contralateral side. Scale bar, 500 μm. (e) Immunohistochemistry for tyrosine hydroxylase (TH) with HRP detection in coronal sections, showing degeneration of dopaminergic neurons in the substantia nigra pars compacta at the ipsilateral side, 2 weeks after 6-OHDA injection, by comparison to the intact contralateral side. Scale bar, 500 μm. The insets in the substantia nigra pars compacta are also shown at higher magnification. Scale bar, 100 μm. (f) Immunohistochemistry for the human-specific cytoplasmic antigen (green) in coronal sections of the striatum shows that the transplanted cells have populated the lesion site. Scale bar, 100 μm. Nuclei are shown in blue in (a) and (e). (g) 3D reconstruction of the contacts of grafted cells immunostained for human synaptophysin (yellow) on DARPP32+ host medium spiny neurons (red). Scale bar, 3 μm

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amount of 6-OHDA used and the species under surgery. Usually, 6-OHDA is administered in a unilateral manner which has the advantage that each animal can serve as its own within-subject control (ipsilateral and contralateral side to the lesion) in addition to between-subject effects (lesioned animals and sham-operated controls) [32]. Drug-induced [26] or drug-free assessments of motor behavior [33–35] of the unilateral model have been developed. Bilateral administration has also been used, however, it leads to adipsia, aphagia, and eventually to premature death, due to the animal’s inability to self-sustain [36]. With the advancements in mouse stem cell research, several studies have contributed to behavioral and histochemical characterization of the 6-OHDA model in mouse strains [37–39]. However, a detailed characterization of this model in immunocompromised mice for use in human cell transplantation studies, particularly the NOD/SCID strain that supports xenograft survival [14, 18], had not been performed until recently [16]. The use of this strain is described in the present method for transplantation of human iPSC-derived dopaminergic cells. To differentiate human iPSCs [40] toward the dopaminergic lineage a floor plate induction protocol [14] was applied with some modifications (Fig. 1a). At the end of floor plate induction (11 days in vitro; DIV), LMX1A/FOXA2-positive dopaminergic precursors were derived [16]. At this stage practically all cells were LMX1Apositive floor plate precursors [16] and approximately half were also FOXA2-positive in agreement with a dopaminergic fate (Fig. 1a). The cells further differentiated between DIV 11–28 to express the midbrain dopaminergic marker Nurr1 (Fig. 1a). Further enrichment in PSA-NCAM–positive neuronal cells was achieved at 28 DIV by magnetically activated cell sorting (MACS isolation) on the basis of PSA-NCAM immunoreactivity. This resulted in obtaining at 30 DIV—the day of transplantation—a culture consisting of approximately 80% of cells expressing PSA-NCAM (Fig. 1a). Cells were then transplanted in the striatum of adult NOD/SCID mice that had received a unilateral intrastriatal 6-OHDA injection 3 weeks before, causing a severe depletion of dopaminergic terminals in the striatum and loss of dopamine neurons in the substantia nigra pars compacta (Fig. 1b–e). Three months after cell transplantation, immunohistochemical analysis revealed survival, differentiation and integration of human iPSCderived neurons in the mouse brain [16]. Graft-derived cells populated the lesioned area and formed numerous contacts with the host medium spiny neurons (Fig. 1f, g).

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Materials

2.1 Dopaminergic Differentiation of iPSCs

1. Knockout DMEM (KO DMEM) plus 1% penicillin– streptomycin. 2. Knock Out Serum Replacement (KSR) medium: For 50 ml use 21:21 ml DMEM high glucose/F12 plus 1% penicillin–streptomycin, 7.5 ml KSR, 500 μl GlutaMAX, 50 μl ß-mercaptoethanol. 3. N2 medium: For 50 ml use 24.5:24.5 ml DMEM high glucose/F12 plus 1% penicillin–streptomycin, N2 supplement 500 μl, 500 μl GlutaMAX, 50 μl ß-mercaptoethanol. 4. NB/B27 medium: For 50 ml use Neurobasal plus 1% penicillin–streptomycin 48.5 ml, B27 supplement 1 ml, 500 μl GlutaMAX. 5. Day 0 medium: KSR medium supplemented with 100 nM LDN193189, 10 μM SB431542, 100 ng/ml SHH C24II, 2 μM purmorphamine, 100 ng/ml FGF8, 10 μM Y-27632. 6. Day 1 medium: KSR medium supplemented with 100 nM LDN193189, 10 μM SB431542, 100 ng/ml SHH C24II, 2 μM purmorphamine, 100 ng/ml FGF8. 7. Day 3 medium: KSR medium supplemented with 100 nM LDN193189, 10 μM SB431542, 100 ng/ml SHH C24II, 2 μM purmorphamine, 100 ng/ml FGF8, 3 μM CHIR99021. 8. Day 5 medium: KSR medium and N2 medium (3:1) supplemented with 100 nM LDN193189, 100 ng/ml SHH C24II, 2 μM purmorphamine, 100 ng/ml FGF8, 3 μM CHIR99021. 9. Day 7 medium: KSR medium and N2 medium (1:1) supplemented with 100 nM LDN193189, 100 ng/ml SHH C24II, 2 μM purmorphamine, 100 ng/ml FGF8, 3 μM CHIR99021. 10. Day 9 medium: KSR medium and N2 medium (1:3) supplemented with 100 nM LDN193189 and 3 μM CHIR99021. 11. Day 11 medium: NB/B27 medium supplemented with 20 ng/ml BDNF, 20 ng/ml GDNF, 1 ng/ml TGFβ3, 200 μM ascorbic acid, 0.5 mM dibutyryl cAMP, 10 mM DAPT, and 3 μM CHIR99021. 12. Day 13 medium (differentiation medium): NB/B27 medium supplemented with 20 ng/ml BDNF, 20 ng/ml GDNF, 1 ng/ ml TGFβ3, 200 μM ascorbic acid, 0.5 mM dibutyryl cAMP, and 10 mM DAPT. 13. Matrigel. 14. Polyethylenimine (PLE)/laminin/fibronectin.

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2.2 Magnetically Activated Cell Sorting (MACS)

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1. 1% bovine serum albumin (BSA) in phosphate buffer saline (PBS) solution: For 500 ml use 1 g BSA diluted in 500 ml PBS 1. The solution was then filtered using a 500 ml vacuum filter system with 0.22 μm pore size. 2. Neubauer Chamber. 3. Cell strainer, 40 μm pore size. 4. Anti-PSA-NCAM magnetic microbeads. 5. LS columns. 6. MACS separator.

2.3 6-OHDA Lesions and Cell Transplantation

1. Male NOD.CB17-Prkdcscid/NCrHsd mice 9–10 weeks old (see Note 1). 2. Desipramine hydrochloride solution: 2 mg/ml dissolved in 0.9% saline and administered to the animal at 25 mg/kg. 3. 6-Hydroxydopamine hydrochloride (6-OHDA) solution: 4 μg/μl 6-OHDA dissolved in 0.9% saline containing 0.02% ascorbic acid. 4. Weighing device. 5. Electric razor. 6. Sterile surgical tools: scissors, needle holder, medium and fine forceps. 7. Local antiseptic (e.g., iodine). 8. Local analgesic (e.g., lidocaine). 9. Systemic analgesic (e.g., carprofen, 5 mg/kg). 10. Eye gel. 11. Scalpel (surgical blade, no. 12). 12. 26-G syringe (10 μl). 13. Dental drill with small (1- to 1.5-mm) cutting burrs. 14. Nonabsorbable silk sutures. 15. Mouse stereotactic apparatus with heating plate and a microdrive pump. 16. Inhalation device for anesthesia. 17. Heating plate. 18. 5% glucose in 0.9% sterile saline. 19. Hanks’ Balanced Salt solution without calcium, without magnesium (HBSS).

2.4 Immunocytochemistry, Quantification, and Statistical Analysis

1. 4% Paraformaldehyde (PFA) in PBS. 2. 4% agarose in double distilled (dd) H2O. 3. Vibrating microtome. 4. Circle coverslips 10 mm.

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5. Microscope slides B/50, 76  26 mm. 6. Micro cover glasses, 24  50 mm. 7. Antigen retrieval buffer: 10 mM sodium citrate pH 6: For 500 ml, use 1.47 g trisodium citrate (dihydrate) and 500 ml ddH2O. Adjust pH to 6.0 with 1 N HCl. (For 100 ml 1 N HCl use 8.6 ml 36% HCl and 91.4 ml.) 8. Immunocytochemistry blocking solution: 0.1% Triton X-100, 5% donkey serum in PBS: For 100 ml blocking, use 0.1 ml Triton X-100, 5 ml donkey serum and PBS 1 up to 100 ml. 9. Immunohistochemistry blocking solution: 0.1% Triton X-100, 2 mg/ml bovine serum albumin (BSA), and 1.5% donkey serum in PBS 1. For 100 ml blocking, use 0.1 ml Triton X-100, 0.2 g BSA, 1.5 ml donkey serum, and PBS 1 up to 100 ml. 10. 0.3% H2O2 solution: For 100 ml solution: dilute 1 ml from stock 30% H2O2 in 99 ml PBS 1. 11. VECTASTAIN Elite ABC HRP Kit (Vector, PK-6100). 12. Peroxidase substrate solution: For 1 ml solution, use 10 μl buffer, 20 μl 3,30 -Diaminobenzidine (DAB), 12.5 μl H2O2, 12.5 μl Nickel, and 945 μl PBS 1. 13. Hoechst 33342. 14. Antifade Mountant with DAPI. 15. Mowiol 4-88 mounting medium: For 24 ml add 2.4 g of Mowiol 4-88 to 6 g of glycerol. Stir to mix. Add 6 ml of H2O and leave overnight at RT. Add 12 ml of 0.2 M Tris– HCl (pH 8.5) and heat to 50  C for 10 min with occasional mixing. After the Mowiol dissolves, clarify by centrifugation at 5000  g for 15 min. Aliquot and store at 20  C. 16. Confocal laser scanning microscopy platform TCS SP8 (Leica). 17. Image processing program ImageJ (NIH). 18. Adobe Photoshop. 2.5 Primary and Secondary Antibodies

1. Rabbit polyclonal anti-Ki67 (1:200, Abcam, ab15580). 2. Rabbit polyclonal anti-nestin (1:500, Merck Millipore, ABD69). 3. Mouse monoclonal anti-FOXA2 (HNF-3β, RY-7; 1:100, Santa Cruz, sc-101060). 4. Mouse monoclonal anti-PSA-NCAM (1:100; Merck Millipore, MAB5324). 5. Mouse monoclonal anti-Nurr1 (1:100, Santa Cruz, sc-81345). 6. Mouse monoclonal anti-MAP2 (1:200, Merck Millipore, MAB3418).

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7. Mouse monoclonal anti-human cytoplasmic STEM121 (1:500, Clontech, Y40410).

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antigen

8. Mouse monoclonal anti-synaptophysin clone EP10 (1:200, Thermo Fisher, 14-6525-80). 9. Rabbit monoclonal ab40801).

anti-DARPP32

(1:1000,

Abcam,

10. Rabbit polyclonal anti-tyrosine hydroxylase (1:500, Merck Millipore, AB152). 11. Donkey anti-mouse Alexa Fluor 488 (green) and donkey antirabbit polyclonal Alexa Fluor 546 (red) secondary antibody (1:1000, Molecular Probes). 12. Biotinylated goat anti-rabbit IgG antibody (1:200, Vector, BA-1000).

3

Methods

3.1 Dopaminergic Differentiation of Human iPSCs

Human iPSCs are cultured and expanded as previously described [40] whilst dopaminergic differentiation is performed as described below: 1. Day 0 of differentiation: human iPSC colonies are dissociated into a single cell suspension by incubation with Accutase for 20 min. 2. The dissociated cells are centrifuged at 500  g for 5 min and resuspended in KO DMEM and counted using a Neubauer chamber. Human iPSCs are plated at a concentration of 40,000 cells per cm2 on Matrigel-coated 6-well dishes (for coating procedure see below) in Day 0 medium. 3. Medium is changed the next day (Day 1 medium) and every other day from then on. From Day 13 onward, differentiation medium is used. 4. At 20 days in vitro (DIV), cells are dissociated using Accutase and replated at high density (300,000 cells per cm2) on dishes containing coverslips precoated with polyethylenimine/laminin/fibronectin (for coating procedure see below) in differentiation medium until the desired maturation stage. 5. Matrigel coating: Matrigel is thawed on ice and then diluted in serum-free medium (e.g., DMEM/F12, 1:20) using cooled tips. Diluted Matrigel is added to the plates and incubation follows at room temperature in the laminar hood for 1 h. Right before use excess material is aspirated and the plates are washed gently using serum-free medium.

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6. Polyethylenimine/laminin/fibronectin coating: The day of cell passage, PLE is thawed on ice and diluted in sterile double distilled (dd) H2O in a concentration of 15 mg/ml. Incubation at 37  C (cell incubator) follows for 1 h. In parallel, laminin and fibronectin are thawed on ice. After 1 h of incubation, the plates are washed three times with ddH2O. Laminin and fibronectin are diluted in sterile PBS (1 and 2 mg/ml, respectively) using cooled tips and incubated at 37  C for at least 2 h. The plates are then washed twice with sterile PBS right before cell plating. 3.2 Magnetically Activated Cell Sorting (MACS) of PSA-NCAMPositive Cells

Magnetic labeling and separation are performed as described below, based on the manufacturer’s instructions (www.miltenyibiotech. com) (see Note 2). 1. At 28 DIV, human iPSC-derived dopaminergic cells are incubated with 10 mM Y-27632 (ROCK inhibitor) diluted in differentiation medium for 1 h to prevent cell death. 2. Cells are dissociated into single cell suspension by incubation with Accutase for 10 min. 3. The dissociated cells are centrifuged at 500  g for 5 min, resuspended in NB and passed through a 40 μm cell strainer to ensure generation of single cell suspension (see Note 3). 4. Cell number is determined using a Neubauer Chamber. 5. Cell suspension is centrifuged at 300  g for 10 min and supernatant is completely removed. 6. Cells are resuspended in 60 μl of buffer per 107 cells, and incubation for 10 min in the refrigerator (2–8  C) follows (see Note 4). 7. 20 μl of anti-PSA-NCAM MicroBeads per 107 total cells are added, mixed well and incubated for 15 min in the refrigerator (2–8  C). 8. Cells are washed with 1–2 ml of buffer per 107 cells and centrifuged at 300  g for 10 min. 9. The supernatant is completely removed and up to 108 cells are resuspended in 500 μl of buffer. 10. An LS column is placed in the magnetic field of a MACS separator and prepared by rinsing with 3 ml of buffer. 11. The cell suspension is applied onto the column and unlabeled cells pass through. 12. The column is washed with 3  3 ml of buffer. 13. The column is then removed from the separator and placed in a 15 ml collection tube.

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14. 5 ml of differentiation medium are pipetted onto the column and labeled cells are immediately flushed out by firmly pushing the plunger into the column. 15. Cells are counted and replated at high density as above. 16. After 2 days (30 DIV) cells are dissociated to single cell suspension and are used for immunocytochemistry or for transplantation. 3.3 Immunocytochemistry

1. At 11–28 DIV or 2 days after MACS (30 DIV), cells plated on coverslips are fixed with 4% paraformaldehyde for 20 min at room temperature. They are washed three times with PBS 1, blocked for 30 min and are subsequently incubated with primary antibodies (diluted in 1:1 blocking solution and PBS 1) at 4  C overnight. 2. The next day coverslips are washed three times with PBS 1 and incubated for 2 h with the secondary antibody (diluted in 1:1 blocking solution and PBS 1). 3. Following incubation with the secondary, coverslips are washed three times with PBS 1 and once with ddH2O and then mounted on microscope slides with Antifade Mountant containing DAPI.

3.4 6-OHDA Lesions in Immunocompromised Mice

In this protocol, a unilateral intrastriatal 6-OHDA lesion was made and extensive post-operative care was given to the mice (see Note 5). 1. 6-OHDA solution is prepared immediately prior to injection and kept on ice protected from light during the procedure. 2. Desipramine is administered to each mouse 30 min before the 6-OHDA injection (see Note 6). 3. The mouse is first placed in an anesthesia chamber using gaseous isoflurane (2% in 2:1 oxygen to nitrous oxide). 4. After ensuring that the mouse has been anesthetized, the head is shaved and the mouse is stabilized into the stereotaxic apparatus, which is connected to an inhaled anesthesia mask. 5. The skin at the top of the head is cleaned using a cotton swab soaked with local antiseptic (e.g., iodine). 6. A local anesthetic (e.g., lidocaine) is applied on the skin. 7. Using a scalpel, an incision to expose the skull is made and connective tissue is removed. 8. After locating bregma, the drill is placed on it to zero the coordinates according to [41] and a small hole is drilled in the following coordinates: AP ¼ +0.5 mm, ML ¼ +1.8 mm. 9. The appropriate amount of fresh 6-OHDA solution is loaded on the syringe. Infusion is made with a rate of 0.5 μl/min, in the mid-right striatum (two injections of 1 μl: AP ¼ +0.5 mm

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to bregma, ML ¼ +1.8 mm, DV ¼ 3.0 and 3.5 mm). The syringe is left in place for a further 2 min, before being slowly removed. 10. The syringe is slowly removed to prevent drawback of the toxin solution, the mouse head is cleaned and the skin is carefully sutured. 11. Each mouse is left to recover in a cage with paper towels placed on a heating plate and is returned to its cage after recovering. 12. Analgesic (e.g., carprofen) is provided subcutaneously immediately after surgery and in the following day. 3.5 Intrastriatal Cell Transplantation

Three weeks after 6-OHDA injection, human iPSC-derived PSANCAM-enriched dopaminergic cells (30 DIV) are stereotactically transplanted in mice. 1. Cells are dissociated into single cell suspension by incubation with Accutase for 10 min, resuspended in DMEM/F12 and passed through a 40 μm cell strainer to ensure generation of single cell suspension. Their number is determined using a Neubauer counting chamber. 2. Cells are centrifuged, resuspended in cold HBSS at a density of 100,000 cells/μl and maintained in ice during the procedure. 3. Each mouse is anesthetized and stabilized in the stereotactic apparatus, as described in Subheading 3.4. 4. Each mouse receives 2  1 μl cells at the same coordinates as the 6-OHDA lesions were made at a rate of 0.5 μl/min. The syringe is left in place for a further 2 min, before being slowly removed. 5. Each mouse is left to recover in a cage with paper towels placed in a heating plate and returned in its cage after recovering. Analgesic is provided immediately after surgery and the next day.

3.6 Euthanasia and Immunohistochemistry

1. Three months after cell transplantation, animals are transcardially perfused with 30 ml PBS 1 followed by 30 ml 4% PFA, their brains are removed and postfixed overnight at 4  C in the same fixative. 2. Perfused brains are embedded in 4% agarose, sectioned into 40 μm-thick slices on a vibrating microtome and serially collected. 3. For immunofluorescence staining, free-floating vibratome sections are incubated with sodium citrate buffer brought to boiling, for 30 min at 37  C for antigen retrieval. Sections are left to cool down at room temperature (RT) and washed three times with PBS 1.

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4. Sections are subsequently incubated with blocking solution for 2 h to saturate nonspecific binding sites and then with primary antibodies diluted in blocking solution at 4  C for three nights with gentle agitation. 5. Following primary antibodies, sections are washed five times (1 h each wash) with blocking solution and are incubated overnight at 4  C with secondary antibodies (diluted 1:1000 in blocking solution) with gentle agitation. 6. The following day sections are washed five times (1 h each wash) with blocking solution and incubated for 30 min with Hoechst 33342 for fluorescent DNA staining (diluted in 0.1% Triton X-100, PBS) with gentle agitation. 7. For HRP detection of tyrosine hydroxylase, after blocking and incubation with the primary antibody (as described above), sections are washed three times (10 min each wash) with PBS 1, are incubated with 0.3% H2O2 for 10 min at RT to kill endogenous peroxidase enzyme, with gentle agitation. Sections are then washed with PBS 1 once and incubated with a biotinylated secondary antibody (diluted in blocking solution) for 2 h with gentle agitation. After three washes with PBS 1, incubation with Vectastain Kit for 30 min follows. The sections are then washed three times with PBS 1 and incubated with peroxidase substrate solution until the desired stain intensity develops. 8. Sections are then mounted on microscope slides using either Antifade Mountant containing DAPI (immunofluorescence staining) or Mowiol (HRP detection) and covered with coverslips (see Note 7). 3.7 Confocal Microscopy, Cell Counting, and Statistical Analysis

4

Digital images, both after fluorescence immunocytochemistry and immunohistochemistry, are acquired using a Leica TCS-SP8 confocal microscope. For HRP detection, images are acquired using an upright light microscope. Quantification of positive cells for each marker is performed using ImageJ on 20 randomly selected fields from each experiment. For 3D reconstruction, images are processed using the surface contact area XTention of Imaris Software v9.2.1. All experiments are replicated at least three times. Statistical analysis is performed using the Student’s t test.

Notes 1. Mice older than 9 weeks old are used, weighing more than 25 g (www.envigo.com). Mice are expected to lose weight after the 6-OHDA injection since they do not feed properly, and, according to our experience, younger mice weighing less than 25 g have fewer chances to survive during the weeks following 6-OHDA injection.

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2. Pilot in vivo transplantation with unsorted cells resulted in cellular overgrowth within the grafts. We therefore reasoned that an enriched population of neuronal cells differentiated to the dopaminergic lineage, consisting of progenitors and early neurons with restricted proliferative capacity, would be appropriate for transplantation studies. To this end, we applied MACS isolation on the basis of PSA-NCAM immunoreactivity at 28 DIV. This is in agreement with other studies pointing toward the necessity of employing sorting methods to eliminate nonneuronal and, in particular, highly proliferating cells using fluorescence-activated cell sorting (FACS) or magnetically activated cell sorting (MACS) [17, 19, 42, 43]. 3. This step is important in order to remove cell clumps. 4. As indicated by the manufacturer (www.miltenyibiotec.com), it is important to work fast, to keep cells cold, and to use precooled solutions. In this way, capping of antibodies on the cell surface and nonspecific cell labeling will be prevented. 5. We started by introducing lesions in the Median Forebrain Bundle that led to high mortality rates in the NOD/SCID mouse strain (over 85%). For this reason we proceeded with intrastriatal 6-OHDA injections and, using extensive postoperative care, we managed to reduce the mortality rate to 40%. Immediately after surgery and daily during the first week following 6-OHDA injection, mice received 5% glucose solution in sterile saline subcutaneously to prevent dehydration. Additionally, once a day for the first 2 weeks postinjection, food pellets soaked in water were placed in a petri dish inside the cage to facilitate feeding. In order to avoid competition for food and any other aggressive behaviors, mice were kept in separate cages after surgery. 6. 6-OHDA exhibits a high affinity for several catecholaminergic transporters including the dopamine and norepinephrine transporters [44]. For this reason, desipramine which is a selective noradrenaline reuptake inhibitor, is administered prior to 6-OHDA injection to spare the noradrenergic neurons from damage. 7. Care should be taken to avoid air bubbles while mounting the sections onto slides and covering with coverslips. No mechanical pressure should be introduced over sections and they should be left for 12 h in RT before imaging, allowing the immunofluorescence mountant to fully solidify.

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Acknowledgments This work was supported by a grant from Stavros Niarchos Foundation to the Hellenic Pasteur Institute as part of the Foundation’s initiative to support the Greek Research Center ecosystem; the projects MIS 5002486 and MIS 5002755 funded by the Operational Program “Competitiveness, Entrepreneurship and Innovation” (NSRF 2014-2020) and cofinanced by Greece and the European Union (European Regional Development Fund). References 1. Prince M, Bryce R, Albanese E, Wimo A, Ribeiro W et al (2013) The global prevalence of dementia: a systematic review and metaanalysis. Alzheimers Dement 9:63–75.e2 2. Gan L, Cookson MR, Petrucelli L, La Spada AR (2018) Converging pathways in neurodegeneration, from genetics to mechanisms. Nat Neurosci 21:1300–1309 3. Barker RA, Go¨tz M, Parmar M (2018) New approaches for brain repair—from rescue to reprogramming. Nature 557:329–334 4. Barker RA, Parmar M, Studer L, Takahashi J (2017) Human trials of stem cell-derived dopamine neurons for Parkinson’s disease: dawn of a new era. Cell Stem Cell 21:569–573 5. Lees AJ, Hardy J, Revesz T (2009) Parkinson’s disease. Lancet 373:2055–2066 6. Stoker TB, Torsney KM, Barker RA (2018) Emerging treatment approaches for Parkinson’s disease. Front Neurosci 12:693–693 7. Kordower JH, Freeman TB, Snow BJ, Vingerhoets FJ, Mufson EJ et al (1995) Neuropathological evidence of graft survival and striatal reinnervation after the transplantation of fetal mesencephalic tissue in a patient with Parkinson’s disease. N Engl J Med 332:1118–1124 8. Li W, Englund E, Widner H, Mattsson B, van Westen D et al (2016) Extensive graft-derived dopaminergic innervation is maintained 24 years after transplantation in the degenerating parkinsonian brain. Proc Natl Acad Sci U S A 113:6544–6549 9. Barker RA, Drouin-Ouellet J, Parmar M (2015) Cell-based therapies for Parkinson disease-past insights and future potential. Nat Rev Neurol 11:492–503 10. Chang C-Y, Ting H-C, Liu C-A, Su H-L, Chiou T-W et al (2018) Induced pluripotent stem cells:a powerful neurodegenerative disease modeling tool for mechanism study and drug discovery. Cell Transplant 27:1588–1602

11. Grealish S, Diguet E, Kirkeby A, Mattsson B, Heuer A et al (2014) Human ESC-derived dopamine neurons show similar preclinical efficacy and potency to fetal neurons when grafted in a rat model of Parkinson’s disease. Cell Stem Cell 15:653–665 12. Kikuchi T, Morizane A, Doi D, Magotani H, Onoe H et al (2017) Human iPS cell-derived dopaminergic neurons function in a primate Parkinson’s disease model. Nature 548:592–596 13. Kirkeby A, Grealish S, Wolf DA, Nelander J, Wood J et al (2012) Generation of regionally specified neural progenitors and functional neurons from human embryonic stem cells under defined conditions. Cell Rep 1:703–714 14. Kriks S, Shim JW, Piao J, Ganat YM, Wakeman DR et al (2011) Dopamine neurons derived from human ES cells efficiently engraft in animal models of Parkinson’s disease. Nature 480:547–551 15. Steinbeck JA, Choi SJ, Mrejeru A, Ganat Y, Deisseroth K et al (2015) Optogenetics enables functional analysis of human embryonic stem cell-derived grafts in a Parkinson’s disease model. Nat Biotechnol 33:204–209 16. Zygogianni O, Antoniou N, Kalomoiri M, Kouroupi G, Taoufik E et al (2019) In vivo phenotyping of familial Parkinson’s disease with human induced pluripotent stem cells: a proof-of-concept study. Neurochem Res 44 (6):1475–1493 17. Doi D, Samata B, Katsukawa M, Kikuchi T, Morizane A et al (2014) Isolation of human induced pluripotent stem cell-derived dopaminergic progenitors by cell sorting for successful transplantation. Stem Cell Rep 2:337–350 18. Hargus G, Cooper O, Deleidi M, Levy A, Lee K et al (2010) Differentiated Parkinson patient-derived induced pluripotent stem cells grow in the adult rodent brain and reduce

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motor asymmetry in Parkinsonian rats. Proc Natl Acad Sci U S A 107:15921–15926 19. Sundberg M, Bogetofte H, Lawson T, Jansson J, Smith G et al (2013) Improved cell therapy protocols for Parkinson’s disease based on differentiation efficiency and safety of hESC-, hiPSC-, and non-human primate iPSC-derived dopaminergic neurons. Stem Cells 31:1548–1562 20. Kikuchi T, Morizane A, Doi D, Okita K, Nakagawa M et al (2017) Idiopathic Parkinson’s disease patient-derived induced pluripotent stem cells function as midbrain dopaminergic neurons in rodent brains. J Neurosci Res 95:1829–1837 21. Hargus G, Ehrlich M, Arauzo-Bravo MJ, Hemmer K, Hallmann AL et al (2014) Origin-dependent neural cell identities in differentiated human iPSCs in vitro and after transplantation into the mouse brain. Cell Rep 8:1697–1703 22. Behrstock S, Ebert AD, Klein S, Schmitt M, Moore JM et al (2008) Lesion-induced increase in survival and migration of human neural progenitor cells releasing GDNF. Cell Transplant 17:753–762 23. Bove´ J, Perier C (2012) Neurotoxin-based models of Parkinson’s disease. Neuroscience 211:51–76 24. Glinka Y, Tipton KF, Youdim MB (1996) Nature of inhibition of mitochondrial respiratory complex I by 6-hydroxydopamine. J Neurochem 66:2004–2010 25. Blum D, Torch S, Lambeng N, Nissou M, Benabid AL et al (2001) Molecular pathways involved in the neurotoxicity of 6-OHDA, dopamine and MPTP: contribution to the apoptotic theory in Parkinson’s disease. Prog Neurobiol 65:135–172 26. Ungerstedt U (1968) 6-Hydroxy-dopamine induced degeneration of central monoamine neurons. Eur J Pharmacol 5:107–110 27. Mandel RJ, Randall PK (1985) Quantification of lesion-induced dopaminergic supersensitivity using the rotational model in the mouse. Brain Res 330:358–363 28. Dowd E, Dunnett SB (2005) Comparison of 6-hydroxydopamine-induced medial forebrain bundle and nigrostriatal terminal lesions in a lateralised nose-poking task in rats. Behav Brain Res 159:153–161 29. Yuan H, Sarre S, Ebinger G, Michotte Y (2005) Histological, behavioural and neurochemical evaluation of medial forebrain bundle and striatal 6-OHDA lesions as rat models of Parkinson’s disease. J Neurosci Methods 144:35–45

30. Blandini F, Armentero MT, Martignoni E (2008) The 6-hydroxydopamine model: news from the past. Parkinsonism Relat Disord 14 (Suppl 2):S124–S129 31. Sauer H, Oertel WH (1994) Progressive degeneration of nigrostriatal dopamine neurons following intrastriatal terminal lesions with 6-hydroxydopamine: a combined retrograde tracing and immunocytochemical study in the rat. Neuroscience 59:401–415 32. Smith GA, Heuer A (2011) Animal models of movement disorders: 6-OHDA toxin models of PD in mice, vol 61. Springer/Humana, New York 33. Schallert T, Fleming SM, Leasure JL, Tillerson JL, Bland ST (2000) CNS plasticity and assessment of forelimb sensorimotor outcome in unilateral rat models of stroke, cortical ablation, parkinsonism and spinal cord injury. Neuropharmacology 39:777–787 34. Tillerson JL, Cohen AD, Philhower J, Miller GW, Zigmond MJ et al (2001) Forced limbuse effects on the behavioral and neurochemical effects of 6-hydroxydopamine. J Neurosci 21:4427–4435 35. Schallert T, Whishaw IQ, Ramirez VD, Teitelbaum P (1978) Compulsive, abnormal walking caused by anticholinergics in akinetic, 6-hydroxydopamine-treated rats. Science 199:1461–1463 36. Ungerstedt U (1971) Adipsia and aphagia after 6-hydroxydopamine induced degeneration of the nigro-striatal dopamine system. Acta Physiol Scand Suppl 367:95–122 37. Boix J, Padel T, Paul G (2015) A partial lesion model of Parkinson’s disease in mice—characterization of a 6-OHDA-induced medial forebrain bundle lesion. Behav Brain Res 284:196–206 38. Bagga V, Dunnett SB, Fricker RA (2015) The 6-OHDA mouse model of Parkinson’s disease—terminal striatal lesions provide a superior measure of neuronal loss and replacement than median forebrain bundle lesions. Behav Brain Res 288:107–117 39. Virgone-Carlotta A, Uhlrich J, Akram MN, Ressnikoff D, Chretien F et al (2013) Mapping and kinetics of microglia/neuron cell-to-cell contacts in the 6-OHDA murine model of Parkinson’s disease. Glia 61:1645–1658 40. Kouroupi G, Taoufik E, Vlachos IS, Tsioras K, Antoniou N et al (2017) Defective synaptic connectivity and axonal neuropathology in a human iPSC-based model of familial Parkinson’s disease. Proc Natl Acad Sci U S A 114: E3679–E3688

Human Neural Precursors for Brain Repair 41. Paxinos G, Franklin KBJ (2013) Paxinos and Franklin’s The mouse brain in stereotaxic coordinates. Academic Press, Amsterdam 42. Kim DS, Lee DR, Kim HS, Yoo JE, Jung SJ et al (2012) Highly pure and expandable PSANCAM-positive neural precursors from human ESC and iPSC-derived neural rosettes. PLoS One 7:e39715

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Chapter 4 In Vitro Direct Reprogramming of Mouse and Human Astrocytes to Induced Neurons Katerina Aravantinou-Fatorou and Dimitra Thomaidou Abstract Direct neuronal reprogramming, rewiring the epigenetic and transcriptional network of a differentiated cell type to neuron, apart from being a very promising approach for the treatment of brain injury and neurodegeneration, also offers a prime opportunity to investigate the molecular underpinnings of neuronal cell fate determination, as the precise molecular mechanisms that establish neuronal fate and diversity at the transcriptional and epigenetic level are incompletely understood. Recent studies from a number of groups, including ours, have shown that astrocytes can be directly reprogrammed into functional neurons in vitro and in vivo following ectopic overexpression of combinations of transcription factors, neurogenic proteins, miRNAs, and small chemical molecules. In this chapter we describe the protocols for in vitro converting primary cortical astrocytes of mouse and human origin to induced neurons, through forced expression of two neurogenic molecules, either each one alone or in combination: the master regulatory bHLH proneural transcription factor NEUROGENIN2 (NEUROG2) and the neurogenic protein CEND1. Forced expression of each one of the two neurogenic proteins in primary astrocytes via retroviral gene transfer results in their direct conversion to subtypespecific induced neurons, while simultaneous coexpression of both molecules drives them predominantly toward acquisition of a neural precursor cell (NPC) state. Although mouse and human astrocytes exhibit differences in their reprogramming rate and particular characteristics, they can both get efficiently in vitro transdifferentiated to NPCs and induced neurons upon NEUROG2 or/and CEND1 forced expression using the reprogramming protocols described in the chapter, presenting valuable cellular platforms for mechanistic studies and in vivo applications to restore neurodegeneration. Key words Astrocytes, Reprogramming, Transdifferentiation, Induced neurons, Neural progenitor cells, Cend1, Neurogenin2

1

Introduction Brain damage results in loss of specific populations of neurons and/or glial cells and is accompanied by the development of defined neurological symptoms. In all cases CNS damage results in the activation of a CNS cell population that plays pivotal role in neuronal network integrity as well as brain immune response, that is, astrocytes. Following brain injury and neurodegenerative

Chrissa Kioussi (ed.), Stem Cells and Tissue Repair: Methods and Protocols, Methods in Molecular Biology, vol. 2155, https://doi.org/10.1007/978-1-0716-0655-1_4, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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diseases, astrocytes surrounding the injury area acquire or reactivate stem cell potential, therefore highlighting a cell source outside the stem cell niche being directed toward a neurogenic fate [1, 2]. To this end recent studies from a number of groups, including ours, demonstrate that astrocytes can be directly reprogrammed to neural stem cell-like cells and functional neurons both in vitro [3, 4] and to some extent in vivo [5–7] following brain injury, provided they overexpress certain neurogenic factors, and can thus be used as the cell source that will restore neuronal damage and lost connectivity following brain injury. 1.1 What Can the Astrocytes Offer to Regeneration Therapies?

The adult cerebral cortex has limited ability to regenerate lost neural tissue after brain damage, partly due to lack of a resident population of neural stem cells (NSCs) responsive to injury-derived signals. Limited compensatory cortical neurogenesis has been reported following stroke [8] or induced apoptotic degeneration [9], but the number of neurons produced is insufficient to replenish neuronal loss after injury and restore cortical function [10, 11]. To overcome this limitation, efforts have been made to stimulate the endogenous NSCs population residing in the neighboring subventricular zone (SVZ) with growth factors, in order to recruit a population of neural precursor cells (NPCs) to the lesioned cortex. Yet adequate recruitment of SVZ NPCs to successfully replace damaged cortical neurons has so far not been achieved [12– 14]. Transplantation of suitable cell types into the adult central nervous system (CNS) has also attracted considerable interest as an alternative strategy to overcome the regenerative limitations of the lesioned brain [15–17]. Astrocytes constitute an abundant glial cell type that endogenously possess a neuroptotective and neuromodulating activity in the healthy brain and the tendency to proliferate following brain trauma [7, 18]. After a brain injury, astrocytes revert to an activated stage characterized by multiple and complex changes in their morphology, gene expression and function, a process referred to as astrogliosis. Astrogliosis is characterized by high proliferating activity, hypertrophy, and changes in astrocytic morphology, that ultimately result in the creation of the astroglial scar to seal off the injured tissue and restrict inflammation and neuronal death [19– 21]. Astroglia activation occurs in many neurodegenerative or neuroinflammatory conditions, including acute invasive brain injury or stroke [2, 22] and its overall effect is time and context dependent and can be either neuroprotective or neurotoxic, resulting in restriction or progression of pathology [23–25]. Interestingly, besides the above mentioned morphological and functional changes, reactive astrocytes start sharing morphological and molecular phenotype hallmarks of neural stem cells (NSCs) and developmental radial glia, including expression of NESTIN, VIMENTIN, and brain lipid-binding protein (BLBP) [1, 2], a fact suggesting

Direct Reprogramming of Astrocytes to iNs and NPCs In Vitro

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that they are a widespread source of cells that, when stimulated, can be driven toward a neurogenic fate. In specific brain areas in particular, such as the striatum, the neurogenic capacity of astrocytes is much enhanced following neurodegeneration [18], highlighting the existence of an endogenous cell source capable of restoring connectivity and function. On that direction, the idea of direct reprogramming of astrocytes to the neuronal subtype needed in situ, depending on the area that has been affected, has appeared as a pioneer approach for the design of brain regeneration strategies. Astrocytes are almost everywhere within the CNS and get activated to some extent in all neurodegenerative disorders, they have therefore appeared to be a key cell source to replace lost neurons through their transdifferentiation. Hence the reprogramming potential of both mouse and human astrocytes is being extensively studied both in vitro and in vivo in animal models of neurodegeneration [25–31]. Chouchane and Costa [32] have highlighted four advantages of using astrocytes instead of adult neural stem cells residing in the subependymal zone (SEZ), or the subgranular zone (SGZ) of the hippocampus neurogenic areas [33] as cell sources for regenerative approaches in the future: (a) astrocytes are located within the lesioned site, eliminating the need of their relocation; (b) when activated by the insult, they have the ability to proliferate in large numbers [34]; (c) they can be quickly and efficiently reprogrammed into neurons using simple molecular manipulations [26, 28, 35, 36] and (d) they are involved in the formation of the glial scar, which contributes to the generation of an anti-neurogenic environment [1]. Thus utilizing endogenous reactive astrocytes, which have the intrinsic characteristic to proliferate after trauma and, most importantly, possess a neuronal lineage epigenetic memory [37], as a neuron-producing cell source seems one of the most direct and applicable therapeutic options. 1.2 Direct Reprogramming of Astrocytes to Induced Neurons

A major challenge in regenerative medicine is to unravel the most effective way to obtain customized functional cell types. Direct lineage reprogramming has emerged as a promising, fast approach for manipulating cell fate toward the desired cell type, avoiding the major risk of teratoma formation associated with the use of induced pluripotent stem cells (iPSCs). A step further, utilization of endogenous brain cells that can be directly reprogrammed into neurons, remains a challenge with important clinical applications for the treatment of neurodegeneration. In that context, a growing number of studies from a number of groups have shown that mouse and human astrocytes can be directly reprogrammed into functional neurons in vitro and in vivo following ectopic overexpression of combinations of proneural transcription factors, neurogenic proteins, miRNAs, and small molecules [26, 27, 38, 39, 40, 41] (Table 1). The very first successful efforts of in vitro

Source

Sox2 + VPA/BDNF/ NOG

Adult striatum

NEUROG2/ASCL1 Cerebral cortex of postnatal mice

NeuroD1

Cerebral cortex,

Plasmids

LV

RV

Synaptic input

Mature excitability, synaptic input (at 4 wpi)

TUJ1/MAP2/NeuN

Glutamatergic and GABAergic iNS with spontaneous electrical activity

Niu et al. (2015) [43] Chouchane et al. (2017) [44]

4 wpi

Guo et al. (2014) [5]

Guo et al. (2014) [5]

40 wpi

14 dpi

8 wpi

Niu et al. (2013) [6]

24 month

Spontaneous synaptic currents indicating the presence of postsynaptic receptors

Heinrich et al. (2011) [27]

32 dpi

Glutamatergic or GABAergic synapses

Buffo et al. (2005) [42]

30 dpi

Max time investigated References

n.d.

Functional outcome

Calretinin interneurons Mature excitability, synaptic input (through Ascl1 expression)

NeuN

DCX/NeuN/Tbr1/ Ctip2

NeuroD1

Cerevral cortex

RV

DCX

LV ASCL1, BRN2, KLF4, MYC, MYT1L, OCT4, SOX2, and ZFP521, and miR9, miR124, miR125, and miR128 supplemented with BDNF and noggin

Adult striatum

DCX

TUJ1/MAP2/Tbr1/ Tbr2/vGaT/ vGluT1/CamKIIα

RV

dominant negativeOlig2 (Olig2VP16)

Type of iN

Neurog2 or Mash1 and/or RV distal-less homeobox 2 (Dlx2)

Vector used

Reprogramming factor

Cerebral cortex

Mouse Cerebral astrocytes cortex, stab wound

Cell type

Table 1 Combination of factors used for direct neuronal reprogramming of mouse and human astrocytes

44 Katerina Aravantinou-Fatorou and Dimitra Thomaidou

Neurog2 or Ascl1

Sox2 or Olig2

Cerebral cortex

TUJ1

Functional synapses, glutamatergic and GABAergic neurons 21 dpi

14 DPT

SOX2, PAX6, NESTIN TUJ1+, MAP2+, GAD67 21 dpi + neurons, GFAP+ astrocytes and oligodendrocytes (in vivo experiment)

n.d. PBAE-based Neurons (TUJ1 and nanoparticles MAP2) or premature oligodendrocytes (PDGFRα and O4)

RV

OCT4, SOX2, or NANOG LV

Cerebral cortex

Human Cerebral astrocytes cortex

Li et al. (2016) [47]

Masserdotti et al. (2015) [46]

Corti et al. (2012) [45]

Direct Reprogramming of Astrocytes to iNs and NPCs In Vitro 45

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reprogramming of astrocytes to induced subtype specific neurons were achieved through forced expression of transcription factors known to instruct neurogenesis in embryonic development, among which the proneural transcription factors such as Pax6, Neurog2, Mash1, Dlx2 and NeuroD1, ASCL1, LMX1B, and NURR1 neurons [16, 28, 48, 49]. More recently reprogramming protocols have also focused on the use of combinations of small molecules to induce neuronal conversion with or without the use of neurogenic transcription factors [14, 30]. The most widely used miRNAs are miR9/9∗-miR124 [50], that act through posttranscriptional regulation of their multiple neuronal and glial mRNA targets, while the small chemical factors used are blockers or enhancers of certain pathways (WNT, Notch, Ca+), chromatin remodeling factors, mimics of normal proteins or RNAs, or cell cycle controllers [51]. A pioneer transcription factor which triggers neurogenesis through transdifferentiation of mouse activated astrocytes is the proneural bHLH transcription factor NEUROGENIN2, known to induce the formation of glutamatergic and dopaminergic neurons from primary astrocytes in vitro [16]. Previous studies of our group have shown that the neurogenic protein CEND1 participates in the pathway of neuronal differentiation through its activation by the bHLH preneural genes NEUROGENIN 1 and 2 [52, 53]. To this end we have used these two neurogenic proteins to induce direct neuronal reprogramming of primary mouse cortical astrocytes and have shown that CEND1 alone or in combination with NEUROGENIN2 induces reprogramming of mouse astrocytes toward in vitro production of functional induced neurons [4]. Mouse and human astrocytes share common molecular and phenotypical characteristics, but they exhibit distinguishable differences in their morphology and gene expression levels [2, 22, 54, 55], a fact suggesting that different reprogramming protocols are required for their efficient neuronal conversion. To this end we have modified the protocol we use for direct reprogramming of mouse astrocytes and have recently managed to convert human adult cortical astrocytes into neuronal cells using the same set of neurogenic factors (CEND1 and NEUROGENIN2) in combination with the small chemical molecules forskolin and valproic acid. Valproic acid (VPA) is a histone deacetylase (HDAC) inhibitor capable of increasing reprogramming efficiency by modifying chromatin accessibility to transcription factors [31], while forskolin has been reported to promote neuronal conversion efficiency [7, 56] and enhance cell morphology changes [30]. In this chapter we describe the methodology used to in vitro transdifferentiate mouse and human astrocytes toward the neuronal lineage using the neurogenic factors CEND1 and NEUROG2 (Fig. 1). According to these protocols, overexpression of each one of the two neurogenic molecules or both using retroviral vectors results in in vitro transdifferentiation of either mouse postnatal

Reprogramming Methods A. Primary pure astrocytic cultures’ reprogramming using viral vectors Primary astrocytes Cerebral Cortex dissociation

C57/Bl6 mice Brain isolation

Viral particles

1 month

20h

Analysis

B. Mouse Astrosphere Protocol Proliferation protocol -> Confocal Analysis

Day 1

Day 2

Day 3

Day 4

Plate mouse astrocytes

Cend1/ Neurog2 transduction

Neural Stem Cell Medium

Floating Astrospheres

Differentiation protocol -> Confocal Analysis

C. Primary pure human astrocytic cultures’ transductions using viral vectors Human Cerebral Cortex

Primary human astrocytic culture Expand, Freeze and astroglial marker analysis

Provided by ScienceCell

IF for molecular characterization

Day 1

Day 2

Day 3

Plate human astrocytes

Astrocytic transduction

Reprogramming media

Day 10

Differentiation media

Day 30

Induced Neuronal cultures

D. Human Astrosphere Protocol

Day 1

Day 2

Day 3

Day 7

Plate human astrocytes

Cend1 / Neurog2 tranduction

Human Neural Stem Cell Medium

Floating Astrospheres

Proliferation protocol -> Confocal Analysis Differentiation protocol -> Confocal Analysis

Fig. 1 Differentiation protocols for NPC and neuron induction upon reprogramming of mouse and human astrocytes. (a) Primary pure cortical mouse astrocytic cultures, derived from the brain of p5 C57/Bl6 mice are transduced with the retroviruses overexpressing the genes of interest and the cultures are kept for up to

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cortical astrocytes or human adult cortical astrocytes to multipotent, expandable neural precursor cells (NPCs) and differentiated neurons possessing either GABA+, TH+, or GLUT+ subtype specificity (Fig. 2). Moreover, long-term live cell imaging proved that astrocytic transdifferentiation to neurons is not always direct, but depending on the reprogramming factor being overexpressed, cells may undergo a limited number of divisions before giving rise to postmitotic neurons.

2

Materials 1. Mouse astrocytic growth medium: DMEM low glucose, 1% penicillin–streptomycin (P/S), 10% fetal bovine serum. 2. Human adult astrocytes: isolated from adult human cerebral cortex, catalogue number #1800, ScienCell Research Laboratories, https://www.sciencellonline.com/human-astrocytes. html. 3. Human astrocytic growth medium: 1:1 DMEM high glucose/ F12, 1% P/S, 100 N2, 50 B27, 100 NEAA, 10 ng/ml FGF2, 10% FBS, 0.2 mM ascorbic acid (AA), 0.2% β-mercaptoethanol, 2 mM GlutaMAX. 4. HEK-293T medium: DMEM high glucose, 1% P/S, 10% fetal bovine serum. 5. T/E neutralization solution: DMEM high glucose, 10% FCS, 1% P/S. 6. Cell culture washing solution: DMEM high glucose, 1% P/S. 7. Trypsin/EDTA (T/E) solution: 0.025% trypsin and 0.01% EDTA in Phosphate Buffered Saline (PBS).

ä Fig. 1 (continued) 1 month in vitro. Immunocytochemical or mRNA analysis experiments follow. (b) Following double transduction with the retroviruses overexpressing CEND1-IRES-GFP and NEUROG2-IRES-DSRED, the culture medium is replaced with the Neural Stem Cells Medium and the floating mouse astrospheres are propagated for several passages and analyzed for their molecular, multipotent, and phenotypical properties in the presence or absence of growth factors. (c) Primary pure human astrocytic cultures are transduced with the retroviruses overexpressing CEND1-IRES-GFP or NEUROG2-IRES-DSRED. The day after the transduction, the medium is replaced with the reprogramming medium and 1 week later the differentiation medium is induced to the culture dishes. Then the cells are kept under culture for 1 month. PCR and confocal analysis are performed in several time points during the experiment. (d) Human astrospheres are formed following double transduction with the retroviruses overexpressing CEND1-IRES-GFP and NEUROG2-IRES-DSRED. The day after the transduction the culture medium is replaced with the human Neural Stem Cells Medium and the floating astrospheres can be propagated for several passages. Human astrospheres are analyzed for their molecular, multipotent, and phenotypical characteristics after their exposure to Neural Stem Cells Medium in the presence or absence of growth factors

Direct Reprogramming of Astrocytes to iNs and NPCs In Vitro

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Fig. 2 (a, b) Mouse induced neurons are formed upon overexpression of either CEND1-IRES-GFP or NEUROG2-IRES-DSRED. CEND1 overexpression directs reprogrammed neurons toward acquisition of GABA+ identity (a), whereas NEUROG2 overexpression directs reprogrammed neurons toward TH+ identity (b). The functional marker SYNAPSIN, indicating presynaptic activity, is present in the synaptic membranes of the reprogrammed cells. (c, d). Induced human neurons also appear upon overexpression of either CEND1-IRES-GFP or NEUROG2-IRES-DSRED with well differentiated neuronal morphology, subtypespecific neuronal phenotype relevant to the mouse reprogramming results and synaptic marker expression. GABA+ neurons are formed upon CEND1 overexpression (c) and TH+ neurons upon NEUROG2 overexpression (d), whereas synaptic molecules, such as SYNAPSIN, are also present. (e–h). Live mouse and human astrospheres on a dish form 3D spheres (e–g). High NESTIN expression (f–h) in tenth passaged mouse and human astrospheres. Nuclei are stained by TOPRO-III (blue). Scale bars, 70 mm

8. Mouse differentiation medium: Neurobasal medium, 1% P/S supplemented with 3.5 mM glucose, 1 B27, 20 μl BDNF. 9. Human reprogramming medium: 1:1 DMEM high glucose/ Neurobasal, 1% P/S supplemented with 1 B27, 1 N2,

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20 ng/ml IGF, 0.2 μM AA, 100 μM dibutyryl-cAMP, small molecules: 0.5 mM VPA, 10 μM forskolin. 10. Human differentiation medium: Neurobasal medium, 1% P/S, 1 B27, 1 N2, 20 ng/ml BDNF, 20 ng/ml GDNF, 20 ng/ ml IGF, 0.2 μM ascorbic acid, 100 μM cAMP, 1 μg/ml laminin. 11. Mouse astrosphere medium: 1:1 DMEM/F12, 1 M HEPES, 1% P/S, 20 ng/ml epidermal growth factor (EGF), 20 ng/ml basic fibroblast growth factor (bFGF), supplemented with 1 B27. 12. Human astrosphere medium: 1:1 DMEM/F12, 1% P/S supplemented with 1 B27, 20 ng/ml EGF, 20 ng/ml bFGF. 13. Polylysine (PLL) stock solution: 0.015 mM polylysine in PBS. 14. Polyornithine (PLO) stock solution: 1 mg/ml PLO in PBS. 15. Laminin: 1 mg/ml laminin. 16. CaCl2: 2.5 M. 17. HBS solution. 18. pUMVC3-gag-pol: the packaging plasmid. 19. pHCMVG: the plasmid producing the pseudotyping envelope vesicular stomatitis virus glycoprotein. 20. pCAG-IRES-GFP: the expression plasmid of the protein GFP. 21. pCAG- IRES-DsRed: the expression plasmid of the protein DSRED. 22. pCAG-Cend1-IRES-GFP: the expression plasmid of the proteins CEND1 and GFP, constructed using Gateway recombination cloning technology (Invitrogen) [4]. 23. pCAG-Neurog2-IRES-DsRed: the expression plasmid of the proteins NEUROG2 and DSRED. 24. Paraformaldehyde (PFA): 4% PFA in PBS. 25. Blocking buffer: 5% goat or donkey serum in PBS, 0.1% Triton X-100. 26. Dilution buffer: 1% goat or donkey serum in PBS, 0.05% Triton X-100. 27. Mounting medium: 10% MOWIOL 4-88 diluted in 6 g of glycerol, 6 ml ddH2O and 12 ml of 0.2 M Tris (pH 8.5). 28. Single cell tracking and movie generation: Timm’s Tracking Tool (TTT) [4], Image J, Cell profiler (National Institute of Health, MD, USA) software. 29. RNA extraction: TRIzol Reagent. 30. cDNA synthesis: ImProm-II Kit (Promega) and RT2 First Strand Kit (Qiagen).

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31. qRT-PCR array kit: Mouse Neurogenesis RT2 Profiler TM PCR Array (Version 4.0, Qiagen). 32. DNA polymerase: Platinum-Taq DNA Polymerase. 33. SYBR Green. 34. Data Analysis of RT-PCR arrays: web portal of Qiagen (https://www.qiagen.com/us/shop/pcr/primer-sets/rt2profiler-pcr-arrays/?catno¼PAMM404Z#orderinginformation). 35. Image analysis software: Imaris, Olympus Cell R, Image-Pro Plus, ICY, and Image J/Fiji.

3

Methods

3.1 Mouse Astrocytic Culture

1. Primary astrocytes are derived from postnatal day 5 (P5) cortical astroglia. Postnatal day 5 (P5) C57BL/6J pups are deeply anesthetized, and under a stereoscope their brain is dissected out, the meninges removed and cerebral hemispheres are isolated and dissociated chemically using 1% trypsin/DNase and mechanically triturated with a fire-polished pasteur pipette. 2. Dissociated cells are centrifuged at 950 rpm, resuspended in mouse astrocytic growth medium and plated in a 75 cm flask coated with 0.015 mM polylysine (see Note 1). 3. Upon confluency in 3–4 days, the flask is shaken overnight at 1000  g for removal of contaminating oligodendrocyte precursor cells, microglia, and few neurons. 4. After 5 days cells are triturated using T/E and plated on polyD-lysine 10 mm glass coated coverslips at a density of 50,000 cells per coverslip (in 6- or 24-well plates) in the same medium. 5. Cells are cultured at 37  C and a CO2 concentration of 5%. The vast majority of the cells in these cultures is positive for the astrocytic marker glial fibrillary acidic protein (GFAP).

3.2 Human Astrocytic Culture

1. Human astrocytes are cultured in T-75 flasks coated with 0.015 mM PLL in human astrocytes growth medium. 2. To prepare PLL-coated T-75 flasks, 10 ml of sterile water are added to a T-75 flask, followed by 15 μl of PLL stock solution. The flask is left in a 37  C incubator overnight (or for a minimum of 1 h). The flasks are rinsed twice and 15 ml of basic growth medium is added 1 h before using them. 3. One vial of human astrocytes in quickly thawed in a 37  C water bath and cells are gently resuspended in 1 ml of basic human astrocytes growth medium and transferred into the

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equilibrated, PLL-coated flask at a density of 5000 cells/cm2 (see Notes 2 and 3). 4. Culture medium is replaced the next day by fresh one to remove residual DMSO and unattached cells, then every other day thereafter until the culture reaches 90% confluency (see Note 4). 5. About 97% of the cells express the astrocytic marker GFAP, while they are negative for the neuronal marker βIII-tubulin, DCX, the microglial marker Iba11, the glial progenitors’ marker NG2, or the oligodendrocyte marker O4, indicating that the human cell population is pure astrocytic. 3.2.1 Subculturing of Human Astrocytes

1. For subculturing human astrocytes, growth medium is removed and the cells are rinsed with the washing solution. 2. 1 ml of T/E solution is added into the T-75 flask and is gently shaken for complete coverage of cells by T/E solution. 3. The flask is incubated at 37  C for 4 min or until cells completely round up and detach from flask bottom. The side of the flask is then gently tapped to remove any remaining cells from the bottom and checked under a microscope to certify that all cells detach from flask bottom. 4. 9 ml of neutralization solution is added to the flask and detached cells are centrifuged in a 15 ml centrifuge tube at 600  g for 5 min. 5. The astrocytes are counted and replated at a density of 5000 cells/cm2 in warm complete medium.

3.3 Recombinant VSVG Pseudotyped Retrovirus Production

1. HEK-293T cells are diluted 1:3 or 1:4 in order not to form aggregates during the transformation phase. 2. The next day, the cells are plated at a density of 3  106 cells in a 100 mm diameter round plastic dish and in a volume of 7 ml of HEK-293T medium. 3. 18–24 h later and at least 1 h before transfection, the cell medium is discarded and 6 ml of fresh HEK-293T medium is added to the dishes. 4. Transfection takes place by adding the mix of 2.5 M CaCl2 and the cDNA of three plasmids simultaneously in subconfluent HEK-293T cells using the calcium chloride method: (a) 5 μg pHCMVG. (b) 10 μg pUMVC3-gag-pol. (c) 10 μg pCAG-Cend1-IRES- GFP, or pCAG-Neurog2IRESDsRed, or pCAG-IRES-GFP, or pCAG-IRESDsRed in 2 M CaCl2–ddH2O to a final volume of 500 μl. 5. Next, 500 μl of 2 HBS solution are added into the previous mixture, while bubbling in the solution with a Pasteur glass

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pipette. Immediately thereafter (within 1–2 min) the solution with the DNA is added to the cells, drop by drop. The plasmid– CaCl2–H2O solution becomes opaque at this stage. This is the most critical step for forming DNA–calcium phosphate coprecipitate. 6. The medium is replaced after 16–18 h and the culture supernatant containing viral particles is collected twice at 48 and 72 h post-transfection. 7. The collected viral supernatant is centrifuged at 1000  g for 5 min to remove dead cells and cell debris and stored at 80  C until used to transform eukaryotic cells or proceeded to ultracentrifugation. 8. Viral titer is determined in NIH 3T3 cells transduced with serial dilutions of non-concentrated viral supernatant. Additionally, transgene expression in NIH 3T3 cells is estimated 48 h post-transduction by immunocytochemistry (see Note 5). 3.4 Direct Reprogramming Protocols 3.4.1 Direct Reprogramming of Mouse Astrocytes

1. Retroviral transduction of astrocytes cultured as adherent astroglia is performed by adding the viral supernatant 1–2 days after plating them on glass coverslips coated with PLL, using retroviruses overexpressing CEND1 or NEUROG2, together with GFP or DsRED, respectively, located after an internal ribosomal entry site (IRES). Control cultures are transduced with retroviral vectors encoding IRES-GFP or IRES-DsRed only. 2. 24 h after transduction, the medium containing the viral particles is removed and replaced by the growth medium of fresh astrocytes and the cells are allowed to transdifferentiate (see Note 6). 3. 5 days posttransduction the astrocytic media is replaced by mouse differentiation and the coating of plates or coverslips is changed to PLO/laminin (PLO is added to the plates for overnight incubation at 37  C. The next morning PLO is washed three times with PBS and laminin is added for 2 h at 37  C) (see Note 7). 4. The cells are cultured at 37  C and a CO2 concentration of 5% for different time periods (1, 2, 3, 4, 7, 10, 14 days following transduction) and are further used either for immunocytochemistry and confocal imaging or live cell imaging.

3.4.2 Direct Reprogramming of Human Astrocytes

1. The human primary astrocytes are seeded at a density of 50,000 cells/cm2. 2. The following day, viral transduction is performed using the retroviruses RV-IRES-GFP, RV-IRES-DSRED, RV-CEND1IRES-GFP, and RV-NEUROG2-IRES-DSRED.

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3. Two days after seeding, the cells are over 90% confluent. At this time point, the growth medium is replaced by the reprogramming medium. The reprogramming medium containing small molecules is half changed every 2 days. 4. The second week after transduction, the culture medium is replaced by the differentiation medium. The differentiation medium is also half changed every 3 days (see Note 8) and the coating of plates or coverslips is changed to PLO/laminin (preparation of the coating the same as mentioned previously). 5. The reprogrammed cultures are kept for at least 4 weeks posttransduction and they are then fixed with paraformaldehyde 4% and processed for immunocytochemistry and confocal imaging. 3.5 Astrosphere Formation

Double overexpression of CEND1 and NEUROGENIN2 on both mouse and human astrocytic cultures results in the formation of colonies of small round cells loosely attached to the culture dish 48 h following transduction. Generation of multipotent mouse or human astrospheres from double transduced colonies is performed with the following protocol: 1. Cell colonies are collected at the bottom by centrifuging at 500  g for 5 min and the supernatant is discarded. 2. The cell pellet containing the colonies is gently resuspended in Neural Stem Cell medium. 3. The colonies are plated in uncoated 48-well plates and placed in the incubator (see Note 9). 4. The NSC medium is changed every other day (see Note 10). 5. Astrospheres are detected 21 h after transduction of mouse astrocytes and 100 h after transduction of human astrocytes and at the end of the protocol amount to 100% of the whole culture. 6. When the spheres are big enough, they are dissociated as single cells, using T/E for 5 min at 37  C, T/E neutralization solution, and washing solution as previously described. 7. Mouse astrospheres can be propagated for more than 20 passages and human astrospheres for 10 passages. They both exhibit proliferation and differentiation characteristics similar to NPCs, expressing NESTIN, whereas following withdrawal of growth factors (EGF and bFGF), they give rise to bIIITUBULIN+ neurons, GFAP+ astrocytes, and O4+ oligodendrocytes.

3.6

Live Cell Imaging

Time-lapse video microscopy of transduced astrocytic cultures is performed 12 h after transduction using the wide-field time-lapse Olympus IX-81 microscope at constant conditions of 37  C and 7% CO2. Phase-contrast videos are acquired as follows:

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1. The microscope, the cell chamber system for control of conditions, the camera, and the cell^R software are turned on 2 h before the initiation of the experiment (see Note 11). 2. The 6-well experimental plate is then transferred to the chamber and left in there for 1/2 h. 3. Nine different fields per well are chosen to be followed, saving their x, y, z coordinates in cell^R software. 4. The experiment is programmed to acquire phase-contrast images of each chosen field every 5 min for 3–10 days using 10 or 20 low-magnification phase-contrast objectives, using a Hamamatsu ORCA camera (CCD ORCA/AG) (see Note 12). 5. By the end of the experiment, immunocytochemistry for βIIITUBULIN is performed inside the plate in order to identify induced neurons. The plate is then placed back on the microscope stage and the coordinates of each induced neuron during time lapse are marked using cell^R software. 6. Single-cell tracking is performed using the TTT computer program. 7. Movies are assembled using TTT, Image J, or Cell profiler software and are run at a speed of 15 frames/s, to track the lineage trees of transduced cells. 8. Live-cell imaging starting as soon as 12 h after double CEND1/NEUROG2 transduction combined with lineage tracing is also performed to visualize the timing of the first divisions resulting in mouse astrosphere formation. 3.7 Immunofluorescence and Confocal Imaging

1. Cells on coverslips are fixed with 4% PFA for 20 min and blocked for unspecific binding of the antibodies with the blocking buffer for 1 h at room temperature. 2. Cells are incubated overnight at 4  C with the following antibodies:, mouse monoclonal glial fibrillary acidic protein (antiGFAP), rabbit polyclonal anti-GFAP, mouse monoclonal antiBrdU, mouse monoclonal anti-neuronal β-III TUBULIN, rabbit monoclonal anti-NEUROG2, rabbit polyclonal antineuronal β-III TUBULIN, rabbit polyclonal anti-GABA, rat polyclonal anti-DsRED, rabbit polyclonal anti-GFP, mouse monoclonal anti-GFP, guinea pig anti-glutamate transporter I, goat polyclonal anti-Ki67, mouse monoclonal anti-CEND1 (homemade), rabbit polyclonal anti-CEND1 (homemade), chicken polyclonal anti-NESTIN, rabbit polyclonal anti-NG2, mouse monoclonal anti-O4, rabbit polyclonal anti-OLIG2, rabbit polyclonal anti-SYNAPSIN-1, and rabbit polyclonal anti-TH, which are diluted into the dilution buffer. 3. The following day, cells are incubated with the appropriate secondary goat or donkey antibodies conjugated with Alexa

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Fluor 488, 546, or 647, which are diluted into the dilution buffer, and mounted in the mounting medium. 4. DAPI or TopRoIII is used to stain nuclei at room temperature simultaneously with the secondary antibodies. 5. Confocal or epifluorescence microscopy is performed using a Leica TCS-SP5II or SP8 confocal microscope or a wide-field time-lapse Olympus IX-81 Cell-R imaging system (10 and 20 airy objectives or 40 and 60 oil immersion objectives). 6. Images are analyzed using Imaris, Olympus Cell R, Image-Pro Plus, ICY, and Image J/Fiji software. 7. For quantification and statistical analysis, the total numbers and percentages of all different molecular phenotypes are counted in 30 randomly selected 20 visual fields. This sampling is repeated in three independent experiments and the average number was used to calculate the mean number of positive cells/visual field. 8. Quantification of process length and number is performed using the Cell profiler open source software. Total process length is determined by tracing each individual neuron, and the number of end processes was counted manually for the same cells. 9. Difference between mean values is analyzed using unpaired t tests. Student’s t-test comparison within pairs of groups with p-value 0.05 is considered as statistically significant (∗p-value 0.05; ∗∗p-value 0.01, ∗∗∗p-value 0.001, ∗∗∗∗ p-value 0.0001). 3.8 Real-Time qPCR Analysis

1. Total RNA is collected using the TRIzol RNA isolation protocol and cDNA synthesis is performed using ImProm-II Kit with hexamer primers for neurogenesis-related genes (Tables 2 and 3). 2. Real-time Quantitative PCR is performed in triplicate in a Roche LightCycler 96 PCR machine with Platinum-Taq DNA Polymerase and SYBR Green. 3. Quantification is carried out with the ΔΔCt method using GAPDH as a reference gene.

3.9

RT-PCR Arrays

1. Total RNA is collected using the TRIzol RNA isolation protocol and the RT2. First Strand Kit is used for the cDNA synthesis. 2. qRT-PCR array analysis for 84 key factors related to the processes of neurogenesis is performed on 96-well plates using the Mouse Neurogenesis RT2 Profiler TM PCR Array. 3. Data analysis is performed by converting Raw Ct Data to FoldChange. Results are generated via the web portal of Qiagen.

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Table 2 List of primer sequences for the detection of mouse mRNA expression level Oligo name

Sense/Antisense

cDNA

Sequence (50 –30 )

Cend1 F

Sense

Genomic

GGCTAACTAGGGAACCCACTG

Cend1 R

Antisense

Genomic

GCTAGAGATTTTCCACACTGACTAA

Neurogenin2 F

Sense

Genomic

AGAAGACCCGCAGGCTCAAG

Neurogenin2 R

Antisense

Genomic

CGTGGAGTTGGAGGATGACG

β-Catenin F

Sense

Genomic

ATGGAGCCG GACAGAAAAGC

β-Catenin R

Antisense

Genomic

CTTGCCACTCAGGGAAGGA

RLP5 F

Sense

Genomic

GAAGAGAAGGCGAGTGACCG

RLP5 R

Antisense

Genomic

AAGGACAGAACTCTGTGGCG

Brn2 F

Sense

Genomic

GCAGCGTCTAACCACTACAGC

Brn2 R

Antisense

Genomic

GCGGTGATCCACTGGTGA

Frizzled4 F

Sense

Genomic

CAGCTGACAACTTTCACGCC

Frizzled4 R

Antisense

Genomic

CCGAACAAAGGAAGAACTGC

GAPDH F

Sense

Genomic

AACTCCCTCAAGATTGTCAGCAA

GAPDH R

Antisense

Genomic

ATGTCAGATCCACAACGGATACA

Table 3 List of primer sequences for the detection of human mRNA expression level Oligo name

Sense/Antisense

cDNA

Sequence (50 –30 )

MASH1 F

Sense

Genomic

AAGAGCAACTGGGACCTGAGTCAA

MASH1 R

Antisense

Genomic

AGCAAGAACTTTCAGCTGTGCGTG

NCAM F

Sense

Genomic

ACATCACCTGCTACTTCCTGA

NCAM R

Antisense

Genomic

CTTGGACTCATCTTTCGAGAAGG

MAP2 F

Sense

Genomic

GAGAATGGGATCAACGGAGA

MAP2 R

Antisense

Genomic

CTGCTACAGCCTCAGCAGTG

NGN2 F

Sense

Genomic

TCAGACATGGACTATTGGCAG

NGN2 R

Antisense

Genomic

GGGACAGGAAAGGGAACC

GFAP F

Sense

Genomic

GGTTGAGAGGGACAATCTGG

GFAP R

Antisense

Genomic

GGGTGGCTTCATCTGCTTC

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Notes 1. It is important that the astrocytes, mouse or human, are plated in poly-L-lysine or poly-D-lysine–coated flasks to promote cell attachment. 2. When human astrocytes get unfrozen from liquid nitrogen, 106 cells should be placed in a T-25 PLL or PDL coated flask. They will be confluent in about a week, maximum 10 days in the case of human astrocytes. If not, the culture should be discarded. 3. The flask’s cap should be loosened, if it is not a filtered one, to allow gas exchange. 4. For best results, the culture must not be disturbed for at least 24 h after its initiation. The astrocytes will grow better and faster if they are not removed frequently out of the incubator for the first few days of the experiment. 5. The produced viral particles have the ability to transfect the target cells; however, they cannot proliferate within them, since the gag, pol, and env genes necessary for their proliferation are not included in their genome. 6. During the reprogramming, cells’ morphology and confluence should be thoroughly examined in a daily basis under the microscope, to observe their morphological changes and proliferation rate during the transdifferentiation process, as these events happen fast. Multiple wells should be used for each experimental condition, in order to stop the experiment at various time-points according to the scientific question raised or depending on the transduction efficiency, the reprogrammed population, the presence of neurons in the exact time points, the cellular confluency, and so on. 7. As soon as the first cells with neuronal morphology appear in culture, cells should be transferred to PLO/Laminin-coated coverslips or plates, since the newborn neurons grow better on that type of coating and they survive for longer time periods under culture. 8. Reprogrammed cultures should contain a high number of astrocytes. The astrocytes support the newborn neurons, promote their differentiation and synapse formation and for this reason the more differentiated newborn neurons lie on top of astrocytes. If the transformation efficiency is extremely good and the culture is out of mature astrocytes, astrocytes can be seeded on top to enrich the culture. 9. The first isolation of the astrospheres might be tricky, as in order to grow healthy robust spheres, a good density of cells is required. If the spheres are few in a big well, they will die

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shortly after their isolation. If a small number of new astrospheres is produced, they should be initially transferred in a 96-well plate, to increase their density. 10. The Neural Stem Cell medium must be freshly made every time before being used, for best results. A fresh batch should be prepared every 3–4 days, and bFGF and EGF must be added to the amount used just before each experiment. 11. Before the start of the long-term live cell imaging experiment, it is extremely important to switch on the system and the conditions of temperature, humidity, and CO2, at least 2 h before the plate is placed in the microscope stage, so as all microscope metal surfaces adjust to those conditions and no shifts and loss of focus occur during imaging. 12. During long-term live cell imaging, warm medium should be added whenever it is needed, without removing the plate of the microscope stage. References 1. Pekny M, Nilsson M (2005) Glia 50 (4):427–434. https://doi.org/10.1002/glia. 20207 2. Sofroniew MV, Vinters HV (2010) Astrocytes: biology and pathology. Acta Neuropathol 119 (1):7–35. https://doi.org/10.1007/s00401009-0619-8 3. Heinrich C, Spagnoli FM, Berninger B (2015) In vivo reprogramming for tissue repair. Nat Cell Biol 17(3):204–211. https://doi.org/10. 1038/ncb3108 4. Aravantinou-Fatorou K, Ortega F, ChroniTzartou D et al (2015) CEND1 and NEUROGENIN2 reprogram mouse astrocytes and embryonic fibroblasts to induced neural precursors and differentiated neurons. Stem Cell Rep 5(3):405–418. https://doi.org/10. 1016/j.stemcr.2015.07.012 5. Guo Z, Zhang L, Wu Z et al (2014) In vivo direct reprogramming of reactive glial cells into functional neurons after brain injury and in an Alzheimer’s disease model. Cell Stem Cell 14 (2):188–202. https://doi.org/10.1016/j. stem.2013.12.001 6. Niu W, Zang T, Zou Y et al (2013) In vivo reprogramming of astrocytes to neuroblasts in the adult brain. Nat Cell Biol 15 (10):1164–1175. https://doi.org/10.1038/ ncb2843 7. Torper O, Pfisterer U, Wolf DA et al (2013) Generation of induced neurons via direct conversion in vivo. Proc Natl Acad Sci U S A 110 (17):7038–7043. https://doi.org/10.1073/ pnas.1303829110

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Direct Reprogramming of Astrocytes to iNs and NPCs In Vitro 39. Karow M, Sa´nchez R, Schichor C et al (2012) Reprogramming of pericyte-derived cells of the adult human brain into induced neuronal cells. Cell Stem Cell 11(4):471–476. https://doi. org/10.1016/j.stem.2012.07.007 40. Biswas D, Jiang P (2016) Chemically induced reprogramming of somatic cells to pluripotent stem cells and neural cells. Int J Mol Sci 17 (2):226. https://doi.org/10.3390/ ijms17020226 41. Liu ML, Zang T, Zou Y et al (2013) Small molecules enable neurogenin 2 to efficiently convert human fibroblasts into cholinergic neurons. Nat Commun 4:2183. https://doi. org/10.1038/ncomms3183 42. Buffo A, Milan RV, Erturk D et al (2005) Expression pattern of the transcription factor Olig2 in response to brain injuries: Implications for neuronal repair. PNAS. 102 (50):18183–18188. https://doi.org/10. 1073/pnas.0506535102 43. Niu W, Zang T, Smith DK, Vue TY et al (2015) SOX2 Reprograms resident astrocytes into neural progenitors in the adult brain. Stem Cell Rep 4(5):780–794. https://doi.org/10. 1016/j.stemcr.2015.03.006 44. Chouchane M, Melo de Farias AR, Sousa Moura DM et al (2017) Lineage reprogramming of astroglial cells from different origins into distinct neuronal subtypes. Stem Cell Rep 9(1):162–176. https://doi.org/10.1016/j. stemcr.2017.05.009 45. Corti S, Nizzardo M, Simone C et al (2012) Direct reprogramming of human astrocytes into neural stem cells and neurons. Exp Cell Res 318(13–16):1528–1541. https://doi. org/10.1016/j.yexcr.2012.02.040 46. Masserdotti G, Gillotin S, Sutor B et al (2015) Transcriptional mechanisms of proneural factors and REST in regulating neuronal reprogramming of astrocytes. Cell Stem Cell 17 (1):74–88. https://doi.org/10.1016/j.stem. 2015.05.014 47. Li X, Kozielski K, Cheng Y et al (2016) Nanoparticle-mediated conversion of primary human astrocytes into neurons and oligodendrocytes. Biomater Sci 4(7):1100–1112. https://doi.org/10.1039/c6bm00140h

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Chapter 5 Preparation of Bioscaffolds Delivering Stem Cells for Neural Regeneration Li Yao and Ashley DeBrot Abstract Bioscaffolds have been proven for their feasibility in neural repair. Neural conduits have been investigated in the repair of wounded peripheral nerve and spinal cord. These conduits support axonal growth by providing structural guidance. Induced pluripotent stem cells (iPSCs) that are induced from a patient’s own somatic cells have demonstrated significant neural cell differentiation capability and can circumvent immune system rejection. The combinatorial implantation of neural conduits and iPSCs may significantly enhance neural regeneration. The repair of nerves and spinal cords using biodegradable multichannel collagen conduits has been reported in our previous studies. In this review, we describe a method to fabricate a collagen neural conduit containing iPSC-derived neural cells. Key words Bioscaffolds, Stem cells, iPSC cells, Conduit, Axon, Spinal cord injury, Nerve injury, Regeneration

1

Introduction In the development of therapeutic strategies, bioscaffolds have been used to repair wounded neural tissue. In a number of studies, biomaterial scaffolds have been implanted in wounded spinal cord, including compression, hemisection, and complete transection spinal cord injury (SCI) models. Biomaterial scaffolds that fill the defect of neural injury conduct axonal regeneration. The wounded spinal cord of a complete transection model is normally repaired by a nerve conduit that provides structural guidance for axonal growth [1–5]. In the majority of peripheral nerve injuries, the nerve ends cannot be directly sutured, and a nerve conduit is needed to connect the nerve end in the nerve repair. A number of synthetic and natural biomaterials have been fabricated as neural conduits and used to repair a wounded nerve [6–11]. However, these studies have shown limited success since the repair outcome using these nerve conduits is inferior to the autologous nerve repair. In previous studies, we generated collagen-based nerve conduits and

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investigated them for peripheral nerve and spinal cord repair [12– 14]. These bioscaffolds can potentially serve as a carrier for stem cell delivery into wounded spinal cord or peripheral nerve. Induced pluripotent stem cells (iPSCs) that are induced from a patient’s own somatic cells offer several benefits in neural repair because they are free of the ethical concerns associated with embryonic stem cells (ESCs) and can circumvent immune system rejection. In one study, IPSCs were grafted into wounded spinal cord, and the outcome showed that the iPSC-derived neurons formed synapses with host neurons and improved the functional recovery [15]. We have developed a method to combine the application of a nerve conduit and iPSC-derived neural stem cells. In this chapter, we introduce the preparation of a nerve conduit containing iPSC-derived neural stem cells for implantation into wounded peripheral nerve or spinal cord.

2

Materials

2.1 Fabrication of Conduit

1. Cylindrical four-channel negative molds with four or seven channels having an internal diameter of 530 μm. 2. 50 mM acetic acid solution: add 2.85 ml acetic acid solution into 1000 ml of distilled water, and keep the solution in a refrigerator at 4  C. 3. 10 mg/ml type I collagen solution: dissolve 1 g type I collagen in 100 ml 50 mM acetic acid solution, and keep the solution in a refrigerator at 4  C. 4. 2-Morpholinoethanesulfonic (MEF) acid solution (50 mM): dissolve 9.75 g MEF in 1000 ml distilled water. 5. 1-Ethyl-3-(3-dimethylaminopropyl) solution (0.877 g/ml).

carbodiimide

(EDC)

6. N-Hydroxysuccinimide (HNS) solution. 7. NaH2PO4 (0.1 M) solution: dissolve 11.9 g NaH2PO4 in 1000 ml distilled water. 8. Stainless steel wires. 2.2 Differentiation of i-HNSCs into Neurons

1. Induced pluripotent stem cell-derived human neural stem cells (i-HNSCs) (Cell Applications, Inc.). 2. i-HNSC culture medium (Cell Applications, Inc.). 3. Neuron differentiation medium (Cell Applications, Inc.). 4. Poly-L-ornithine (20 μg/ml). 5. Accutase cell detachment reagent. 6. Laminin (10 μg/ml). 7. Sterile Dulbecco’s phosphate-buffered saline (DPBS).

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2.3 Live/Dead Cell Viability Assay

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1. Reagents for LIVE/DEAD® assay: ethidium homodimer-1 (Ethd-1) and Calcein AM. 2. Matrigel.

2.4 Immunocytochemistry

1. Anti-neurofilament antibody (TUJ1). 2. Alexa Fluor® 488 donkey anti-mouse secondary antibody. 3. Blocking solution: a mixture of 10 ml fetal bovine serum and 90 ml phosphate-buffered saline (PBS). 4. 0.01% Triton X-100 solution: a mixture of 0.01 ml Triton X-100 and 100 ml PBS. 5. 4% paraformaldehyde. 6. Phosphate-buffered saline.

3

Methods

3.1 Fabrication of Multichannel Collagen Nerve Conduits (See Fig. 1)

1. Insert two stainless steel wires into two opposite channels of the molds. 2. Ensure that the distance between the edges of the two molds is 2 mm (see Note 1). 3. Coat the top of the two stainless steel wires with the collagen solution. 4. Air-dry the collagen on the wires. 5. Insert the additional two wires into the two adjacent channels. 6. Coat the entire surface of the four wires with the collagen solution. 7. Air-dry the collagen on the wires. 8. Insert one wire into the remaining channel of the two molds. 9. Place the collagen solution around the wires. 10. Air-dry the collagen on the wires. 11. Transfer the air-dried collagen conduits with the stainless steel wires into a plastic tube containing the cross-linking solution of EDC (20 mM) and NHS (10 mM) in an MEF solution (50 mM; pH 5.5) (see Note 2). 12. Cross-link the conduits for 6 h. 13. Rinse the cross-linked conduit with the stainless steel wires in distilled water. 14. Transfer the conduits to an NaH2PO4 (0.1 M) solution and leave the conduits in solution for at least 1 h. 15. Wash the conduits with distilled water repeatedly. 16. Freeze the collagen conduits with stainless steel wires in a freezer at 80  C.

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Fig. 1 Collagen conduits: (a) collagen conduit fabricated on molds with stainless steel wires inserted; (b) nerve conduits for peripheral nerve repair; (c, d) seven-channel collagen conduit for spinal cord repair. (Figure reproduced from Yao et al. [13, 14])

17. Freeze-dry the conduits with wires in a freeze dryer. 18. Remove the stainless steel wires and molds from the collagen conduits. 3.2 Characterization of i-HNSC Differentiation by Immunocytochemistry

1. Coat a 24-well plate with poly-L-ornithine for 1 h at 37  C and with laminin for 2 h at 37  C. 2. Seed 5000 i-HNSCs in each of the 24 wells, and culture the cells with an i-HNSC culture medium in a 37  C 5% CO2 incubator (see Note 3). 3. After 2 days, change the medium with a neuron differentiation medium, and change this medium every 3 days. 4. After culturing for about 2–3 weeks, study the cells for immunostaining or for 3D culturing in Matrigel. 5. To perform immunostaining, fix the cells with 4% paraformaldehyde for 30 min. 6. Wash the cells with PBS three times. 7. Permeabilize the cells with 1 Triton X-100 for 20 min.

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Fig. 2 Growth of i-HNSCs and i-HNSC-derived neurons in Matrigel: (a) bright field image of i-HNSC-derived neurons, (b) i-HNSC-derived neurons labeled with anti-neurofilament antibody (TUJ1), (c, d) growth of i-HNSCs in Matrigel, (d) live i-HNSCs in Matrigel labeled with Calcein AM, (e, f) growth of i-HNSC-derived neurons in Matrigel, (f) live i-HNSC-derived neurons labeled with Calcein AM

8. Wash the cells with PBS three times. 9. Treat the cells with 10% FBS to prevent nonspecific binding for 1 h. 10. Wash the cells with PBS three times. 11. Label the neurons with anti-neurofilament antibody and place the cell plate in a refrigerator (4  C) overnight. 12. Wash the cells with PBS three times. 13. Treat the cells with secondary antibody and keep the plate at room temperature for 2 h. 14. Wash the cells with PBS three times. 15. Observe the labeled cells and take images using fluorescent microscope (Fig. 2a, b). 3.3 Growth of Stem Cells in Matrigel Within Neural Conduit

1. Culture the i-HNSCs in a 6-well culture plate according to the protocol of Cell Applications, Inc. 2. When the cells are confluent, remove the cell culture medium and wash the cell culture layer with sterile DPBS. 3. Add 0.5 ml Accutase into the cell culture plate grown with i-HNSCs to detach the cells. 4. Add cell culture medium into the cell culture wells, and collect all cells and medium in a centrifuge tube. 5. Centrifuge the cells at 200  g for 5 min.

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6. Remove the cell culture medium and add fresh medium. 7. Perform cell counting. 8. Transfer the cell culture medium with 100,000–500,000 million cells into a centrifuge tube. 9. Centrifuge the cells at 200  g for 5 min. 10. Transfer the Matrigel from the freezer to room temperature to defrost it. 11. Transfer about 50 μl of Matrigel solution into a centrifuge tube and leave it at room temperature for 5–10 min before mixing cells with the gel. 12. Remove the medium and transfer the Matrigel solution into the centrifuge tube. Then mix the Matrigel with cell pellets using a pipette to disperse the cells in the gel. 13. Using a pipette, inject the Matrigel into the channels of the conduit. 14. Transfer the conduits into a well of 24-well cell culture plate and place the plate in a 37  C, 5% CO2 incubator for 5–10 min. 15. Transfer 1 ml of the cell culture medium into each cell culture well. 3.4 Cell Viability Assay for Cells Grown in Matrigel

1. Collect i-HNSCs and i-HNSC–derived neurons as described in Subheading 3.3. 2. Bring the Matrigel to room temperature to defrost it. 3. Transfer 200 μl of the Matrigel solution into one well of the 48-well plate and leave the Matrigel at room temperature for 5–10 min before adding cells to the gel. 4. Mix 100,000 cells into 200 μl of Matrigel and stir the cells and Matrigel solution with pipette tips. 5. Transfer the plate into an incubator and leave it there for 5–10 min to allow the Matrigel to form. 6. Add 800 μl of cell culture medium to the well. 7. After culturing the cells in Matrigel for various days, perform the live/dead cell viability assays to those cells grown in the Matrigel. 8. Remove solutions of the assay from the freezer and allow them to warm to room temperature. 9. Transfer 2 μl EthD-1 solution and 0.5 μl Calcein AM solution into 1 ml of the sterile PBS solution in a centrifuge tube. 10. Vortex the centrifuge tube. 11. Remove the cell culture medium from the culture well and transfer 300 μl of the solution into each well.

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Fig. 3 Migration of i-HNSC-derived neurons in hydrogels (lines indicate tracks of migration of cells in Matrigel)

12. Incubate the cells with solution for 20 min in an incubator. 13. Visualize the labeled cells under a fluorescent microscope (Fig. 2c–f). 3.5 Time-Lapse Recording of i-HNSC Migration in Matrigel

1. Grow the i-HNSCs in Matrigel as described in Subheading 3.4. 2. After the cells are cultured for various days, transfer the cell culture plate into a time-lapse microscope placed in a plastic incubator supplied with 5% CO2, and control the incubator temperature at 37  C by warm air generated by a heater. 3. Record the migration of cells for 3 h, and capture the images every 5 min, as was the reported method for cell migration in a 3D hydrogel [16, 17]. 4. Analyze the time-lapse images using NIH ImageJ software and the Chemotaxis and Migration Tool (Fig. 3).

4

Notes 1. The distance between the two molds determines the length of the conduits that varies according to the need for nerve or spinal cord repair. 2. The concentration of EDC cross-linking solution can modulate the cross-linking level of the collagen conduit. 3. The culture method of i-HNSCs is described in Subheading 3.3.

References 1. Teng YD, Lavik EB, Qu X, Park KI, Ourednik J, Zurakowski D, Langer R, Snyder EY (2002) Functional recovery following traumatic spinal cord injury mediated by a unique polymer scaffold seeded with neural stem cells. Proc Natl Acad Sci U S A 99(5):3024–3029

2. Straley KS, Foo CW, Heilshorn SC (2010) Biomaterial design strategies for the treatment of spinal cord injuries. J Neurotrauma 27 (1):1–19 3. Olson HE, Rooney GE, Gross L, Nesbitt JJ, Galvin KE, Knight A, Chen B, Yaszemski MJ,

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Windebank AJ (2009) Neural stem cell- and Schwann cell-loaded biodegradable polymer scaffolds support axonal regeneration in the transected spinal cord. Tissue Eng Part A 15 (7):1797–1805 4. De Laporte L, Yan AL, Shea LD (2009) Local gene delivery from ECM-coated poly(lactideco-glycolide) multiple channel bridges after spinal cord injury. Biomaterials 30 (12):2361–2368 5. Chen BK, Knight AM, Madigan NN, Gross L, Dadsetan M, Nesbitt JJ, Rooney GE, Currier BL, Yaszemski MJ, Spinner RJ, Windebank AJ (2011) Comparison of polymer scaffolds in rat spinal cord: a step toward quantitative assessment of combinatorial approaches to spinal cord repair. Biomaterials 32(32):8077–8086 6. Rui J, Dadsetan M, Runge MB, Spinner RJ, Yaszemski MJ, Windebank AJ, Wang H (2012) Controlled release of vascular endothelial growth factor using poly-lactic-co-glycolic acid microspheres: in vitro characterization and application in polycaprolactone fumarate nerve conduits. Acta Biomater 8(2):511–518 7. Tang X, Xue C, Wang Y, Ding F, Yang Y, Gu X (2012) Bridging peripheral nerve defects with a tissue engineered nerve graft composed of an in vitro cultured nerve equivalent and a silk fibroin-based scaffold. Biomaterials 33 (15):3860–3867 8. de Ruiter GC, Spinner RJ, Malessy MJ, Moore MJ, Sorenson EJ, Currier BL, Yaszemski MJ, Windebank AJ (2008) Accuracy of motor axon regeneration across autograft, single-lumen, and multichannel poly(lactic-co-glycolic acid) nerve tubes. Neurosurgery 63(1):144–153; discussion 153–5 9. Daly WT, Yao L, Abu-rub MT, O’Connell C, Zeugolis DI, Windebank AJ, Pandit AS (2012) The effect of intraluminal contact mediated guidance signals on axonal mismatch during peripheral nerve repair. Biomaterials 33 (28):6660–6671 10. Daly W, Yao L, Zeugolis D, Windebank A, Pandit A (2012) A biomaterials approach to

peripheral nerve regeneration: bridging the peripheral nerve gap and enhancing functional recovery. J R Soc Interface 9(67):202–221 11. Pawelec KM, Koffler J, Shahriari D, Galvan A, Tuszynski MH, Sakamoto J (2018) Microstructure and in vivo characterization of multi-channel nerve guidance scaffolds. Biomed Mater 13(4):044104 12. Yao L, Billiar KL, Windebank AJ, Pandit A (2010) Multichanneled collagen conduits for peripheral nerve regeneration: design, fabrication, and characterization. Tissue Eng Part C Methods 16(6):1585–1596 13. Yao L, de Ruiter GC, Wang H, Knight AM, Spinner RJ, Yaszemski MJ, Windebank AJ, Pandit A (2010) Controlling dispersion of axonal regeneration using a multichannel collagen nerve conduit. Biomaterials 31 (22):5789–5797 14. Yao L, Daly W, Newland B, Yao S, Wang W, Chen BK, Madigan N, Windebank A, Pandit A (2013) Improved axonal regeneration of transected spinal cord mediated by multichannel collagen conduits functionalized with neurotrophin-3 gene. Gene Ther 20 (12):1149–1157 15. Nori S, Okada Y, Yasuda A, Tsuji O, Takahashi Y, Kobayashi Y, Fujiyoshi K, Koike M, Uchiyama Y, Ikeda E, Toyama Y, Yamanaka S, Nakamura M, Okano H (2011) Grafted human-induced pluripotent stem-cellderived neurospheres promote motor functional recovery after spinal cord injury in mice. Proc Natl Acad Sci U S A 108 (40):16825–16830 16. Kaphle P, Li Y, Yao L (2019) The mechanical and pharmacological regulation of glioblastoma cell migration in 3D matrices. J Cell Physiol 234(4):3948–3960 17. Seyedhassantehrani N, Li Y, Yao L (2016) Dynamic behaviors of astrocytes in chemically modified fibrin and collagen hydrogels. Integr Biol (Camb) 8(5):624–634

Chapter 6 Improved Isolation of Human Vascular Wall–Resident Mesenchymal Stem Cells Diana Klein Abstract Niches for tissue-resident mesenchymal stem cells (MSCs) have been identified in many adult tissues. In particular, MSCs residing in the vascular stem cell niche came into focus: the so-called vascular wall–resident MSCs (VW-MSCs) were, based upon their anatomic location, (1) distributed throughout the adult organism, and (2) supposed to be the first line cells which could be addressed in response to a pathologic trigger acting on or in close vicinity to the vascular system. Like tissue-resident MSCs in general, VW-MSC contribute to organ integrity and harbor the capacity to suppress inflammation and promote repair during normal vessel homeostasis, although resident MSCs present in the healthy situation of an individual seems not to bear sufficient for protection or repair following injury. In contrast, injury affected MSCs could contribute to disease induction and progression. A detailed understanding of the molecular repertoire as well as of the signaling pathways controlling stem cell fate of VW-MSCs is prerequisite to understand how (1) endogenous VW-MSCs contribute to normal vessel homeostasis as well as diseases that include the vascular system, (2) a potential on-site manipulation of these cells directly within their endogenous niche could be used for therapeutically benefits, and (3) isolated and therapeutically applied VW-MSCs in terms of exogenous MSCs with superior repair capabilities might be logically more efficient to address vascular diseases than MSCs derived from other tissues. This chapter describes a straightforward protocol for the improved isolation of human VW-MSCs. Key words Tissue-resident, Stem cell, MSC, Vascular wall, Vascular stem cell, Adventitia, Vasculogenic zone

1

Introduction Tissue-specific stem cells have been shown to regulate maintenance and regeneration of postnatal tissues [1, 2]. Among these adult stem cells, multipotent mesenchymal stem cells (MSCs) are phenotypically characterized as plastic adherent cells with a fibroblast-like morphology that express the surface markers CD90, CD105, CD73, CD44, and CD146 while being lineage negative (CD45, CD31, CD34, CD11b, CD79a, CD19, HLA-DR). MSCs bear clonogenic capabilities and were able to differentiate into mesenchymal (and also non-mesenchymal) tissues in vitro and in vivo

Chrissa Kioussi (ed.), Stem Cells and Tissue Repair: Methods and Protocols, Methods in Molecular Biology, vol. 2155, https://doi.org/10.1007/978-1-0716-0655-1_6, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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[3, 4]. Furthermore, MSCs possess the ability to modulate the immune system [5, 6]. Isolated and therapeutically applied (exogenous) MSCs lack ethical concerns and are well tolerated as these cells lack histocompatibility as well as teratoma-formation issues [7, 8]. Thus, MSCs are supposed to be the most promising stem cell type for cell-based therapies [9–11]. Although MSCs have been described to home at the site of tissue damage once therapeutically applied , the current view is that MSCs act indirectly through a paracrine mechanism of action. Herein, MSCs transiently provide a local source of trophic factors such as protective growth factors and cytokines in the local environment and thereby preserve existing functional tissue from further destruction or support regenerative processes [12–14]. The most frequent source for obtaining MSCs is the bone marrow; respective cells have also been succesfully isolated from umbilical cord blood, placenta, blood, fetal liver, lung, adipose tissue, and blood vessels [15–20]. In particular, MSCs residing in blood vessels, the so-called vascular wall–resident MSCs (VW-MSCs), turned out to be decisive for vascular morphogenesis, repair, and selfrenewal of vascular wall cells and for the local capacity of neovascularization in disease processes [19–23]. VW-MSCs, together with endothelial and hematopoietic progenitor cells, reside within the vasculogenic zone of large and mid-sized blood vessel, an adventitial stem cell niche that is closely located to the smooth muscle cell layer [19, 21–23]. A basic understanding of VW-MSCs and their adventitial niche as well as their fate during vascular diseases is a prerequisite to improve autologous therapies, as well as drug interventions, for example, to target delivery of small molecules or growth factors required for (VW-)MSC activation, proliferation, migration and finally differentiation within and out of their respective vessel niche. Nowadays it has become clear that tissue-specific stem cells do not differentiate with the same efficacy in lineage-specific cell types because tissue-specific stem cells are generally more effective when differentiating toward cell types typical for their tissue of origin. Therefore, it was speculated that VW-MSCs are particularly wellsuited to address vascular damage [21, 24–26]. Affirmative, VW-MSCs turned out to mediate vascular protection more efficiently than bone marrow–derived MSCs (BM-MSCs) [24, 25]. Earlier reports already suggested that the classically BM-MSC were less effective for MSC therapy as compared to other stem cell sources, for example, as compared to adipose tissue–derived or fetal MSCs [27–32]. Thus, VW-MSCs offer a promising therapeutic option for the prevention and treatment of diseases associated with vascular damage and remodeling, for example hypertension, ischemic diseases, congenital vascular lesions (aneurysms, fibromuscular hyperplasia, stenosis in collaterals), shear stress, or irradiation [33– 35]. Here a detailed protocol for the relatively simple extraction of

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Fig. 1 Scheme for the improved isolation of human vascular wall–resident mesenchymal stem cells. CD44(+) VW-MSC can be isolated from large and mid-sized arteries as well as from greater saphenous vein fragments (obtained after bypass surgery; remaining/sparse material). Vessel fragments, mechanically cleared from contaminating muscle and fatty tissue, will be subjected for collagenase digestion (generation of a crude cell extract). Highly pure VW-MSCs depleted of contaminating cell types can be yielded using MACS-technology in combination with a monoclonal CD44 antibody (positive selection) followed by a selective adherence on plastic dishes in appropriate MSC medium (medium selection) for further expansion. The purity of these cell preparations is routinely >95% as to be analyzed by expression of a marker panel including STRO1, CD105, CD73, CD44, CD90, and CD29 in the absence of lineage markers (lack of CD31, CD34, CD45, as well as CD68, CD11b, CD19) via flow cytometry. Cultured VW-MSCs will show a flattened, fibroblast-like pattern typical for MSCs and form clonally cell aggregates upon culturing. In order to access clonogenicity and multipotency of isolated CD44(+) VW-MSCs, single-cell-derived clones should be established (by limited dilution in a 96-well plate). In vitro, these cells will differentiate into adipocytes, chondrocytes and osteocytes, as well as into pericytes and smooth muscle cells that in turn stabilize newly formed vascular sprouts [20, 26, 37]

CD44(+)CD90(+)CD73(+)CD34( )CD45( ) VW-MSCs out of adult human blood vessels is provided (Fig. 1) [20].

2

Material

2.1 Cell Isolation Components

1. Cell culture phosphate-buffered saline (cell culture 1 PBS pH 7.2, sterile), supplemented with 100 U penicillin/100 μg streptomycin (P/S). 2. Collagenase solution: Opti-MEM™ I Reduced Serum Medium (no phenol red) containing 0.2% type 2 collagenase (see Note 1). 3. PBS containing 5% FCS (see Note 2). 4. 70 μm cell strainer, sterile. 5. Cell culture medium: Mesenchymal Stem Cell Growth Medium 2 (MSC-GM2).

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6. Tissue culture dishes/flasks for adherent cell culture (see Note 3). 2.2 Immunomagnetic Separation Components

1. Purified CD44 monoclonal antibody (clone IM7; rat IgG2b, κ; BioLegend, San Diego, CA, USA) (see Note 4). 2. Anti-Rat IgG MicroBeads (see Note 4). 3. Prepare separation buffer containing phosphate-buffered saline (PBS), pH 7.2, 0.5% bovine serum albumin (BSA), and 2 mM EDTA (diluted from 0.5 M EDTA, pH 8.0; (see Note 5). 4. MACS Columns and MACS Separator (see Note 6).

2.3 Culture Components

1. Cell culture medium: Mesenchymal Stem Cell Growth Medium 2 (MSC-GM2) (see Note 7). 2. Tissue culture dishes/flasks for adherent cell culture (see Note 3). 3. Trypsin–EDTA (0.05% trypsin–EDTA–PBS) (see Note 8).

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Methods

3.1 Isolation and Culture of Mesenchymal Stem Cells from Human Internal Thoracic Arteries (hITA)

1. Wash obtained tissue material containing the human internal thoracic artery (hITA) by twice with 5 mL PBS (with P/S) (see Note 9). 2. Place specimens of hITA under a dissection microscope and remove contaminating fatty and muscle tissue. 3. Wash resulting vessels again (2–3 times) with 5 mL PBS (with P/S). 4. Cut vessels/vessel fragments into small (0.25 cm) long pieces (see Note 10). 5. Transfer vessel pieces into 5 mL (precooled) Opti-MEM I medium containing 0.2% type 2 collagenase and place the tube for 30 min (maximum) at 37  C using a water bath. During that time (starting 10 min after placing the tube at 37  C), vessels were mechanically minced and dissociated by up-and-down pipetting (2–4 times each) every 10 min using a 5 mL pipette (see Note 11). 6. On dissociation, cells are washed twice in PBS containing 5% FCS (centrifuge cell suspension at 300–400  g, 10 min, 4  C). 7. Cellular suspensions were passed through 70 μm cell strainer and count the cells (see Note 12). 8. Centrifuge cell suspension at 300–400  g, 10 min, 4  C. 9. Resuspend the cell pellet MSC-GM2 medium on plastic culture dishes (passage 0, P0) for expansion in a humidified atmosphere using a standard cell incubator at 37  C and 5% CO2 (see

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Note 3). Allow MSCs to adhere for 24 h. Wash 1 with PBS, replace with fresh MSC-GM2 medium and allow the cells to expand prior to immunomagnetic separation (see Note 13). For direct immunomagnetic separation, keep the cell pellet (counted cells) obtained from the step before (step 7) shortly on ice and directly proceed with the following improved isolation of VW-MSCs (Subheading 3.2). 3.2 Improved Isolation of VW-MSCs (Highly Pure Cell Cultures) Using Immunomagnetic Separation

The isolation is performed according to the manufacturer’s instructions (MACS Cell Separation Miltenyi Biotec GmbH). 1. Resuspend cell pellet obtained from step 7 (Subheading 3.1) in 100 μL (volume minimum) of separation buffer per 107 total cells. 2. Add 10 μL of rat anti-human CD44 antibody (BioLegend) per 107 total cells (see Note 14). 3. Mix once by up-and-down pipetting and incubate for 15 min on ice (see Note 15). 4. Wash cells by adding 5 mL of separation buffer (per 107 cells), centrifuge at 300–400  g for 10 min and aspirate supernatant. 5. Resuspend cell pellet in 90 μL of separation buffer (per 107 total cells). 6. Add 10 μL of anti-rat MicroBeads per 107 total cells. 7. Mix once by up-and-down pipetting and incubate for 15 min on ice (see Note 15). 8. Wash cells by adding 5 mL of separation buffer, centrifuge at 300–400  g for 10 min and aspirate supernatant. 9. Resuspend cell pellet in 500 μL of separation buffer and proceed with the immunomagnetic separation. 10. Place MS column in the magnetic field of the MiniMACS (magnet). 11. Wash the column by rinsing with 2–3 mL of separation buffer. 12. Apply the labeled cell suspension onto the column and collect the flow-through (containing unlabeled cells) (see Note 16). 13. Wash column with 2–3 2 mL of separation buffer. Discard the flow through (unlabeled cells) if not otherwise needed (e.g., isolation of endothelial cells). 14. Remove MS column from the magnet and place it on a 15 mL cell culture tube (sterile). 15. Pipette 5 mL of separation buffer onto the column. Collect the CD44-selected VW-MSCs by gravity flow (first fraction). 16. Place MS column on a new 15 mL cell culture tube (sterile).

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17. Pipette additional 5 mL of separation buffer onto the column. Immediately flush out the residual magnetically labeled cells by firmly pushing the plunger into the column (second fraction). 18. Count the cells (fraction 1 and 2 independently) and centrifuge cell suspensions at 300–400  g, 10 min, 4  C (see Note 17). 19. Resuspend the cell pellet MSC-GM2 medium on plastic culture dishes (P0) for expansion in a humidified atmosphere using a standard cell incubator at 37  C and 5% CO2 (see Note 3). Allow MSCs to adhere for 24 h. Wash 1 with PBS, replace with fresh MSC-GM medium, and allow the cells to expand (see Note 18). 3.3 Culture of VW-MSCs

1. Remove the medium from the well (usually P0 starts at from a 24-well, until 85–95% confluency) and wash 1 with PBS. 2. Trypsinize cells with 0.5 mL of 0.05% trypsin–EDTA–PBS solution for 1–3 min, resuspend cells in 10 mL of MSC medium and centrifuge cell suspension at 300–400  g at room temperature (see Note 8). 3. Replate cells at the desired density in MSC-GM2 (see Note 7). 4. For immunophenotypical characterization (MSC marker expression via flow cytometry) the protocol described by Faca et al. can be used [36]. 5. In order to access functional characterization (e.g., clonogenicity and multipotency) of isolated CD44(+) VW-MSCs, singlecell-derived clones were established by limited dilution in a 96-well plate [20, 26, 37]. VW-MSCs differentiation into adipogenic, osteogenic o and chondrogenic lineages can be performed according to Ciuffreda et al. [38] or using the ready-touse differentiation media (Mesenchymal Stem Cell Adipogenic/Chondrogenic/Osteogenic Differentiation Medium) [20, 26, 37].

4

Notes 1. Collagenase, Type 2 (CLS-2, activity: 125 units per mg dry weight; contains higher relative levels of protease activity, particularly clostripain). After dissolving the required amount of CLS-2 in Opti-MEM I medium, the solution is filtered through a 0.22-μm membrane and stored on ice prior to use. 2. Alternatively, Opti-MEM™ I medium supplemented with 5% FCS can be used. 3. Best performance was obtained using tissue culture dishes/ flasks from Greiner Bio-One (Kremsmu¨nster, Austria): 24-well/ surface area: 1.9 cm2, seeding density: 0.05  106;

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6-well/9.5 cm2, 0.3  106; T-25/25 cm2, 0.7  106; T-75/ 75 cm2, 2.1  106). 4. Alternatively, the anti-human monoclonal CD44 Antibody (clone 8E2F3; mouse IgG1; antibodies-online Inc., Limerick, PA, USA) in combination with Anti-Mouse IgG MicroBeads can be used. 5. As recommended by the manufacturer this solution should be degassed before use, as air bubbles could block the column. 6. MS columns for a maximum selection of 107 labeled cells out of 2  108 total cells were used in combination with the MiniMACS separator. 7. Best results were obtained using the MSC-GM2 medium. After the first 2–3 passages the medium can also be replaced by the MSC medium: Dulbecco’s Modified Eagle Medium, high glucose (DMEM with 4.5 g/L glucose), supplemented with sterile-filtered, heat-inactivated 20% fetal calf serum (FCS), 1% penicillin–streptomycin (P/S) (without significant loss of MSC characteristics). Usually experiments were performed with isolated cells in the early passages (until P2-6) to ensure primary cell characteristics. 8. Alternatively, detachment of VW-MSCs from the tissue culture plasticware can be achieved by incubating PBS-washed cells 2–3 min in Accutase® solution (in Dulbecco’s PBS without calcium or magnesium salts). This procedure turned out to be less stressful for the cells. 9. Vessel fragments of hITA, the radial artery, or the saphenous vein can be used. Vessel fragment are usually 2–5 cm long. 10. Optional: “open” obtained vessel longitudinally using a scissors prior to cutting. This may improve the mechanical cell loosening with a 5 mL pipette during collagenase treatment (see Note 11). 11. Prolonged incubation might result in smooth muscle cells contamination, as elastic membranes might be digested too. A longitudinal opening of the vessel fragments prior to collagenase digestion is usually not necessary as the VW-MScs will be yielded from the adventitia. However, longitudinal cutting will improve the yield of endothelial cells if this isolation in parallel is desired. 12. Red blood cells can be removed prior to culturing or further purification using the ammonium chloride–based BD Pharm Lyse™ lysing solution (10 stock solution diluted 1/10 in aqua ad iniectabilia). Therefore, the cell pellet is resuspended in 1 mL BD Pharm Lyse™ lysing solution (1), incubated for 1 min at room temperature, the tube is filled with PBS

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containing 5% FCS and centrifuged again (300–400  g, 10 min, 4  C). 13. Depending on the obtained (sparse) starting material, which can sometimes be quite small (less than 1 cm of vessel length), an in vitro expansion prior to immunomagnetic separation might be useful to have enough donor material. However, based on the selective adherence on plastic and the MSC medium selection, expanded cells can already have a high purity 80–95% as analyzed by expression of a marker panel including Stro1, CD105, CD73, CD44, CD90 and CD29 via flow cytometry. However, for a direct downstream analysis (e.g., RNA and/or DNA isolation) directly isolated highly pure VW-MSCs without culturing might be desired. In that case, directly proceed to the immunomagnetic separation and collect purified cells for the direct analysis. 14. The suggested use of this antibody is 0.5 μg per million cells in 100 μL volume. Depending on the different antibody charges/lots and the MSCs used, it would be worth that the antibody be titrated for optimal performance for each application. 15. Alternatively incubate for 15 min in the dark in the refrigerator (2–8  C). 16. Although the column will not dry up, passing the liquids (cell suspension and washing) should be performed in a timely manner. 17. In general, both fractions can be combined. However, unlabeled cells passing through the column might be trapped due to a simple filtration effect. In addition, pushing the plunger into the column is a quite stressful procedure for the retained cells, although it will increase the yield. For optimal cell isolation the protocol here recommends to plate both fractions independently with a focus on the first fraction for further culture expansions. 18. For a direct downstream analysis (e.g., RNA and/or DNA isolation for microarrays) directly isolated highly pure VW-MSCs without culturing might be desired. In that case, pellet purified VW-MSCs for further downstream processing. One aliquot of the cell suspension should be used to confirm the purity of isolated cells (expression of a MSC marker panel including STRO1, CD105, CD73, CD44, CD90, and CD29 in the absence of CD31, CD34, CD45, as well as CD68, CD11b, CD19 expressing cells via flow cytometry (see Note 13).

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Acknowledgments The respective work was supported by grants of the DFG (GRK1739/2), the BMBF (ZISS 02NUK024-D, ZISStrans 02NUK047D), the Brigitte und Dr. Konstanze Wegener-Stiftung, and the Ju¨rgen Manchot-Stiftung (Du¨sseldorf, Germany). References 1. Le Blanc K, Davies LC (2018) MSCs-cells with many sides. Cytotherapy 20(3):273–278. https://doi.org/10.1016/j.jcyt.2018.01.009 2. Le Blanc K, Frassoni F, Ball L, Locatelli F, Roelofs H, Lewis I, Lanino E, Sundberg B, Bernardo ME, Remberger M, Dini G, Egeler RM, Bacigalupo A, Fibbe W, Ringden O, Developmental Committee of the European Group for B, Marrow T (2008) Mesenchymal stem cells for treatment of steroid-resistant, severe, acute graft-versus-host disease: a phase II study. Lancet 371(9624):1579–1586. https://doi.org/10.1016/S0140-6736(08) 60690-X 3. Dominici M, Le Blanc K, Mueller I, SlaperCortenbach I, Marini F, Krause D, Deans R, Keating A, Prockop D, Horwitz E (2006) Minimal criteria for defining multipotent mesenchymal stromal cells. The International Society for Cellular Therapy position statement. Cytotherapy 8(4):315–317. https:// doi.org/10.1080/14653240600855905 4. Fernandez Vallone VB, Romaniuk MA, Choi H, Labovsky V, Otaegui J, Chasseing NA (2013) Mesenchymal stem cells and their use in therapy: what has been achieved? Differentiation 85(1–2):1–10. https://doi.org/10. 1016/j.diff.2012.08.004 5. Wang S, Qu X, Zhao RC (2012) Clinical applications of mesenchymal stem cells. J Hematol Oncol 5:19. https://doi.org/10.1186/17568722-5-19 6. Mariani E, Facchini A (2012) Clinical applications and biosafety of human adult mesenchymal stem cells. Curr Pharm Des 18 (13):1821–1845 7. Otto WR, Wright NA (2011) Mesenchymal stem cells: from experiment to clinic. Fibrogenesis Tissue Repair 4:20. https://doi.org/10. 1186/1755-1536-4-20 8. Sharma RR, Pollock K, Hubel A, McKenna D (2014) Mesenchymal stem or stromal cells: a review of clinical applications and manufacturing practices. Transfusion 54 (5):1418–1437. https://doi.org/10.1111/ trf.12421

9. Brown C, McKee C, Bakshi S, Walker K, Hakman E, Halassy S, Svinarich D, Dodds R, Govind CK, Chaudhry GR (2019) Mesenchymal stem cells: cell therapy and regeneration potential. J Tissue Eng Regen Med 13 (9):1738–1755. https://doi.org/10.1002/ term.2914 10. Naji A, Eitoku M, Favier B, Deschaseaux F, Rouas-Freiss N, Suganuma N (2019) Biological functions of mesenchymal stem cells and clinical implications. Cell Mol Life Sci 76(17):3323–3348. https://doi.org/10. 1007/s00018-019-03125-1 11. Andrzejewska A, Lukomska B, Janowski M (2019) Concise review: mesenchymal stem cells: from roots to boost. Stem Cells 37 (7):855–864. https://doi.org/10.1002/ stem.3016 12. Conese M, Carbone A, Castellani S, Di Gioia S (2013) Paracrine effects and heterogeneity of marrow-derived stem/progenitor cells: relevance for the treatment of respiratory diseases. Cells Tissues Organs 197(6):445–473. https://doi.org/10.1159/000348831 13. De Becker A, Riet IV (2016) Homing and migration of mesenchymal stromal cells: how to improve the efficacy of cell therapy? World J Stem Cells 8(3):73–87. https://doi.org/10. 4252/wjsc.v8.i3.73 14. Leibacher J, Henschler R (2016) Biodistribution, migration and homing of systemically applied mesenchymal stem/stromal cells. Stem Cell Res Ther 7:7. https://doi.org/10. 1186/s13287-015-0271-2 15. Jin HJ, Bae YK, Kim M, Kwon SJ, Jeon HB, Choi SJ, Kim SW, Yang YS, Oh W, Chang JW (2013) Comparative analysis of human mesenchymal stem cells from bone marrow, adipose tissue, and umbilical cord blood as sources of cell therapy. Int J Mol Sci 14(9):17986–18001. https://doi.org/10.3390/ijms140917986 16. Kern S, Eichler H, Stoeve J, Kluter H, Bieback K (2006) Comparative analysis of mesenchymal stem cells from bone marrow, umbilical cord blood, or adipose tissue. Stem Cells 24 (5):1294–1301. https://doi.org/10.1634/ste mcells.2005-0342

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17. Zhu Y, Yang Y, Zhang Y, Hao G, Liu T, Wang L, Yang T, Wang Q, Zhang G, Wei J, Li Y (2014) Placental mesenchymal stem cells of fetal and maternal origins demonstrate different therapeutic potentials. Stem Cell Res Ther 5(2):48. https://doi.org/10.1186/scrt436 18. Gotherstrom C, West A, Liden J, Uzunel M, Lahesmaa R, Le Blanc K (2005) Difference in gene expression between human fetal liver and adult bone marrow mesenchymal stem cells. Haematologica 90(8):1017–1026 19. Ergun S, Tilki D, Klein D (2011) Vascular wall as a reservoir for different types of stem and progenitor cells. Antioxid Redox Signal 15 (4):981–995. https://doi.org/10.1089/ars. 2010.3507 20. Klein D, Weisshardt P, Kleff V, Jastrow H, Jakob HG, Ergun S (2011) Vascular wallresident CD44+ multipotent stem cells give rise to pericytes and smooth muscle cells and contribute to new vessel maturation. PLoS One 6(5):e20540. https://doi.org/10.1371/ journal.pone.0020540 21. Klein D (2016) Vascular Wall-resident multipotent stem cells of Mesenchymal nature within the process of vascular remodeling: cellular basis, clinical relevance, and implications for stem cell therapy. Stem Cells Int 2016:1905846. https://doi.org/10.1155/ 2016/1905846 22. Klein D, Hohn HP, Kleff V, Tilki D, Ergun S (2010) Vascular wall-resident stem cells. Histol Histopathol 25(5):681–689. https://doi.org/ 10.14670/HH-25.681 23. Worsdorfer P, Mekala SR, Bauer J, Edenhofer F, Kuerten S, Ergun S (2017) The vascular adventitia: an endogenous, omnipresent source of stem cells in the body. Pharmacol Ther 171:13–29. https://doi.org/10.1016/j. pharmthera.2016.07.017 24. Klein D, Schmetter A, Imsak R, Wirsdorfer F, Unger K, Jastrow H, Stuschke M, Jendrossek V (2016) Therapy with multipotent Mesenchymal stromal cells protects lungs from radiation-induced injury and reduces the risk of lung metastasis. Antioxid Redox Signal 24 (2):53–69. https://doi.org/10.1089/ars. 2014.6183 25. Klein D, Steens J, Wiesemann A, Schulz F, Kaschani F, Rock K, Yamaguchi M, Wirsdorfer F, Kaiser M, Fischer JW, Stuschke M, Jendrossek V (2017) Mesenchymal stem cell therapy protects lungs from radiation-induced endothelial cell loss by restoring superoxide dismutase 1 expression. Antioxid Redox Signal 26(11):563–582. https://doi.org/10.1089/ars.2016.6748

26. Steens J, Zuk M, Benchellal M, Bornemann L, Teichweyde N, Hess J, Unger K, Gorgens A, Klump H, Klein D (2017) In vitro generation of Vascular Wall-resident multipotent stem cells of Mesenchymal nature from murine induced pluripotent stem cells. Stem Cell Reports 8(4):919–932. https://doi.org/10. 1016/j.stemcr.2017.03.001 27. Wegmeyer H, Broske AM, Leddin M, Kuentzer K, Nisslbeck AK, Hupfeld J, Wiechmann K, Kuhlen J, von Schwerin C, Stein C, Knothe S, Funk J, Huss R, Neubauer M (2013) Mesenchymal stromal cell characteristics vary depending on their origin. Stem Cells Dev 22(19):2606–2618. https://doi. org/10.1089/scd.2013.0016 28. Prasanna SJ, Gopalakrishnan D, Shankar SR, Vasandan AB (2010) Pro-inflammatory cytokines, IFNgamma and TNFalpha, influence immune properties of human bone marrow and Wharton jelly mesenchymal stem cells differentially. PLoS One 5(2):e9016. https://doi. org/10.1371/journal.pone.0009016 29. Ribeiro A, Laranjeira P, Mendes S, Velada I, Leite C, Andrade P, Santos F, Henriques A, Graos M, Cardoso CM, Martinho A, Pais M, da Silva CL, Cabral J, Trindade H, Paiva A (2013) Mesenchymal stem cells from umbilical cord matrix, adipose tissue and bone marrow exhibit different capability to suppress peripheral blood B, natural killer and T cells. Stem Cell Res Ther 4(5):125. https://doi.org/10. 1186/scrt336 30. Zhang ZY, Teoh SH, Chong MS, Schantz JT, Fisk NM, Choolani MA, Chan J (2009) Superior osteogenic capacity for bone tissue engineering of fetal compared with perinatal and adult mesenchymal stem cells. Stem Cells 27 (1):126–137. https://doi.org/10.1634/ste mcells.2008-0456 31. Montesinos JJ, Flores-Figueroa E, CastilloMedina S, Flores-Guzman P, HernandezEstevez E, Fajardo-Orduna G, Orozco S, Mayani H (2009) Human mesenchymal stromal cells from adult and neonatal sources: comparative analysis of their morphology, immunophenotype, differentiation patterns and neural protein expression. Cytotherapy 11 (2):163–176. https://doi.org/10.1080/ 14653240802582075 32. Wang X, Kimbrel EA, Ijichi K, Paul D, Lazorchak AS, Chu J, Kouris NA, Yavanian GJ, Lu SJ, Pachter JS, Crocker SJ, Lanza R, Xu RH (2014) Human ESC-derived MSCs outperform bone marrow MSCs in the treatment of an EAE model of multiple sclerosis. Stem Cell Reports 3(1):115–130. https://doi. org/10.1016/j.stemcr.2014.04.020

MSC Isolation from Human Vessels 33. Renna NF, de Las HN, Miatello RM (2013) Pathophysiology of vascular remodeling in hypertension. Int J Hypertens 2013:808353. https://doi.org/10.1155/2013/808353 34. Korshunov VA, Schwartz SM, Berk BC (2007) Vascular remodeling: hemodynamic and biochemical mechanisms underlying Glagov’s phenomenon. Arterioscler Thromb Vasc Biol 27(8):1722–1728. https://doi.org/10.1161/ ATVBAHA.106.129254 35. Gibbons GH, Dzau VJ (1994) The emerging concept of vascular remodeling. N Engl J Med 330(20):1431–1438. https://doi.org/10. 1056/NEJM199405193302008 36. Faca VM, Orellana MD, Greene LJ, Covas DT (2016) Proteomic analysis of mesenchymal

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Chapter 7 In Vitro Generation of Vascular Wall–Typical Mesenchymal Stem Cells (VW-MSC) from Murine Induced Pluripotent Stem Cells Through VW-MSC–Specific Gene Transfer Jennifer Steens, Hannes Klump, and Diana Klein Abstract Among the adult stem cells, multipotent mesenchymal stem cells (MSCs) turned out to be a promising option for cell-based therapies for the treatment of various diseases including autoimmune and cardiovascular disorders. MSCs bear a high proliferation and differentiation capability and exert immunomodulatory functions while being still clinically safe. As tissue-resident stem cells, MSCs can be isolated from various tissue including peripheral or umbilical cord blood, placenta, blood, fetal liver, lung, adipose tissue, and blood vessels, although the most commonly used source for MSCs is the bone marrow. However, the proportion of MSCs in primary isolates from adult tissue biopsies is rather low, and therefore MSCs must be intensively expanded in vitro before the MSCs find particular use in therapies that may require extensive and repetitive cell replacement. Therefore, more easily accessible sources of MSCs are needed. Here, we present a detailed protocol to generate tissue-typical MSCs by direct linage conversion using transcription factors defining target MSC identity from murine induced pluripotent stem cells (iPSCs). Key words Mesenchymal stem cells, Vascular wall, Lineage conversion, In vitro generation, HOX, Differentiation, iPSC

1

Introduction Among the tissue-resident stem cells, mesenchymal stem cells (MSCs), also termed mesenchymal stromal cells, comprise a heterogeneous pool of multipotent stem cells that regulate tissue homeostasis by their ability to secrete cytokines locally and/or to replace affected cells and thus support repair and healing processes of affected tissues [1–3]. These unique properties have promoted numerous applications of MSCs in clinical trials to treat a wide range of diseases [4–9]. For a therapeutically application, exogenous MSCs, either ex vivo isolated or in vitro generated and expanded, could foster tissue regeneration either (1) directly by homing in on particular anatomical sites after transplantation and differentiating into

Chrissa Kioussi (ed.), Stem Cells and Tissue Repair: Methods and Protocols, Methods in Molecular Biology, vol. 2155, https://doi.org/10.1007/978-1-0716-0655-1_7, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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specific cell types to locally replace the damaged tissue or (2) indirectly through a “hit and run” mechanism, whereby MSCs are temporary localized to the targeted tissue, stimulating recovery of injured cells and limiting inflammation by secretion of bioactive molecules [10–14]. Every organ seems to contain MSCs. Therefore, it was suggested that MSCs were distributed through the postnatal organism via the vascular system [15–17]. The vascular wall serves here as niche for resident MSCs (VW-MSCs) among other residing stem and progenitor cells. And in terms of manufacturing exogenous MSCs with superior repair capabilities, VW-MSCs itself might be logically more efficient when therapeutically applied than MSCs derived from other tissues for the treatment of diseases that are characterized by vascular damage [14, 18–20]. However, variations of the quality of obtained MSCs and their respective tissue sources, as well as subsequent cell culture have caused numerous inconsistencies in the reported in vivo effectiveness of MSCs [21–24]. Replicative senescence of ex vivo isolated and in vitro expanded MSCs could lead to a decline of MSCs’ plasticity and in vivo potency over time [25–29]. In addition, these MSCs might have accumulated many DNA abnormalities during a lifetime [30, 31]. To circumvent these potential drawbacks, MSCs can be alternatively generated in vitro from rejuvenated induced pluripotent stem cells (iPSCs) [21, 32, 33]. Several studies have reported the successful generation of MSCs from iPSCs classically using media that contains a high serum concentration or MSC-typical growth factors (such as basic fibroblast growth factor) after dissociation of embryoid bodies (EB) [34–36]. However, a direct forward programming approach of iPSCs using target cell-specific and in particular MSC-specific transcription factors was hampered by the lack of identity of those cell type (MSC)-specific factors. In general, tissue-resident stem cells differentiate mainly into tissue types that are typical for the tissue which they were derived from, which implicates that there must be a certain code or priming which is determined by tissue of origin [13, 14, 34]. Analyzing the molecular repertoire of VW-MSCs, certain homeodomaincontaining master regulators (HOX-gene family members) came into focus that are selectively expressed in VW-MSCs [37]. A comparison of the expression patterns of the 39 human HOX genes in these VW-MSCs with other terminally differentiated vascular cells (endothelial cells and smooth muscle cells) and undifferentiated embryonic stem cells revealed that the HOX genes HOXB7, HOXC6, and HOXC8 (the “VW-MSC-specific HOX code”) were differentially regulated and exclusively upregulated in VW-MSCs [37]. This HOX code was used then to directly program mouse iPSCs toward vascular wall–typical multipotent stem cells of mesenchymal nature by ectopic lentiviral expression of the VW-MSC specific HOX-code encompassing HOXB7, HOXC6,

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Fig. 1 Scheme of the lentiviral SIN vector coexpressing the coding sequences of HOXB7, HOXC6, and HOXC8 separated by 2A esterase elements together with the gene encoding the reporter Turquoise2 (cyan) fluorescent protein [20]. If silencing of the introduced transgenes (HOX code) is a problem, the lentiviral vector containing the sequence of a CBX element can be used (see Note 15)

and HOXC8 [20]. In brief, a lentiviral vector expressing the so-called Yamanaka factors was first used to reprogrammed tail dermal fibroblasts from transgenic mice containing the GFP-gene integrated into the nestin-locus (NEST iPSCs) to facilitate lineage tracing after subsequent MSC differentiation. A lentiviral vector expressing the VW-MSC-specific HOX-code then induced MSC differentiation (Fig. 1). This direct programming approach successfully mediated the generation of VW-typical MSCs with classical MSC characteristics, both in vitro and in vivo [20]. Here we provide now the detailed protocol (Fig. 2).

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Materials All solutions are prepared freshly before use and maintained at indicated temperatures. Ultrapure (sterile) water (Millipore; Burlington, MA, USA) should be used for preparation of solutions.

2.1 Tail-Tip Fibroblast Isolation

1. Mice: 8-week-old Nestin-GFP (NEST-GFP) transgenic mice [38, 39]. 2. Cell culture phosphate-buffered saline (cell culture 1 PBS pH 7.2, sterile), supplemented with 100 U penicillin/100 μg streptomycin (P/S). 3. Fibroblast medium (nTTF medium): Dulbecco’s modified Eagle’s medium (DMEM, high glucose) with 10% fetal calf serum (FCS), 1% sodium pyruvate, 1% (1) penicillin–streptomycin (Pen/Strep). 4. Tissue culture dishes/flasks for adherent cell culture (see Note 1). 5. 0.1% gelatin dissolved in 1 PBS.

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Fig. 2 Experimental design for the induction of vascular wall–typical mesenchymal stem cells (VW-MSCs) from murine induced pluripotent stem cells (iPSCs). In order to facilitate lineage tracing of induced MSCs, primary dermal tail tip fibroblasts derived from a transgenic mouse in which the gene encoding GFP is expressed under the regulatory control of the nestin promoter (NEST-GFP) are reprogrammed. Transduction of fibroblasts is performed with a lentiviral vector coexpressing the four Yamanaka factor genes (OCT4, KLF4, SOX2, and MYC) together with the coding sequence of the red fluorescent tdTomato protein. For quality and potency characterization of the generated bona-fide iPSCs, isolated cells that will have silenced the reprogramming vector (dtTomato) and express SSEA1+ will be sorted by flow cytometric and subsequently expanded to different NEST iPSC clones, and finally be established as independent cell lines. Quality testing will include flow cytometric analysis of SSEA1 expression, alkaline phosphatase staining, and immunofluorescence staining of pluripotency-associated markers (SSEA1, SOX2, OCT4, NANOG). As the most stringent test for pluripotency of iPSCs, teratoma assays will be performed. Obtained feeder-free cultured NEST iPSCs are then transduced with a lentiviral SIN vector coexpressing the coding sequences of HOXB7, HOXC6, and HOXC8, and Turquoise2 (Cyan), which are all co-translationally separated by 2A esterase moieties [20]. Two to four days after transduction the cells will be sorted for cyan fluorescence and cultured in MSC medium. Generated MSCs can be characterized 14 days after MSC induction when cells were sufficiently expanded 2.2 Reprogramming of Mouse NEST-GFP Tail-Tip Fibroblasts

1. Gelatin-coated cell culture plates. 2. Trypsin–EDTA (0.05% trypsin–EDTA–PBS) (see Note 2). 3. Feeder layer cells: mouse embryonic fibroblasts [CF-1; ATCC® SCRC-1040™; American Type Culture Collection (ATCC)]. 4. Lentiviral supernatant (lentiviral vector coexpressing the four Yamanaka factor genes and the red-fluorescent reporter gene tdTomato (see Note 3). 5. Protamine sulfate 4 μg/mL.

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6. Mouse iPSC medium: KnockOut™ DMEM (KO-DMEM), 15% FCS (sterile filtered), LIF 10 ng/mL, 1 P/S, 1% Lglutamine, monothioglycerol (MTG) 1.5  104 M. 7. 70 μm cell strainer, sterile. 2.3 Virus Stock Production (HOX Code) and Titration

1. Virus production with HEK293T/17 cells (CRL-11268). 2. HEK293T medium: DMEM high glucose (4.5 g/L glucose), 10% FBS, 1 mM sodium pyruvate. 3. Titration with HT1080 cells (CCL-121, ATCC). 4. HT1080 medium: DMEM high glucose, 10% FCS, 1 mM sodium pyruvate, 1% MEM NEAA (nonessential amino acids), 2 mM (1%) L-glutamine. 5. DMEM–10% FCS. 6. Chloroquine diphosphate, 25 μM. 7. Trypsin–EDTA 1: dilution 1/10 with PBS. 8. DMEM w/o FCS (for transfection). 9. Polyethylenimine (PEI) solution: 2 mg/mL PEI, stored at 4  C. 10. HEPES buffer: HEPES (1 M) pH 7.3, prepared in water (filtered sterile and stored at room temperature). 11. HEPES medium: DMEM (4.5 g/L glucose), 10% FCS, 1% sodium pyruvate, 1% P/S, 20 mM HEPES. 12. Protamine sulfate–containing medium: DMEM (4.5 g/L glucose) supplemented with 10% FCS and 4 μg/mL protamine sulfate. 13. Flow cytometry (FACS) buffer: PBS containing 10% FCS.

2.4 Viral Transduction (HOX Code) of Mouse NEST iPSCs

1. MSC medium: DMEM with 4.5 g/L glucose, 20% (filtered) FCS, 1% P/S or PAN-MSC medium. 2. Mouse iPSC medium: KnockOut™ DMEM (KO-DMEM), 15% FCS (sterile filtered), LIF 10 ng/mL, 1 P/S, 1% Lglutamine, monothioglycerol (MTG) 1.5  104 M. 3. Gelatin-coated cell culture plates (see Note 4).

3

Methods

3.1 Tail-Tip Fibroblast Isolation

1. Incubate desired plates (6-well plates) with gelatin solution (2 mL per well) for 45 min at room temperature. 2. Isolate 1 cm length of tail-tip from 8-week-old Nestin-GFP (NEST-GFP) transgenic donor mice 3. Wash 1 with 2–3 mL with ethanol (70%) and 2 with 2–3 mL PBS containing P/S.

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3. Remove the superficial dermis (with a scalpel and pincette) and cut the remaining tissue into 1-mm pieces using a scalpel. 4. Remove gelatin-solution from the plastic culture dishes. 5. Transfer five to six pieces into a gelatin-coated six-well. Add 1 drop of fibroblast medium to each tail tip peace and ensure that the tail tip piece is still in contact with the plate (and not swimming). 6. Incubate plate in a humidified atmosphere using a standard cell incubator at 37  C and 5% CO2. 7. The next day, complete each well with 2 mL of fibroblast medium. Ensure that the tail tip piece stays in contact with the plate. 8. Culture the tissue pieces/cells for 5–7 days; perform regularly a medium change (every 2 days) (see Note 5). 9. Optional: trypsinize cells with 0.5 mL of 0.05% trypsin– EDTA–PBS solution fo1 1–3 min, resuspend cells in 10 mL of fibroblast medium, centrifuge cell suspension at 300–400  g at room temperature and replate fibroblasts in fibroblast medium (passage 1) (see Note 5). 3.2 Reprogramming of Mouse NEST-GFP Tail-Tip Fibroblasts

1. Wash fibroblasts with 2 mL 1 PBS. 2. Trypsinize primary fibroblasts by incubating for 2–3 min with 2 mL prewarmed (37  C) trypsin–EDTA solution, harvest the cells in 5 mL DMEM–10% FCS. 3. Centrifuge cell suspension at 300–400  g, 5 min, room temperature, resuspend the pellet in 5 mL and count the cells. 4. Adjust primary fibroblasts to a density of 1  105 cells in 150 μL fibroblast medium. 5. Add 4 μg/mL protamine sulfate and viral supernatant of a multiplicity of infection (MOI) of 10 to the cell suspension. 6. Replate cells at a density of 4000 cells per cm2 in a gelatincoated 6-well plate. 7. Perform a medium change after 24 h. 8. Prepare the cells for flow cytometry sorting (FACS) (48 h after transduction): harvest cells by adding 1 mL 1 trypsin–EDTA, incubate for 5 min at 37  C, add 1 mL DMEM/FCS medium to the cells, centrifuge, and resuspend respective pellet in 500 μL medium. Transfer cells by filtering through a 70 μm cell strainer into an FACS tube. 9. Sort red-fluorescent cells and replate sorted cells at a density of 1500 cells/cm2 on gamma-irradiated MEFs in 6-well plates using 2 mL in mouse iPSC medium per well (see Note 6).

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10. Two weeks after transduction, colonies with an iPSC-like morphology can be picked and transferred to a 15 mL tube. 11. Dissociate picked colonies by a short (2 min) trypsin–EDTA incubation followed by adding 5 mL iPSC medium. 12. Centrifuge and resuspend respective pellet in 500 μL medium. Add fluorescently labeled SSEA1 antibody (e.g., SSEA1-APC) in the concentration recommended by the manufacturer and incubate for 10 min on ice. Wash cells one with 5 mL PBS/FCS. 13. Transfer cells by filtering through a 70 μm cell strainer into an FACS tube and sort red-dtTomato–negative (cells lacking red fluorescence) and SSEA1 positive cells as single cells by flow cytometry. 14. Single dtTomato() and SSEA1(+) NEST iPSCs should be plated as single cells in gelatin-coated 96-well plates (without feeder layer) in mouse iPSC medium (containing LIF) and allow to grow (see Note 6). 15. For NEST iPSC expansion, wash cells with a confluency of 70–90% once with 2 mL PBS (per well of a 6-well plate). Detach cells by adding 1 mL trypsin–EDTA (per 6-well), rinse the cell layer by tilting the plate gently and then directly remove the trypsin solution. Incubate for 2 min at 37  C. Harvest the cells by applying 5 mL iPSC medium and count the cells. Replate the cells in the desired amounts on gelatin- or Vitronectin-coated cell culture plates (see Note 7). 16. Quality testing of single NEST iPSC clones shall include flow cytometric analysis of SSEA1 expression, alkaline phosphatase staining, and immunofluorescence staining of pluripotencyassociated markers (SSEA1, SOX2, OCT4, NANOG). As the most stringent test for pluripotency of iPSCs, teratoma assays shall be performed [20, 40] (see Note 8). 3.3 Virus Stock Production (HOX Code) and Virus Titration

1. Harvest HEK293T cells at 80–90% confluency with 2 mL 1 trypsin–EDTA solution, incubate the cells for 5 min at 37  C, add 2 mL medium to cells, centrifuge and resuspend pellet in 5 mL HEK293T medium. 2. Count cells (using trypan blue) and plate 5  106 cells per 10 cm2 in DMEM + 10% FBS. 3. Let cells attach at least for 4–6 h at 37  C/5% CO2 in a standard humidified cell culture incubator. 4. Thaw all plasmids on ice, reestimate the plasmid concentration and calculate the volume of plasmids needed for the transfection (see Note 9). 5. Prepare tubes (sterile) containing 500 μL 2 HBSS (2 mL reaction tube).

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6. Calculate the volume of plasmids and water ad 450 μL (total volume). 7. Mix in the 2 mL reaction tube: volumes of plasmids and water. 8. Add 50 μL of CaCl2 (final volume of 500 μL) and mix well. 9. Add Plasmid-CaCl2 mix to HBSS and mix strongly (final volume: 1 mL); incubate mixture for 20 min at room temperature. 10. Remove the medium from the 293T cells and add 10 mL of DMEM–10% FCS medium containing 25 μM chloroquine diphosphate (freshly prepared) per 10 cm2. (For 10 mL of DMEM medium, add 10 μL of 25 mM chloroquine diphosphate). 11. Add plasmid-CaCl2-HBSS mixture (1 mL per plate) dropwise to 293T cells by using a 200 μL pipette (to avoid big drops). 12. The next day (day 2): perform a medium change using DMEM supplemented with 10% FCS (Add the new medium slowly to the cells to avoid detaching of cells). 13. The next day (day 3): harvest viral supernatant (1. SN) by using a 10 mL pipette and filter the supernatant through a 45 μm filter into a 50 mL tube, add 400 μL HEPES buffer per 10 mL supernatant. Store the viral supernatant at 4  C (until the next day). 14. Refill plates containing virus-producing 293T with 10 mL fresh medium (DMEM–10% FBS) (see Note 10). 15. The next day (day 4): harvest viral supernatant (2. SN) by using a 10 mL pipette and filter the supernatant through a 45 μm filter into a 50 mL tube, add 400 μL HEPES buffer per 10 mL supernatant. Combine both supernatants (see Note 10). 16. Perform an ultracentrifugation of the viral supernatants to get concentrated supernatant at 27,000 rpm (> 55,000  g) and 4  C for 90 min. 17. During ultracentrifugation: harvest cells of mock transfection (293T cells treated with CaCl2) as well as of HOX code/ control transfection (293Tcells + plasmid mixture + CaCl2) by adding 2 mL 1 trypsin–EDTA to the cells, incubate 5 min at 37  C, stop the reaction by adding 2 mL FACS buffer and transfer cells into a 15 mL tube. Centrifuge cells at 1200 rpm (max. 800  g), resuspend the pellet in 200 μL FACS buffer. Determine via flow cytometry cyan fluorescence (successful transfection). 18. After ultracentrifugation discard the supernatant and add 400 μL HEPES medium to the pellet. Incubate over night at 4  C (without resuspension by pipetting).

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19. The next day (day 5): Actively resuspend pellet in HEPES medium and make aliquots of concentrated viral supernatant. Freeze aliquots at 80  C. 20. Virus titration: harvest HT1080 cells with 2 mL 1 trypsin– EDTA solution (incubate for 5 min at 37  C), add 2 mL medium, centrifuge and resuspend pellet in 5 mL medium. Count cells and plate 1  105 cells per well in a 24-well plate in DMEM + 10% FCS. 21. Let cells attach at least for 4–6 h at 37  C. 22. Make a dilution series of concentrated (1 μL, 0.3 μL, 0.1 μL, 0.03 μL) and unconcentrated viral supernatant (20 and 200 μL), each dilution is used for two wells (duplicates) (see Note 11). 23. Remove medium of from the HT1080-containing wells and add 500 μL protamine sulfate medium per well. Add 500 μL of the virus–DMEM mixture to each well. 24. Incubate cells overnight at 37  C. 25. After 24 h (day 2), perform a medium change with DMEM– 10% FCS. 26. After additional 24 h (day 3), trypsinize cells by adding 200 μL 1 trypsin–EDTA per well, incubate cells for 5 min at 37  C, add 300 μL FACS buffer to each well and transfer samples into FACS tubes. Determine via flow cytometry cyan fluorescence (see Note 12). 3.4 In Vitro Generation of VW-MSCs from NEST iPSCs (HOX Code Transduction)

1. For each condition/viral supernatant (control versus HOX, Fig. 1), plate 1  104 cells/cm2 in a gelatin-coated 6-well plate in iPSC medium. 2. Thaw viral supernatant at room temperature. Add concentrated viral supernatant with a multiplicity of infection (MOI) of 5 (or 100–200 μL unconcentrated viral supernatant dropwise to the cells. Shake the plate carefully. 3. The next day (after 24 h) change the medium to remove viral particles. 4. After additional 24 h (48 h after transduction) generate embryoid bodies (EB) using the hanging drop method: Detach cells by adding 1 mL trypsin–EDTA per 6 well on the PBS washed cells. Directly remove the trypsin solution and incubate for 2 min at 37  C. Harvest the cells by applying 5 mL iPSC medium and count the cells (see Note 13). 5. Plate single cell drops (30–40 μL per drop) containing 2000 cells/20 μL on the lid of a plastic dish. Invert the plate and culture the drops “hanging” for 48 h.

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6. Harvest aggregated EBs by turning the plate around and collect aggregates by rinsing the plate with PBS using a 5 mL pipette. Centrifuge collected EBs and replate in MSC medium (see Note 14). 7. Sorting of transduced cells can be performed 10–14 days after transduction (8–10 days after plating the EBs in MSc medium): harvest cells by adding 1 mL 1 trypsin–EDTA, incubate the cells for 5 min at 37  C, add 1 mL MSC medium to cells, centrifuge and resuspend pellet in 500 μL medium. Transfer cells by filtering (70 μm cell strainer) into an FACS tube. CFP and GFP-positive cells can be sorted out and replated in MSC medium (see Note 6). 8. Let the cells allow to expand in MSC medium. Generated VW-MSCs are ready now for a detailed in vitro and in vivo analysis [20, 40] (see Note 15).

4

Notes 1. Best performance is obtained using tissue culture dishes/flasks from Greiner Bio-One (Kremsmu¨nster, Austria): 24-well/ surface area: 1.9 cm2, seeding density: 0.05  106; 6-well/ 9.5 cm2, 0.3  106; T-25/25 cm2, 0.7  106; T-75/75 cm2, 2.1  106). 2. Alternatively, detachment of cells from the tissue culture plasticware can be achieved by incubating PBS-washed cells 2–3 min in Accutase® solution (in Dulbecco’s PBS without calcium or magnesium salts; Sigma, St. Louis, MO, USA). This procedure turned out to be less stressful for the cells. 3. Transduction of fibroblasts is performed with a lentiviral vector coexpressing the four Yamanaka factor genes (OCT4, KLF4, SOX2, and MYC) together with the coding sequence of the red fluorescent tdTomato protein (Takahashi and Yamanaka, 2006). Vector particle production is performed as previously described [40]. Expression plasmids encoding lentiviral gag-pol (pcDNA3.GP.CCC), HIV-rev (239_RSV_Rev) and the reprogramming cassette (pRRL.PPT.SF.OKSM.I.GFP. Pre) were transiently transfected into HEK293 cells together with pMDG_VSVG for pseudotyping [40]. Culture supernatants are collected 24 and 48 h posttransfection. The produced virus is concentrated by ultracentrifugation of the supernatants at 27,000 rpm (>55,000  g), 1.5 h, 4  C and titrated on HT1080 cells. 4. Alternatively, Vitronectin XF™ (STEMCELL Technologies; Cambridge, MA, USA) coated plates can be used. Desired plates (6-well plates) were incubated with Vitronectin solution

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(2 mL per well) for 2 min at room temperature. Plates were stored then at 4  C until use. Directly before the use the remaining coating solution is removed. 5. Fibroblasts start to migrate out 1–2 days after plating (passage 0, P0). After 5–7 days dense cell “plaques” around the tissue pieces can be observed. These cells can be trypsinized and expanded into T75 flasks (P1). However, primary fibroblasts (P0) cells will be used for the reprogramming approach. 6. ZellShield™ (an antibiotics combination ready-to-use for protecting cell cultures from a broad range of common contaminants; BioChrom, Berlin, Germany) can be added to the medium to avoid contaminations after FACS. 7. Alternatively, for the generation of iPSCs from fibroblasts, a self-inactivating lentiviral reprogramming vector, which contains two FRT sites in the ΔU3 regions within the LTRs that allow for excision of the viral vector by Flp recombinase after successful reprogramming [41, 42]. To eliminate the risk of reactivation of the integrated vector, which could lead to inhibition of differentiation [43], the delivered Flp recombinase into respective NEST iPSCs by lentiviral-mediated protein transfer to excise the reprogramming vector can be performed [40]. 8. For potency testing, expression of pluripotency markers can be used by different methods (alkaline phosphatase activity; immunofluorescence of OCT4, NANOG, SOX2, cell surface antigens SSEA4, TRA1-60, TRA1-81, and flow cytometry. The pluripotent mouse embryonic stem cells derived from the 129/Sv/Ev strain (line CCE) is recommended as a reference for pluripotency [20]. 9. Amounts of plasmids needed for the transfection. Generated NEST-GFP iPSCs will be transduced using a lentiviral SIN vector coexpressing the coding sequences of HOXB7, HOXC6, and HOXC8 separated by 2A esterase elements together with the gene encoding Turquoise2 (cyan) fluorescent protein (Fig. 1). Vector containing supernatants will be collected from HEK293 cells transfected with 5 μg of pRRL. PPT.SF.HOXB7.2A.C6L.2A.C8.2A.Turq plasmid or 5 μg of control plasmid (same vector without the HOX genes) mixed with 15 μg of wild-type Gag-Pol plasmid (MLV_SynGag/ pCDNA3.GP.CCCC), 5 μg pRSV_Rev, and 2 μg of pMDGVSVG plasmid (pMDG/pMD2G) encoding the envelope protein. Vectors are concentrated by ultracentrifugation. 10. Make some aliquots of unconcentrated viral supernatant (1–2 aliquots of 1–2 mL 1. SN) and freeze aliquots at 80  C. 11. Pipetting scheme for the virus–DMEM mixture (total volume 500 μL) (Table 1).

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Table 1 Sequential virus dilutions 1 μL

0.3 μL

0.1 μL

0.03 μL

20 μL

200 μL

5 μL viral SN concentrated

150 μL of 1 μL dilution

166 μL of 0.3 μL dilution

150 μL of 0.1 μL dilution

40 μL viral 400 μL SN viral SN

495 μL DMEM

350 μL DMEM

334 μL DMEM

350 μL DMEM

960 μL 600 μL DMEM DMEM

200 μL dilution into new tube

200 μL dilution into new tube

200 μL dilution into new tube

200 μL dilution into new tube

800 μL DMEM

800 μL DMEM

800 μL DMEM

800 μL DMEM

12. The titer calculation follows the formula: TU/mL ¼ (# of cells at transduction)  [MOI/(mL of lentiviral stock used at transduction)]. With # ¼ total number of cells in culture when viral particles were added (at transduction), MOI (multiplicity of infection) ¼ number of integrations per cell, TU ¼ transduction unit. Thus, via flow cytometry the percent of transduced cells after transduction (the transduction efficiency) is estimated. The number of cells at transduction as well as the volume of the viral stock used to transduce the cells is known. Calculate the TU via: (TU/mL) ¼ (N  P)/(V  D) with N(#) ¼ cell number per well used, P ¼ percentage of cyan-positive cells (should be around 10–20%), V ¼ virus volume used in each well, D ¼ dilution-fold. 13. Alternatively, at this stage the cells can be sorted via flow cytometry based on their cyan fluorescence. 14. Alternatively, the EBs might be dissociated prior to plating. At this stage then an FACS-analysis for cyan fluorescence can be performed in order to determine the number of transduced cells. 15. In order to prevent transgene silencing and variegation in cell lines, multipotent and pluripotent stem cells, and their differentiated progeny that bear methylation-prone chromatin environments, a minimal 0.7 kb element of A2UCOE (ubiquitous chromatin opening elements (UCOEs); a 1.5 kb sequence derived from the human HNRPA2B1-CBX3 locus) containing merely the CBX3 promoter in the context of a lentiviral vector (Fig. 2) can be used to protect linked heterologous regulatory elements from methylation, thereby conferring stable transgene expression (antisilencing effect of CBX3) [44–46].

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iPSC-Derived VW-MSCs functional MSCs and repair bone defects. Stem Cells Transl Med 5(11):1447–1460. https:// doi.org/10.5966/sctm.2015-0311 36. Chen YS, Pelekanos RA, Ellis RL, Horne R, Wolvetang EJ, Fisk NM (2012) Small molecule mesengenic induction of human induced pluripotent stem cells to generate mesenchymal stem/stromal cells. Stem Cells Transl Med 1 (2):83–95. https://doi.org/10.5966/sctm. 2011-0022 37. Klein D, Benchellal M, Kleff V, Jakob HG, Ergun S (2013) Hox genes are involved in vascular wall-resident multipotent stem cell differentiation into smooth muscle cells. Sci Rep 3:2178. https://doi.org/10.1038/srep02178 38. Yamaguchi M (2005) Analysis of neurogenesis using transgenic mice expressing GFP with nestin gene regulatory regions. Chem Senses 30(Suppl 1):i117–i118. https://doi.org/10. 1093/chemse/bjh142 39. Yamaguchi M, Saito H, Suzuki M, Mori K (2000) Visualization of neurogenesis in the central nervous system using nestin promoterGFP transgenic mice. Neuroreport 11 (9):1991–1996 40. Stanurova J, Neureiter A, Hiber M, de Oliveira KH, Stolp K, Goetzke R, Klein D, Bankfalvi A, Klump H, Steenpass L (2016) Angelman syndrome-derived neurons display late onset of paternal UBE3A silencing. Sci Rep 6:30792. https://doi.org/10.1038/ srep30792 41. Voelkel C, Galla M, Maetzig T, Warlich E, Kuehle J, Zychlinski D, Bode J, Cantz T, Schambach A, Baum C (2010) Protein transduction from retroviral gag precursors. Proc Natl Acad Sci U S A 107(17):7805–7810. https://doi.org/10.1073/pnas.0914517107 42. Warlich E, Kuehle J, Cantz T, Brugman MH, Maetzig T, Galla M, Filipczyk AA, Halle S,

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Chapter 8 Analysis of Tooth Innervation in Microfluidic Coculture Devices Pierfrancesco Pagella and Thimios A. Mitsiadis Abstract Innervation plays a key role in the development, homeostasis, and regeneration of organs and tissues. However, the mechanisms underlying these phenomena are not well understood yet. In particular, the role of innervation in tooth development and regeneration is neglected. Cocultures constitute a valuable method to investigate and manipulate the interactions between nerve fibers and teeth in a controlled and isolated environment. Microfluidic systems for allow cocultures of neurons and different cell types in their appropriate culture media, while permitting the passage of axons from one compartment to the other. Here we describe how to isolate and coculture developing trigeminal ganglia and tooth germs in a microfluidic coculture system. This protocol describes a simple and flexible way to coculture ganglia/nerves and their target tissues and to study the roles of specific molecules on such interactions in a controlled and isolated environment. Key words Developmental biology, Orofacial development, Tooth, Innervation, Trigeminal ganglion, Microfluidics, Coculture systems

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Introduction Innervation plays a key role in the development, homeostasis, and regeneration of organs and tissues. Furthermore, innervation is involved in the regulation of stem cell proliferation, mobilization, and differentiation [1–4]. In spite of the rich innervation of adult teeth, and in contrast to all other organs and tissues of the body, developing teeth start to be innervated at the earliest postnatal stages [5, 6]. Sensory nerves from the trigeminal ganglia and sympathetic nerves from the superior cervical ganglia innervate the adult teeth [5, 6]. Teeth develop as a result of sequential and reciprocal interactions between the oral ectoderm and cranial neural crest-derived mesenchyme. These interactions give rise to epithelial-derived ameloblasts and mesenchyme-derived odontoblasts that are responsible for the formation of enamel and dentin, respectively [7]. During embryogenesis, nerve fibers projecting

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from the trigeminal ganglia progressively surround the developing tooth germs but they do not contact them neither penetrate them. Nerve fibers enter the dental pulp mesenchyme at more advanced developmental stages that are associated with odontoblast differentiation and dentin matrix deposition events [8]. Dental pulp innervation is completed soon after tooth eruption in the oral cavity [6]. Different studies investigated the role of innervation in tooth development [1, 9–13]. Nevertheless, the role of innervation in tooth formation and regeneration is still highly controversial in mammals [1]. Cocultures constitute a valuable method to investigate and manipulate the interactions between nerve fibers and teeth in a controlled and isolated environment [14–17]. At the same time, coculturing involves important technical adjustments. For example, nerves and specific dental tissues (e.g., dental pulp, dental follicle, dental epithelium) often require different culture media in order to guarantee survival and physiological behavior of the tissues for long periods of time [18]. Microfluidics systems allow for cocultures of neurons and different cell types in their appropriate culture media. In these devices, dental tissues and neurons are separated in different compartments, while allowing for the growth of axons from the neural cell bodies through microchannels toward the compartment containing their target tissue [18–20]. We have recently demonstrated that trigeminal ganglia (TG) and teeth are able to survive for long periods of time when cocultured in microfluidic devices [18]. Moreover, we have demonstrated that teeth from different developmental stages maintain in these in vitro conditions the same repulsive or attractive effects on trigeminal innervation that they show in vivo [18]. This protocol provides information about a simple, powerful, and flexible way to coculture ganglia/nerves and target tissues and to study the roles of specific molecules on such interactions in a controlled and isolated environment.

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Materials

2.1 Material for Dissection

1. Dissection scissors and forceps, dissection needles (insulin needles). 2. Dissection glass dish. 3. Stereoscope.

2.2

Solutions

1. Phosphate-buffered saline (PBS): 137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 1.8 mM KH2PO4, pH 7.4. 2. Poly-D-lysine: stock 1 mg/ml, dilute to 0.1 mg/ml in distilled, sterile H2O to obtain the working solution.

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3. Laminin: stock 100 μg/ml (Sigma-Aldrich, L2020), dilute to 5 μg/ml in neurobasal medium to obtain the working solution. 4. Paraformaldehyde 4% in PBS (PFA 4%). 2.3

Culture Media

1. Tooth culture medium: DMEM-F12, 20% fetal bovine serum, 10 U/ml penicillin–streptomycin, 2 mM L-glutamine, 150 μg/ml ascorbic acid. 2. Trigeminal ganglia culture medium: Neurobasal medium, B27, 50 ng/ml recombinant nerve growth factor (β-NGF), 10 U/ ml penicillin–streptomycin, 2 mM L-glutamine.

2.4 Microfluidic Coculture Chamber Components

1. AXIS Axon Isolation Device: Millipore, AX15010-TC (microchannels of different length are available). 2. Glass coverslips 24 mm  24 mm. 3. 6-wells plate or 35 mm petri dish. 4. Biopsy punch (diameter: 1–4 mm). 5. Vacuum chamber and pump.

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Methods

3.1 Preparation of Dissection Material, Culture Media, Microfluidic Devices

1. Autoclave microdissection forceps and scissors (121  C, sterilization time: 20 min) and store them in a sterile container. 2. Sterilize glass coverslips (24 mm  24 mm) by incubating them in 1 M HCl for 24 h at 37  C. Wash them three times with sterile, distilled H2O and three times with ethanol 99%. Dry then the coverslips under sterile flow hood. Finally, autoclave the coverslips to complete the sterilization. Coverslips can be stored in ethanol 70%. 3. Remove carefully the AXIS Axon Isolation Devices from the package using sterile forceps and place them in a sterile petri dish. 4. Using a sterile biopsy punch, create one hole per sample to be cultured in correspondence of the culture chambers (Fig. 1; see Note 1). 5. Sterilize the AXIS Axon Isolation Devices by immerging them in ethanol 70%. Dry then AXIS Axon Isolation Devices and coverslips completely under a sterile flow hood. Wait a minimum of 3 h before proceeding (see Note 2). 6. Place each coverslip into a 35 mm petri dish or into a well within a 6-wells plate (see Note 3). 7. Place the AXIS Axon Isolation Device onto the coverslip and press gently but firmly with a forceps with bent ends in order to allow full adhesion between the isolation device and the glass coverslip (Fig. 1).

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Fig. 1 Schematic representation of the protocol. Left: structure and main components of the microfluidic coculture device. Right: localization of trigeminal ganglia and tooth germs in the mouse embryo head and their placement in the microfluidic coculture device

8. In each culture chamber, pipet 150 μl of poly-D-lysine (0.1 mg/ml). Place the microfluidic devices under vacuum for 5 min, in order to remove all the air from the culture chambers (see Note 4). Incubate the devices with poly-D-lysine overnight at 37  C. 9. Wash chambers three time with sterile, distilled H2O. Fill chambers with 150 μl laminin working solution (5 μg/ml) and incubate for 2 h at 37  C (see Note 5). 10. Prepare 10 ml of medium for trigeminal ganglia cultures and 10 ml for tooth organ cultures. 3.2 Mouse Embryo Generation and Dissection

Animal treatments have to be performed in accordance with the Animal Welfare Law and with the regulations of the deputed institution. 1. Determine embryonic age according to vaginal plug (vaginal plug: embryonic day of development 0.5: E0.5) and confirm it via morphological criteria. For this protocol we generally use E14.5–E17.5 mouse embryos.

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2. Clean the dissection area and the stereoscope with ethanol 70%. 3. Sacrifice the pregnant mother via cervical dislocation. Block the neck of the mouse with the first and second finger onto a grid and pull with decision the tail. 4. Dissect the skin around the lower abdomen and open the abdomen using scissors. Locate the uterus: during such late stages of pregnancy, the uterus fills the abdominal cavity and the embryos are clearly visible. 5. Dissect out the uterus and place in a tube filled with PBS on ice. When on ice, the tissue can be left for several hours. Discard the corpse of the mother according to the guidelines of your institution. 6. Dissect out the embryos from the uterus and free them from their extraembryonic tissues. Place the embryos in PBS on ice. 7. Decapitate the embryos using scissors and separate the lower jaw from the rest of the head using microdissection scissors. Remove precisely the lower jaw without damaging the trigeminal ganglia (see Note 6). Preserve the lower jaw and the rest of the head in cold PBS on ice. 8. To dissect TG, take the head and place it onto a dissection glass petri dish, previously filled with cold PBS. Using the forceps remove the skin and the skull. Remove then the telencephalon and the cerebellum by placing forceps below the telencephalon and lift. The telencephalon and the cerebellum will flip together, leaving the bottom of the skull exposed. 9. Localize the trigeminal ganglia. Use the forceps to isolate the TG. Eliminate the remnants of the trigeminal projections using the dissection needles as knives. Place the dissected TG in a petri dish filled with cold PBS and keep them on ice. 10. To dissect embryonic teeth, place the lower jaw, previously separated from the skull, onto a dissection glass petri dish filled with cold PBS. Remove using dissection needles as knives the tongue and the skin surrounding the jaw. Separate the left and the right hemijaws by cutting along the midline of the jaw. The tooth germs are easily visible. Isolate the tooth germs using dissection needles and remove the excess of nondental tissues. Place the dissected tooth germs in a petri dish filled with cold PBS and keep them on ice (Fig. 1). 3.3 Microfluidic Cocultures

1. After dissection remove laminin from the microfluidic devices. Fill the chambers with 200 μl of the respective media. 2. Transfer gently with forceps the dissected TG and tooth germs into the holes created by punching (Fig. 1, see Note 7). Culture the samples in incubator at 37  C with 5% CO2. Change the culture medium every 48 h (see Note 8).

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3. Cocultures can be easily imaged by time-lapse microscopy during the culture period by placing the culture dishes in an environmental chamber (37  C, 5% CO2). Focus on the extending neurites in order to follow the progress of innervation. Cocultures can be maintained for over 10 days. 4. After the culture period different analysis can be performed (see Note 9). In order to fix the cultures wash the chambers by pipetting 150 μl of PBS into one well per chamber and letting PBS flow through the chambers three times. Remove the PBS and fix the samples by pipetting 150 μl of PFA 4% in one well per chamber. Incubate the samples at room temperature for 15 min. Wash the chambers twice with PBS as described above.

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Notes 1. Do not perform the punch too close ( Color> Split Channels to grayscale for each specific channel (RGB) and measure the blue area. 6. Analyze the pyoktanin blue stained area surrounding the epiglottis, aryepiglottic fold, and interarytenoid fold in the axial direction.

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Notes 1. To avoid further nerve damage during SLN clipping, we conduct the operation under a stereomicroscope. 2. The surgical field is kept clean and clear after transecting the SLN. 3. The cages are changed every 3 days to avoid infection to the rats. 4. We collect SHEDs from exfoliated deciduous teeth (from individuals 6–12 years old) and adult third molars (extracted from individuals 18–30 years old). 5. The animals are evaluated with 2 min observation sessions by two observers who were not in charge of the surgeries or data acquisition. 6. Video analysis shows that some rats drink water while holding the bottle with their hands. Those data are excluded from further analysis.

References 1. Jafari S, Prince RA, Kim DY, Paydarfar D (2003) Sensory regulation of swallowing and airway protection: a role for the internal superior laryngeal nerve in humans. J Physiol 550 (Pt 1):287–304. https://doi.org/10.1113/ jphysiol.2003.039966 2. Kitagawa J, Nakagawa K, Hasegawa M, Iwakami T, Shingai T, Yamada Y, Iwata K

(2009) Facilitation of reflex swallowing from the pharynx and larynx. J Oral Sci 51 (2):167–171 3. Tsuruta T, Sakai K, Watanabe J, Katagiri W, Hibi H (2018) Dental pulp-derived stem cell conditioned medium to regenerate peripheral nerves in a novel animal model of dysphagia. PLoS One 13(12):e0208938

Peripheral Nerve Regeneration 4. Gronthos S, Mankani M, Brahim J, Robey P, Shi S (2000) Postnatal human dental pulp stem cells (DPSCs) in vitro and in vivo. Proc Natl Acad Sci U S A 97(25):13625–13630 5. Miura M, Gronthos S, Zhao M, Fisher L, Robey P, Shi S (2003) SHED: stem cells from human exfoliated deciduous teeth. Proc Natl Acad Sci U S A 100(10):5807–5812 6. Wakayama H, Hashimoto N, Matsushita Y, Matsubara K, Yamamoto N, Hasegawa Y, Ueda M, Yamamoto A (2015) Factors secreted from dental pulp stem cells show multifaceted

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benefits for treating acute lung injury in mice. Cytotherapy 17(8):1119–1129 7. Sakai K, Yamamoto A, Matsubara K, Nakamura S, Naruse M, Yamagata M, Sakamoto K, Tauchi R, Wakao N, Imagama S, Hibi H, Kadomatsu K, Ishiguro N, Ueda M (2012) Human dental pulp-derived stem cells promote locomotor recovery after complete transection of the rat spinal cord by multiple neuro-regenerative mechanisms. J Clin Invest 122(1):80–90

Chapter 10 Healing of Full-Thickness Murine Skin Wounds Containing Nanofibers Using Splints for Efficient Reepithelialization and to Avoid Contracture Nilika Bhattacharya, Arup K. Indra, and Gitali Ganguli-Indra Abstract Wound healing process is the outcome of a series of actions and combined with collaborative process involving concerted efforts of multiple cell types. The dynamic series of events constituting each of these overlapping rather than discrete stages of wound healing increases its complexity and the necessity to understand it. The contrasting mechanisms of wound healing employed by mouse (via wound contraction) and humans (via reepithelialization) puts forth the need of a model closely mimicking human woundhealing and hence comes the applicability of the mouse excisional wound splinting model. Use of siliconebased splints has demonstrated their effectiveness in aptly resembling the human reepithelialization mediated wound healing by preventing contraction during healing. The rising popularity of nanofiberbased treatments for wound healing through sustained release of factors/molecules promoting wound closure can be potentially implemented in association with this model to determine its efficacy in wound management in a more humanized way. Key words Skin, Wound healing, Mice, Biopsy, Analgesic, Nanofibers, Silicone splints, Histology, Immunohistochemistry, Protein assay

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Introduction The complex and actively changing nature of the wound healing process involves an elaborate and intricate interplay of a large number of diverse cell types present in the skin with the epidermal keratinocytes, resident dermal endothelial cells, fibroblast and immune cells participating in a range of paracrine signaling mechanisms mediated through soluble growth factors such as chemokines and cytokines. In addition, the efficient migration of hair follicular stem cells (HFSC), cytoskeletal system, cell adhesion, and extracellular matrix (ECM) molecules make it increasingly difficult to study wound healing owing to huge cohort of variables being involved in the process [1–3]. Not only this but intrinsic factors such as age, mental health status, site of the wound, and one’s

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immune background can significantly affect the outcome of a wound. Besides, extrinsic factors such as environment, hygiene, and nutritional status can cause a significant variation in the healing time from one subject to another [3]. Hence there is a dire need of a method that will allow for proper characterization of the four stages of wound healing for therapeutic intervention. Given the multifaceted role of skin in serving not only as a protective or defense barrier but also in the regulation of the body temperature (via sweat glands) or sensory perception or in vitamin D absorption (crucial for teeth and bone development) clearly explains the need for restoring its integrity following a wound or an abrasion [4]. The wound healing process in humans are characterized by a four-stage process namely hemostasis; inflammation; proliferation and remodeling. The various stages comprising the wound healing process in humans have been shown to strongly mimic the steps involved in repairing a damaged house [3]. The first stage, also referred to as hemostasis involves resealing of the damaged blood vessels very similar to the role played by the utility workers in sealing off the water and gas pipelines following damage to a house before starting upon any kind of new construction work. The utility workers in context to wound healing essentially represent platelets. Following this, the unwanted broken parts of the house needs to be cleared off before execution of any form of the mending process. The inflammation stage of the wound healing process harbors the execution of two crucial events—one being involved in removal of unwanted cell debris or broken tissues through recruitment of neutrophils (the most common form of polymorpholeukocytes (PMN)) and the other being the commencement of the repair process under the guidance of tissue resident as well as infiltrated macrophage mediated signaling [5]. The third stage is essentially the reconstruction phase wherein the fibroblast or the “framer” cells help in collagen deposition to serve as a framework for dermal regeneration to take place, thereby leading to granulation tissue formation. The other types of cells involved in this particular stage include endothelial cells which help in the formation of the new blood vessels aimed at restoring the vascular supply to the wounded region. Keratinocytes are yet another crucial player in this stage marking the reepithelialization process to form the outermost covering atop the wound followed by the contraction phase. The final stage is the “finishing touch” or the “remodeling phase” wherein the fibroblasts cells mediate the replacement of the collagen type 3 comprising the scar tissue by collagen type 1 aimed at imparting higher tensile strength and apoptosis of additional wanted cells, thereby restoring the original unwounded skin morphology and architecture [3]. Ethical considerations often limit our ability to understand various diseases employing human subjects. Over the past decade, mouse has emerged to be the most sought-after mammal to serve as

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disease model given its short life cycle, easily amendable via genetic manipulation, large litter size, and relatively noncumbersome maintenance as compared to other higher mammalian forms [6, 7]. However significant anatomical differences between the skin types in human versus mouse have raised several questions about the appropriateness of it being used as an in vivo model to study wound healing. Further studies have shown the wound healing mechanism in mouse to be significantly different from that employed in humans. Mouse wound healing occurs primarily via the contraction mechanism as opposed to humans where the healing is mediated via the reepithelialization process or “growth of new tissues” Hence translating those findings to develop therapies for human might not be justifiable [8]. Herein lies the importance of incorporating a splint around the wound, the detailed protocol of which has been discussed later in this chapter. The idea behind placing splints around the wounds is to mimic the human wound healing process in mouse. Placing splints around the wounds ensures stitching of the adjacent skin, thereby restricting its mobility. As a result, the contractive mechanism of wound healing is prevented from occurring, thereby mediating healing via angiogenesis, granulation tissue formation, and reepithelialization perfectly resembling the biological mechanism of wound healing in humans [9]. It has been shown that splinting of the wounds with silicone splints can mimic the human wound healing process without any contraction [10]. This method can indeed help translate the findings made in mice to humans to get a better understanding of the fine intricacies of this multistage process. Previous unpublished microarray studies in Indra lab have shown significant downregulation of certain factors in mice displaying delayed wound healing process. Ongoing wound healing studies aim at using nanofibers loaded with those depleted factors to analyze the efficiency of wound healing in a mice model of delayed wound healing. Nanofibers are essentially fibers made from polymers (natural or synthetic) having their diameter in the nanometer range. Their large surface area-to-volume ratio, high porosity, controlled drug release potential and appreciable mechanical strength have raised their popularity as a potential drug delivery mechanism. Electrospinning serves as the most widespread method of generating nanofibers owing to their ability to produce fibrous network models of a wide range of diameters (thin, ultra-thin) closely resembling the in-vivo extracellular matrix thus serving as a perfect scaffold in tissue engineering and wound healing studies [11]. Different studies have shown employment of different types of nanofibers in the wound healing process. Studies involving nanodiamond loaded PCL (poly-ε-caprolactone) nanofibers have shown enhanced wound healing rates by promoting proliferation of the epithelial cells as well as reduced attachment of Staphylococcus aureus species

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and thereby restriction microbial infiltration of the wounds [12]. Yet another study showed use of chitosan-based nanofibers in promoting wound healing owing to their bioactive role in mediating scar tissue formation [13]. Chitosan is also known for its role against a wide range of microorganisms subject to the interaction of the charged groups present in their backbone with the bacterial cell wall components, and hence using chitosan based nanofibers have also been shown to circumvent wound related infections [14]. Curcumin based nanofibers have been shown to have potent antioxidant and anti-inflammatory roles [15]. A study in 2018 showed a new variant of nanofiber employing 1α, 25-dihydroxyvitamin D3 with its potential role in promoting wound healing by bolstering the innate immune response [16]. All of the above studies show the importance of using electro spun generated nanofibers in wound management. However, in the absence of splints the nanofibers often fail to remain secured to their respective wounds created on either skin side, thereby limiting the use to one type of nanofiber (control or experimental) per mouse to avoid erroneous results. Use of splints can exclude the need of including additional mouse as controls, thereby reducing the animal number. Splints can help in securely fixing the nanofibers placed on the wound and hence the two wounds created on either side of the skin can serve as control and experiment respectively further reducing the experimental variability (owing to the presence of multiple subjects) as well as the associated cost incurred [9]. Depending on the mouse strain being used, the splint may detach prior to the complete wound closure owing to hair regrowth in the clean-shaven area. An important consideration while using splints is to determine the appropriate wound size for different mouse strains to ensure that the splint remains attached during the entire course of healing. Splint detachment prior to complete healing can lead to enhanced mobility of the skin, thereby promoting the inherent wound contraction mode of healing instead of reepithelialization mediated healing [9]. Lingering wounds or in other words chronic wounds affect a significant proportion of the population in the USA with the increased incidence resulting from the increasing proportion of elderly population along with the diabetic and obesity epidemic. The mouse excisional model of wound splinting along with nanofibers loaded with different factors such as growth factors, chemokines, and cytokines [17–20] can indeed serve as an appropriate model to study the various stages of wound healing, more so in the context of humans owing to their ability to resemble the human reepithelialization mediated healing mechanism.

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Materials 1. Biosafety hood. 2. Small animal anesthesia system or anesthetic ketamine– xylazine. 3. Clippers for shaving. 4. 70% ethanol, Betadine, chlorhexidine. 5. Sterile drapes and surgical tools such as fine iris scissors regular and curved, forceps, gloves, gowns, drapes, mask and cap. All items must be surgical grade. 6. Disposable biopsy punches 5 mm or size of choice as needed, wound stents 14 mm OD  7 mm ID. 7. Nanofibers loaded with drugs/compounds/growth factors, cytokines, chemokines can be inserted into the wounds and followed through different days of healing. 8. Krazy glue. 9. Sterile synthetic absorbable suture (5-0, 1800 ). 10. Tegaderm to protect the wounds. 11. Tracing paper. 12. High resolution digital camera. 13. Digital Vernier caliper. 14. Phosphate-buffered saline (PBS): 137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, and 1.76 mM KH2PO4 in ddH2O, pH 7.4 (with HCl). 15. 4% paraformaldehyde (PFA) in PBS. 16. RNAlater. 17. Image analysis software such as Photoshop or Image J. 18. Standard supplies required for paraffin embedding the biopsied tissue samples, followed by sectioning and immunohistochemistry. 19. For skin sample collection wound tissue with 2–3 mm border can be excised at different time points for histological and histopathological evaluation [21].

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3.1 Steps of Skin Wounding Using Splints and Nanofibers

1. Mice for the wound healing studies must be housed in the standard housing conditions at the satellite animal facility. 2. Before starting the experiment, the Animal Care and User Protocol (ACUP) must be approved by the Institutional Animal Care and Use Committee (see Note 1).

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3. Based on the research subject, age of the mice can be chosen. Mouse models such as mice which are aged, ulcerated, and diabetic are some examples. Mice are first anesthetized and then shaved the hairs using electric clipper on the dorsal side of the mice (see Note 2). Wipe the area with 70% ethanol and allow it to dry. 4. To reduce pain and sufferings and to avoid interference with the data analyses low doses of analgesic such as buprenorphine SR can be given. The skin must be prepared using three sets of alternating scrubs (Betadine or chlorhexidine followed by 70% alcohol). All surgeries must be performed in a clean environment, using aseptic techniques (see Note 3). 5. To generate two symmetrical full-thickness excisional wounds, the dorsal skin from the mice midline are pulled together and the resulting folded skin is punched with a 5 mm (size of the punch depends on the study needs) sterile biopsy punch to create two wounds on the back of mice (see Note 4). Using forceps remove both the circle pieces of the skin. Insert the nanofibers into the wounds using the forceps (see Note 5). Since mice wound heals by contraction splinting the wounds is a great way to mimic human wounds. 6. Splinting procedure: Once the wounds are generated, donut shaped silicone splints 14 mm OD  7 mm ID are placed on the wound and Krazy glue (quick bonding adhesive) is used to secure the splints on the wound (see Note 5). They are further secured by interrupted 6-0 nylon sutures. Once the suturing is complete a semiocclusive dressing such as Tegaderm must be used to wrap the mice. Upon completion of the procedure mice can be placed in individual cages and returned back to the designated room for recovery (Fig. 1). 7. The experimental mice must be monitored every day, 7 days a week, with no exception on holidays or weekends (see Note 6). The wound closure process must be monitored every other day. The wounds are traced on a clear transparency sheets with a sharpie marker (see Note 7) and then mice are photographed in batches for example wild type vs mutant or compound versus no compound. Using a digital Vernier caliper (see Note 8), the larger and the minor diameter of the wounds are measured. The formula for the wound area calculation and the wound closure percentage is given below. Calculation of wound area formula Diameter A=2 multiplied by diameter B=2  PieðπÞ Calculation of percentage of wound closure: fðarea of original wound  area of actual woundÞ=area of original woundg  100

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Fig. 1 Full-thickness wound healing procedure using nanofiber and silicone splints. (a) Full-thickness wound generated using 5 mm punch biopsy. (b) Inserted Nanofiber in the wound with the help of the forceps. (c) Small amount of glue was spread outside the circular wound and splints were placed the on the glued area for the splints to stick on to the skin. (d) Three sutures were done to make sure the splints remain in place. (e) Examples showing mice with inserted nanofibers and splints sutured. (f) Taking measurement using digital Vernier caliper. (g) Wound healing work station

8. Samples for histology, RNA, and protein analysis can be collected in PFA, RNAlater, and liquid nitrogen, respectively, or as required for specific studies as described [7]. 3.2

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Murine models of full thickness wounds are routinely used for testing nanofibers with various ingredients such as vitamin D3, curcumin, growth factors, combination of a grafted chitosan and an antioxidant agent are able to help fight/reduce infection, and promote wound healing process and will be beneficial for the patients with chronic wounds.

Notes 1. An approved ACUP is required prior to initiation of any mice work. 2. To reduce work load and to avoid the messy shaving on the day of the procedure mice can be shaved the previous evening.

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3. For surgeries use sterilized surgical instruments, gowns, gloves, masks, hats in a ventilated work station unit. 4. Make sure to use fixed sized punches for creating uniform size full thickness wounds so the errors can be minimized for wound closure analysis. 5. Using forceps make sure to push the nanofibers inside the wound securely and moisten them with a drop of sterile PBS using a sterile syringe and a needle. 6. Make sure the silicone splints remain stuck to the skin after using the Krazy glue before making the interrupted sutures. 7. This stage is important since the wounds are traced on a tracing paper. Make sure to separate your experimental and the test mice group to avoid any confusion. 8. The formula for the wound area calculation is given above and the wound closure percentage should be calculated according to the formula. Very carefully take the measurement with the Vernier caliper. Also, two independent blindfolded evaluations must be done for accuracy and consistency. Make sure to use an Excel spreadsheet for your measurements.

Acknowledgments This work was supported by NIH grant 1R15AR068584-01 (PI: GI, OSU) and 1R01GM123081-01 (PI: Xie, UNMC). References 1. Broughton G 2nd, Janis JE, Attinger CE (2006) The basic science of wound healing. Plast Reconstr Surg 117(7 Suppl):12S–34S. https://doi.org/10.1097/01.prs. 0000225430.42531.c2 2. Gonzalez AC, Costa TF, Andrade ZA, Medrado AR (2016) Wound healing—a literature review. An Bras Dermatol 91(5):614–620. https://doi.org/10.1590/abd1806-4841. 20164741 3. Keast D, Orsted H (1998) The basic principles of wound care. Ostomy Wound Manage 44:24–28 4. Lee SH, Jeong SK, Ahn SK (2006) An update of the defensive barrier function of skin. Yonsei Med J 47(3):293–306. https://doi.org/10. 3349/ymj.2006.47.3.293 5. Koh TJ, DiPietro LA (2011) Inflammation and wound healing: the role of the macrophage. Expert Rev Mol Med 13:e23. https://doi. org/10.1017/S1462399411001943

6. Perlman RL (2016) Mouse models of human disease: an evolutionary perspective. Evol Med Public Health 2016(1):170–176. https://doi. org/10.1093/emph/eow014 7. Ganguli-Indra G (2014) Protocol for cutaneous wound healing assay in a murine model. Methods Mol Biol 1210:151–159. https:// doi.org/10.1007/978-1-4939-1435-7_12 8. Zomer HD, Trentin AG (2018) Skin wound healing in humans and mice: challenges in translational research. J Dermatol Sci 90 (1):3–12. https://doi.org/10.1016/j. jdermsci.2017.12.009 9. Wang X, Ge J, Tredget EE, Wu Y (2013) The mouse excisional wound splinting model, including applications for stem cell transplantation. Nat Protoc 8(2):302–309. https://doi. org/10.1038/nprot.2013.002 10. Dunn L, Prosser HC, Tan JT, Vanags LZ, Ng MK, Bursill CA (2013) Murine model of wound healing. J Vis Exp 75:e50265. https://doi.org/10.3791/50265

Wound Healing in Mice with Nanofibers and Silicone Splints 11. Huang Z-M, Zhang Y, Kotaki M, Ramakrishna S (2003) A review on polymer nanofibers by electrospinning and their applications in nanocomposites. Compos Sci Technol 63:2223–2253. https://doi.org/10.1016/ S0266-3538(03)00178-7 12. Houshyar S, Kumar GS, Rifai A, Tran N, Nayak R, Shanks RA, Padhye R, Fox K, Bhattacharyya A (2019) Nanodiamond/poly-epsilon-caprolactone nanofibrous scaffold for wound management. Mater Sci Eng C Mater Biol Appl 100:378–387. https://doi.org/10. 1016/j.msec.2019.02.110 13. Guo G, Mei L, Fan R, Li X, Wang Y, Han B, Gu Y, Zhou L, Yu Z, Tong A (2017) Nanofibers for improving wound repair process: the combination of grafted chitosan and antioxidant agent. Polym Chem 8. https://doi.org/ 10.1039/C7PY00038C 14. Arkoun M, Daigle F, Heuzey MC, Ajji A (2017) Mechanism of action of electrospun chitosan-based Nanofibers against meat spoilage and pathogenic bacteria. Molecules 22(4). https://doi.org/10.3390/ molecules22040585 15. Fereydouni N, Darroudi M, Movaffagh J, Shahroodi A, Butler AE, Ganjali S, Sahebkar A (2019) Curcumin nanofibers for the purpose of wound healing. J Cell Physiol 234 (5):5537–5554. https://doi.org/10.1002/ jcp.27362 16. Jiang J, Zhang Y, Indra AK, Ganguli-Indra G, Le MN, Wang H, Hollins RR, Reilly DA, Carlson MA, Gallo RL, Gombart AF, Xie J (2018)

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1alpha,25-dihydroxyvitamin D3-eluting nanofibrous dressings induce endogenous antimicrobial peptide expression. Nanomedicine (Lond) 13(12):1417–1432. https://doi.org/ 10.2217/nnm-2018-0011 17. Behm B, Babilas P, Landthaler M, Schreml S (2012) Cytokines, chemokines and growth factors in wound healing. J Eur Acad Dermatol Venereol 26(7):812–820. https://doi.org/10. 1111/j.1468-3083.2011.04415.x 18. Greenhalgh DG, Sprugel KH, Murray MJ, Ross R (1990) PDGF and FGF stimulate wound healing in the genetically diabetic mouse. Am J Pathol 136(6):1235–1246 19. Tsuboi R, Shi CM, Sato C, Cox GN, Ogawa H (1995) Co-administration of insulin-like growth factor (IGF)-I and IGF-binding protein-1 stimulates wound healing in animal models. J Invest Dermatol 104(2):199–203 20. Park SA, Covert J, Teixeira L, Motta MJ, DeRemer SL, Abbott NL, Dubielzig R, Schurr M, Isseroff RR, McAnulty JF, Murphy CJ (2015) Importance of defining experimental conditions in a mouse excisional wound model. Wound Repair Regen 23(2):251–261. https://doi.org/10.1111/wrr.12272 21. Liang X, Bhattacharya S, Bajaj G, Guha G, Wang Z, Jang HS, Leid M, Indra AK, Ganguli-Indra G (2012) Delayed cutaneous wound healing and aberrant expression of hair follicle stem cell markers in mice selectively lacking Ctip2 in epidermis. PLoS One 7(2): e29999. https://doi.org/10.1371/journal. pone.0029999

Chapter 11 Enrichment and Characterization of Human and Murine Pulmonary Mesenchymal Progenitor Cells (MPC) Megan Summers, Karen Helm, and Susan M. Majka Abstract Tissue resident mesenchymal progenitor cells (MPC) are important regulators of tissue repair or regeneration, remodeling, inflammation, and angiogenesis. Here we describe a technology used to define, isolate, and characterize a population of resident lung MPC in both human and mouse explanted tissue. The definition of this population using a defined set of markers facilitates the repeatable isolation of a mesenchymal subpopulation population by flow cytometry and the subsequent translational study of this specific cell type and function. Key words Mesenchymal progenitor cell, Lung MPC, Flow cytometry, Isolation of MPC, Abcg2 MPC, Mouse MPC, Human MPC, Colony-forming unit analysis, CFU-F

1

Introduction The first mesenchymal stem/progenitor cells were identified by Friedenstein as bone marrow derived (BM-MPC) [1, 2]. Subsequent research defined MPC populations as present in additional adult tissues, prompting the International Society of Cellular Therapy to define an MPC by specific conditions: (1) adherence to plastic (2) expression of specific cell markers (CD73, CD90, CD105) while lacking other markers (CD45, CD34, CD14, CD11b) and (3) multilineage differentiation into mesenchymal lineages including osteoblasts, adipocytes, and chondroblasts [3]. Lung MPCs were initially identified in patient samples and were selected on the basis of their ability to adhere to plastic [4, 5]. Other groups adapted a method initially used to identify BM hematopoietic cells, to isolate a specific population of resident lung MPC, via visualization of a side population (SP) phenotype by flow cytometry [6–11]. These MPC populations, independent of their origin, demonstrated multilineage mesenchymal differentiation potential to osteocyte, adipocyte, and chondrocyte lineages,

Chrissa Kioussi (ed.), Stem Cells and Tissue Repair: Methods and Protocols, Methods in Molecular Biology, vol. 2155, https://doi.org/10.1007/978-1-0716-0655-1_11, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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and in varying combinations express the characteristic mesenchymal cell surface determinants ABCG2, CD90, CD105, CD106, CD73, CD44, Stro-1, and Sca1. In addition, they lacked the hematopoietic markers c-kit and CD34 [6, 7, 12–14]. Analysis of gene expression of lung MPCs compared to BM-MPCs indicated organ-specific gene signatures [15]. To date, additional populations of MPC have been lineage labeled and characterized in adult murine lung during both tissue homeostasis and disease [16–19]. Understanding the role MPC play during tissue homeostasis, repair, and disease is paramount to the identification of novel targets to facilitate pulmonary regeneration. The ability to repeatedly isolate and characterize a lung MPC population prompted the identification of Abcg2 as a determinant of one lung MPC population, which facilitated lineage tracing and analysis in murine model systems as well as isolation of a corresponding population from human tissue explants [16, 20]. Thus, we present methods for human and murine MPC isolation and characterization based on the expression of Abcg2.

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Materials All reagents and surgical dissection tools must be sterilized prior to performing the procedures. Tissue collection and reagent handling should be done using sterile technique in a certified Class II biological safety cabinet. All universal biosafety guidelines for processing and handling human and murine tissue should be followed.

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Tissue Digestion

1. One cubic inch of explanted distal lung tissue or explanted murine lungs: collected using sterile technique. Tissue should be processed quickly or stored in growth medium (alpha MEM with 20% fetal bovine serum (FBS), antibiotic/antimycotic) at 4  C for no longer than 6 h. 2. Type 2 Collagenase (filtered): 125 U/mg reconstituted with 35 mL HBSS (see Note 1). 3. Hanks balanced saline solution (HBSS). 4. Growth medium (alpha MEM containing nucleosides with 20% fetal bovine serum (FBS), 1% antibiotic–antimycotic). 5. Scalpels, forceps, and scissors. 6. Disposables: 50 mL conical tubes, 100 mm dishes, serological pipettes, 100 and 40 μM cell strainers. 7. Hybridization oven or other suitable equipment which can provide agitation at a constant 37  C. 8. Red blood cell lysis buffer.

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1. Vented cap T75 tissue culture flasks. 2. Attachment factor or other suitable gelatin solution (for coating flasks). 3. Growth medium. 4. 0.25% trypsin/EDTA. 5. Dulbecco’s phosphate buffered saline (DPBS) without calcium and magnesium.

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1. Staining Buffer: PBS with 2% FBS. 2. 40 ,6-diamidino-2-phenylindole (DAPI). 3. 100 and 40 μM cell strainers. 4. CD45 (APC), Ter119(Pacific Blue), and other fluorescent conjugated primary antibodies. 5. Sample tubes appropriate for your cytometer. 6. Growth medium. 7. Cell counter or hemocytometer. 8. Microcentrifuge tubes.

2.4 Colony Forming Unit Assay

1. 100 mm tissue culture dishes. 2. 0.1% gelatin. 3. Dulbecco’s phosphate buffered saline (DPBS) without calcium and magnesium. 4. 0.25% trypsin/EDTA. 5. Growth medium. 6. 4% Paraformaldehyde (see Note 2). 7. Giemsa stain: Dilute Giemsa stock solution at a ratio of 1:20 in DI-H2O (see Note 3). 8. Cell counter or hemocytometer.

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Methods All reagents used for tissue culture should be prewarmed to 37  C. Reagents for flow cytometry should be at 4  C, chilled on ice. Fluorescent reagents should be refrigerated and kept in the dark.

3.1 Human Lung Tissue Digestion

1. Using dissection tools, remove any pleura and mince lung tissue to a paste. 2. Place the minced lung tissue (~0.2 g/20 g mouse) into a 50 mL conical tube with 5–7 mL reconstituted type 2 collagenase (450 U/mL).

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3. Place the closed tube into the hybridization oven and agitate at 37  C for 1.5 h (see Note 4). 4. Add an equal volume of HBSS and triturate well. 5. Centrifuge for 10 min at 500  g to pellet cells and remove supernatant. 6. Resuspend in 10–20 mL of culture medium. 7. Whole lung cell suspensions may be cultured in filter capped T-75 flasks at this time. 3.2 Murine Tissue Digestion

1. Spray mouse with Ethanol (keep hair wet and out of the prep). 2. Grip the abdominal skin with forceps and make an incision from mid abdomen to under the mouse chin. Pull back skin from area where incision will be made. 3. Spray with ethanol. Grip the sternum with forceps and cut through the peritoneum (abdominal membrane). 4. Just below the ribcage/diaphragm cut side to side. 5. Make a small incision in the diaphragm just below the heart. The lungs will deflate and pull away from the diaphragm. 6. Surgically remove the diaphragm by cutting through the diaphragm side to side making sure not to nick the lungs. Expose the chest cavity. 7. Open up the chest cavity by cutting the ribcage laterally on each side of the mouse. Cut the ribs very carefully making sure to not nick the lungs and expose the lungs and heart. 8. Cut the adhesions/vessels between lungs and liver (toward back of mouse). 9. Hold the heart with curved forceps. To flush the blood from the lungs, insert a 20G yellow needle on a 10 mL syringe filled with sterile HBSS into the right ventricle and gently push 3–5 mL of HBSS through until the lungs appear white. 10. Dissect the lung lobes removing the trachea and large bronchi and place in a petri dish filled with Hanks buffered saline (HBSS). 11. Following collection of all lung lobes forceps are used to place tissue onto the lid of the dish. Tissue is then minced into tiny pieces (or a paste) using disposable scalpels with enough liquid to keep them moist but not so much that they float and move. 12. Mince lungs using disposable scalpels to a paste. 13. Place the minced lung tissue into a 50 mL conical tube with 7 mL reconstituted type 2 collagenase (450 U/mL). 14. Place the closed tube into a hybridization oven and digest at 37  C for 30 min.

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15. After 30 min triturate the solution until it flows easily through a 10 mL pipette (approximately 5 times) [21]. 16. Place the solution back into the rotating hybridization oven to digest for 15 more minutes at 37  C. 17. Triturate and add an equal volume of HBSS to dilute the solution. 18. Filter the solution through a 100 μM cell strainer. 19. Centrifuge for 10 min at 500  g to pellet cells and remove supernatant. 20. Resuspend the solution in 10 mL of room temperature red blood cell lysis buffer. 21. Incubate the solution for 15 min at room temperature. 22. Add an equal volume of HBSS. 23. Filter the solution through a 40 μM cell strainer. 24. Centrifuge for 10 min at 500  g to pellet cells and remove supernatant. 25. Whole lung single cell suspensions may be cultured or sorted to enrich for MPC at this time. 3.3 Human and Murine Tissue Culture

1. Gelatin-coat the T75 flasks for 20 min at room temperature prior to use (see Note 5). 2. Resuspend and plate the cell pellet in aMEM growth medium. Human cell preparations should be resuspended in 30 mL of growth medium and separated into three ventilated T-75 flasks. Murine cell isolations should be resuspended in 10–15 mL of growth medium and plated in one ventilated T-75 flask. 3. After 24 h, remove media and gently rinse the inside of the flask well with prewarmed DPBS to remove dead cells and debris. 4. Replace with prewarmed aMEM growth medium. 5. When cells reach 75% confluence, passage the cells at 1:2 or 1:3 ratio. First, rinse with DPBS to remove serum and then apply 0.25% trypsin–EDTA for no longer than 2 min (see Note 6). 6. Inactivate the trypsin with serum containing medium in a 1:1 ratio, wash the cells via brief centrifugation, resuspend the pellet and divide the cells into new flasks containing growth medium.

3.4 Enrichment of Human and Murine MPCs by Flow Cytometry

1. Trypsinize cultured adherent lung cells for no longer than 2 min and inactivate with growth medium in a 1:1 ratio as described above (see Note 7). 2. Centrifuge at 500  g for 10 min and pour off supernatant. 3. Resuspend cells in cold staining buffer so that each antibody reaction microcentrifuge tube has a volume of 300 μL.

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4. Filter the cells through a 100 μM cell strainer. 5. Count the cells. 6. The final concentration of cells in each tube should not exceed 1  106 cells/mL. If samples are very concentrated, they may be diluted using additional staining buffer (see Note 8). 7. For both mouse and human add 0.5–3 μL of fluorescently conjugated CD45 antibody (0.2 mg/mL) to each sample staining tube. For human samples add a fluorescently conjugated ABCG2 or other antibody of interest. For mouse add a Ter119 (0.2 mg/mL) fluorescently conjugated antibody (Figs. 1 and 2) (see Note 9). 8. Add 300 μL of cells to each of the staining tubes. 9. Vortex gently and incubate samples in the dark on ice for 10 min. 10. Centrifuge at 500  g for 10 min and remove supernatant. 11. Resuspend in staining buffer and transfer to a tube appropriate for your particular flow cytometer (see Note 10). 12. Add DAPI (1–2 μg/mL final concentration) to discriminate live versus dead cells (see Note 11). 13. Store samples on ice and in the dark until analysis or sorting. 14. For human cells, we typically combine CD45-APC and ABCG2-PE but any compatible combination of fluorophores may be selected and optimized for each cytometer [16, 20]. 15. Murine MPC isolation has been optimized using our lineage reporter mouse model in which Abcg2pos cells are labeled with membrane enhanced GFP (eGFP) [16, 22, 23]. All nonrecombined cells are membrane tomato positive, which can be visualized and compensated for in the PE channel if the instrument is equipped with a yellow (561 nm) laser. Nonhematopoietic cells are separated as CD45-APCneg. Any mesenchymal driver with a fluorophore reporter may be utilized. 16. Instrument controls consist of no color (cells alone) and single fluorophore labeled control samples, using either cells or compensation beads. Compensation beads are particularly useful because they are clearly positive for the antibodies used in the assay, while the experimental cells may represent a minority of the cells and thus are insufficient for color compensation. Ideally, positive controls of highly enriched murine eGFP and tomato expressing cells and tomato alone controls (i.e., an established cell line) would be included [24]. 17. These samples may be utilized to set the standard gates for analysis and sorting. First cell populations are identified by properties of light scatter, forward versus side scatter (Figs. 1a and 2a).

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18. Next side scatter height versus width followed by forward scatter height versus width are gated to isolate single cells and eliminate doublets. The best parameters for detecting cell singlets often vary between manufacturers (Figs. 1b, c and 2b). 19. Viability is next identified by plotting DAPI in a histogram (DAPI versus side scatter area is also an option) (Figs. 1d, and 2c) (see Note 11). 20. Gates are next set to distinguish CD45 populations (Figs. 1e and 2d). 21. Gates to visualize cells of interest, the CD45neg MPC, are next drawn (Figs. 1e and 2e) and optimized using the following controls. 22. FMO (fluorescence minus one) controls are also essential to differentiate which population of cells will be considered positive or negative. 23. Following instrument setup (compensation and voltage adjustment) a small volume of sample is analyzed to determine where your cells of interest fall. 24. Save a presort file. 25. Begin analysis/sorting. 26. For human samples, a visible population of CD45neg ABCG2pos cells should be visible once approximately 10,000 events are analyzed. The ABCG2pos-PE positive population varies in intensity and visualization may be improved by plotting side scatter area versus ABCG2-PE (Fig. 1f). 27. For murine samples, the membrane eGFP is bright. A subpopulation of eGFP cells will likely express membrane tomato, as loss of tomato following recombination requires cell division. 28. Save a postsort file for analysis. 29. Collect MPC into growth medium. 30. A small aliquot of sorted cells may be used in a purity check following cleaning of the instrument to remove any residual sample in the instrument. 31. Plate onto a gelatin-coated flask and culture as described in Subheading 3.3. 32. MPC may be visualized within 48 h as large adherent cells with multiple processes (Fig. 5a). 3.5 Cell Surface Marker Characterization by Flow Cytometry

1. Determine how many staining reaction tubes you will need to test your entire panel of relevant markers and prepare all needed tubes. 2. Trypsinize cells at 37  C for no longer than 2 min (see Note 7). 3. Inactivate trypsin with serum containing medium.

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4. Filter through sterile 100 μM filter and count the cells. 5. Centrifuge samples at 500  g for 10 min and pour off supernatant. 6. Resuspend cells in staining buffer at 300 μL per antibody reaction (see Notes 8 and 9). The resuspended cell concentration can be less than but no more than 3  106 cells/mL. 7. To each sample staining tube add 0.5–3 μL of an appropriate antibody (see Notes 12–14). 8. Vortex gently and incubate samples for 10 min on ice, in the dark. 9. Centrifuge at 500  g for 10 min and remove the supernatant. 10. Resuspend each sample in 500 μL of staining buffer. 11. Transfer samples to flow cytometry tubes and add DAPI to discriminate live versus dead cells. 12. Store on ice in the dark until analysis. 13. Set compensation, voltage parameters and initial gates for size as described in Subheading 3.4 (Figs. 3 and 4). 3.6 Colony Forming Unit Assay

1. CFU-F should be performed in duplicate or triplicate per cell line. Coat 100 mm culture dishes with 0.1% gelatin for up to an hour. 2. Trypsinize cells and inactivate as described in previous sections. 3. Filter cells through a 100 μM cell strainer. 4. Count the cells and add 3  103 human cells per 100 mm culture dish. Add an equal amount of fresh media to each dish. 5. Culture cells for 5–10 days, observing the formation of colonies daily. If the assay must run longer than 7 days to visualize slow growing cell lines, the media should be changed (see Note 15). 6. Wash the plates with Dulbecco’s phosphate buffered saline (DPBS) without calcium and magnesium. 7. Fix the cells using 4% paraformaldehyde (PFA) for 20 min at room temperature. 8. Remove PFA. 9. Wash the plates with DPBS three times. 10. Add 10 mL Giemsa stain diluted to the proper concentration. This can be left to stain overnight. 11. Remove Giemsa staining solution and wash the plates gently with distilled water to remove unbound stain. Rinse continuously until the water is clear (see Note 16). 12. Leave the plates uncovered and allow them to air-dry. 13. Colonies may be enumerated by eye or using a microscope (Fig. 5b) [22, 25, 26].

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Notes 1. The contents of the sealed vial may be under pressure, causing the powder to disperse upon opening. Remove the stopper very slowly and carefully. The typical enzyme concentration is approximately 16,000 U/vial resulting in a final concentration of 450 U/mL. For additional details please refer to [21]. 2. PFA should be disposed of as a bio/chemical hazard. 3. Giemsa stain should be disposed of as a bio/chemical hazard.

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Fig. 4 Characterization of primary murine lung MPC lines by flow cytometry. Gating strategies utilized to characterize cell surface marker expression of CD45neg Abcg2/eGFPpos murine lung MPC. (a) The population is visualized by plotting side scatter area (SSC-A) versus Forward scatter area (FSC-A). (b) Singlet gates. (c) Singlet gates. (d) The DAPI histogram distinguishes live versus dead cells. (e) Expression of eGFP is confirmed. (f) Singlecolor gates may be drawn as SSC-A versus the fluorophore or multiple parameters may be analyzed at once

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Fig. 5 Culture and enumeration of human and murine MPC colony forming potential (CFU-F). (a) Representative bright field images of human and murine ABCG2pos MPC. (b) Representative images of human and murine CFU-F Giemsa stained assays to enumerate clonal potential of MPC

4. Volumes can be scaled up depending on the number of lungs isolated. Tube lids should be sealed with Parafilm to reduce leakage when incubated horizontally. 5. Gelatin coated plastic increases cell adhesion and is most useful immediately after collagenase digest of flow sorting to establish the culture. 6. Cultures must be passaged promptly when areas reach 75–80% confluence. To passage cultures quickly, limiting the cell exposure to trypsin, visualize the cell edges lifting and tap the flask on a hard surface to release the cells. Longer exposures to trypsin may decrease cell viability. 7. Longer trypsinization can cleave surface determinants resulting in a false negative. 8. Antibodies may be used as single-color samples or combined with compatible antibody sets to reduce the number of samples and determine relationships between markers. Compensation between each fluorophore should be determined using positive and negative compensation beads. An FMO including all stains minus the one of interest can be used to set positive and negative gates. 9. Any compatible combination of fluorophores optimal for each particular cytometer may be used. Ter119 marks late stages of the murine erythroid lineage. 10. Cell strainer snap caps may be utilized as an additional filter step at this stage, immediately prior to sorting/analysis. Filtering is recommended if the preparation was stored on ice.

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11. For murine cell isolation/analysis DAPI and Ter119 can be combined into a “dump gate” to eliminate dead and remaining red blood cells from analysis. 12. Preliminary titration analysis is necessary to determine the optimal antibody concentration. 13. Dispensing antibodies into tubes prior to the addition of cell suspension decreases the risk of contamination of stock solutions. 14. To determine true positive or negative populations relative to a low positive population, a known high and low positive control is helpful. 15. The assay should be concluded when colonies have formed but before they are contiguous. 16. Do not scrape colonies while removing stain and water.

Acknowledgments The authors would like to extend their greatest appreciation for the expert technical assistance provided by Kelsey Chow and Karen Helm at the University of Colorado, Christa Gaskill of Vanderbilt University Medical Center and Catherine E. Alford of the Flow Cytometry Special Resource Center, Department of Pathology and Laboratory Medicine, Veterans Affairs Tennessee Valley Healthcare System, Nashville, TN. This work was funded by grants to S.M. Majka from the NIH/NHLBI R01HL116597 and NIH/NHLBI R01HL136449. The project was also supported in part by the University of Colorado Cancer Center shared flow cytometry resource (P30-CA046934 NCI). References 1. Bianco P, Robey PG, Simmons PJ (2008) Mesenchymal stem cells: revisiting history, concepts, and assays. Cell Stem Cell 2 (4):313–319. https://doi.org/10.1016/j. stem.2008.03.002 2. Charbord P (2010) Bone marrow mesenchymal stem cells: historical overview and concepts. Hum Gene Ther 21(9):1045–1056. https://doi.org/10.1089/hum.2010.115 3. Dominici M, Le Blanc K, Mueller I, SlaperCortenbach I, Marini F, Krause D, Deans R, Keating A, Dj P, Horwitz E (2006) Minimal criteria for defining multipotent mesenchymal stromal cells. The International Society for Cellular Therapy position statement. Cytotherapy 8(4):315–317

4. Lama VN, Smith L, Badri L, Flint A, Andrei A-C, Murray S, Wang Z, Liao H, Toews GB, Krebsbach PH, Peters-Golden M, Pinsky DJ, Martinez FJ, Thannickal VJ (2007) Evidence for tissue-resident mesenchymal stem cells in human adult lung from studies of transplanted allografts. J Clin Invest 117(4):989–996 5. Hennrick KT, Keeton AG, Nanua S, Kijek TG, Goldsmith AM, Sajjan US, Bentley JK, Lama VN, Moore BB, Schumacher RE, Thannickal VJ, Hershenson MB (2007) Lung cells from neonates show a mesenchymal stem cell phenotype. Am J Respir Crit Care Med 175 (11):1158–1164. https://doi.org/10.1164/ rccm.200607-941OC 6. Summer R, Fitzsimmons K, Dwyer D, Murphy J, Fine A (2007) Isolation of an adult

Isolation of Lung MPC mouse lung mesenchymal progenitor cell population. Am J Respir Cell Mol Biol 37 (2):152–159. https://doi.org/10.1165/ rcmb.2006-0386OC 7. Morigi M, Introna M, Imberti B, Corna D, Abbate M, Rota C, Rottoli D, Benigni A, Perico N, Zoja C, Rambaldi A, Remuzzi A, Remuzzi G (2008) Human bone marrow mesenchymal stem cells accelerate recovery of acute renal injury and prolong survival in mice. Stem Cells 26(8):2075–2082. https:// doi.org/10.1634/stemcells.2007-0795 8. Irwin D, Helm K, Campbell N, Imamura M, Fagan K, Harral J, Carr M, Young KA, Klemm D, Gebb S, Dempsey EC, West J, Majka S (2007) Neonatal lung side population cells demonstrate endothelial potential and are altered in response to hyperoxia-induced lung simplification. Am J Physiol Lung Cell Mol Physiol 293:L941–L951 9. Fatima S, Zhou S, Sorrentino BP (2012) Abcg2 expression marks tissue-specific stem cells in multiple organs in a mouse progeny tracking model. Stem Cells 30(2):210–221. https://doi.org/10.1002/stem.1002 10. Tadjali M, Zhou S, Rehg J, Sorrentino BP (2006) Prospective isolation of murine hematopoietic stem cells by expression of an Abcg2/ GFP allele. Stem Cells 24(6):1556–1563. https://doi.org/10.1634/stemcells.20050562 11. Zhou S, Schuetz JD, Bunting KD, Colapietro A-M, Sampath J, Morris JJ, Lagutina I, Grosveld GC, Osaw M, Nakauchi H, Sorrentino BP (2001) The ABC transporter Bcrp1/ABCG2 is expressed in a wide variety of stem cells and is a molecular determinant of the side-population phenotype. Nat Med 7:1028–1034 12. Jun D, Garat C, West J, Thorn N, Chow K, Cleaver T, Sullivan T, Torchia EC, Childs C, Shade T, Tadjali M, Lara A, Nozik-Grayck E, Malkoski S, Sorrentino B, Meyrick B, Klemm D, Rojas M, Wagner DH, Majka SM (2011) The pathology of bleomycin-induced fibrosis is associated with loss of resident lung mesenchymal stem cells that regulate effector T-cell proliferation. Stem Cells 29 (4):725–735. https://doi.org/10.1002/ stem.604 13. Chateauvieux S, Ichante´ J-L, Delorme B, Frouin V, Pie´tu G, Langonne´ A, Gallay N, Sensebe´ L, Martin MT, Moore KA, Charbord P (2007) Molecular profile of mouse stromal mesenchymal stem cells. Physiol Genomics 29 (2):128–138. https://doi.org/10.1152/ physiolgenomics.00197.2006 14. McQualter JL, Brouard N, Williams B, Baird BN, Sims-Lucas S, Yuen K, Nilsson SK,

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Simmons PJ, Bertoncello I (2009) Endogenous fibroblastic progenitor cells in the adult mouse lung are highly enriched in the Sca-1 positive cell fraction. Stem Cells 27 (3):623–633. https://doi.org/10.1634/ste mcells.2008-0866 15. Walker NM, Badri LN, Wadhwa A, Wettlaufer S, Peters-Golden M, Lama VN (2011) Prostaglandin E2 as an inhibitory modulator of fibrogenesis in human lung allografts. Am J Respir Crit Care Med 185(1):77–84. https://doi.org/10.1164/rccm.2011050834OC 16. Gaskill CF, Carrier EJ, Kropski JA, Bloodworth NC, Menon S, Foronjy RF, Taketo MM, Hong CC, Austin ED, West JD, Means AL, Loyd JE, Merryman WD, Hemnes AR, De Langhe S, Blackwell TS, Klemm DJ, Majka SM (2017) Disruption of lineage specification in adult pulmonary mesenchymal progenitor cells promotes microvascular dysfunction. J Clin Invest 127(6). https://doi.org/10.1172/ JCI88629 17. Xie T, Liang J, Liu N, Huan C, Zhang Y, Liu W, Kumar M, Xiao R, DArmiento J, Metzger D, Chambon P, Papaioannou VE, Stripp BR, Jiang D, Noble PW (2016) Transcription factor TBX4 regulates myofibroblast accumulation and lung fibrosis. J Clin Invest 126(8):3063–3079. https://doi.org/10. 1172/JCI85328 18. Kramann R, Schneider RK, DiRocco DP, Machado F, Fleig S, Bondzie PA, Henderson JM, Ebert BL, Humphreys BD (2015) Perivascular Gli1+ progenitors are key contributors to injury-induced organ fibrosis. Cell Stem Cell 16(1):51–66. https://doi.org/10.1016/j. stem.2014.11.004 19. El Agha E, Kramann R, Schneider RK, Li X, Seeger W, Humphreys BD, Bellusci S (2017) Mesenchymal stem cells in fibrotic disease. Cell Stem Cell 21(2):166–177. https://doi.org/ 10.1016/j.stem.2017.07.011 20. Marriott S, Baskir RS, Gaskill C, Menon S, Carrier EJ, Williams J, Talati M, Helm K, Alford CE, Kropski JA, Loyd J, Wheeler L, Johnson J, Austin E, Nozik-Grayck E, Meyrick B, West JD, Klemm DJ, Majka SM (2014) ABCG2(pos) lung mesenchymal stem cells are a novel pericyte subpopulation that contributes to fibrotic remodeling. Am J Physiol Cell Physiol 307(8):C684–C698. https://doi.org/10.1152/ajpcell.00114.2014 21. Chow KS, Jun D, Helm KM, Wagner DH, Majka SM (2011) Isolation & characterization of hoechst(low) CD45(negative) mouse lung mesenchymal stem cells. J Vis Exp 56:3159. https://doi.org/10.3791/3159

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22. Chow K, Fessel JP, KaoriIhida S, Schmidt EP, Gaskill C, Alvarez D, Graham B, Harrison DG, Wagner DH, Nozik-Grayck E, West JD, Klemm DJ, Majka SM (2013) Dysfunctional resident lung mesenchymal stem cells contribute to pulmonary microvascular remodeling. Pulm Circ 3(1):31–49. https://doi.org/10. 4103/2045-8932.109912 23. Gaskill C, Marriott S, Pratap S, Menon S, Hedges LK, Fessel JP, Kropski JA, Ames D, Wheeler L, Loyd JE, Hemnes AR, Roop DR, Klemm DJ, Austin ED, Majka SM (2016) Shared gene expression patterns in mesenchymal progenitors derived from lung and epidermis in pulmonary arterial hypertension: identifying key pathways in pulmonary vascular disease. Pulm Circ 6(4):483–497. https://doi. org/10.1086/688314

24. Alvarez DF, Helm K, DeGregori J, Roederer M, Majka S (2010) Publishing flow cytometry data. Am J Physiol Lung Cell Mol Physiol 298(2):L127 25. Alt E, Yan Y, Gehmert S, Song Y-H, Altman A, Gehmert S, Vykoukal D, Bai X (2011) Fibroblasts share mesenchymal phenotypes with stem cells, but lack their differentiation and colony-forming potential. Biol Cell 103 (4):197–208. https://doi.org/10.1042/ bc20100117 26. Kuznetsov SA, Mankani MH, Bianco P, Robey PG (2009) Enumeration of the colonyforming units-fibroblast from mouse and human bone marrow in normal and pathological conditions. Stem Cell Res 2(1):83–94. https://doi.org/10.1016/j.scr.2008.07.007

Chapter 12 Isolation and Culture of Quiescent Skeletal Muscle Satellite Cells Francisco Herna´ndez-Torres, Lara Rodrı´guez-Outeirin˜o, and Amelia Ara´nega Abstract It has been shown that freshly isolated satellite cells from adult muscle constitute a stem cell-like population that exhibits more efficient engraftment and self-renewal activity in regenerating muscle than myoblast. Thus, purification of pure populations of quiescent satellite cells from adult skeletal muscle is highly necessary, not only for understanding the biology of satellite cells and myoblasts but also for improving cell-based therapies for muscle regeneration. This chapter describes a basic protocol used in our laboratory to isolate quiescent muscle satellite cells from adult skeletal muscle by enzymatic dissociation followed by a sequential magnetic-activated cell sorting (MACS). This method is cheap and fast providing and alternative procedure to other purification methods that require fluorescence-activated cell sorting (FACS) machines. Freshly isolated quiescent satellite cells purified by this method can be used in a broad range of experiments including cell transplantation for satellite cell self-renewal experiments or cell therapies. Key words Skeletal muscle, Muscle satellite cells, Myogenesis, Myogenic differentiation, Cell culture, Magnetic-activated cell sorting (MACS) cell separation

1

Introduction Skeletal muscle is a heterogeneous tissue that represents between 30% and 38% of the human body mass [1]. This tissue retains a highly adaptive and robust capacity to regenerate throughout most of life [2]. The maintenance as well as the repair of adult muscle tissue are both directed by satellite cells [2, 3]. These cells were originally identified via electron microscopy in 1961 by Alexander Mauro, located underneath the basal lamina and adjacent to the plasma membrane of the skeletal muscle myofiber [4]. In its quiescence state, satellite are characterized by the expression of the transcription factor Pax7 and represent a genuine somatic stem cell population indispensable for skeletal muscle repair [5, 6]. Within a context of physiological stimuli (physical exercise or pathological conditions) satellite cells become activated, leave their quiescent

Chrissa Kioussi (ed.), Stem Cells and Tissue Repair: Methods and Protocols, Methods in Molecular Biology, vol. 2155, https://doi.org/10.1007/978-1-0716-0655-1_12, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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state and enter the cell cycle to expand their progeny, termed myogenic precursor cells or myoblasts [7, 8]. During this activation the basic helix–loop–helix (bHLH) factors Myf5, Myod1, Myog, and Myf6 act sequentially to advance satellite cells and myoblast toward myogenic differentiation and fusion to form multinucleated myofibers in order to repair the damaged muscle [7, 8]. There are different stable myoblast cell lines commonly used in the field of myogenesis (e.g., C2C12, Sol8, L6, or MM14). These cell lines are able to differentiate into myotubes upon incubation in differentiation medium thus being suitable for the study of myogenic process in vitro. However, exist some degree of variability in results obtained with them, mainly due to the origin of cells, the side-effects characteristic of the immortalization process, specific culture conditions and/or passage number. Thus, in order to avoid these sources of variability, the use of primary myogenic cells is highly recommended. Since the first protocol for mouse myoblast purification was developed [9] several adapted protocols has emerged [10, 11]. All these protocols allow investigators to get yields of primary myoblast enough for culturing and perform myogenic differentiation studies in vitro. As we mentioned before, myoblasts arise as consequence of satellite cell activation. In this regard, it has been shown that freshly isolated satellite cells from adult muscle constitute a stem cell-like population that exhibits more efficient engraftment and self-renewal activity in regenerating muscle than myoblast [12–15]. Therefore, in order to take advantage of this ability, purification of pure populations of quiescent satellite cells from adult skeletal muscle is highly necessary, not only for understanding the biology of satellite cells and myoblasts, but also to improve cell-based therapies for muscle regeneration. This chapter describes a basic protocol used in our laboratory to isolate quiescent muscle satellite cells from adult skeletal muscle by enzymatic dissociation followed by a sequential magneticactivated cell sorting (MACS). First, mouse satellite cells are isolated by depletion of nontarget cells directly labeled with a cocktail of monoclonal antibodies conjugated with magnetic MACS MicroBeads (Satellite Cell Isolation Kit). These cells are retained within a MACS Column in the magnetic field of a MACS Separator, while the unlabeled satellite cells pass through the column. Soon after, this mouse satellite cells fraction is enriched by the use of Anti-Integrin α-7 MicroBeads that magnetically label integrin α-7+ cells, a specific marker for satellite cells [14]. Finally, the cell suspension is loaded onto a second MACS Column, which is placed in the magnetic field of a MACS Separator. The magnetically labeled integrin α-7+ cells are retained within the column while the unlabeled cells run through. After removing the column from the magnetic field, the magnetically retained integrin α-7+ cells can be eluted as the positively selected cell fraction. This method is a faster, more economical, and more reliable alternative than purification

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methods of quiescent satellite cells that require the use of expensive fluorescence-activated cell sorting (FACS) machines. Freshly isolated quiescent satellite cells purified by this method can be used in a broad range of experiments such as gene and protein expression profiles, oligonucleotide or plasmid transient transfection, viral transduction, cell transplantation for satellite cell selfrenewal experiments or cell therapies.

2 2.1

Materials Buffers

1. Phosphate-buffered saline (PBS): 137 mM NaCl, 2.7 mM KCl, 4.3 mM Na2HPO4·7H2O, 1.4 mM KH2PO4, pH 7.3. 2. MACS buffer: PBS, 0.5% bovine serum albumin (BSA), 2 mM EDTA. Filter the buffer through 0.22 μm filter.

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1. Dulbecco’s Modified Eagle’s Medium (DMEM). 2. Dulbecco’s Modified Eagle Medium: F12 (DMEM:F12). 3. DMEM, high glucose, GlutaMAX™ Supplement, pyruvate. 4. Fetal bovine serum (FBS), Research Grade. 5. Horse serum (HS). 6. Ultroser™ G serum substitute. Ultroser™ G serum substitute is available as a lyophilized powder. Resuspend it in 20 mL of sterile cell culture water and filter the serum through 0.22 μm filter. Aliquot and freeze at 20  C. 7. Penicillin–streptomycin. 8. Satellite Cell Growth Medium: DMEM, high glucose, GlutaMAX™ Supplement, pyruvate 37.5%, DMEM: F12 37.5%, FBS 20%, Ultroser™ G 2%, Penicillin–Streptomycin 3%. 9. Differentiation Medium: DMEM 97%, HS 2%, Penicillin– Streptomycin 1%.

2.3 Solutions and Reagents

1. 1 Trypsin solution from porcine pancreas. 2. Collagenase D solution. Resuspend 500 mg of Collagenase D in 50 mL of DMEM: F12 Medium to get a final concentration of 10 mg/mL. Filter the solution through 0.22 μm filter. Aliquot and freeze at 20  C. 3. Gelatin Solution 0.1%. Dilute 0.5 mL of Gelatin Solution 2% in 9.5 mL of sterile cell culture water to get 10 mL of Gelatin Solution 0.1%. 4. Enzymatic Solution: DMEM: F12 86%, Collagenase D solution 10%, Trypsin solution 4%. 5. Satellite Cell Isolation Kit mouse. 6. Anti-Integrin α-7 MicroBeads mouse.

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7. Red Blood Cell Lysis Solution 10. 8. LS columns. 9. MS Columns. 2.4 Plasticware and Glassware Supplies

1. Standard Plastic Pasteur pipettes. 2. 5 mL serological glass pipettes. 3. 10 mL serological glass pipettes. 4. 25 mL serological glass pipettes. 5. Polypropylene conical centrifuge tubes, sterile (15, 30, and 50 mL). 6. Plastic petri dishes, 100-mm. 7. Tissue culture dishes, 35-mm. 8. Cell strainer, 100 μm nylon mesh. 9. Cell strainer, 40 μm nylon mesh.

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1. Autoclave oven. 2. Dissection tools: scissors (Surgical, Straight Extra Fine Bonn, Curved Extra Fine Bonn and Vannas Spring Scissors) and forceps (Straight Dumont #5 and curved Dumont #7 Forceps) (see Fig. 1a). 3. Stereo dissecting microscope with transmitted light base. 4. Biosafety laminar flow cabinet. 5. Pipette controller. 6. Benchtop refrigerated centrifuge for 15, 30 and 50 mL tubes. 7. Standard humidified tissue culture incubator (37  C, 5% CO2 in air). 8. Water bath. 9. Inverted phase contrast microscope for monitoring cell culture. 10. Hemacytometer.

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3.1 Muscle Dissection and Digestion

1. Sacrifice 1 adult mice (3–8 weeks) by cervical dislocation (see Note 1). 2. Spray abdomen, back, and hind limbs with 70% ethanol. Pinch the skin of the abdomen, slit with surgical scissors, and peel off skin to completely show hind limb muscles by pulling the skin in opposing directions (see Note 2). 3. With a fine Vannas Spring scissors, cut through the thin fascia without damaging the underlying hind limb muscles. With the help of fine Dumont forceps and Curved Extra Fine Bonn

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Fig. 1 Isolation of muscle satellite cells from skeletal muscle tissue. (a) Dissection tools from left to right, scissors (Surgical, Straight Extra Fine Bonn, Curved Extra Fine Bonn and Vannas Spring Scissors) and forceps (Straight Dumont #5 and curved Dumont #7 Forceps). (b) Mincing tissue into a smooth pulp in DMEM:F12 (1:1) with Straight Extra Fine Bonn Scissors. (c) Triturating muscle pieces in Enzymatic Solution trough an 18G needle. (d) Transferring supernatant–FBS mixture containing the dissociated cells onto a 10 μm cell strainer. (e) Collecting flow-through MACS containing unlabeled cells from Satellite Cell Isolation Kit MicroBeads by LS Column. (f) Applying cell suspension labeled with Anti-Integrin α-7 MicroBeads onto MS Column. (g) Flush out labeled integrin α7–positive cells from MS Column with 1 mL of buffer by quickly pushing the plunger. (h) Colonies of satellite cells and activated myoblast after 24 h with Satellite Cell Growth Medium, 20

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scissor, remove all leg skeletal muscles (tibialis anterior, extensor digitorum lateralis, extensor digitorum longus, peroneus longus, gastrocnemius, plantaris, soleus, and quadriceps). Then transfer muscles to ice-cold sterile PBS in a plastic 15 mL tube (see Note 3). 4. From this moment onward, all steps must be carried out in a sterile biosafety laminar flow cabinet. Wash blood off muscles in PBS and transfer muscles to a new sterile 100-mm plastic petri dish with 10 mL of DMEN:F12 (1:1) (see Note 4). 5. With the help of fine Dumont forceps and fine Vannas Spring scissors, remove connective tissue, blood vessels, nerve bundles, and adipogenic tissue under a dissection inverted microscope (see Note 5). 6. Using Straight Extra Fine Bonn scissors, cut and mince the tissue into a smooth pulp (see Fig. 1b). 7. Transfer minced muscles with Pasteur pipette into a Falcon 50 mL tube and centrifuge at 200  g at 4  C for 5 min, aspirate and discard the supernatant. 8. Resuspend minced muscles with 5 mL of Enzymatic Solution and incubate at 37  C for 60 min in a water bath shaker with at low rotation speed (10–30 oscillations/min) (see Notes 6–8). 9. Centrifuge at 1300 rpm at 4  C for 5 min, aspirate and discard the supernatant. 10. Resuspend the pellet with 5 mL of Enzymatic Solution and triturate (up and down with a 14G needle) to homogenize the mixture (see Note 9). 11. Let the mixture to decant for 5 min. Recover the supernatant and transfer it to a new 50 mL with 5 mL of FBS and keep it in the refrigerator (2–8  C). 12. Resuspend the pellet with 5 mL of Enzymatic Solution and incubate at 37  C for 20 min in a water bath shaker. 13. Triturate again to homogenize the mixture to dissociate into single cell suspension (up and down with an 18G needle) (see Fig. 1c). 14. Let the mixture to decant for 5 min. Recover the supernatant and transfer it to the Falcon 50 mL tube with FBS used in step 11. Discard the pellet (mostly tendon and tissue debris). 15. Place a 100 μm cell strainer onto a 50 mL tube. Transfer the supernatant–FBS mixture containing the dissociated cells onto the cell strainer. Pipet the cell suspension up and down on the filter until it passes through (see Fig. 1d). 16. Place a 40 μm cell strainer onto a 50 mL tube. Transfer the flow-through recovered from previous step containing the

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dissociated cells onto the cell strainer. Pipet the cell suspension up and down on the filter until it passes through. 17. Transfer the flow-through into a 30 mL centrifuge tube and centrifuge at 1300 rpm at 4  C for 10 min, aspirate and discard the supernatant. 18. In order to erase red blood cells, resuspend the pellet with 1 mL PBS, mix it with 10 mL of Red Blood Cell Lysis Solution 1 and incubate at room temperature for 2 min (see Note 10). 19. Centrifuge at 1300 rpm at 4  C for 10 min, aspirate and discard the supernatant. At this point, cells are ready for magnetic labeling and separation. 3.2 Magnetic Labeling and Separation

1. Resuspend cell pellet in 160 μL of MACS buffer + 40 μL of Satellite Cell Isolation Kit MicroBeads. Mix well and incubate for 15 min in the refrigerator (2–8  C) (see Note 11). 2. Meanwhile place an LS Column in the magnetic field of a suitable MACS Separator and equilibrate it by rinsing with 3  1 mL of MACS buffer (see Note 12). 3. Remove cell–MicroBeads suspension from the refrigerator and adjust volume to 500 μL using MACS buffer. 4. For depletion of nonsatellite cells, apply cell suspension onto the column and collect flow-through containing unlabeled cells (see Fig. 1e). 5. Wash column with 3  1 mL of MACS buffer. Collect unlabeled cell that pass through and combine with the flowthrough from step 4. 6. Centrifuge at 1300 rpm at 4  C for 10 min; aspirate and discard the supernatant. 7. Resuspend cell pellet in 160 μL of MACS buffer + 40 μL of Anti-Integrin α-7 MicroBeads. Mix well and incubate for 15 min in the refrigerator (2–8  C). 8. Meanwhile place an MS Column in the magnetic field of a suitable MACS Separator and equilibrate it by rinsing with 3  500 μL of MACS buffer. 9. Remove cells–MicroBeads suspension from the refrigerator and adjust volume to 500 μL using MACS buffer. 10. Apply cell suspension onto the column. Discard the flowthrough (Integrin α7–negative cells) (see Fig. 1f). 11. Wash column with 3  500 μL of MACS buffer. 12. Remove column from the separator and place it on a suitable collection tube.

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13. Pipet 1 mL of buffer onto the column. Immediately flush out the magnetically labeled cells (Integrin α7–positive cells) by firmly pushing the plunger into the column (see Fig. 1g). 14. Centrifuge at 1300 rpm at 4  C for 5 min; carefully aspirate and discard the supernatant. 15. Resuspend purified cell pellet in 500 μL of Satellite Cell Growth Medium and count cell number by hemocytometer. Starting from an adult mouse, each animal yields 1–2  105 pure satellite cells. At this point satellite cells are ready for cell culture. 3.3

Cell Culture

3.3.1 Grown and Maintenance

1. For maintenance, dilute cells with Satellite Cell Growth Medium at desired cell density and seed them on 0.1% gelatin-coated dishes (see Notes 13 and 14). Usually 1–2  105 cells are seeded in 35-mm dishes but, alternatively, 1–2  104 cells/well can be seeded in 24-well trays. 2. Culture the cells undisturbed in the incubator for 3 days at 37  C and 5% CO2. Quiescent satellite cells in culture activate and enter the cell cycle within 24 h after isolation to undergo myogenic precursor cells or myoblasts that star to proliferate. At this point small colonies of small and round shape satellite cells and activated myoblast are visible in the plate (see Fig. 1h). 3. Let the colonies growing in the incubator at 37  C and 5% CO2 by feeding the cells every other day with Satellite Cell Growth Medium. Once proper myoblasts confluence is reached (50–70% confluence/colony or when starting cell fusion), cells can be switched to Differentiation Medium for myoblast fusion and differentiation into myofibers.

3.3.2 Differentiation

1. Replace Satellite Cell Growth Medium by Differentiation Medium. Let the cells growing in the incubator at 37  C and 5% CO2 and refeed with this medium every other day. 2. In the Differentiation Medium, myoblasts exit the cell cycle, fuse with each other, and become multinucleated myotubes, which express MHC. Typically, this takes 3–5 days.

4

Notes 1. In order to minimize animal stress, place the animal in a narcotic chamber and apply a flow of isoflurane 5% to anesthetize the animal before cervical dislocation. 2. The isolation of hind limb muscles from mice is performed outside a biosafety cabinet. For this step, no sterile techniques are necessary except autoclaved dissection tools and spray of ethanol 70%.

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3. Autoclave dissection tools in sterilization pouches before star the procedure. 4. Muscle collection from both limbs should be performed within 5 min to preserve viability. 5. This step is critical in order to improve later enzymatic digestion and to avoid cell strainer obstruction. The cleaner is the muscle tissue the higher are satellite cells yields. 6. All media as well as Enzymatic Solution should be warmed at 37  C before use. 7. High rotation speed could result in premature satellite cells activation. In case that your laboratory has not water bath shaker, place the sample in a water bath and slightly move it by hand every 10 min. 8. At the end of this step observe the appearance of the digesting muscle mixture. Muscle pieces should look smaller and the digestion medium should start to appear cloudy. Enzymatic Solution incubation time may need to be adjusted depending on collagenase and trypsin activity. Longer or shorter incubation might be required depending on the size, age, and/or muscle condition (e.g., fibrotic muscles from old or mdx mice models need longer digestion time); however, avoid muscle overdigestion as this inevitably results in cell death. 9. Do not crush the sample through the needle, as high pressure could induce cell death. In case the tissue clumps still remain let the sample in the water bath for additional 10 min and repeat the step. 10. Use ice-cold sterile PBS and Red Blood Cell Lysis Solution 1. 11. Always use freshly prepared MACS buffer and keep it cold (2–8  C). 12. Perform washing steps by adding buffer aliquots only when the column reservoir is empty. Avoid air bubbles formation since they could block the column. 13. For coating tissue culture dishes with gelatin, distribute 300–500 μL of gelatin solution 0.1% into 35-mm culture dishes (150–200 μL into each well in case of use of 24-well plates). Swirl gently the dishes to allow even coating of the plating surface. Allow the gelatin-coated dishes to sit at room temperature for at least 1 h. Remove the entire volume of gelatin solution from the dishes (it will leave a thin coat of gelatin at the bottom of the dishes) and, finally, let the gelatincoated dishes sit in the tissue culture hood for at least 30 min before plating the isolated cells. Aliquots of diluted gelatin can be reused multiple times for coating purpose if stored at 4  C.

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14. For the mouse strain (C57BL/6) and hind limb muscles used for the protocol described herein, each preparation typically yields 1–2  105 cells. However, cell yields can vary depending on the mouse strain and/or the age of the animal. Thus, muscles from neonatal and young mice (1-month old or less) yield considerably more myogenic progenitors than muscles from adult mice. References 1. Janssen I, Heymsfield SB, Wang ZM et al (2000) Skeletal muscle mass and distribution in 468 men and women aged 18-88 yr. J Appl Physiol 89:81–88 2. Almada AE, Wagers AJ (2016) Molecular circuitry of stem cell fate in skeletal muscle regeneration, ageing and disease. Nat Rev Mol Cell Biol 17:267–279 3. Vallejo D, Herna´ndez-Torres F, LozanoVelasco E et al (2018) PITX2 enhances the regenerative potential of dystrophic skeletal muscle stem cells. Stem Cell Reports 10:1398–1411. https://doi.org/10.1016/j. stemcr.2018.03.009 4. Mauro A (1961) Satellite cell of skeletal muscle fibers. J Biophys Biochem Cytol 9:493–495 5. Seale P, Sabourin LA, Girgis-Gabardo A et al (2000) Pax7 is required for the specification of myogenic satellite cells. Cell 102:777–786 6. Seale P, Ishibashi J, Scime` A et al (2004) Pax7 is necessary and sufficient for the myogenic specification of CD45+:Sca1+ stem cells from injured muscle. PLoS Biol 2:E130 7. Zanou N, Gailly P (2013) Skeletal muscle hypertrophy and regeneration: interplay between the myogenic regulatory factors (MRFs) and insulin-like growth factors (IGFs) pathways. Cell Mol Life Sci 70:4117–4130

˜ o L, 8. Hernandez-Torres F, Rodrı´guez-Outeirin Franco D et al (2017) Pitx2 in embryonic and adult myogenesis. Front Cell Dev Biol 5:46 9. Rando TA, Blau HM (1994) Primary mouse myoblast purification, characterization, and transplantation for cell-mediated gene therapy. J Cell Biol 125:1275–1287 10. Hindi L, McMillan JD, Afroze D et al (2017) Isolation, culturing, and differentiation of primary myoblasts from skeletal muscle of adult mice. Bio Protoc 7 11. Shahini A, Vydiam K, Choudhury D et al (2018) Efficient and high yield isolation of myoblasts from skeletal muscle. Stem Cell Res 30:122–129 12. Collins CA, Olsen I, Zammit PS et al (2005) Stem cell function, self-renewal, and behavioral heterogeneity of cells from the adult muscle satellite cell niche. Cell 122:289–301 13. Montarras D, Morgan J, Collins C et al (2005) Direct isolation of satellite cells for skeletal muscle regeneration. Science 309:2064–2067 14. Sacco A, Doyonnas R, Kraft P et al (2008) Selfrenewal and expansion of single transplanted muscle stem cells. Nature 456:502–506 15. Conboy MJ, Cerletti M, Wagers AJ et al (2010) Immuno-analysis and FACS sorting of adult muscle fiber-associated stem/precursor cells. Methods Mol Biol 621:165–173

Chapter 13 Isolation, Culture, Cryopreservation, and Identification of Bovine, Murine, and Human Spermatogonial Stem Cells Pedro M. Aponte Abstract Spermatogonial stem cells (SSCs) are the germ cells at the basis of spermatogenesis in adult mammals. SSCs offer many biotechnological possibilities and are fundamental cells in the study of spermatogenesis (Aponte, World J Stem Cells 7:669–680, 2015). This chapter describes detailed procedures for SSC isolation, culture, cryopreservation, and characterization in bovine, murine, and human models. Key words Spermatogenesis, Spermatogonial stem cells, Testis, Isolation, Cell culture, Characterization, Cryopreservation, Functional test

1

Introduction Spermatogenesis is the complex and highly coordinated process leading to sperm generation in the testis. As a stem cell system, spermatogenesis in adult mammals is based on germ stem cells called spermatogonial stem cells (SSC) [1, 2]. They belong to the male germ line and generate the first wave of spermatogenesis in prepubertal animals [3]. After puberty, they maintain the steady state of sperm production throughout life. SSCs are located in the basal lamina of seminiferous tubules close to the blood vessels present at the interstitium. They are intermingled with their morphologically identical and committed to differentiation progenitors type A spermatogonia. SSCs offer many biotechnological possibilities and are key in the study of spermatogenesis [1]. This chapter shows procedures for isolation, culture, cryopreservation, and identification of SSCs in three models: bovine, mice, and humans.

Chrissa Kioussi (ed.), Stem Cells and Tissue Repair: Methods and Protocols, Methods in Molecular Biology, vol. 2155, https://doi.org/10.1007/978-1-0716-0655-1_13, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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Materials All solutions are prepared by using ultrapure water and analytical/ cell culture grade reagents. 1. MEM including nonessential amino acids, 0.13% sodium bicarbonate, 4 mM L-glutamine, penicillin (100 iU/mL)–streptomycin (100 μg/mL), 0.04 mg/mL gentamicin, 15 mM HEPES (see Note 1). 2. MEM 1 with DNase. 3. Enzyme mix I: collagenase I 2 mg/mL, hyaluronidase 2 mg/ mL, and trypsin 2 mg/mL (see Note 2). 4. Enzyme mix II: collagenase I 2 mg/mL and hyaluronidase 2 mg/mL (see Note 2). 5. MEM 1 (90% v/v), BSA (1% w/v), DNase (2 mg/mL) (see Note 3). 6. Percoll discontinuous gradient preparation. Starting solution: 82.2% Percoll, BSA (0.7%), DNase I (0.05 mg/mL) in MEM 10 (final MEM 10 concentration 9.76% v/v). Filter solution with a 0.22 μm syringe filter. Diluent solution: 45.25 mL MEM 1 (final concentration 90.5% v/v), BSA (0.7%), DNase I (0.05 mg/mL). Follow the directions of the Table 1, starting from the most concentrated solution (82.2%).

2.1

MEM-BSA

2.2 Mouse Type A Spermatogonia Isolation Enzymes

MEM and 0.1% BSA. 5 μg/mL DNase I, 1 mg/mL hyaluronidase, 1 mg/mL trypsin, and 1 mg/mL collagenase (see Note 5).

2.2.1 Mix I 2.2.2 Mix II

2.3

SSC Medium

5 μg/mL DNase I, 1 mg/mL hyaluronidase, and 1 mg/mL collagenase (see Note 5). 1. A defined SSC medium was originally designed by KanatsuShinohara et al. [4]. The medium consists of StemPro-34 serum free medium (SFM) supplemented with StemPro supplement, 25 μg/mL insulin, 100 μg/mL transferrin, 60 μM putrescine, 30 nM sodium selenite, 6 mg/mL D-(1)-glucose, 30 μg/mL pyruvic acid, 1 μL/mL DL-lactic acid, 5 mg/mL bovine albumin, 2 mM L-glutamine, 5  105 M 2-mercaptoethanol, MEM Vitamin Solution (100), MEM nonessential amino acid solution, 104 M ascorbic acid, 10 μg/mL D-biotin, 30 ng/mL β-estradiol, and 60 ng/mL

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Table 1 Preparation of Percoll solutions for the generation of a discontinuous gradient (see Note 4) %

Percoll (mL)

Diluent (mL)

1

65

15.8 (of 82.2%)

4.2

2

50

15.4 (of 65%)

4.6

3

40

16 (of 50%)

4

4

36

14.4 (of 40%)

1.6

5

34

12.3 (of 36%)

0.7

6

32

9.4 (of 34%)

0.6

7

30

6.6 (of 32%)

0.4

8

28

4.6 (of 30%)

0.4

9

20

2.1 (of 28%)

0.9

progesterone. Growth factors are added at the following concentrations: epidermal growth factor (EGF) 20 ng/mL, basic fibroblast growth factor (bFGF2) 10 ng/mL, leukemia inhibitory factor (LIF) 100 ng/mL, and glial cell line-derived neurotrophic factor (GDNF) 40 ng/mL (all human recombinant) [5]. 2. There are some species-specific modifications on the use of growth factors with respect to the above general protocol for bulls. In mice, growth factors should be of rodent origin whenever possible and GDNF concentration can be 10 ng/ mL [4]. For humans, growth factors of human origin can easily be found. GDNF can be added at 10 ng/mL and LIF at 10 ng/mL [6]. 3. Mix the constituents with Stem cell Pro and filter through a bottle-top filter unit. 2.4 Bouin’s Fluid (Cell Fixation)

3

70% saturated picric acid, 25% formaldehyde (37%), 5% acetic acid (glacial) (see Note 6).

Methods

3.1 Type A Spermatogonia Primary Isolation

As SSCs are a subpopulation of type A spermatogonia, protocols should aim to first isolate the later. A good yield of type A spermatogonia can be obtained from prepubertal animals, during spermatogonial proliferation, before spermatocytogenesis (meiosis) of the first wave of spermatogenesis. As the wave progress, more differentiated cells start to populate the testis and “dilute” the SSCs. This depends on animal species, breed, and, obviously, age (Table 2).

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Table 2 Recommended age for type A spermatogonia isolation Species

Developmental premeiosis phase

Mouse

8–9 days postnatal [7]

Bulls (Bos taurus)

12–16 weeks [8]

Bulls (Bos indicus)

36–44 weeks [9]

Humans

6 months–9 years [10]

3.2 Criteria for Type A Spermatogonia Morphological Identification

Type A spermatogonia are large, round cells with a spherical nucleus, a sharp nuclear membrane, several (1–3) relatively large nucleoli, a high nucleus–cytoplasm ratio, and many cytoplasmic inclusions mostly concentrated at one side of the cell [11].

3.3 Type A Spermatogonia Isolation in Bovine Models

Testis material can be obtained from slaughterhouses or through castration. 1. After excision from the animal, testes should be transported to the laboratory in an ice box (~4  C). This temperature will preserve most germ cells for at least 3 h. 2. In the lab, testes should be rinsed in tap water in order to remove gross contaminating material.

3.3.1 Testis Dissection

Work with gloves in a clean area. 1. For the dissection, use scissors to open and remove the external investments of the testis (from outer to inner: external spermatic fascia, parietal layer of vaginal tunic and visceral layer). 2. Rinse the testes in sterile 0.9% NaCl isotonic solution to further remove blood and debris. At this time, each testis should be placed on a sterile petri dish (usually 100  15 mm, depending on testis size) and weighed. 3. Transfer the testis to the culture room in a closed petri dish. 4. Preliminary to further steps, thaw DNase I and turn on the laminar flow and water bath. DNase I will prevent cells or tissue pieces to stick to each other or to the walls of the tubes or vials. Thoroughly mix all frozen-thawed reagents. The protocol for type A isolation is performed in 2 days.

3.3.2 Isolation Part I, DAY 1 (See Note 7)

1. Weight about 20 g of testis for further processing and take a sample for histological examination from the remaining testis material (fix in neutral buffered formalin or Bouin’s fluid, as preferred). After weighing transfer, the testis material into a sterile petri dish and cut it in 2 similar-sized pieces. Place each

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piece in a separate new sterile petri dish and add 12.5 mL MEM + DNase. 2. Remove the tunica albuginea with the help of a pincette and a surgical blade and mince the testis with two surgical blades while trying to remove all the connective tissue present in the mediastinum and prepare the seminiferous tubules as much as possible. This process should take no longer than 10 min to avoid excessive autolysis of the tissue. 3. After mincing all the material (20 g), transfer all of it into two 50 mL conical tubes (10 g each) and add 12.5 mL of enzyme Mix I to each tube. (There should be a volume of near 25 mL of tissue + enzyme mixture I on each tube). 4. Close the tubes and after tightly wrapping the cap with Parafilm, transfer them into a shaking water bath, 140 cycles/min for 60 min at 32  C. Avoid that the tubes float on the surface of the water bath by lowering them deep into the water with a heavy object (see Note 8). 5. Stop the water bath and centrifuge the tubes for 1 min at 300 rpm. Remove the supernatant and rinse with MEM + DNase. Repeat the process three times to yield a clear supernatant. 6. Remove the supernatant with a pipette controlling device or aspiration pump device until the 12.5 mL mark in the tube and add 12.5 mL enzyme Mix II to each tube. 7. Close the tubes and tightly cover the cap with Parafilm. Similarly as in the first enzymatic digestion, transfer the tubes into the shaking water bath at 32  C and 140 cycles/min, this time for 45 min. At this time a tube containing FCS can be thawed to be used in further steps. 8. After the second enzymatic digestion, centrifuge for 2 min at 400 rpm. Collect the supernatant from both tubes through 77 (first) and 55 μm (second) nylon filters (strainers) into an individual 50 mL tube and centrifuge it for 5 min at 900 rpm. 9. Aspirate the supernatant as much as possible and resuspend the pellet in 4 mL MEM + DNase+BSA and place on ice. 10. Measure the purity (Nomarski microscope, add 10 μL of cell suspension to a slide and cover with a coverlid and register type A spermatogonia in 100 cell counts to obtain the percentage), viability (commercial live and dead kit, according to manufacturer instructions) and concentration (with a Neubauer counting chamber using 7 μL of cell suspension). 11. Dilute the cell suspension in 10% FCS in MEM 1, add 5 mL FCS and 4 mL cell suspension in a tube with 41 mL MEM.

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12. Fill each of four 75 cm2 culture flasks with 12.5 mL of cell suspension. Gently shake the flasks in different directions to evenly spread the cells in the bottom culture surface, avoiding leaving dry areas. Loosely cap the flasks (unless the caps are equipped with ventilation openings) and take them inside CO2 incubators (with humid atmosphere at 37 C, 5% CO2) overnight. 3.3.3 Isolation Part II, DAY 2

1. After overnight culture, shake each flask by sliding it back and forth over the laminar flow cabinet work surface and collect the cells with a pipette controller (using 10 mL pipettes). Rinse the cultured surface of the flask with its own medium 2 or 3 times before transferring them to 50 mL conical tubes and then centrifuge 5 min at 900 rpm (see Note 9). 2. Aspirate the supernatant as much as possible and resuspend the pellet in 0.6–0.7 mL MEM/DNase/BSA on a vial and place it on ice. 3. Measure the purity, viability, and concentration as previously). 4. Prepare one Percoll gradient for each cell suspension (see Note 10). 5. Take nine 1.5-mL centrifuge plastic vials and number them from 1 to 9. Fill 1 mL of each Percoll solution concentration into the vials (from higher concentration to lower, starting with tube 1 up to tube 9). The Percoll solutions should have been prepared in advance. The concentrations are: 65%, 50%, 40%, 36%, 34%, 32%, 30%, 28%, and 20%. The gradient can be built up with a peristaltic pump or manually, gently letting the solutions slide through the tube walls, with 1 mL of each Percoll concentration/density. 6. Use a 10 mL polycarbonate tube and sequentially load the Percoll solutions from the 1.5-mL vials to the gradient tube, from the highest concentration (bottom ¼ tube 1) to the lowest (top ¼ tube 9), as shown in Fig. 1. If using a pump, wash the tubing system with 1 mL ethanol followed by 1 mL distilled water before building the gradient. After use, wash in the opposite sense (1 mL distilled water followed by 1 mL ethanol). 7. For the Percoll separation, gently load the cell suspension (0.6–0.7 mL) on the top of the column, through the tube wall using a pipette (see Note 11) and centrifuge the column for 30 min at 800  g. at a refrigerated centrifuge (4  C). 8. Gently, take the tube out of the centrifuge and mark the fractions in the tube with a marker for easier visualization. Cells found in the interface between the density solutions should be considered as fractions 1–10. Gently collect fractions 1–5 in different 1.5 mL vials using a 100–1000 μL micropipette.

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Fig. 1 Fractions in the column of the Percoll discontinuous gradient and most probable location of the population of type A spermatogonia (including the morphologically identical subpopulation of spermatogonial stem cells—SSCs) after centrifugation

9. Add MEM–BSA to each cell suspension until de 1.5 mL mark on the vial, mix and centrifuge for 5 min at 7168  g. 10. Aspirate the supernatant with a micropipette and leave about 100 μL to avoid disturbing the pellet. Resuspend in 1 mL MEM–BSA and centrifuge for 5 min at 2800  g. 11. Carefully aspirate all the supernatant with a micropipette, being careful not to lose the pellet and depending of the size of the pellet, resuspend in 150–500 μL of MEM–BSA. 12. Assess the purity of the different fractions using the Nomarski microscope. Only fractions with more than 50% purity will be counted for concentration and viability assessment and used for culture. Usually type A spermatogonia will be in fraction 3 (Fig. 1). 3.4 Type A Spermatogonia Isolation in Mouse Models

Mouse are castrated or sacrificed to obtain the testes. Because of their small sizes, testes from at least 2–4 animals should be used. All washing steps and enzymatic digestions are done in MEM (see MEM recipe in the Subheading 2). 1. Decapsulate the testes, and dissect the tubules apart with forceps in a petri dish with MEM containing 5 μg/mL DNase I. 2. After washing three times by letting the suspension to sediment (in a 50 mL tube), removing the supernatant and adding new MEM, the tubule fragments are incubated in proteolytic enzymes. Place the testes in 20 mL MEM containing mouse proteolytic enzyme mixture I (see Subheading 2) for 15 min.

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3. As mentioned before, wash the tubular fragments (three times) by repeatedly pipetting them in fresh medium (MEM) and letting them sediment. 4. Perform a second incubation in mouse proteolytic enzyme mixture II in MEM for 30 min. 5. Pipette the tubule fragment suspension in and out several times, and centrifuge at 30  g for 2 min. 6. Collect the supernatant and filter it through 77- and 55-μm cell strainers in one 50 mL-tube (as described for bull type A spermatogonia isolation procedure). Perform all incubations (enzymatic digestions) in a shaking water bath (60–80 cycles/ min) at 32  C (more details in the bull type A spermatogonia isolation procedure). 7. Build a Percoll gradient as described for bull type A spermatogonia isolation. The tube with the column should be centrifuged at 800  g for 30 min at 18  C. Mouse type A spermatogonia will be located in fractions 3 and 4 (Fig. 1). Evaluate purity, survival and concentration of the obtained cell suspensions. 3.5 Type A Spermatogonia Isolation in Human Models

1. Human testes can be obtained from donations (castration or intact testes from corpses after accidents). Exercise appropriate biosafety steps during cell manipulation as contagious human pathogens can be located in the human testis (see Note 12). 2. The isolation procedure is similar to that of bulls and mice. Small tissue pieces (100–200 mg) are subject to enzymatic digestion similarly as with bovine testicular tissues.

3.6 Type A Spermatogonial Culture 3.6.1 Type A Spermatogonia Culture in Bovine, Mouse, and Human Models

Final cell suspensions (after isolation) are temporarily kept in MEM–DNaseI–BSA at 4  C. 1. Estimate viability and purity. With a 7 μL cell suspension sample estimate general cell suspension concentration and through the cell suspension volume estimate the total number of viable type A spermatogonia (total number of viable type A spermatogonia ¼ volume  concentration in cells/μL, corrected for purity and viability). Only cell suspensions with more than 50% viable type A spermatogonia are advised to be used for culturing. 2. Estimate the number of wells to be seeded. For regular differentiation essays basic culture medium is MEM + 10% FCS and amphotericin B 1:1000. Seeding density is (viable type A spermatogonia) for bulls 4  104 spermatogonia/cm2 (short term) and 2  104 spermatogonia/cm2 (long term). For mice, seed at 12  104 cells cm2. For humans, use 10  102 to 20  102 cells/cm2. Cells can be seeded in uncoated dishes. For optimal

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maintenance of SSCs a special medium should be used that includes SSC self-renewal basic medium with growth factors, hormones, and FCS (or serum replacement agent). FCS is added at 1–2.5% for SSC self-renewal and higher (typically 10%) for SSC differentiation (this will also stimulate the proliferation of contaminating somatic cells) or alternatively KO serum replacement. For appropriate maintenance SSCs should be cultured in an SSC medium (see Subheading 2). 3. Culture the cells in 4-well chamber slides of 2 cm2/well at a species-specific under-body temperature (bovine 32–37  C mouse and humans 37  C in a humidified atmosphere with 5% CO2 and the medium refreshed twice a week (see Note 13). Under the culture conditions presented in this protocol, contaminating somatic cells (mostly Sertoli cells) will form a monolayer and contact inhibition will not allow the cells to be cultured for more than 2 weeks (short term culture). During the 2 weeks period germ–somatic cells colonies may appear. These colonies can be assessed by aspirating them with a 100 μL micropipette with a slightly bent tip, fixing them and histologically processing them in histo-cassettes with small gauge net (biopsy grade). Long-term culture requires subculturing. Passaging is done with trypsin-EDTA (0.25%). Cells should be passaged 1:2–1:4 every 3–5 days, depending on the rate of somatic cell proliferation. Cells need to be subcultured just before 100% confluency to avoid contact inhibition and subsequent detachment of the monolayer from the culture plate [5]. Mouse spermatogonia may require longer passaging times (i.e., 10 days) [4]. For human type A spermatogonia cultures can be passaged every 7–10 days at 80–90% confluency starting at 1:1 [6]. For mouse, BSA is added (5 mg/mL) to the SSC culture medium [4]. Somatic cells support the maintenance of SSCs. If somatic cells disappear from the culture SSCs can be maintained on a mitomycin C–inactivated mouse embryonic fibroblasts (MEF) feeder layer [4]. For human SSC propagation, passaged germ cell clusters are placed in cultured plates coated with laminin (20 μg/mL) [6]. 3.7 SSC Functional Assays

In order to know a close estimate of the SSCs present in a cell suspension, a functional assay should be done that involves SSC transplantation. Type A spermatogonial population containing the SSC subpopulation is transplanted to an immunocompromised mouse strain previously cleared from endogenous germ cells via local irradiation or chemotherapy. Only vera SSC are able to home in (attach to seminiferous tubules basal lamina, in the basal compartment) and if SSC from donor and host are related, colonization and spermatogenic outburst will occur along the seminiferous

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tubules of transplanted animals. Individual cells or colonies can be counted through unbiased stereological counting methods (i.e., the dissector) [12]. Cell suspensions resulting from isolation or frozen-thawed cells are transplanted into testes of NMRI nude mouse or other, immune compromised mouse strain via the efferent duct. 1. Deplete endogenous spermatogenesis through radiation. Mice receive a fractionated dose of 1.5 and 12 Gy of X-rays directed to the testis area (mice should be anesthetized by a veterinarian or other licensed professional) with an interval of 24 h. Alternatively, use Busulfan. This drug can be dissolved in DMSO and diluted 1:1 in distilled sterile water for injection at a dose of 30 mg/kg (0.3 mg/10 g) IP [13] or alternatively injected intratesticular at a dose of 6 mg/kg (0.06 mg/10 g) [14]. Dissolve with sterile distilled water as needed to reach the appropriate concentration. 2. One month after irradiation or cytotoxic chemical treatment, testes of recipient mice require a surgery (veterinarian or other licensed professional) in which the testes are exteriorized through an incision of the abdomen midline. 3. Transplant the donor cells. A cell suspension containing 5  103 to 20  103 cells/μL or 105 to 4  105 cells in 20 μL is transplanted per testis (doses per testis ¼ 20 μL). If cells come from cell culture plates are collected by treating them with trypsin-EDTA (0.25%) for approximately 15 min at 37  C. After repeated pipetting to break cell attachments, trypsin must be inactivated by addition of 10% FCS (in MEM). The cell suspension to be injected contains trypan blue to better visualize the cell suspension progress into the seminiferous tubules. They are injected with pulled siliconized glass capillary tubes (using a micropipette puller, according to the manufacturer’s directions). After pulling, the capillaries are autoclaved and before transplantation, the tip of the pulled capillary tube is broken with tweezers and the tip stained with a permanent marker for easier visualization. The nonpulled end of the cell suspension loaded glass capillary can be attached to a 1 mL syringe with a syringe to glass capillary accessory available in the market. A comprehensive description of the procedure is described by Ogawa et al. [15]. The contralateral nontransplanted testis can be used as the negative control. 4. Subjects are sacrificed or castrated after the time of specific duration of spermatogenesis to guarantee the occurrence of one round of spermatogenesis (duration of spermatogenesis in bull ¼ 61 days [16], mouse ¼ 34.5 days [17], human ¼ 74 days [18]).

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5. Testes must be immediately fixed with paraformaldehyde 4% in PBS at 4  C for 24 h and/or Bouin’s fluid for 48 h. Gently rocking of the samples during fixation is advised whenever possible. After the fixation period, samples can be kept in ethanol 70% (v/v) at 4  C. 6. Characterize transplanted/colonizing SSCs through specific markers will demonstrate their presence at the end of functional tests. Animals with great phylogenetic distance with mice (i.e., humans or bulls) will have SSCs that only home in on the mouse seminiferous epithelium but do not proliferate. SSCs from none related animals (i.e., mice and rats) will produce spermatogenic colonies in the transplanted mouse testis. Each colony can be assumed to be originated from one SSC. Therefore, counting colonies will provide an indication of the original SSC number. Cell characterization can be done through immunohistochemistry or PCR. Some specific markers are mentioned in the following section. 3.8 Molecular SSC Characterization

1. Cells in culture may change their shape. Therefore, morphological recognition can be complex. Table 3 summarizes markers that can be used to characterize cells appearing in testicular cell cultures. 2. Immunohistochemistry can be performed directly in the culture plate wells or alternatively cells can be grown in round coverlids inside the wells or chamber slides. 3. Medium has to be removed and the cells rinsed 2–3 times with PBS to remove serum (in case it has been used).

Table 3 Markers for cells likely to be present in testicular cell culturesa Origin

Cell type

Marker

Somatic

Sertoli cells

Vimentin Sox9 Sulfated glycoprotein 2 GATA1 Alpha smooth muscle actin 3-beta-HSD

Peritubular myoid cells Leydig cells Germinal

Germ cells (general) Type A spermatogonia Spermatogonial stem cells

a

Markers can be used for immunohistochemistry or qPCR Specific for bovine species

b

DDX4 (VASA) DBA (lectin)b PGP9.5 Gfr-alfa1 Zbtb16 (PLZF) NEUROG3

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4. Cells should be immediately fixed with paraformaldehyde 4% in PBS at 4  C for 10 min. If immunohistochemistry is not to be performed immediately, culture plates or chamber slides can be filled with ethanol 70% v/v and stored at 4  C. 3.9 SSC Cryopreservation

1. Freezing and thawing protocols for bovine spermatogonia have been described [19]. Bovine type A spermatogonia population (containing SSCs) can be cryopreserved in MEM medium containing 20% FCS, 20% DMSO, and 0.14 M sucrose using a non–controlled rate freezing protocol. 2. Vials containing cells are kept on ice. Cells are added the freezing medium dropwise divided in 3 aliquots (waiting 5 min between aliquots, mixing manually and gently every 3 drops) always on ice or inside a cool room. 3. For human or mice, sucrose can be left out with no effect on type A spermatogonial survival. 4. Cells are placed in cryovials (3  106 cells/mL/vial, from which approximately 50% will survive after thawing) and stored at 80  C for 72 h (or at least one night). After that time, cells can be stored in liquid nitrogen at 196  C.

4

Notes 1. From the reagents above, L-glutamine and penicillin–streptomycin should be aliquoted in sterile conic tubes (10 mL and 5 mL, respectively) and kept at 20  C. Gentamicin can be stored at RT. The rest of the reagents should be kept at 4–10  C. 2. Weight in sterile weighing paper. Keep the enzyme mixtures preweighted and identified in a sterile 50 mL tube at 18  C until 15 min before the time of use. 3. BSA can be prepared in advance. Add 10% w/v BSA in sterile MQ water. Let it sit for about 5 min. Mix with magnetic stirring until completely dissolved. Pass the solution through a 0.22 μm syringe filter, aliquot in vials and freeze (18  C). 4. Always preload the tubes (50 mL capacity) with diluents (quantities in Table 1 column “diluent”). If necessary, solutions can be stored overnight at 4  C. 5. Keep the enzyme mixture preweighted in a sterile 15 mL tube at 18  C. 6. For better morphological results, acetic acid glacial (that enhances details in nuclear chromatin) should be added fresh. The use of commercially available Bouin’s fixative is perfectly all right.

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7. Work with gloves in the laminar flow. 8. The temperature of the water bath should always be lower than the species body temperature (in this case bull’s 32  C vs. body temperature of 39  C) not to disturb spermatogenesis). 9. This step is called differential plating. Most somatic cells attach to the bottom of the flask while germ cells remain loose and are collected to proceed with the isolation procedure. 10. It is important to use polycarbonate tubes with Percoll (the constituent silica particles will not adhere to the walls of these tubes) to ensure the best possible purification. Recommended tube features are as follows: round bottom; 100 mm long; outer diameter 17 mm, inner diameter 14 mm. 11. Avoid disturbing the column. Hold the tube tight to a supporting rack with Parafilm plugs. 12. The testis is an immune sanctuary and several pathogens are able to elude the immune surveillance locally in the testis. 13. At the start of the cultures, during refreshing, aspiration of the medium has to be very gentle since germ cells are not yet adherent and can be easily lost. Gelatin coating (0.2%, w/v) is optional.

Acknowledgments This work was supported by the Chancellor Grant 2016, project 4347, Universidad San Francisco de Quito, USFQ. References 1. Aponte PM (2015) Spermatogonial stem cells: current biotechnological advances in reproduction and regenerative medicine. World J Stem Cells 7:669–680. https://doi.org/10.4252/ wjsc.v7.i4.669 2. de Rooij DG, Griswold MD (2012) Questions about spermatogonia posed and answered since 2000. J Androl 33:1085–1095. https:// doi.org/10.2164/jandrol.112.016832 3. Sung WK, Komatsu M, Jagiello GM (1986) The kinetics of the first wave of spermatogenesis in the newborn male mouse. Gamete Res 14:245–254. https://doi.org/10.1002/mrd. 1120140308 4. Kanatsu-Shinohara M, Ogonuki N, Inoue K et al (2003) Long-term proliferation in culture and germline transmission of mouse male germline stem cells. Biol Reprod 69:612–616. https://doi.org/10.1095/biolreprod.103. 017012

5. Aponte PM, Soda T, Teerds KJ et al (2008) Propagation of bovine spermatogonial stem cells in vitro. Reproduction 136:543–557. https://doi.org/10.1530/REP-07-0419 6. Sadri-Ardekani H, Mizrak SC, van Daalen SKM et al (2009) Propagation of human spermatogonial stem cells in vitro. JAMA 302:2127–2134. https://doi.org/10.1001/ jama.2009.1689 7. Bellve´ AR, Cavicchia JC, Millette CF (1977) Spermatogenic cells of the prepuberal mouse. Isolation and morphological characterization. J Cell Biol 74:68–85. https://doi.org/10. 1083/jcb.74.1.68 8. Curtis SK, Amann RP (1981) Testicular development and establishment of spermatogenesis in Holstein bulls. J Anim Sci 53:1645–1657. https://doi.org/10.2527/jas1982.5361645x 9. Aponte PM, de Rooij DG, Bastidas P (2005) Testicular development in Brahman bulls.

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Theriogenology 64:1440–1455. https://doi. org/10.1016/j.theriogenology.2005.03.016 10. Nistal M, Paniagua R, Gonza´lez-Peramato P et al (2015) Perspectives in pediatric pathology, chapter 3. Testicular development from birth to puberty: systematic evaluation of the prepubertal testis. Pediatr Dev Pathol 18:173–186. https://doi.org/10.2350/12-09-1255-PB.1 11. van Pelt AM, Morena AR, van Dissel-Emiliani FM, Boitani C et al (1996) Isolation of the synchronized A spermatogonia from adult vitamin A-deficient rat testes. Biol Reprod 55:439–444. https://doi.org/10.1095/ biolreprod55.2.439 12. Geuna S, Herrera-Rincon C (2015) Update on stereology for light microscopy. Cell Tissue Res 360:5–12. https://doi.org/10.1007/s00441015-2143-6 13. Wang D-Z, Zhou X-H, Yuan Y-L et al (2010) Optimal dose of busulfan for depleting testicular germ cells of recipient mice before spermatogonial transplantation. Asian J Androl 12:263–270. https://doi.org/10.1038/aja. 2009.67 14. Qin Y, Liu L, He Y et al (2016) Testicular Busulfan injection in mice to prepare recipients

for spermatogonial stem cell transplantation is safe and non-toxic. PLoS One 11:e0148388. https://doi.org/10.1371/journal.pone. 0148388 15. Ogawa T, Are´chaga JM, Avarbock MR et al (1997) Transplantation of testis germinal cells into mouse seminiferous tubules. Int J Dev Biol 41:111–122 16. Staub C, Johnson L (2018) Review: spermatogenesis in the bull. Animal 12:s27–s35. https://doi.org/10.1017/ S1751731118000435 17. Oakberg EF (1956) Duration of spermatogenesis in the mouse and timing of stages of the cycle of the seminiferous epithelium. Am J Anat 99:507–516. https://doi.org/10.1002/aja. 1000990307 18. Heller CH, Clermont Y (1964) Kinetics of the germinal epithelium in man. Recent Prog Horm Res 20:545–575 19. Izadyar F, Matthijs-Rijsenbilt JJ, den Ouden K et al (2002) Development of a cryopreservation protocol for type A spermatogonia. J Androl 23:537–545

Chapter 14 Long-Term Ex Vivo Expansion of Murine Spermatogonial Stem Cells in a Simple Serum-Free Medium Hiroshi Kubota and Kazue Kakiuchi Abstract Spermatogonial stem cells (SSCs) possess both self-renewal and differentiation abilities to sustain lifelong production of enormous numbers of spermatozoa in males. SSCs hold a unique position among tissuespecific stem cells in adults because of their ability to transmit the genetic information to subsequent generations. Ex vivo expansion of SSCs in conjunction with their transplantation is highly invaluable to study SSCs and develop new reproductive technologies for therapeutic applications. In this chapter, we describe a culture system involving a simple serum-free medium for mouse SSCs. Elimination of the serum from the culture is important to enhance the effects of exogenous factors, which are rather masked by the serum, and to avert the serum-induced inflammatory responses of testicular mesenchymal cells, which cause adverse effects on SSC proliferation. Consequently, using this culture system has proven for the first time that glial cell line–derived neurotrophic factor (GDNF) was found to be the key factor to drive the selfrenewing proliferation of SSCs, and fibroblast growth factor 2 enhanced the GDNF-dependent proliferation of SSCs. Besides determining these two key cytokines, the simplicity of the system enabled individual modification of its components to develop long-term cultures of rat and rabbit SSCs. The basics of these culture systems will enable development of the culture conditions for human and other mammalian SSCs in the near future. Key words Spermatogonial stem cells, Germline stem cells, Stem cell spermatogonia, Serum-free medium, Feeder cells, Stem cell culture, Spermatogonial transplantation, Self-renewal, Spermatogenesis, Testis

1

Introduction The role of spermatogenesis is to generate male gametes for fertilization, resulting in reproduction, which ensures the continuation of the species. Spermatogenesis is highly complex, but wellorganized, and is thought to be one of the most productive cellrenewing systems in adult tissues [1, 2]. This high productivity relies on spermatogonia, which comprise a stem cell subpopulation called spermatogonial stem cells (SSCs) or stem cell spermatogonia, and a transit-amplifying (TA) cell subpopulation, which is composed of several spermatogonia subtypes [3]. SSCs can self-renew

Chrissa Kioussi (ed.), Stem Cells and Tissue Repair: Methods and Protocols, Methods in Molecular Biology, vol. 2155, https://doi.org/10.1007/978-1-0716-0655-1_14, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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themselves and produce differentiation-committed spermatogonia that continuously supply and replenish the TA cell compartment. Although there are very few SSCs in the seminiferous tubules, development of the SSC transplantation technique allowed researchers to identify SSCs unequivocally [4]. By this technique, the biological competence of a single germ cell can be assessed and it can be concluded that the cell is an SSC if it can both self-renew and give rise to spermatozoa. The SSC transplantation technique was first reported in 1994 using mice [5, 6]. Considerable research has been performed in the last 25 years using this technique as a functional assay for SSCs, and most of our knowledge about SSCs, including their niche, has been obtained from these research [7]. One of the most remarkable achievements with the help of the SSC transplantation technique is the development of culture systems that support ex vivo expansion of SSCs. Successful SSC culture in addition to transplantation has made a tremendous impact in the field of germ cell biology. Notably, SSCs are the only cells that undergo self-renew and transmit genetic information to subsequent generations; thus, they are an ideal target cell to manipulate the germline of organisms, including laboratory and domestic animals [8]. Furthermore, SSCs can be cryopreserved. Even after 14 years of storage in liquid nitrogen, mouse SSCs can be transplanted with complete reconstitution of spermatogenesis, producing functional sperms that generate normal offspring [9]. This observation suggests that cryopreservation of SSCs can immortalize the germline of an individual male. One of the possible clinical applications of SSCs is restoration of fertility after cancer therapy in prepubertal boys with cancer [10]. For such patients, testisbiopsy containing SSCs is cryopreserved before the anticancer treatment, such as chemotherapy or irradiation, which could destroy SSCs leading to infertility. After successful treatment, SSCs could be isolated from the testisbiopsy and transplanted into the testes of the patients. Ex vivo expansion of SSCs would be crucial to increase their number and eliminate the cancer cells before transplantation; however, long-term culture systems for human SSCs remain to be developed. There are mainly two types of culture methods for mouse SSC; one uses a complex medium containing animal serum, serumreplacements, or proprietary serum-derived supplements [11], and the other uses a simple serum-free medium, in which all components are known [12]. Although SSCs of some mouse strains do not proliferate by the former method, the latter supports long-term proliferation of SSCs isolated from all the strains examined. In this chapter, we describe a method for continuous expansion of mouse SSCs in a simple serum-free medium. The culture system consists of a serum-free medium and mitotically-inactivated feeder cells [13]. Elimination of the serum from the medium has two benefits: First, compared with the serum-supplemented conditions, serum-

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free conditions enhance the effects of exogenous factors, such as hormones and cytokines, which are rather masked by the serum [14]. Second, because serum induces production of inflammatory signals from the cells in culture, removal of serum averts the seruminduced inflammatory responses of testicular mesenchymal cells, which cause adverse effects on self-renewal and proliferation of SSCs [15]. The key for self-renewing expansion of SSCs in culture is supplementation of glial cell line–derived neurotrophic factor (GDNF) [12]. Although the beneficial effect of GDNF on spermatogonial proliferation was originally demonstrated using transgenic mice [16], conclusive evidence for its direct role on SSC selfrenewal has been demonstrated by the serum-free culture system in conjunction with transplantation [12]. In addition, SSCs from some mouse strains require a low amount of fibroblast growth factor 2 (FGF2) for their efficient proliferation in addition to GDNF [12]. The two factors are expressed in seminiferous tubules, and especially GDNF is crucial to maintain spermatogonial population in vivo [16, 17]; therefore, they likely function as key components of the SSC niche. In addition to the soluble compartment of the culture system, feeder cells are the last crucial piece. The SNL76/7 (SNL) cell line, a derivative of the mouse embryonic fibroblast cell line STO [18], was used as the feeder layer instead of primary mouse embryonic fibroblasts (MEFs), which are broadly used for various stem cell cultures, including the other SSC culture systems [11]. Using feeders of a cloned cell line minimizes variability of the culture conditions. The remarkable feature of our culture system is its simplicity, which enables individual modification of the culture components, and it has thus become possible to develop long-term rat and rabbit SSC cultures [19, 20]. Interestingly, the culture system was originally developed for rodent and human somatic stem/progenitor cells [21–23]. Thus, a common rule may underlie the culture systems for ex vivo self-renewal of stem cells. Functional understanding of the culture system’s constituents will provide invaluable information for the prospective long-term cultures of SSCs derived from other mammals, including valuable livestock, endangered animals, and humans.

2 2.1

Materials SNL Feeder Cells

1. SNL76/6 cells (European Collection of Authenticated Cell Cultures, 07032801; ATCC; SCRC-1049; Cell_Biolabs, CBA-316) (see Note 1). 2. SNL expansion medium (10F-DMEM): High glucose Dulbecco’s Modified Eagle Medium (DMEM), 10% fetal bovine

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serum (FBS), 2 mM L-glutamine, 50 units/mL penicillin, 50 μg/mL streptomycin. 3. Mitomycin C (MMC): Dissolve MMC powder with sterilized MilliQ water to make 0.5 mg/mL solution. Divide in sterile cryovials and store at 80  C. 4. Dulbecco’s phosphate buffered saline without Mg and Ca (PBS). 5. Trypsin–EDTA (0.05%): Dilute 0.5% trypsin–4.8 mM EDTA (10) with PBS. 6. Freeze medium (2): 80% 10F-DMEM, 20% DMSO. 7. Freezing container (e.g., Nalgen, 5100-0001). 8. CELLBANKER® 1 plus (Zenoaq, CB021). 9. Gelatin solution (0.1%): Dissolve gelatin (from porcine skin), Type A (Sigma-Aldrich, G2500) in MilliQ water and sterilize by autoclaving. 10. Methylene blue staining solution: 2% methylene blue in 60% methanol. 2.2 Preparation of Testicular Cells and Culture of Spermatogonial Stem Cells

1. 0.25% trypsin–EDTA (1) (Thermo Fisher Scientific, 25200). 2. DNase I solution (7 mg/mL): Dissolve DNase I (SigmaAldrich, DN25) in PBS and sterilize by filtration with a 0.22μm membrane. 3. Serum-free medium for mouse SSCs (mSFM): Tables 1 and 2. 4. GDNF (10 μg/mL): Reconstitute GDNF (R&D Systems, 212-GD) with PBS containing 0.1% bovine serum albumin (BSA), 50 units/mL penicillin, and 50 μg/mL streptomycin and store at 80  C. 5. FGF2 (10 μg/mL): Reconstitute FGF2 (Corning, 354060) with PBS containing 0.1% BSA, 50 units/mL penicillin, and 50 μg/mL streptomycin and stored at 80  C. Dilute 10 μg/ mL FGF2 tenfold to make a 1 μg/mL solution for supplementation into SFM.

2.3 Preparation of GDNF-Conditioned Medium

1. GDNF-expressing HEK293 cells (GDNF-293): Two GDNFexpressing HEK293 cell lines, pGDNF-293 and hGDNF-293, can be used to prepare GDNF-conditioned medium (GDNFCM) (see Note 2). 2. Glycoprotein Denaturing Buffer (10) (New England Biolabs, P0704S): 5% SDS, 400 mM DTT. 3. GlycoBuffer 2 (10) (New England Biolabs, P0704S): 500 mM sodium phosphate, pH 7.5 at 25  C.

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Table 1 Serum-free medium composition for mouse spermatogonial stem cellsa Stock solution

Final concentration

MEMα (Thermo Fisher Scientific, 12000-022)



1

Sodium bicarbonate (Sigma-Aldrich, S5761)



2.2 g/L

Penicillin Streptomycin (Thermo Fisher Scientific, 15140)

1  104 units/mL 1  104 mg/mL

50 units/mL 50 μg/mL

BSA (Sigma-Aldrich, A7030b)



2 mg/mL

FFA mixture (Table 2)

100 meq/L

7.6 μeq/L

Insulin (FUJIFILM Wako, 099-06473)

4 mg/mL

5 μg/mL

Transferrin (FUJIFILM Wako, 208-18971)

20 mg/mL

10 μg/mL

Putrescine (Sigma-Aldrich, P5780)

100 mM

60 μM

2-ME (Sigma-Aldrich, M7522)

14.4 M

50 μM

Sodium selenite (Sigma-Aldrich, 21448)

300 μM

30 nM

GlutaMAX™ (Thermo Fisher Scientific, 35050)

200 mM

2 mM

HEPES (Sigma-Aldrich, H0887)

1M

10 mM

Add BSA, FFA mixture, and penicillin–streptomycin to sterile MEMα with sodium bicarbonate. Incubate at 4  C overnight. The next day, add the remaining compounds to the medium followed by filtration through a 0.22-μm membrane for sterilization (see Note 8) b The lot of BSA needs to be tested for its effectiveness on SSC proliferation a

Table 2 Free fatty acid mixture composition Free fatty acid (FFA)

Stock solutiona

100 meq/L FFA mixtureb

Linolenic acid (Sigma-Aldrich, L2376)

1M

5.6 μL (5.6 mM)

Oleic acid (Sigma-Aldrich, O1008)

1M

13.4 μL (13.4 mM)

Palmitoleic acid (Sigma-Aldrich, P9417)

1M

2.8 μL (2.8 mM)

Linoleic acid (Sigma-Aldrich, L1012)

1M

35.6 μL (35.6 mM)

Palmitic acid (Sigma-Aldrich, P0500)

1M

31.0 μL (31.0 mM)

Stearic acid (Sigma-Aldrich, S4751)

151 mM

76.9 μL (11.6 mM)

Ethanol



834.7 μL

Final volume a

1000 μL

Each FFA is dissolved in ethanol to make its corresponding stock solutions. The stock solutions of palmitic and stearic acids are solid at room temperature. To prepare the mixture at room temperature, dissolve these two FFA stock solutions by warming up to 45–50  C. Divide into 100-μL aliquots in 0.5-mL microfuge tubes, purge with N2, seal with Parafilm®, and store at 80  C b To make a final concentration of 7.6 μeq/L FFA mixture in the serum-free medium, add 76 μL of 100 meq/L FFA mixture to 1000 mL of medium. Numbers in parentheses indicate the final concentration of each FFA in 100 meq/L FFA mixture

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4. Sample buffer (4): 4% SDS, 0.25 M Tris–HCl pH 6.8, 28% glycerol, 0.02% Bromophenol blue, 20% 2-ME. 5. Phosphate buffered saline with Tween 20 (PBST): Add Tween 20 to a final concentration of 0.05% in PBS. 6. Rabbit anti-human GDNF antibody (Santa Cruz Biotechnology, sc-328). 7. Horseradish peroxidase-conjugated donkey anti-rabbit IgG (GE Healthcare, NA934). 8. Recombinant human GDNF (Prospec, CYT-305): Reconstitute GDNF with PBS containing 0.1% BSA, 50 units/mL penicillin, and 50 μg/mL streptomycin and store at 80  C. 9. ECL Select Western Blotting Detection Reagent (GE Healthcare, RPN2235). 2.4

Transplantation

1. Busulfan (Sigma-Aldrich, B2635). 2. Borosilicate glass capillary (World Precision Instruments, TW100-3). 3. Microelectrode MPH6S15).

2.5 LacZ Staining of Recipient Testes

holder

(World

Precision

Instruments,

1. 4% paraformaldehyde in PBS 2. LacZ rinse buffer: 0.2 M sodium phosphate pH 7.3, 2 mM magnesium chloride, 0.01% sodium deoxycholate, 0.02% NP40. 3. LacZ staining solution: 5 mM potassium ferricyanide (SigmaAldrich, P3667), 5 mM potassium ferrocyanide (SigmaAldrich, P3289), 1 mg/mL 5-bromo-4-chloro-3-indolyl-β-Dgalactoside (X-gal) in LacZ rinse buffer. Add 25 mg/mL X-gal in DMSO to potassium ferricyanide–potassium ferrocyanide solution. 4. 10% neutral buffered formalin (NBF).

3

Methods

3.1 Preparation of the Frozen SNL Feeder Stocks

1. Thaw one vial of early passage SNL cells and seed 1  106 of them in a 100-mm dish with 10 mL of 10F-DMEM. Culture the cells at 37  C in a humidified 5% CO2 incubator. When the cells reach 90% confluence in the dish (approximately 3 days after seeding), dissociate them with 0.05% trypsin–EDTA and subculture. In general 4–6  106 SNL cells at an early passage are sufficient to cover 90% of a 100-mm dish (see Note 3). 2. Prepare five 100-mm dishes containing SNL cells at 90% confluence. Add 0.2 mL of 0.5 mg/mL MMC into 10 mL of the

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culture medium, thereby making a final concentration of 10 μg/mL of MMC. 3. Incubate the cells with the MMC-containing medium for 3 h at 37  C. Hourly rotate the dishes to stir the culture medium. 4. Remove the MMC-containing medium from the dishes and rinse the MMC-treated SNL cells 3 times with 5 mL/dish of PBS. 5. Add 2 mL/dish of 0.05% trypsin–EDTA and incubate the dishes at 37  C for 5 min. 6. Add 2 mL/dish of 10F-DMEM to stop the tryptic digestion. 7. Collect the dissociated cells and centrifuge at 600  g for 5 min at 4  C. 8. Resuspend the MMC-treated SNL cells in 10F-DMEM at a density of 6–12  106 cells/mL. 9. Dropwise add an equal volume of ice-cold 2 freeze medium to the cell suspension, obtaining a final concentration of 3–6  106 cells/mL MMC-treated SNL cells with 10% DMSO in 10F-DMEM. 10. [Alternative to steps 8 and 9] Directly add an ice-cold proprietary freeze medium (e.g., CELLBANKER® 1 plus), instead of 10F-DMEM and an equal volume of the 2 homemade freeze medium, so as to obtain 3–6  106 cells/mL. 11. Aliquot 1 mL of the MMC-treated SNL cell suspension into sterile cryovials and freeze them at 80  C using a freezing container. 12. Store the cryovials at 80  C. 13. For a leak test of the MMC-treated SNL cells, thaw one vial and plate on 0.1% gelatin-coated 100-mm dish with 10 mL of 10F-DMEM. Following 3 weeks of culture, stain the cells with methylene blue staining solution, rinse with tap water, and check whether any deep blue–stained colonies are formed. 3.2 Preparation of SNL Monolayer Feeders

1. Add 0.25 mL of 0.1% gelatin solution into each well of 24-well plates. 2. Incubate the plates at 37  C for at least 30 min. 3. Thaw one vial of MMC-treated SNL cells in a water bath at 37  C and immediately put the cell suspension into 9 mL of ice-cold 10F-DMEM in a 50-mL conical centrifugation tube (see Note 4). 4. Centrifuge at 600  g for 5 min at 4  C. 5. Resuspend the cell pellet with 5 mL of 10F-DMEM and count the cells.

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6. Add 10F-DMEM to make a cell suspension of 2  105 cells/ mL. 7. Remove the 0.1% gelatin solution from the wells and put 0.5 mL (1  105 cells) of the MMC-treated SNL cell suspension per well of the 24-well plates. 8. Use the SNL feeder cells within 7 days. If SNL feeders need to be used 5–7 days after they are seeded for SSC culture, change the medium to fresh 10F-DMEM 1 day before starting the SSC culture. 3.3 Preparation of Testis Cell Suspension

1. Prepare three 35-mm dishes and add 2 mL of PBS to each dish. 2. Sacrifice a mouse and separate two testes with fine forceps using sterile procedures. Place testes into the first 35-mm petri dish. 3. Transfer testes to the second 35-mm petri dish and remove tunica albuginea under a dissecting microscope, uncovering seminiferous tubules. Place the testes with exposed tubules to the third 35-mm petri dish. 4. Using a micropipette with a 200-μL filter tip, suck the testes without tunica and transfer them to a 5-mL round-bottom polypropylene tube containing 1.8 mL of 0.25% trypsin– EDTA and 0.2 mL of 7 mg/mL DNase I solution. 5. Pipet up and down by a micropipette with a 1000-μL filter tip until seminiferous tubules disperse into small pieces. 6. Incubate the dispersed tissues at 37  C for 3 min. 7. Add 0.25 mL of 7 mg/mL DNase I solution and 0.25 mL of FBS to stop the enzymatic digestion. 8. Pipet well with the 1000-μL filter tip to make a single-cell suspension. If DNA from dead cells remains, add 0.1 mL of 7 mg/mL DNase I solution and pipet again until it disappears. 9. Centrifuge the cell suspension at 600  g for 5 min at 4  C. 10. Remove the supernatant with a graduated pipette, and resuspend the cell pellet with 1 mL of mSFM. If DNA from dead cells appears again, add a small volume (~0.05 mL) of 7-mg/ mL DNase I solution and pipet until it disappears. 11. Centrifuge the cell suspension at 600  g for 5 min at 4  C. 12. Remove the supernatant, and resuspend the cells in 1 mL of mSFM. 13. Repeat steps 11 and 12. 14. Count the cells. Typical cell number of a testis from a 4–5-days post-partum (dpp) and 6–7-dpp C57BL/6 mouse is 1.0–1.5  106 cells and 1.5–2.0  106 cells, respectively.

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Fig. 1 Initiation and long-term expansion of spermatogonial clumps. (Upper left, day 1) One day after seeding of 7-dpp mouse testicular cells on SNL feeder cells. Some spermatogonia, which can be identified as pale round large cells with a smooth surface, start dividing. (Upper middle, day 2) After passaging onto a fresh feeder layer, many dividing spermatogonia can be identified. (Upper right, day 3) Tightly attached small spermatogonial clumps are formed. (Lower left, day 15) Spermatogonial clump-forming cells continue proliferating, and the size of the clumps increases. Arrowheads indicate spermatogonial cells or clumps. (Lower middle, day 96) Spermatogonia clumps steadily proliferate, retaining the SSC activity that can be assessed by the SSC transplantation assay (Fig. 3). (Lower right) Immunocytochemical analysis of spermatogonial clumps. The clumps were fixed with paraformaldehyde and stained with anti-DDX4 and anti-ZBTB16 antibodies, followed by the secondary antibodies conjugated with Alexa 568 and Alexa 488. The germ cellspecific marker DDX4 (red) and undifferentiated spermatogonia marker ZBTB16 (green) were strongly expressed in most of the clump-forming cells. Bar ¼ 50 μm 3.4 Initiation of Spermatogonial Clump Formation in Testicular Cell Cultures

1. Add an appropriate volume of mSFM to prepare 5  105/mL of testicular cell suspension. 2. Remove the culture medium (10F-DMEM) from the SNL feeder layers in a 24-well plate and rinse each well with 1 mL of PBS twice to wash out the residual medium containing serum (see Note 5). 3. Place 5  105 testicular cells (1 mL) in mSFM onto the SNL feeder cells. 4. Add GDNF and FGF2 at the final concentrations of 10 ng/mL and 0.5 ng/mL, respectively. 5. Culture the testicular cells overnight (for 16–20 h) at 37  C in a humidified 5% CO2 incubator (Fig. 1 D1).

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6. Collect the weakly attached cells by gently pipetting using a graduated pipette (1 mL or 2 mL) or a micropipette with a 1000-μL filter tip. Transfer the cells collected from each well onto fresh SNL feeder layers in a same-sized well as the original culture (1:1). 7. Change the medium the next day. Dividing spermatogonia can be identified (Fig. 1 D2). 8. Change the medium every other day. Dividing spermatogonia form small clumps with tight intercellular contacts (Fig. 1 D3, D15). The spermatogonial clump-forming cells continue proliferating and form large clumps (see Note 6). 9. [Optional procedure] When some adherent somatic cells are detached by the blowing-off procedure at step 6, contaminated non-germ cells such as testicular fibroblasts will outgrow and overwhelm proliferating spermatogonia in culture. In such occasion, repeat subculturing by the blowing-off procedure using a 1 mL or 2 mL graduated pipette or a micropipette with a 1000-μL filter tip, because spermatogonial clumps can be readily removed from the feeder layers. Transfer the collected clumps onto fresh SNL feeder layers. 3.5 Long-Term Ex Vivo Expansion of Spermatogonial Stem Cells

1. After 2–3 weeks in culture, spermatogonial clump-forming cells appear to show stable proliferation and can be subcultured following dissociation with 0.25% trypsin–EDTA. 2. For dissociating the spermatogonial clumps with 0.25% trypsin–EDTA, remove the culture medium and save it in a 15-mL conical centrifugation tube. 3. Add 0.2 mL of 0.25% trypsin–EDTA solution into each well and incubate the plate in a 37  C incubator for 3–5 min to dissociate the spermatogonial clump-forming cells. 4. Add 0.05 mL (1 drop) of FBS into each well to neutralize trypsin. 5. Add 1 mL of saved culture medium and gently pipet dissociated cells several times with a graduated pipette. 6. Pool the dissociated cells from all the wells into the 15-mL conical centrifugation tube that contained the saved culture medium (from step 2). 7. Rinse wells with 2 mL of PBS to collect the residual cells from the wells and transfer the PBS with the collected remaining cells into the same 15-mL centrifugation tube used in steps 2 and 6. 8. Centrifuge at 600  g for 5 min at 4  C. 9. Remove the supernatant and resuspend the cell pellet with 1 mL of mSFM.

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Fig. 2 Flow cytometric analysis of spermatogonial clump-forming cells. The cultured cells (day 106) were collected with 0.25% trypsin–EDTA and stained with anti-mouse THY1 or anti-mouse KIT conjugated with PE/CY5. Histograms with red lines represent the stained cells, whereas the filled histograms represent the unstained cells. The cells expressed the undifferentiated spermatogonial marker THY1, whereas the expression of the differentiation marker KIT was negligible. The cell surface phenotype is identical to SSCs in the testis

10. Repeat steps 8 and 9. 11. Count the germ cells, which can be identified as cells with fewer organelles and a smooth spherical cell surface. Place the cells onto fresh SNL feeders in a 24-well plate. Do not put more than 1  105 cells per well. 12. The proliferation speed of spermatogonial clump-forming cells gradually increases, and they become the dominant cell population (Fig. 1 D96). Fibroblastic cells gradually disappear. Usually, spermatogonial clump-forming cells can be subcultured every 7 days at the plating density of 1  105 cells/well in a 24-well plate after 1 month in culture. In 1 week, 1  105 cells proliferate to 3–4  105 cells. Spermatogonial clumps express the evolutionally conserved germ cell marker DDX4 and undifferentiated spermatogonial marker ZBTB16 (Fig. 1). In addition, the cells express the undifferentiated spermatogonial surface marker THY1, but do not express the differentiation marker KIT (Fig. 2). These phenotypes indicate that the mouse spermatogonial clump-forming cells retain the unique characteristics of undifferentiated spermatogonia [15, 24]. 3.6 Preparation of GDNF-Conditioned Medium for Spermatogonial Stem Cell Culture

Serum-free GDNF-CM derived from GDNF-293 cells can be used for SSC culture. 1. Expand GDNF-293 in 100-mm dishes with 10 mL of 10F-DMEM per dish at 37  C in a humidified 5% CO2 incubator. When the cells proliferate to 90% confluence in the dishes, harvest them with 0.05% trypsin–EDTA. 2. Seed GDNF-293 at the density of 1  106 cells/well in 12-well plates coated with 0.1% gelatin in 10F-DMEM and culture for 2 days (see Note 7).

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3. Two days after seeding, remove the 10F-DMEM medium and rinse the cells with 1 mL PBS/well 3 times. 4. Add 0.7 mL mSFM into each well. 5. After 4 days of culture with mSFM, carefully collect the conditioned medium from the wells using a graduated pipette. Avoid touching or peeling off the cells by the tip of the pipette. 6. For the second round collection, add 0.7 mL of fresh mSFM into each well. Culture the plates for another 4 days. 7. Centrifuge the collected medium at 600  g for 5 min at 4  C. 8. Filtrate the supernatant through a 0.22-μm filter (see Note 8) and store at 4  C. 9. Collect the conditioned medium from the wells of the second round culture with mSFM for 4 days. 10. Repeat steps 7 and 8. 11. Evaluate the GDNF level in the conditioned media by western blot analysis (see Subheading 3.7). 12. Aliquot the supernatant into 15-mL conical tubes and store at 35  C. 13. To use for SSC culture, thaw a tube at 4  C, and centrifuge at 600  g for 5 min at 4  C (see Note 9). 14. Use the GDNF-CM to yield 10 ng/mL GDNF in mSFM. Typically ~30 μL of the supernatant is added into 1 mL mSFM for a regular SSC culture. 3.7 Biochemical Evaluation of GDNF in Conditioned Media from GDNF-293

Western blot analysis can be used to measure the protein content of GDNF in the conditioned medium derived from GDNF-293 [25]. A purified recombinant GDNF (standard GDNF) is electrophoresed side by side for constructing a standard curve. 1. Mix 10 μL of the GDNF-CM or mSFM containing the GDNF standards (1 ng, 3 ng, and 5 ng) and 1.1 μL of 10 Glycoprotein Denaturing Buffer in 1.5-mL microtubes, and boil for 10 min. 2. Spin down the boiled mixture and add 1.4 μL of 10% NP-40 and 1.4 μL of 10 GlycoBuffer 2. 3. Mix with 4.3 μL of 4 sample buffer and boil for 3 min. 4. Electrophorese the samples by a 5–15% gradient SDS-PAGE at 20 V constant voltage for 60 min. 5. Transfer the electrophoresed proteins onto a polyvinylidene difluoride (PVDF) membrane at 25 V constant voltage for 35 min. 6. Block the membrane with 2% skim milk in PBS for 1 h at room temperature on a rocker.

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7. Incubate the membrane with the rabbit anti-human GDNF antibody (1: 10,000) overnight at 4  C on a rocker. 8. After washing with PBST 3 times for 5 min, incubate the membrane with the horseradish peroxidase-conjugated donkey anti-rabbit IgG (1: 20,000) for 1 h at room temperature on a rocker. 9. After washing with PBST 5 times for 5 min each, detect the signal using the ECL select system. 10. Quantify the signals by ImageJ and prepare a standard curve by the gray values of the GDNF standards (1 ng, 3 ng, and 5 ng). 11. Determine the GDNF protein level per 10-μL GDNF-CM using the standard curve. 3.8 Biological Evaluation of GDNF-CM by the Spermatogonial Proliferation Assay

After measuring the GDNF protein level in the GDNF-CM by western blot analysis, it is important to assess the proliferative activity of the GDNF-CM on spermatogonial clump-forming cells. 1. Seed 1  105 cells/well of spermatogonial clump-forming cells on SNL feeder layers in 8 wells of a 24-well plate. 2. Culture the cells with 1 mL of mSFM containing 0.5 ng/mL FGF2 and the supernatant (~30 μL) or GDNF (1 ng, 3 ng, and 10 ng) for 7 days. Set two wells (duplicate) for each condition. Change the medium every other day. 3. After 7 days of culture under each condition, harvest the spermatogonial clump-forming cells from the duplicates and count them. If the number of the cells cultured with GDNF-CM is lower than that with 10-ng/mL GDNF, increase the volume of the GDNF-CM in mSFM for SSC culture.

3.9 Reconstitution of Spermatogenesis by Transplantation of Clump-Forming Spermatogonia

When the spermatogonial clump-forming cells are transplanted into seminiferous tubules devoid of endogenous spermatogenesis, donor cells can colonize to the SSC niche and reconstitute donorderived spermatogenesis. There are two types of recipient mice for efficient colonization of transplanted SSCs. One of them is the white-spotting (W) mouse strains that carry defective mutations in the Kit gene, which encodes a receptor tyrosine kinase required for spermatogonial proliferation and differentiation, and thus lack spermatogenesis [5, 6]. The second one is prepared by injecting the alkylating agent busulfan, which is known as a chemotherapeutic drug. Busulfan induces DNA damage, and its administration to male mice eliminates SSCs and impairs spermatogenesis [5, 6]. The following protocol is used for SSC transplantation into busulfan-treated recipient mice. In addition, to identify the donor cells in recipient seminiferous tubules unequivocally, donor cells are preferably derived from transgenic mice expressing a reporter gene, such as GFP (green fluorescent protein) [e.g.,

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Japan SLC, C57BL/6-Tg(CAG-EGFP), GFP mouse] or LacZ (β-galactosidase) [e.g., Jackson Laboratory, B6.129S7-Gt(ROSA) 26Sor/J, ROSA mouse]. 1. Dissolve busulfan in warm DMSO (~50  C) at 8 mg/mL. 2. Add an equal volume of warm distilled water to make a final concentration of 4 mg/mL busulfan. Keep the solution at ~40  C until injection. 3. Intraperitoneally inject 4 mg/mL busulfan to adult C57BL/6 males at 44 mg/kg. Typically, 0.25–0.3 mL of busulfan solution is administrated per mouse. 4. Maintain the mice for at least 5 weeks to ensure that their spermatogenesis is impaired. 5. For transplantation into the seminiferous tubules, prepare single-cell suspensions of spermatogonial clump-forming cells by enzymatic digestion with 0.25% trypsin–EDTA as described above. Filter the cells through a 30-μm nylon mesh and suspend 2–3  106 cells per 1 mL of mSFM containing 0.7 mg/ mL DNase. Store the cells on ice. 6. Anesthetize the recipient mice and shave the abdomen. Make a midline incision and pull out the testis alongside the fat pad. 7. Dissect efferent ducts out using fine forceps under a dissecting microscope. 8. Mix 4 μL of 2–3  106 cells/mL cell suspension with 4 μL of 0.08% of trypan blue solution to make 8 μL of 1.0–1.5  106 cells/mL cell suspension. 9. Fill the injection pipette with 8 μL of the cell suspension from the rear end of the pipette (see Note 10). 10. Manually insert the micropipette into an efferent duct exposed and thread almost to the rete testis. 11. Increase the pressure in the pipette, resulting infusion of the donor cell suspension into the rete first and then the seminiferous tubules. Seven to eight microlitre of the donor cells can fill 75–95% of the tubules (see Note 11). 12. Two months after transplantation, sacrifice the recipient mice, collect their testes and put them into PBS in 12-well plates. 13. Weigh each testis. If the weight is >50 mg, endogenous spermatogenesis has started to resume. 14. Remove tunica albuginea and directly analyze each testis under a fluorescent stereomicroscope for donor cells from a GFP mouse. Stain the testes with X-gal for donor cells from a ROSA mouse (see Subheading 3.10). 15. Count the spermatogenic colonies consisting of GFP+ green cells or LacZ-stained blue cells to determine the SSC activity

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Fig. 3 Spermatogenic colony formation and reconstitution of spermatogenesis in recipient testes transplanted with SSCs derived from GFP and ROSA mice. (a) Spermatogonial clump-forming cells derived from GFP-expressing mice were transplanted into busulfan-treated recipient mouse testes. Two months after transplantation, the recipient testis was analyzed. A green stretch of cells in the testis indicates the spermatogenic colonies arising from a single SSC. (b) Spermatogonial clump-forming cells derived from ROSA mice were transplanted into infertile W/Wv recipient mouse testes. ROSA mice express LacZ gene (β-galactosidase) in all cells, and thus the recipient testis was analyzed by X-gal staining. A blue stretch of cells in the testis indicates the spermatogenic colonies arising from a single SSC. (c) Histological analysis of the recipient testes transplanted with LacZ-expressing spermatogonial clump-forming cells. The recipient testis was stained with X-gal, embedded in paraffin wax, sectioned, and stained with hematoxylin and eosin. Donor LacZ-expressing SSCs completely reconstituted spermatogenesis (stained blue). The seminiferous tubules where SSCs were not colonized remain devoid of spermatogenic cells. Bar ¼ 1 mm (a, b), 50 μm (c)

that represents the SSC number in the donor cell suspension (Fig. 3a, b). 16. Normalize to 1  105 cells transplanted into the recipient testes. In general, 10–30 colonies will be generated in a recipient testis, which is estimated at 150–250 colonies/ 1  105 cells transplanted. 17. Spermatogenesis derived from spermatogonial clump-forming cells transplanted in the busulfan-treated or W recipients will continue in the host male throughout its life after transplantation. 3.10 LacZ-Staining of the Recipient Testes Transplanted with Spermatogonial Stem Cells

1. Sacrifice recipient mice, collect their testes and put them into PBS in 12-well plates. 2. Weigh each testis. If the weigh is >50 mg, endogenous spermatogenesis has started resuming. 3. Remove tunica albuginea under a dissecting microscope and slightly loosen the seminiferous tubules using fine forceps. This will facilitate to count colonies. 4. Remove PBS and add 2 mL/well of 4% PFA.

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5. Agitate the plates for 1.5–2 h at 4  C on a rocker. 6. Remove the fixative and add 2 mL/well of LacZ rinse buffer. 7. Agitate the plates for 1 h at 4  C on a rocker. 8. Replace the LacZ rinse buffer. Repeat 3 times. 9. Remove the LacZ rinse buffer and add 2 mL/well of LacZ staining solution. 10. Seal the plates with Parafilm® and agitate them overnight at 30–37  C on a rocker. 11. Remove the LacZ staining solution and rinse the testes with tap water twice. 12. Remove the water and add 2 mL/well of 10% NBF. 13. Analyze the testes under a dissection microscope. Count the donor-derived spermatogenic colonies, which are stained blue (Fig. 3b, c).

4

Notes 1. SNL76/7 has clonally been established from Sandoz inbred mouse (SIM) embryo–derived thioguanine-resistant and ouabain-resistant (STO) fibroblast cell line transfected with neomycin resistance (Neo) and murine leukemia inhibitory factor (LIF) genes [18]. SNL76/7 stands for STO-Neo-LIF clone 76/7. 2. The GDNF-expressing stable cell lines pGDNF-293 and hGDNF-293 have clonally been established from HEK293 cells by transfection with pEPI-CAG-pGDNF and pEPICAG-hGDNF plasmids, respectively [25]. The pEPI-CAG plasmid is an S/MAR-based episomal vector containing a chicken β-actin promoter-driven expression unit and a neomycin resistance gene [26]. pGDNF-293 secretes porcine GDNF, whereas hGDNF-293 secretes human GDNF into the culture medium. 3. It is advisable to make frozen stocks of early passage SNL cells. If more than 8–9  105 cells are required for the 90% confluence in a 100-mm dish, stop using and thaw an early passage SNL cells. Testing the FBS lot may help to retain the original characteristics of the cells for a longer period. 4. The cell viability of MMC-treated SNL cells after thawing is usually more than 90%. The frozen SNL feeders can be stored until 2 years without any obvious cell damage, such as low viability. 5. When SNL feeder layers are used with serum-free medium, always rinse twice with PBS (1 mL/well for a 24-well plate).

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6. Reducing the atmospheric oxygen concentration from 21% to 10% improve development of long-term cultures of spermatogonial clump-forming cells [27]. 7. When GDNF-293 cells are seeded into 72 wells (six 12-well plates), approximately 100 mL (0.7 mL/well  72 wells  2) of GDNF-CM is obtained. 8. To rinse away any unknown residuals from the 0.22-μm filter of bottle top filter units, pass at least 100 mL of MilliQ water before filtering the conditioned medium and serum-free medium. 9. After thawing, some precipitants may appear. They seem not to interfere with the proliferation of spermatogonial clumpforming cells in culture. However, if excessive insoluble material appears, centrifuge at 600  g for 5 min and use the supernatant for culture. 10. The injection pipette is made of a borosilicate glass capillary using a micropipette puller (e.g., Sutter, P-97). 11. Pressure for the injection can be generated by an electric microinjector (e.g., Narishige, IM-31) by connecting a 1-mL plastic syringe barrel with flexible tubing. A microelectrode holder is attached to the syringe to couple an injection pipette to the syringe barrel. References 1. Potten CS, Morris RJ (1988) Epithelial stem cells in vivo. J Cell Sci Suppl 10:45–62 2. Kubota H, Brinster RL (2006) Technology insight: in vitro culture of spermatogonial stem cells and their potential therapeutic uses. Nat Clin Pract Endocrinol Metab 2:99–108 3. Meistrich ML, van Beek MEAB (1993) Spermatogonial stem cells. In: Desjardins C, Ewing LL (eds) Cell and molecular biology of the testis. Oxford University Press, New York, pp 266–295 4. Brinster RL (2002) Germline stem cell transplantation and transgenesis. Science 296:2174–2176 5. Brinster RL, Zimmermann JW (1994) Spermatogenesis following male germ-cell transplantation. Proc Natl Acad Sci U S A 91:11298–11302 6. Brinster RL, Avarbock MR (1994) Germline transmission of donor haplotype following spermatogonial transplantation. Proc Natl Acad Sci U S A 91:11303–11307 7. Kubota H, Brinster RL (2018) Spermatogonial stem cells. Biol Reprod 99:52–74

8. Gonza´lez R, Dobrinski I (2015) Beyond the mouse monopoly: studying the male germ line in domestic animal models. ILAR J 56:83–98 9. Wu X, Goodyear SM, Abramowitz LK, Bartolomei MS, Tobias JW, Avarbock MR, Brinster RL (2012) Fertile offspring derived from mouse spermatogonial stem cells cryopreserved for more than 14 years. Hum Reprod 27:1249–1259 10. Brinster RL (2007) Male germline stem cells: from mice to men. Science 316:404–405 11. Kanatsu-Shinohara M, Ogonuki N, Inoue K, Miki H, Ogura A, Toyokuni S, Shinohara T (2003) Long-term proliferation in culture and germline transmission of mouse male germline stem cells. Biol Reprod 69:612–616 12. Kubota H, Avarbock MR, Brinster RL (2004) Growth factors essential for self-renewal and expansion of mouse spermatogonial stem cells. Proc Natl Acad Sci U S A 101:16489–16494 13. Kubota H, Brinster RL (2008) Culture of rodent spermatogonial stem cells, male

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germline stem cells of the postnatal animal. Methods Cell Biol 86:59–84 14. Barnes D, Sato G (1980) Serum-free cell culture: a unifying approach. Cell 22:649–655 15. Kubota H, Avarbock MR, Brinster RL (2004) Culture conditions and single growth factors affect fate determination of mouse spermatogonial stem cells. Biol Reprod 71:722–731 16. Meng X, Lindahl M, Hyvonen ME, Parvinen M, de Rooij DG, Hess MW, Raatikainen-Ahokas A, Sainio K, Rauvala H, Lakso M, Pichel JG, Westphal H et al (2000) Regulation of cell fate decision of undifferentiated spermatogonia by GDNF. Science 287:1489–1493 17. Zhang Y, Wang S, Wang X, Liao S, Wu Y, Han C (2012) Endogenously produced FGF2 is essential for the survival and proliferation of cultured mouse spermatogonial stem cells. Cell Res 22:773 18. McMahon AP, Bradley A (1990) The Wnt-1 (int-1) proto-oncogene is required for development of a large region of the mouse brain. Cell 62:1073–1085 19. Ryu BY, Kubota H, Avarbock MR, Brinster RL (2005) Conservation of spermatogonial stem cell self-renewal signaling between mouse and rat. Proc Natl Acad Sci U S A 102:14302–14307 20. Kubota H, Wu X, Goodyear SM, Avarbock MR, Brinster RL (2011) Glial cell line-derived neurotrophic factor and endothelial cells promote self-renewal of rabbit germ cells with spermatogonial stem cell properties. FASEB J 25:2604–2614 21. Kubota H, Reid LM (2000) Clonogenic hepatoblasts, common precursors for hepatocytic

and biliary lineages, are lacking classical major histocompatibility complex class I antigen. Proc Natl Acad Sci U S A 97:12132–12137 22. Kubota H, Yao HL, Reid LM (2007) Identification and characterization of vitamin A-storing cells in fetal liver: implications for functional importance of hepatic stellate cells in liver development and hematopoiesis. Stem Cells 25:2339–2349 23. Wauthier E, Schmelzer E, Turner W, Zhang L, LeCluyse E, Ruiz J, Turner R, Furth ME, Kubota H, Lozoya O, Barbier C, McClelland R et al (2008) Hepatic stem cells and hepatoblasts: identification, isolation, and ex vivo maintenance. Methods Cell Biol 86:137–225 24. Kubota H, Avarbock MR, Brinster RL (2003) Spermatogonial stem cells share some, but not all, phenotypic and functional characteristics with other stem cells. Proc Natl Acad Sci U S A 100:6487–6492 25. Kakiuchi K, Taniguchi K, Kubota H (2018) Conserved and non-conserved characteristics of porcine glial cell line-derived neurotrophic factor expressed in the testis. Sci Rep 8:7656 26. Kakiuchi K, Tsuda A, Goto Y, Shimada T, Taniguchi K, Takagishi K, Kubota H (2014) Cell-surface DEAD-box polypeptide 4-immunoreactive cells and gonocytes are two distinct populations in postnatal porcine testes. Biol Reprod 90(82):81–11 27. Kubota H, Avarbock MR, Schmidt JA, Brinster RL (2009) Spermatogonial stem cells derived from infertile Wv/Wv mice self-renew in vitro and generate progeny following transplantation. Biol Reprod 81:293–301

Chapter 15 Generating Kidney Organoids from Human Pluripotent Stem Cells Using Defined Conditions Sara E. Howden and Melissa H. Little Abstract The ultimate goal of regenerative medicine is to have access to an unlimited supply of specific cell types on demand, which can be used as effective therapies for a wide range of intractable disorders. With the availability of human pluripotent stem cells (hPSCs) and greatly improved protocols for their directed differentiation into specific cell types, including kidney, this prospect could soon become a reality. We have previously described the generation of kidney organoids from hPSCs. This chapter describes our latest differentiation protocol for generating kidney tissue, which uses a cost-effective and completely defined, xeno-free medium. As with our previous protocol, these complex, multicellular three-dimensional structures are composed of all anticipated kidney cell types including nephrons segmented into the glomerulus, proximal and distal tubule as well as an extensive endothelial network, and renal interstitium. As such, kidney organoids provide useful tools for understanding human development, disease modeling, drug screening/toxicology studies and tissue engineering applications, and may facilitate the development of transplantable hPSC-derived kidney tissue for regenerative medicine purposes in the future. Key words Human pluripotent stem cell, Kidney organoid, Directed differentiation, Nephrogenesis, Kidney development

1

Introduction Chronic kidney disease is one of the fastest growing diseases worldwide and represents a significant economic burden. For over 70 years now, renal failure has been and continues to be treated either with dialysis or organ transplantation. However, with only a subset of patients receiving a donor transplant, and dialysis being associated with low quality of life and high morbidity, there is an urgent need to find alternative treatments. One such approach is to generate replacement renal tissue from human pluripotent stem cells (hPSCs). Pluripotent stem cells offer enormous potential for modeling disease, drug discovery and transplantation medicine due to their ability to differentiate into any given cell type. Induced pluripotent

Chrissa Kioussi (ed.), Stem Cells and Tissue Repair: Methods and Protocols, Methods in Molecular Biology, vol. 2155, https://doi.org/10.1007/978-1-0716-0655-1_15, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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stem cells (iPSCs), generated by introducing defined factors to reprogram terminally differentiated somatic cells [1, 2] are particularly advantageous for the development of autologous or customized cellular therapies to treat or correct many inherited and acquired diseases. Complications associated with immunorejection can be avoided through the generation and subsequent disease correction of patient-specific iPSCs, which can be differentiated into relevant cell types for the repopulation and regeneration of a defective tissue or organ. Based upon our extensive understanding of the molecular and cellular mechanisms governing normal mammalian kidney morphogenesis, we have developed protocols for the directed differentiation of hPSCs to human kidney tissue [3–5]. In 2015, we described an approach for generating kidney organoids containing appropriately patterned and segmented nephrons, surrounding renal stroma, putative ureteric epithelium as well as endogenous endothelial and perivascular cell populations. Since this publication, we have also developed an extensive toolbox of fluorescent iPSC reporter lines which mark specific cell types in the developing kidney, such as nephron progenitors, podocytes, proximal tubule, distal nephron, and ureteric epithelium [6]. These resources are now freely available to other researchers and can be used to facilitate a range of experimental applications, including cellular isolation, time-lapse imaging, and further optimization of kidney differentiation protocols. In this most recent version of our protocol, hPSCs are maintained and differentiated in a completely defined, minimal and xeno-free medium that is substantially more cost-effective than the previously used APEL medium. This simplified medium also provides a much cleaner background for examining specific pathways or growth factors that may influence nephron patterning and/or maturation either during the initial stages of monolayer differentiation or during the organoid stage, when all growth factors are removed from the culture medium. Additionally, cells can also be maintained and differentiated on a recombinant human laminin 521 (LN521) substrate instead of Matrigel, to completely eliminate all animal components from the culture system. Although it is more cost-effective to routinely maintain pluripotent cells on Matrigel, we recommend performing the monolayer stage of differentiation on LN521 to minimize the effects that Matrigel (and any residual growth factors associated with it) may have on differentiation outcomes. As described in the original protocol, hPSCs are first differentiated into posterior primitive streak using the canonical WNT agonist, CHIR99021. The length of the initial CHIR99021 treatment (3–5 days) was interpreted as a means to pattern to anterior (short duration) and posterior (long duration) intermediate mesoderm [4]. We note that this also influences the nephron patterning

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observed in the resulting kidney organoids, in that a shorter period promotes an increased abundance of distal nephron and/or putative ureteric epithelium, whereas a longer period promotes proximal nephron segments, including proximal tubule and podocyte formation. The ratio of distal to proximal nephron can also vary substantially between different hPSC lines, and thus the length and concentration of CHIR99021 treatment should be evaluated for each new hPSC line. Differentiation to kidney requires at least 4 μM CHIR with the concentration varying between 4 and 8 μM. Typically, we find that 7 μM CHIR99021 for 3 days is a reasonable starting point and usually results in an appropriate balance between both distal and proximal nephron segments. At day 7 of differentiation, the culture is dissociated and 2–5  105 cell aggregates are transferred to Transwell plates to facilitate culture in an air–media interface, a format that is known to support the in vitro culture of mouse embryonic kidney explants [7, 8]. To maximize the number of nephrons formed, aggregates are also exposed to a transient CHIR99021 pulse, which stimulates the mesenchyme to undergo an epithelial transition. The organoids are cultured in the presence of FGF9 for an additional 5 days, and then all growth factors are removed from the basal medium until the completion of differentiation at around day 25. Alternatively, growth factors or small compounds may be added at this stage to examine the effects of specific signaling pathways on nephron patterning/maturation. For example, the addition of all-trans retinoic acid (2 μM) between days 13–25 of differentiation consistently promotes podocyte formation and also appears to improve tubular maturation.

2 2.1

Materials Tissue Culture

2.1.1 Equipment

1. Humidified incubator at 37  C with 5% CO2. 2. Laminar flow tissue culture hood. 3. Centrifuge (for 15 mL and 50 mL tubes). 4. Microfuge (for 1.5 ml Eppendorf tubes). 5. Inverted microscope. 6. Filtered glass or plastic 5 mL and 10 mL pipets. 7. Filtered pipette tips. 8. Wide-bore pipette tips. 9. Glass Pasteur pipettes and media waste trap for aspirating media. 10. 15 mL and 50 mL conical centrifuge tubes. 11. 6-well plates.

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12. 6-well Transwell cell culture plate 13. Hemocytometer. 2.1.2 Cell Culture and Differentiation

1. Essential 8 medium. 2. Essential 6 medium. 3. Matrigel. 4. LN521. 5. PBS. 6. 0.5 mM EDTA solution. 7. TrypLE. 8. CHIR99021, made up to 10 mM in DMSO. 9. Y-27632 dihydrochloride, made up to in PBS. 10. FGF9, made up to (100 μg/mL) PBS containing 0.1% (wt/vol) human serum albumin. 11. Fetal bovine serum or recombinant human serum.

3

Methods

3.1 Maintenance and Expansion of Pluripotent Stem Cells

Human pluripotent stem cells can be maintained indefinitely on Matrigel-coated plates in Essential 8 medium and should be passaged every 3–4 days as previously described [9]. Cells should not be allowed to reach >90% confluency and culture media should be changed daily to prevent excessive cell death and/or unwanted differentiation. Prepare Matrigel coated plates at least 30 min in advance. Matrigel should be thawed on ice and divided into small aliquots (typically 200–500 μL per tube). Add 0.2 mg Matrigel (see Note 1) per mL of cold DMEM-F12, mix well and add 1 mL per well of a 6-well plate or 4 mL for each 10 cm dish. Plates can be kept in 37  C incubator for up to 2 weeks. We routinely maintain undifferentiated cells in 6-well plate format. 1. To passage cells, remove spent medium and wash cells with 1 mL room temperature 0.5 mM EDTA (in PBS) solution and aspirate with glass Pasteur pipette. 2. Add 1 mL 0.5 mM EDTA solution and incubate cells for 2–5 min in 37  C incubator. Following EDTA incubation the cells should still be loosely attached to the plate in distinct colonies, but cells should appear somewhat dissociated from one another when observed under the microscope (see Note 2). 3. During the incubation step, remove residual media from a Matrigel coated plate and add 2 mL of fresh Essential 8 medium per well.

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4. Carefully remove EDTA solution and add 3 mL Essential 8 medium to remove cells from the plate. Pipet up and down once or twice to break up cells into smaller clumps and divide evenly over the new 6-well plate. 5. Place cells in 37  C incubator and gently agitate the plate to ensure an even spread of colonies over the entire plate. 3.2 Plating Cells for Differentiation

Prepare LN521-coated plates 1–2 days in advance. LN521 should be thawed on ice and divided into small aliquots (100–200 μL per tube). Add 50 μL (5 μg) per 1 mL of cold DMEM-F12, mix well and add 1 mL to one well of a 6-well plate. Plates should be wrapped in Parafilm for storage and can be kept at 4  C for up to 2 weeks. Cells should be passaged 2 days prior to differentiation. 1. Remove culture media from wells containing cells and add 1 mL of pre-warmed TrypLE and incubate at room temperature for 2–3 min. 2. Remove TrypLE before cells begin lifting from the plate and wash cells from the well with 2–3 mL prewarmed Essential 8 medium. 3. Transfer cells into 15 mL tube and take a small aliquot for counting. 4. Pellet cells by centrifugation at 300  g for 5 min. 5. Resuspend cell pellet in Essential 8 medium and seed 80,000 cells per well of a 6-well LN521-coated plate (see Note 3). 6. Place cells in 37  C incubator and gently agitate the plate to ensure an even spread of colonies over the entire plate.

3.3 Monolayer Differentiation (Days 1–7)

1. The day after plating cells, add 2 mL of Essential 6 medium containing 4–8 μM CHIR99021 to each well of a 6-well plate (Fig. 1). 2. Culture the cells for 3–5 days, refreshing the medium containing CHIR99021 every 2 days. By days 2–3, the pluripotent colonies should become more dispersed (Fig. 2, Day 2). CHIR99021 duration determines the ratio of distal/proximal nephron patterning in the resulting organoid. 3. After the CHIR99021 phase, change culture medium to 2 mL of Essential 6 medium supplemented with 200 ng/mL FGF9 and 1 μg/mL heparin. 4. Refresh with medium containing 200 ng/mL FGF9 and 1 μg/ mL heparin every 2 days until day 7 of the differentiation. By this time point the culture should form a monolayer and cover the vast majority of the well (Fig. 2, Day 7).

Sara E. Howden and Melissa H. Little

Stage

Medium

Pluripotent

Essential 8

Seed cells on LN521

Section 3.1 Steps 1-6

Essential 6 + CHIR99021 (4-8 μm)

Start differentiation

Primative streak

Section 3.3 Steps 1-2

day 1-7

day 0

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Intermediate mesoderm

Essential 6 + FGF9 (200 ng/μl)

Nephrogenesis

day 13-25

Maturation

Endpoint

Essential 6

Procedure

Swith to FGF9 between days 3-5. An ealier switch promotes more distal nephron patternig whereas a later switch promotes more proximal nephron

Method

Section 3.3 Steps 3-4

On day 7 harvest and aggregate cells and transfer to transwell to promote 3D organisation and nephrogenesis. A short (1h) CHIR99021 pulse promotes nephron formation

Section 3.4 Steps 1-13

Remove FGF9 from medium on day 12 and culture organoids for additional ~2 weeks to promote further maturation. Additional factors may be also be added to effects nephron patterning/maturation. For example, retionic acid promotes podocyte formation

Section 3.4 Step 14

Harvest organoids for downstream applications/analyses

Section 3.4 Step 15

Fig. 1 Timeline and overview of the kidney organoid differentiation protocol 3.4 Kidney Organoid Culture (Days 7–25)

1. Aspirate the medium from the day 7 monolayer culture and add 1 mL of TryPLE and place in 37  C incubator for 5 min. 2. Add 5 mL of Essential 6 medium and gently pipette up and down a few times to remove all cells from the plate and break up large clumps. 3. Transfer the cell suspension to a 15 mL falcon tube and centrifuge for 3 min at 300  g to pellet cells. 4. Aspirate the supernatant and resuspend the cells with 3 mL of Essential 6 medium containing 2% fetal bovine serum OR 2% recombinant human serum (see Note 4). 5. Take out 10 μL of cell suspension and perform a cell count with a hemocytometer. 6. Transfer between 200,000 and 500,000 cells into a 1.5 mL microcentrifuge tube. Centrifuge the tube at 300  g for 3 min to aggregate cells. Each aggregate formed represents one kidney organoid. A dissociated day 7 culture can generate up to 20 organoids.

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Day 18

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Day 25

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Fig. 2 Representative bright field images showing the morphological changes that occur during kidney organoid differentiation. After the pluripotent stem cell stage (Day 1), cells become more disperse in response to CHIR99021 treatment (Day 3). The monolayer culture should become confluent and cover the vast majority of the culture surface by Day 7, when the cells are harvested, aggregated by centrifugation and transferred to a Transwell for 3D culture. Epithelial structures typically become apparent after 3–5 days of 3D culture (Day 12) and elongate over the course of differentiation (Day 18–Day 25)

7. Rotate the tube 180 again (back to original starting position) and recentrifuge at 300  g for 2 min. Repeated centrifugation steps ensures the cells from a tightly packed aggregate which facilitates its pipette-mediated transfer in the subsequent step. 8. Rotate the tube 180 and centrifuge the tube again at 300  g for 2 min. 9. During the centrifugation steps, prepare the Transwell plate by adding 1.2 mL of E6 medium containing 5 μM CHIR99021 underneath the Transwell filter insert. 10. Place a wide bore tip over the cell pellet and gently draw up the aggregate into the pipette tip. Gently aspirate the medium along with the aggregate onto the Transwell filter (see Note 5). 11. Carefully remove any excess medium around the organoid using a p20 or p200 pipette before placing into 37  C incubator for 1 h. 12. After the 1 h incubation, remove the medium and add 1.2 mL of Essential 6 medium supplemented with 200 ng/mL FGF9 and 1 μg/mL heparin.

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NPHS1 LTL ECAD GATA3

500 μm

Fig. 3 Representative immunofluorescence image of a typical whole kidney organoid generated using the GATA3mCherry:MAFBmTagBFP2 fluorescent reporter iPSC line [6] and 7 μM CHIR99021 for 3 days in the monolayer differentiation stage. The organoid contains nephrons with podocytes (NPHS1+), proximal tubule (LTL+), distal tubule (ECAD+), and a putative ureteric epithelial (GATA3 + ECAD+) network

13. Refresh medium (Essential 6 containing FGF9 and heparin) every 2 days. The formation of epithelial structures (renal vesicles) can typically be observed after day 10–12 of differentiation (Fig. 2, Day 12). 14. After 5 days, change the culture medium to Essential 6 (without FGF9 and heparin) and refresh medium every 2 days. 15. Harvest kidney organoids for downstream analysis/experiments at or around day 25 (Fig. 2, Day 25). At this stage, kidney organoids are typically 3–5 mm in diameter and should contain nephrons with, glomerulus, proximal tubule, distal tubule and/or ureteric epithelium segments (Fig. 3). Refer to Takasato et al. for a detailed step-by-step guide for analyzing kidney organoids by whole-mount immunofluorescence [5].

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Notes 1. Matrigel concentration varies according to lot number. A lot-specific, product specification sheet with the exact protein concentration is provided by the supplier with each shipment. 2. Timing is critical when passaging cells with EDTA. Cells left too long will dissociate and come away from the plate before addition of the new culture medium, whereas cells that have not been incubated long enough will be very difficult to remove from the plate. If done correctly, the cells should easily come away from the plate upon addition of new culture medium and should not require pipetting up and down more than once or twice with the Pipet-Aid on the fastest setting. Cells that have been incubated too long and become freefloating can be collected by centrifugation before resuspension and plating. 3. Viability of cells following transfer to LN521 after TryPLEmediated dissociation varies substantially from one line to the next. If excessive cell death is observed the day after plating cells for differentiation, a new batch of cells should be plated in medium supplemented with 10 μM Y-27362. 4. The addition of serum at this step aids in cell aggregation by centrifugation. Recombinant human serum can be used as alternative to fetal bovine serum if xeno-free conditions are more desirable by the user. 5. Try to keep the aggregate intact as much as possible. If the cell pellet breaks up excessively during the transfer process, perform another 1–2 centrifugation steps to repellet the cells.

Acknowledgments This work was supported by the National Institutes of Health (DK107344), the National Health and Medical Research Council, Australia (GNT1100970) and the Australian Research Council (DP190101705). M.H.L is a Senior Principal Research Fellow of the NHMRC (GNT1136085). The Murdoch Children’s Research Institute is supported by the Victorian Government’s Operational Infrastructure Support Program. References 1. Yu J, Vodyanik MA, Smuga-Otto K et al (2007) Induced pluripotent stem cell lines derived from human somatic cells. Science 318:1917–1920

2. Takahashi K, Tanabe K, Ohnuki M et al (2007) Induction of pluripotent stem cells from adult human fibroblasts by defined factors. Cell 131:861–872

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3. Takasato M, Er PX, Becroft M et al (2014) Directing human embryonic stem cell differentiation towards a renal lineage generates a selforganizing kidney. Nat Cell Biol 16:118–126 4. Takasato M, Er PX, Chiu HS et al (2015) Kidney organoids from human iPS cells contain multiple lineages and model human nephrogenesis. Nature 526:564–568 5. Takasato M, Er PX, Chiu HS, Little MH (2016) Generation of kidney organoids from human pluripotent stem cells. Nat Protoc 11:1681–1692 6. Vanslambrouck JM, Wilson SB, Tan KS (2019) A toolbox to characterize human induced

pluripotent stem cell-derived kidney cell types and organoids. J Am Soc Nephrol 30:1811. (Epub ahead of print) 7. Giuliani S, Perin L, Sedrakyan S et al (2008) Ex vivo whole embryonic kidney culture: a novel method for research in development, regeneration and transplantation. J Urol 179:365–370 8. Gupta IR, Lapointe M, Yu OH (2003) Morphogenesis during mouse embryonic kidney explant culture. Kidney Int 63:365–376 9. Chen G, Gulbranson DR, Hou Z et al (2011) Chemically defined conditions for human iPSC derivation and culture. Nat Methods 8:424–429

Chapter 16 Painting the Pancreas in Three Dimensions: Whole-Mount Immunofluorescence Method Maricela Maldonado, Jeffrey D. Serrill, and Hung-Ping Shih Abstract The pancreas is composed of different cellular populations, organized into distinct functional units, including acinar clusters, islets of Langerhans, and the ductal system. As a result of research into diabetes, several optical techniques have been developed for the three-dimensional visualization of islet populations, so as to better understand their anatomical characteristics. These approaches are largely reliant on threedimensional whole-mount immunofluorescence staining. In this chapter, we review a revised whole mount immunofluorescence staining method for studying adult pancreatic islet morphology. This method uses smaller samples and combines the blocking and permeabilization steps. This reduces the time needed, relative to existing protocols; the method is compatible with regular confocal microscopy as well. Key words Islet, Pancreas, 3D imaging, Immunofluorescence

1

Introduction Traditionally, analysis of the anatomical characteristics of the pancreas has largely relied on histological analysis on tissue sections. This method, combined with immunofluorescence staining, provides two-dimensional (2D) images that allow us to examine pancreatic tissue and cellular components at single-cell resolution. However, the pancreas is a highly branched organ, with various cell populations distributed unevenly throughout distinct pancreatic tissue domains [1, 2]. For example, in the adult pancreas, the islets of Langerhans (which contain the pancreatic endocrine cells) are connected to the pancreatic duct system [1, 3]. Thus, it is difficult to accurately visualize the volume and spatial position of the islets of Langerhans via 2D histology/imaging techniques. Three-dimensional (3D) whole-mount immunofluorescence (WMIF) staining provides an optical method for labeling and imaging cell populations deep in intact pancreas tissue. This is an ideal tool for studying the anatomical characteristics of pancreatic islets. It has already been applied to studies of whole embryonic

Chrissa Kioussi (ed.), Stem Cells and Tissue Repair: Methods and Protocols, Methods in Molecular Biology, vol. 2155, https://doi.org/10.1007/978-1-0716-0655-1_16, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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mouse pancreas [4, 5], and to adult mouse pancreatic islets [6– 8]. However, adult pancreatic samples are much larger and thicker than embryonic ones; as a result, the incubatory steps (fixative, blocking buffer, antibody, wash buffer, permeabilization, and substrate color development) take much longer. Although these approaches provide unprecedented resolution and cellular details in whole organ scale, they mostly rely on expensive imaging platforms; furthermore, sample preparation can take upward of a week. In many studies, Islet 3D anatomical analysis does not always require single-cell resolution at whole-organ scale7. A rapid and economical WMIF method would be more suitable to different studies. We adapted several existing WMIF protocols [4, 6, 9] to be more compatible with regular confocal microscopy. Our protocol can be combined with various optical tissue clearing (OTC) methods [10], and be modularized with other imaging techniques.

2

Materials All organic solvents should be prepared under a fume hood; and histology-grade reagents are to be used. The BABB solution (benzyl alcohol–benzyl benzoate, 1:2), and either Dent’s fix or bleaching solution, should be prepared and stored in glass containers (e.g., scintillation vials) covered with aluminum foil to block light exposure.

2.1 Glassware and Supplies

1. Double Concave Microscope Slides: 1527-006 Thermo Fisher. Length (Metric)

76 mm

Width (Metric)

26 mm

Thickness (Metric)

2.8–3.2 mm

Diameter (Metric)

16–18 mm diameter well, 1.5 mm deep

Material

Edges: Ground. Corners: 90 .

Product type

Microscope slides

Quantity

12 slides per box

Well count

2 depressed

Dimensions (L  W)

76  26 mm

2. Wheaton™ Liquid Scintillation Glass Vials: 28  61 mm 03-341-25B Fisher Science. 3. Glass coverslips: 24  60 mm 12-545-M Thermo Fisher. 2.2

Reagents

1. 4% paraformaldehyde (PFA) in PBS (see Note 1). 2. 100% methanol (MeOH).

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3. 1PBS. 4. PBS-X: 0.15% Triton X-100 in PBS. 5. PBS-T: 0.15% Tween in PBS. 6. 50%-, 75%-, and 90% MeOH prepared with 1PBS. 7. Dent’s Fix solution: 80% MeOH (40 mL) + 20% DMSO (10 mL) ¼ 50 mL total. 8. Dent’s Bleaching solution: Dent’s Fix (24 mL) + 30% H2O2 (12 mL) ¼ 36 mL total. 9. Blocking buffer: 5% donkey serum + 0.15% Triton X-100 in 1PBS. 10. Rabbit Anti-Chromogranin A antibody (Abcam ab15160). 11. Goat anti-Spp1 (R&D systems AF808). 12. Alexa Fluor® 488 AffiniPure Donkey Anti-Rabbit IgG (Jackson ImmunoResearch 711-545-152). 13. Alexa Fluor® 555 AffiniPure Donkey Anti-Goat IgG (Jackson ImmunoResearch 705-165-147). 14. Under the fume hood prepare Dent’s Bleach as follows: 6 mL of Dent’s Fix solution + 3 mL of 30% H2O2 ¼ 9 mL total (enough for two samples, see Note 2).

3

Methods

3.1 Mouse Pancreas Dissection and Tissue Preparation

The detailed demonstration of the procedures required to successfully remove the pancreas from a mouse by dissection for histological analysis can be found here [11]. Separate the stomach and liver tissues from the pancreas, which are left attached to each other. Remove the pancreas and spread it out for examination (Fig. 1a, see Note 3).

3.2

Fixation

With the pancreas intact, incubate the sample in 25 mL ice-cold 4% PFA at 4  C, on the nutator, for 16 h (overnight). On the second day, wash samples with 25 mL ice-cold 1PBS for 1 h; repeat this washing step twice (total of three times). Again, incubate the sample in 25 mL ice-cold 4% PFA at 4  C, on the nutator, for 16 h (see Note 4).

3.3

Dehydration

1. Using a razor blade, cut the pancreas into approximately cubeshaped samples less than 5 mm on a side; this will ensure a good antibody penetration, and speed up quicker wash procedures (Fig. 1b). Length of sides can range from 2 to 10 mm, depending on the experimental goal. 2. Samples need to be dehydrated by washing in a methanol series of 25%, 50% and 75% methanol–PBS rocking at room

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Fig. 1 Mouse pancreas dissection and tissue preparation. (a) Example of a dissected 6-week-old adult pancreas. The spleen and duodenum are still attached to the pancreas. (b, c) Using a razor blade to cut the pancreas into approximately cube-shaped samples less than 5 mm on a side. Pan pancreas, Sp Spleen, Duo duodenum. Scale bar ¼ 50 mm

temperature. One hour of each wash in the methanol series is sufficient (see Note 5). 3. Wash sample twice in 100% methanol for 1 h gently rocking on the nutator at room temperature. Incubate samples with 25 mL Dent’s fix for 1 h at room temperature. Aspirate the Dent’s fix and wash samples with 25 mL 100% MeOH on the nutator for an hour. Wash one more time with 25 mL 100% MeOH. Once dehydrated to 100% methanol, sample should be stored at 20  C for at least 24 h before continuing with the protocol (see Note 6). 3.4

Bleaching

1. Remove the 100% MeOH from each tube. Add approximately 4 mL of Dent’s Bleach to each tube. Let the samples gently rock in the 4  C room for 2 h. 2. Aspirate the solution and wash with 100% MeOH for 1 h. This step needs to be repeated twice (total of 2 h wash).

3.5 Rehydration and Blocking

1. Remove samples stored in 100% methanol from 20  C. Rehydrate the samples in the reverse methanol series described above: (75%, 50%, 25% methanol–PBS) by rocking gently each solution for 1 h at room temperature. Complete rehydration with two washes of PBS for 1 h each, gently rocking. Remove the 1PBS and place each sample in a 5 mL test tube. Add approximately 4 mL of filtered-blocking buffer (up to 90% of the tube). 2. Seal the top with a cap and Parafilm. Let the samples rock overnight at 4  C.

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1. Remove the blocking buffer from samples using a P1000 pipette or a plastic dropper. 2. Prepare enough primary antibody solution for all of the samples you plan to stain. If we have two samples and we will add 1.5 mL to each. In a 15 mL conical tube prepare the following solution: 3 mL of PBS-X (0.15% Triton-X in PBS, add 3 μL of Goat anti-Spp1 and 6 μL of Rabbit anti-ChrA. 3. Transfer the samples to a new 1.5 mL cryotube. Add approx. 1.5 mL of primary antibody to each cryotube. 4. Parafilm the tube to make sure it is sealed. Leave the samples rocking gently at 4  C overnight (see Note 7). 5. Transfer the samples using a dropper to a 5 mL flow cytometry tube. 6. Wash with 1PBST for 1 h using the rocker at 4  C. This process needs to be repeated at least five times (see Note 8). 7. In this example we have two samples and we will add 1.5 mL to each. In a 15 mL conical tube prepare the following solution: 3 mL of PBS-X (0.15% Triton-X in PBS) þ 6 μL of Donkey anti goat conjugated with Cy3 + 6 μL of Donkey anti rabbit conjugated with Alexa 488 þ 3 μL DAPI. 8. Transfer the samples to a new 1.5 mL cryotube. Add approx. 1.5 mL of secondary antibody to each cryotube. Parafilm the tube to make sure it is sealed. Wrap the samples with aluminum foil. Leave the samples rocking gently at 4  C overnight. 9. Gently transfer the samples using a dropper to a 5 mL flow cytometry tube. Wash with 1PBS for 1 h using the rocker at 4  C. Repeat this step four times (total of 5, see Note 9). 10. Exam how the fluorescence signal–noise ratios using the fluorescence microscope. 11. If the samples look well washed, proceed and add 4% PFA to fix the secondary Abs with the pancreas cells. Fix samples for at least 1 h (see Note 10). 12. After fixation, wash the samples five times with 1PBST for 1 h, each time using the nutator at 4  C. You can either do this the same day or leave the samples in 1PBST overnight. 13. Wash the samples four times with 1PBST (15% Tween) for 1 h each time using the nutator at 4  C (see Note 11).

3.7

Dehydration

1. Dehydrate the samples by washing in ethanol (EtOH) series of 25%, 50%, 75%, 95%, and 100% EtOH–PBS rocking in the 4  C room. One hour of wash for each step in the ethanol series is sufficient. Finally, place the samples in fresh 100% EtOH; they can be used right away, or stored in a 20  C freezer for up to 2 years.

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2. After the dehydration is completed, check to see how the staining looks under the fluorescence microscope; if everything was done correctly, then the islets will be red clusters and the ducts will be branching green lines. 3.8 BABB Clearing Step

1. After completing the dehydration, place 3 mL of BABB solution in a new glass scintillation vial. Add the pancreas sample to the BABB solution; the sample will become transparent, with some yellow-brown color (this may take a few minutes). Note: if the sample does not become transparent after 10 min, replace the BABB solution. If the sample does not become transparent after 1 h, this may indicate that the dehydration was not complete; in this case, wash the samples in 100% EtOH for 1 h, and repeat the BABB cleaning steps. 2. Fill a double concave microscope slide glass with BABB solution; then, place the samples in the dip center. 3. Add a glass coverslip to seal the sample. The glass coverslip should stay on due to surface tension (see Note 12).

4

Notes 1. Detailed protocol for making 4% PFA in PBS can be found here: https://www.rndsystems.com/resources/protocols/pro tocol-making-4-formaldehyde-solution-pbs. 2. H2O2 is corrosive. Be careful! Sometimes adding the H2O2 will produce bubbles. Dent’s Bleach should always be made fresh. Dent’s Fix can be stored longer. All materials that touch the Dent’s Fix or Bleach should be discarded in the Biohazard waste. 3. Leave the spleen and duodenum attached to the pancreas for identification purposes. Rinse off blood with ice-cold PBS. 4. The time intervals mentioned in these protocols are a guideline and can be deviated from if necessary. 5. To determine if samples are sufficiently penetrated and acclimated to each wash, hold the tube upright. Samples that sink to the bottom of the tube are ready for the next wash; samples that float to the top need to have the solution changed, and to have that particular dehydration step repeated. 6. At this point, it is advised to store individual pancreas segments in 15 mL Falcon tubes at 20  C for long-term use; the segments can be used for whole mount IHCs for up to 1 year. 7. Longer incubation times of 1–2 days work too. In our experience, if an antibody has been used successfully on cryosections, then it is likely to work for WMIF.

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8. Sample can be washed for 2 days if necessary. 9. If too much background is present, wash multiple times using 1PBST (15% Tween) for 1 h each wash. Use the fluorescence microscope to check if the background looks reduced. The bigger the sample, the more likely they will require more washes. If necessary, leave the samples washing overnight. If the signal is significantly reduced due to the washes, redo the secondary antibodies staining process and leave them to stain overnight. 10. Make sure to optimize the stain prior to proceeding to fixation. 11. At this point, the samples can be processed with various optical tissue clearing (OTC) methods; these methods can be modularized with other imaging techniques. We had great success combining our WMIF process with the Scale method [12]. However, to reduce the time needed for the process, we recommend using conventional BABA clearing, as follows. 12. At this point, the sample should be ready for visualization/ analysis using a confocal microscope (Fig. 2). The sample will have become transparent upon treatment with BABB solution, so be careful to not lose track of it.

Fig. 2 Confocal whole-mount image of a 6-week-old mouse pancreas tissue. The sample was stained for ductal marker Spp1 (red), endocrine marker Chromogranin A (green), and DAPI (blue). The stained sample was cleared using benzyl alcohol–benzyl benzoate (BABB). Images were acquired using a Zeiss LSM 880 microscope followed by 3D rendering using Zen Blue software

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Acknowledgments We thank Dr. Brian Armstrong from the Light Microscopy Core at City of Hope for assistance in confocal microscopy. This work is supported by JDRF postdoctoral fellowship award P061008 to M. M.; the NIH/NIDDK award DK119590, DMRI start-up funding, and Wanek family funding to H.P.S. References 1. Gittes GK (2009) Developmental biology of the pancreas: a comprehensive review. Dev Biol 326:4–35. https://doi.org/10.1016/j. ydbio.2008.10.024 2. Shih HP, Wang A, Sander M (2013) Pancreas organogenesis: from lineage determination to morphogenesis. Annu Rev Cell Dev Biol 29:81–105. https://doi.org/10.1146/ annurev-cellbio-101512-122405 3. Bertelli E, Regoli M, Orazioli D, Bendayan M (2001) Association between islets of Langerhans and pancreatic ductal system in adult rat. Where endocrine and exocrine meet together? Diabetologia 44:575–584. https://doi.org/ 10.1007/s001250051663 4. Jorgensen MC et al (2007) An illustrated review of early pancreas development in the mouse. Endocr Rev 28:685–705. https://doi. org/10.1210/er.2007-0016 5. Kesavan G et al (2009) Cdc42-mediated tubulogenesis controls cell specification. Cell 139:791–801. https://doi.org/10.1016/j. cell.2009.08.049 6. Borden P, Houtz J, Leach SD, Kuruvilla R (2013) Sympathetic innervation during development is necessary for pancreatic islet architecture and functional maturation. Cell Rep 4:287–301. https://doi.org/10.1016/j.cel rep.2013.06.019

7. Reinert RB et al (2014) Vascular endothelial growth factor coordinates islet innervation via vascular scaffolding. Development 141:1480–1491. https://doi.org/10.1242/ dev.098657 8. Fowler JL et al (2018) Three-dimensional analysis of the human pancreas. Endocrinology 159:1393–1400. https://doi.org/10.1210/ en.2017-03076 9. Serrill JD, Sander M, Shih HP (2018) Pancreatic exocrine tissue architecture and integrity are maintained by E-cadherin during postnatal development. Sci Rep 8:13451. https://doi. org/10.1038/s41598-018-31603-2 10. Richardson DS, Lichtman JW (2015) Clarifying tissue clearing. Cell 162:246–257. https:// doi.org/10.1016/j.cell.2015.06.067 11. Veite-Schmahl MJ, Regan DP, Rivers AC, Nowatzke JF, Kennedy MA (2017) Dissection of the mouse pancreas for histological analysis and metabolic profiling. J Vis Exp (126). https://doi.org/10.3791/55647 12. Hama H et al (2015) ScaleS: an optical clearing palette for biological imaging. Nat Neurosci 18:1518–1529. https://doi.org/10.1038/ nn.4107

Chapter 17 Induction of Osteoblasts by Direct Reprogramming of Mouse Fibroblasts Hui Zhu and Joy Y. Wu Abstract In the tissue culture dish, osteoblast cells can be derived from mesenchymal stem cells (MSCs) and pluripotent stem cells (PSCs) including embryonic stem cells (ESCs) and induced pluripotent stem cells (iPSCs). However, differentiation of osteoblasts from PSCs is time-consuming and low yield. In contrast, we identified four osteogenic transcription factors, Runx2, Osx, Dlx5, and ATF4, that rapidly and efficiently reprogram mouse fibroblasts derived from 2.3 kb type I collagen promoter-driven green fluorescent protein (Col2.3GFP) transgenic mice into induced osteoblast cells (iOBs). iOBs exhibit osteoblast morphology, form mineralized nodules, and express Col2.3GFP and gene markers of osteoblast differentiation. Our method provides a robust system to rapidly generate appropriate and abundant osteoblast cells for osteogenesis and bone regeneration study. Key words Reprogramming, Fibroblast, Osteoblast, Bone

1

Introduction Osteoblasts are bone-forming cells that produce extracellular matrix proteins, including osteocalcin, alkaline phosphatase and type I collagen [1]. A subset of osteoblasts become embedded into bone matrix as osteocytes that account for 95% of cells in the mature bone tissues [1]. Osteoblasts lineage cells also contribute to the bone marrow microenvironment by secreting many of growth factors and cytokines to support hematopoiesis and immune cell development [2, 3]. During development, mesenchymal stem/ progenitor cells differentiate into preosteoblasts, mature osteoblasts, bone-lining cells, and osteocytes [1, 4]. Throughout adulthood, mesenchymal stem cells in the bone marrow continue to give rise to osteoblasts for bone remodeling and regeneration after bone defects [5]. In critical sized bone defects such as those due to trauma and tumors, bone tissue engineering techniques, including the use of differentiated osteoblast lineage cells from MSCs and PSCs,

Chrissa Kioussi (ed.), Stem Cells and Tissue Repair: Methods and Protocols, Methods in Molecular Biology, vol. 2155, https://doi.org/10.1007/978-1-0716-0655-1_17, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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scaffolds and inductive factors, represent a potential therapeutic strategy [6–12]. However, MSCs gradually lose multipotency and are limited by large donor variation [13, 14]. Differentiating osteoblast cells from PSCs is time-consuming, low yield, and carries the risk of teratoma formation if undifferentiated PSCs persist [15]. Therefore, a method that can rapidly and efficiently generate bona fide osteoblasts is essential for bone development and bone regeneration research. Osteoblast marker gene expression and mineralization of the surrounding extracellular matrix are two main assays used to characterize osteogenic differentiation [6]. However, osteogenic differentiation is very heterogeneous and only a small fraction of osteoblasts can yield positive results in assays of osteoblast gene expression and mineralization. We have generated mouse embryonic stem cell (ESC) lines and induced pluripotent stem cell (iPSC) lines from Col2.3GFP transgenic mice, in which GFP expression marks mature osteoblasts [16], and demonstrated that these PSC lines could be successfully differentiated to Col2.3GFP+ cells that are enriched in mature osteoblasts, albeit at low yield [15]. More recently, we harvested mouse fibroblasts from Col2.3GFP transgenic mice and demonstrated that the introduction of four osteoblast-specific transcription factors, Runx2, Osx, distal-less homeobox 5 (Dlx5), and activating transcription factor 4 (ATF4), could rapidly and efficiently convert mouse fibroblasts to induced osteoblasts (iOBs) [17] (Fig. 1A). iOBs form compact mineralized nodules expressing Col2.3GFP and osteoblast marker genes (Fig. 1B). Compared to the differentiation of Col2.3GFP+ osteoblasts from pluripotent stem cells, direct reprogramming of osteoblasts from fibroblasts is much faster (25 days vs 45 days) and more efficient (up to 4–5% vs 0.2% of the Col2.3GFP+ population). Our method provides a rapid and reliable method to study osteoblast differentiation for skeletal regeneration.

2

Materials

2.1 Tail-Tip Fibroblast Isolation and Culture

1. Hanks’ Balanced Salt Solution (HBSS): no calcium, no magnesium, no phenol red. 2. DMEM (high glucose, GlutaMAX™ Supplement, pyruvate). 3. Collagenase II. 4. Collagenase medium (2000 U/ml): Dissolve 30.77 mg Collagenase II (325 U/mg) in 5 ml DMEM or HBSS. Filter through a 0.22 μm filter unit. 5. 0.05% trypsin–EDTA. 6. Fetal bovine serum (FBS). 7. Penicillin–streptomycin.

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A Col2.3GFP mice

Tail-tip Col2.3GFP fibroblasts

Infection with lentivirus containing transcription factor genes

Col2.3GFP+ iOBs

B a

b

1000 um

e

c

1000 um

1000 um

1000 um

f

d

g

1000 um

h

1000 um

Fig. 1 (A) Schematic representation of generation iOBs. (B) At day 25 of reprogramming, iOBs exhibit compact cell clusters (a), cuboidal (b) and nodular (c) osteoblast-like cell morphology and GFP expression (e–g) (live GFP images). iOBs are positive for Alizarin S staining (d) and Von Kossa staining (h)

8. Fibroblast culture medium: DMEM, 10% FBS, 1% penicillin– streptomycin (10,000 U/ml). 9. Dimethyl sulfoxide (DMSO). 10. Freezing medium: 90% FBS, 10% DMSO. 2.2 Lentivirus Production

1. Doxycycline-inducible lentiviral vector containing mouse Runx2, Dlx5, Sp7/Osx, and Atf4 cDNA: pLV-tetO-Runx2; pLV-tetO-Dlx5; pLV-tetO-Sp7/Osx; pLV-tetO-ATF4; Made in house; provided upon request. 2. Doxycycline-inducible lentiviral empty vector for mock transduction: pLV-tetO; Made in house; provided upon request. 3. Lentiviral vector FUdeltaGW-rtTA expressing rtTA (reverse tetracycline-controlled transactivator): Addgene, Cat. No. 19780. 4. Lentiviral packaging vectors psPAX2 and pMAD2.G: Addgene; Cat. No. 16620 and 16619, respectively. 5. Lipofectamine 2000 reagent. 6. EMD Millipore™ Steriflip™ Sterile Disposable Vacuum Filter Units. 7. EMD Millipore™ Amicon™ Ultra-15 Centrifugal Filter Units.

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8. HEK293T culture medium: Same as Fibroblast culture medium. 9. Opti-MEM™ I Reduced Serum Medium (Opti-MEM). 2.3 Direct Reprogramming to Osteoblasts

1. The EmbryoMax 0.1% Gelatin Solution: EMD Millipore North America; Cat. No. ES-006-B. 2. Polybrene: Santa Cruz Biotech, CA; Cat. No. sc-134220. 3. Phosphate Buffered Saline (PBS). 4. L-Ascorbic Acid 2 Phosphate (AA); Dissolve 500 mg AA in 10 ml PBS to make a 1000 stock; sterilize by filtration, aliquot and store at 20  C. 5. β-Glycerol phosphate (β-GP); Dissolve 11.09 g β-GP in 50 ml PBS to make a 100 stock; sterilize by filtration, aliquot and store at 20  C. 6. Dexamethasone (Dex): Sigma, St. Louis, MO; Cat. No. D8893-1MG; Dissolve 1 mg Dex in 1 ml 100% ethanol to make 1 mg/ml (2.5 mM) solution; add 50 vol αMEM to make a 500 stock; aliquot and store at 20  C. 7. Doxycycline (Dox); Dissolve 50 mg Dox in 50 ml PBS to make a 5000 stock; sterilize by filtration, aliquot and store at 20  C. 8. αMEM. 9. Osteogenic medium: αMEM, 10% FBS, 1% penicillin–streptomycin (10,000 U/ml), 50 μg/ml L-ascorbic acid 2 phosphate, 2 μg/ml doxycycline; For calcium deposition and mineralization assays, add 10 mM β-glycerol phosphate and 100 nM dexamethasone.

2.4 Characterization of Induced Osteoblasts

1. 0.25% trypsin–EDTA (1), Phenol Red.

2.4.1 Detection Col2.3GFP+ by Flow Cytometry

3. Staining buffer: 1 PBS supplement with 2% FBS.

2.4.2 Calcium Deposition and Mineralization

1. Formalin Solution, neutral buffered, 10%.

2. Falcon® 70 μm Cell Strainer.

2. UltraPure RNase/DNase-free water (DDW). 3. 5% silver solution: Dissolve 25 g silver nitrate in 500 ml H2O. Keep from light and store at 4  C. 4. 5% sodium thiosulfate: Dissolve 5 g sodium thiosulfate in 10 ml H2O; Keep from light and store in room temperature. 5. Alizarin Red S staining solution: Dissolve 2 g Alizarin Red S in 100 ml H2O, mix and adjust pH to 4.1–4.3 with 0.1% ammonium hydroxide or 2 M hydrochloric acid. Filter through Whatman filter paper, keep from light and store at room temperature.

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1. TRIzol® Reagent. 2. PureLink® RNA Mini Kit. 3. PureLink® DNase Set. 4. iScript™ cDNA Synthesis Kit. 5. iQ™ SYBR® Green Supermix.

3

Methods

3.1 Isolation of Mouse Fibroblasts from Col2.3GFP Transgenic Mice Tail Tips

1. Sterilize mouse tail tip by using 70% ethanol. 2. Snip at least 1 cm of tail close to the tip (see Note 1). 3. Place the tail tip in PBS containing 1% penicillin–streptomycin or HBSS containing 1% penicillin–streptomycin. Bring the tail tip to a laminar flow tissue culture hood. 4. Place the tail tip in a 6 cm tissue culture dish, add 0.5 ml collagenase II (2000 U/ml). Mince the tail tip very well with sterile razor blade/scalpel into small pieces (see Note 2). 5. Add 0.5 ml DMEM or HBSS (final concentration after addition of collagenase II is 1000 U/ml). 6. Transfer 1 ml minced small pieces in diluted collagenase II solution to 15 ml conical vials. Incubate at 37  C in a heat block, shake at 500 rpm for 1 h. 7. Spin at 300  g for 5 min. Carefully discard supernatant. 8. Wash once with 1–3 ml HBSS by mixing and centrifuging as above, discarding the supernatant. 9. Add 1 ml 0.05% trypsin–EDTA, mix thoroughly, and incubate at 37  C in a heat block, shake at 500 rpm for 30 min. 10. Add 1 ml fibroblast culture medium to stop trypsinization. Centrifuge and decant supernatant as above, resuspend pellet in 1 ml fibroblast culture media. 11. Pipet up and down to break up aggregates. Centrifuge and decant supernatant as above. 12. Add 12 ml fibroblast culture medium and plate each tail tip suspension into two 6 cm tissue culture dishes avoiding large pieces of tissue. Each dish has 6 ml fibroblast culture medium. These are passage 0 fibroblasts. 13. Fibroblasts will migrate from the tail tip pieces and expand. Every 2–4 days, feed or split cultures at a ratio of 1:4 to 1:6. Harvest fibroblasts at passage 2, aliquot at 6  105 per frozen vial in freezing medium and freeze down (see Note 3).

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3.2 Lentivirus Production

1. Thaw HEK293T cells, expand and culture in HEK293T culture medium (see Note 4). 2. Seed HEK293T cells in 10 cm dish at density of 4.8–5.5  106/10 cm dish. 3. On the second day of seeding, the cell density should reach 70–80%. 4. Dilute 8 μg lentiviral vector pLV-tetO, pLV-tetO-Runx2, pLV-tetO-Dlx5, pLV-tetO-Sp7/Osx, pLV-tetO-ATF4, or FUdeltaGW-rtTA in 1 ml Opti-MEM. In each 1 ml OptiMEM, add 4 μg packaging vectors pMAD2.G and 6.67 μg psPAX2. 5. Dilute 50 μl Lipofectamine 2000 in 1 ml Opti-MEM. Incubate for 5 min. 6. Combine 1 ml Opti-MEM containing DNA and 1 ml OptiMEM containing Lipofectamine 2000. Incubate for at least 20 min. 7. For HEK293T cells, replace HEK293T culture medium with 6 ml DMEM (no FBS and no penicillin–streptomycin). 8. Add 2 ml DNA and Lipofectamine 2000 complex to 6 ml DMEM in the 10 cm dish, culture cells for 4–6 h. 9. After 4–6 h, change to HEK293T culture medium and culture HEK293T cells for 48 h. 10. After 48 h, the lentiviral particles are ready to harvest. Transfer 293T cell culture supernatant to a 50 ml falcon tube. Spin at 1,200  g for 15 min at room temperature. 11. Transfer the supernatant and filter by using Sterile Disposable Vacuum Filter Units. 12. Transfer filtered supernatant to Ultra-15 Centrifugal Filter Units. 15 ml per Centrifugal Filter Unit. Spin at 2,400  g for 45 min at 4  C. 13. After 45 min of centrifugation, lentiviral particles are concentrated. Transfer concentrated lentiviral particles from the bottom of Centrifugal Filter Units, aliquot, and store at 80  C (see Note 5).

3.3 Direct Reprogramming to Osteoblasts

1. Precoat 6-well tissue culture plate with 0.1% gelatin for 15 min. 1 ml 0.1% gelatin in each well. 2. Thaw a vial of Col2.3GFP fibroblasts (6  105/vial) into a 6-well tissue culture plate in fibroblast culture medium at 1  105 cells per well (day 0). 3. The following day change the medium to fibroblast culture medium containing lentiviral particles and 8 ng/ml polybrene (day 1). Each well in 6-well plate receives 10 μl of rtTA lentiviral particles and 4 μl of each pLV-tetO-Runx2, pLV-tetO-

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Dlx5, pLV-tetO-Sp7/Osx, and pLV-tetO-ATF4 lentiviral particles. For mock transduction, the well receives 10 μl of rtTA lentiviral particles and 16 μl pLV-tetO lentiviral particles. 4. After 24 h (day 2), wash out lentiviruses. Wash cells with PBS twice. Recover cells in fresh fibroblast culture medium. 5. On day 3, trypsinize cells by using 0.05% trypsin–EDTA and split (1:2) to new gelatin-coated 6-well plate. Culture cells in fibroblast culture medium for 48 h. 6. On day 5, replace the fibroblast culture medium by osteogenic medium. For calcium deposition and mineralization assays, add 10 mM β-glycerol phosphate and 100 nM dexamethasone (see Note 6). 7. Change osteogenic medium every 3 days. 3.4 Characterization of Induced Osteoblasts 3.4.1 Detection of Col2.3GFP+ Osteoblasts by Flow Cytometry

Col2.3GFP+ osteoblasts can be detected as early as day 8 after day 1 lentivirus transduction. At day 25, the frequency of Col2.3GFP+ iOB can be reached around 4–5% (Fig. 2). 1. Harvest cells with 0.25% trypsin–EDTA (see Note 7). Stop trypsinization with osteogenic medium. 2. Filter cells through a 70 μm cell strainer into 50 ml falcon tube. 3. Spin cells at 300  g for 5 min at room temperature. 4. Washed cells once with PBS and spin cells as above. 5. Suspend cells in 500 μl staining buffer for flow cytometry analysis.

Mock 10

SSC-A

10

10

10

Runx2-Dlx5-Osx-ATF4

5

10

4

10

3GFP+ Col2.3GFP+ 0

3

10

2

10

0 -10

5

4

3

Col2.3GFP+ 4.95

2

0 2

-10 0

10

3

FITC-A

10

4

10

5

2 0

10

3

10

4

10

5

FITC-A

Fig. 2 Flow cytometry analysis shows at day 25 of reprogramming, iOBs have around 4.95% of Col2.3GFP+ cells. Lentivirus containing pLV-tetO empty vector is used as mock transduction control

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3.4.2 Calcium Deposition by Alizarin Red S Staining

1. Rinsed cells with 1 PBS. 2. Add 10% formalin solution to cover cells, fixing cells for at least 10 min. 3. Carefully aspirate the formalin and wash cells with DDW twice. 4. Carefully aspirate DDW, add enough 2% Alizarin Red S staining solution (1 ml per well in 6-well plate) for 20 min in the dark. 5. Carefully remove Alizarin Red S staining solution. Wash cells with DDW three times. 6. Carefully aspirate DDW. Add 1 PBS and store in 4  C for up to 1 month.

3.4.3 Mineralization Assay by Von Kossa Staining

1. Rinsed cells with 1 PBS. 2. Add 10% formalin solution to cover cells, fixing cells for at least 10 min. 3. Carefully aspirate the formalin and wash cells with DDW twice. 4. Carefully aspirate DDW, add enough 5% silver solution (1 ml per well in 6-well plate). 5. Put cell plate under UV light till black staining is visible. This usually takes 30–45 min. 6. Carefully remove silver solution. Wash cells with DDW three times. 7. Add 5% sodium thiosulfate solution to remove un-reacted silver. Incubate for 5 min at room temperature. 8. Carefully aspirate sodium thiosulfate. Wash cells with DDW three times. 9. Carefully aspirate DDW. Add 1 PBS and store in 4  C for up to 1 month.

3.4.4 Gene Expression Analysis by Quantitative Reverse Transcription PCR (RT-qPCR)

The expression of osteoblast gene markers Col1a1, Bglap, and Ibsp are analyzed by RT-qPCR (Fig. 3). 1. Carefully aspirate cell culture medium. Add TRIzol® Reagent to the cells, 1 ml per well in 6-well plate. Incubate for 5 min in room temperature. 2. Detached cells by pipetting up and down, transfer to 1.5 ml Eppendorf tube. 3. Extract RNA by using PureLink® RNA Mini Kit. (a) Add 0.2 ml chloroform per 1 ml TRIzol® Reagent used. Shake the tube vigorously by hand for 15 s. (b) Incubate at room temperature for 2–3 min. (c) Spin the sample at 12,000  g for 15 min at 4  C.

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Fig. 3 qRT-PCR analysis shows at the day 25 of reprogramming, the expressions of osteoblast markers significantly increase in iOBs compared to mock transduction control. All data represent mean  SEM (One-way ANOVA; ∗ P < 0.05; #P < 0.0001; relative to mock)

(d) Transfer 400 μl of the colorless, upper phase containing the RNA to a fresh RNase-free tube. (e) Add an equal volume 70% ethanol to obtain a final ethanol concentration of 35%. Shake and upside down the tube vigorously by hand for 15 s. (f) Transfer 700 μl of sample to a Spin Cartridge (with a Collection Tube). (g) Spin at 12,000  g for 15 s at room temperature. (h) Discard the flow-through and reinsert the Spin Cartridge into the same Collection Tube. (i) Repeat steps (g) and (h) until the entire sample is processed. (j) Add 350 μl Wash Buffer I (provided by the kit) to the Spin Cartridge containing the bound RNA. Spin at 12,000  g for 15 s at room temperature. Discard the flow-through and the Collection Tube. Insert the Spin Cartridge into a new Collection Tube. (k) Add 80 μl PureLink® DNase mixture containing (8 μl 10 DNase I Reaction Buffer; 10 μl Resuspended DNase (3 U/μl); 62 μl RNase-free water) directly onto the surface of the Spin Cartridge membrane. Incubate at room temperature for 15 min. (l) Add 350 μl Wash Buffer I (provided by the kit) to the Spin Cartridge. Spin at 12,000  g for 15 s at room temperature. Discard flow-through and the Collection

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Tube and insert the Spin Cartridge into a new Collection Tube. (m) Add 500 μl Wash Buffer II with ethanol (provided by the kit) to the Spin Cartridge. (n) Spin at 12,000  g for 15 s at room temperature. Discard flow-through and reinsert the Spin Cartridge into the same Collection Tube. (o) Repeat steps (m) and (n). (p) Centrifuge the Spin Cartridge at 12,000  g for 1 min to dry the membrane with bound RNA. Discard Collection Tube and insert the Spin Cartridge into a Recovery Tube. (q) Add 30 μl RNase–Free Water to the center of the Spin Cartridge. (r) Incubate at room temperature for 1 min. Centrifuge Spin Cartridge and Recovery Tube for 1 min at 12,000  g at room temperature. 4. Measure RNA concentration and purity by NanoDrop. Store RNA at 80  C. 5. Proceed reverse transcription (RT) using iScript Kit. (a) Prepare RT reaction: 4 μl 5 iScript Reaction Mix (provided by the kit); 1 μl iScript Reverse Transcriptase (provided by the kit); 1 μg RNA; and Nuclease-free water (provided by the kit) to make up total 20 μl reaction volume. (b) Incubate the complete reaction mix in a thermal cycler using the program condition provided by the kit. (c) After RT, use 20 μl reaction mix as cDNA for PCR. 6. Quantitative PCR (qPCR). Primer sequences are described in Table 1. Normalize relative mRNA level to the levels of β-Actin. (a) Prepare qPCR reaction: 1 μl cDNA; 0.5 μl forward and reverse primer mix (10 μM); 12.5 μl SYBR mix; and 11 μl DDW. (b) Load to real time qPCR machine and perform real time qPCR using standard program. Use the annealing temperature at 55  C. Table 1 qPCR primer sequences Gene

Forward primer (50 –30 )

Reverse primer (50 –30 )

Col1a1

CACCCTCAAGAGCCTGAGTC

GTTCGGGCTGATGTACCAGT

Bglap

TCTCTCTGCTCACTCTGCTGGCC

TTTGTCAGACTCAGGGCCGC

Ibsp

TACCGGCCACGCTACTTTCTTTAT

GACCGCCAGCTCGTTTTCATCC

Reprogramming Osteoblasts

4

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Notes 1. It is easiest to reprogram fibroblasts from young pups (less than 10 days old). Use the whole tail if the mouse is adult >3 weeks, as it is harder to expand enough fibroblasts from older mice. 2. Prepare collagenase II solution freshly. The final activity concentration is 2000 U/ml. collagenase II digestion activity is usually supplied at around 300 U/mg. Calculate the amount of collagenase II based on the activity of each batch. 3. For better direct reprogramming efficiency, use fibroblast cells within two passages. 4. For better lentivirus production efficiency, use HEK293T cells within seven passages. 5. For each 15 ml HEK293T cell culture supernatant, after centrifuge, lentivirus can be concentrated to 250 μl. Based on this lentivirus concentration methods, we can produce lentivirus stocks with titers of 1  108 to 1  109 infectious units per milliliter. 6. For Col2.3GFP flow cytometry and RNA analysis, we use osteogenic medium. For calcium deposition assay by Alizarin Red S staining and mineralization assay by Von Kossa staining, we use osteogenic medium supplemented with 10 mM β-glycerol phosphate and 100 nM dexamethasone. 7. iOBs form compact cell clusters, cuboidal and mineralized nodules and can be hard to detach from the dish. Treat iOBs with 0.25% trypsin–EDTA for at least 10 min. Use 1 ml pipette tip to detach cells and pipet cells up and down hardly. If needed, use a cell lifter to help detach the cells.

Acknowledgments This work was supported by National Institutes of Health grant DP2OD008466 to J.Y.W. References 1. Long F (2011) Building strong bones: molecular regulation of the osteoblast lineage. Nat Rev Mol Cell Biol 13(1):27–38. https://doi. org/10.1038/nrm3254 2. Panaroni C, Tzeng YS, Saeed H, Wu JY (2014) Mesenchymal progenitors and the osteoblast lineage in bone marrow hematopoietic niches.

Curr Osteoporos Rep 12(1):22–32. https:// doi.org/10.1007/s11914-014-0190-7 3. Panaroni C, Wu JY (2013) Interactions between B lymphocytes and the osteoblast lineage in bone marrow. Calcif Tissue Int 93 (3):261–268. https://doi.org/10.1007/ s00223-013-9753-3

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4. Olsen BR, Reginato AM, Wang W (2000) Bone development. Annu Rev Cell Dev Biol 16:191–220. https://doi.org/10.1146/ annurev.cellbio.16.1.191 5. Prockop DJ (1997) Marrow stromal cells as stem cells for nonhematopoietic tissues. Science 276(5309):71–74. https://doi.org/10. 1126/science.276.5309.71 6. Wu JY (2015) Pluripotent stem cells and skeletal regeneration—promise and potential. Curr Osteoporos Rep 13(5):342–350. https://doi. org/10.1007/s11914-015-0285-9 7. Lou X (2015) Induced pluripotent stem cells as a new strategy for osteogenesis and bone regeneration. Stem Cell Rev Rep 11:645–651. https://doi.org/10.1007/ s12015-015-9594-8 8. Levi B, Hyun JS, Montoro DT, Lo DD, Chan CK, Hu S, Sun N, Lee M, Grova M, Connolly AJ, Wu JC, Gurtner GC, Weissman IL, Wan DC, Longaker MT (2012) In vivo directed differentiation of pluripotent stem cells for skeletal regeneration. Proc Natl Acad Sci U S A 109(50):20379–20384. https://doi.org/ 10.1073/pnas.12180521091218052109 9. Dimitriou R, Jones E, McGonagle D, Giannoudis PV (2011) Bone regeneration: current concepts and future directions. BMC Med 9:66. https://doi.org/10.1186/1741-70159-661741-7015-9-66 10. Both SK, van Apeldoorn AA, Jukes JM, Englund MC, Hyllner J, van Blitterswijk CA, de Boer J (2011) Differential bone-forming capacity of osteogenic cells from either embryonic stem cells or bone marrow-derived mesenchymal stem cells. J Tissue Eng Regen Med 5 (3):180–190. https://doi.org/10.1002/term. 303 11. Bilousova G, Jun du H, King KB, De Langhe S, Chick WS, Torchia EC, Chow KS, Klemm DJ,

Roop DR, Majka SM (2011) Osteoblasts derived from induced pluripotent stem cells form calcified structures in scaffolds both in vitro and in vivo. Stem Cells 29 (2):206–216. https://doi.org/10.1002/ stem.566 12. Li F, Bronson S, Niyibizi C (2010) Derivation of murine induced pluripotent stem cells (iPS) and assessment of their differentiation toward osteogenic lineage. J Cell Biochem 109 (4):643–652. https://doi.org/10.1002/jcb. 22440 13. Siddappa R, Licht R, van Blitterswijk C, de Boer J (2007) Donor variation and loss of multipotency during in vitro expansion of human mesenchymal stem cells for bone tissue engineering. J Orthop Res 25(8):1029–1041. https://doi.org/10.1002/jor.20402 14. Meijer GJ, de Bruijn JD, Koole R, van Blitterswijk CA (2007) Cell-based bone tissue engineering. PLoS Med 4(2):e9. https://doi.org/ 10.1371/journal.pmed.0040009. 06-PLMERIT-0352R2 [pii] 15. Zhu H, Kimura T, Swami S, Wu JY (2019) Pluripotent stem cells as a source of osteoblasts for bone tissue regeneration. Biomaterials 196:31–45. https://doi.org/10.1016/j. biomaterials.2018.02.009 16. Kalajzic I, Kalajzic Z, Kaliterna M, Gronowicz G, Clark SH, Lichtler AC, Rowe D (2002) Use of type I collagen green fluorescent protein transgenes to identify subpopulations of cells at different stages of the osteoblast lineage. J Bone Miner Res 17(1):15–25. https://doi.org/10.1359/jbmr.2002.17.1.15 17. Zhu H, Swami S, Yang P, Shapiro F, Wu J (2019) Direct reprogramming of mouse fibroblasts into functional osteoblasts. J Bone Miner Res. https://doi.org/10.1002/jbmr.3929

INDEX A

I

Adventitia ........................................................................ 77 Astrocytes ..................................................................41–59 Axons ........................................................... 100, 101, 104

Immunofluorescence ...............................................34–36, 55, 56, 89, 93, 105, 190, 193–199 Immunohistochemistry .......................................... 26, 30, 35, 119, 160–162 Induced pluripotent stem cells (iPSC)................... 11, 12, 15–17, 19, 31, 87–90, 184, 202 Innervation ..............................................................99–105 Islets ............................................................. 193, 194, 198

B Biopsy .........................................101, 119, 120, 159, 166 Bioscaffolds................................................................63–69 Bones ........................................2, 72, 116, 125, 201, 202

K

C Clustered regularly interspaced short palindromic repeats \CRISPR-associated protein 9 (CRISPR/Cas9) ............................................12, 13 Co-culture systems..................................................99–105 Colony forming unit analysis .............................. 127, 134 Cryopreservation..........................................151–163, 166

Kidney organoids ................................................. 183–191

D

Germline stem cells ...................................................1, 166

Magnetic-activated cell sorting (MACS) cell separation ............................................................. 75 Mesenchymal progenitor cell (MPC) ................. 125–138 Mesenchymal stem cells (MSC) ....................... 72, 76–78, 84, 85, 92 Mice ............................................................ 1, 3, 7, 12, 24, 25, 27, 29–31, 33–36, 41–59, 77, 84–89, 102, 103, 116–120, 122, 127, 128, 130, 142–144, 148–154, 157–162, 166–169, 172, 175, 177, 178, 180, 185, 194, 195, 201–209 Microfluidics............................................................99–105 Mouse embryoid bodies ............................................... 2, 6 Mouse embryonic fibroblasts (MEF).......................... 2–4, 86, 159, 167 Mouse embryonic stem cells .........................1–8, 93, 202 Muscle satellite cells (MuSC) .............................. 141–150 Myogenesis .................................................................... 142

H

N

Histology .............................................................. 121, 193 HOX ........................................................................ 84, 87, 89, 90, 93 Human pluripotent stem cells (hPSCs) ................. 24, 25, 183–191

Nanofibers ............................................................ 115–122 Nerve injuries .................................................................. 63 Neurons ............................................................. 24–27, 36, 41–59, 64, 66–69, 100 Nucleofection ..................................................... 13–18, 20

Directed differentiation ......................................... 25, 184 Dysphagia ............................................................. 107–112

F Feeder cells ..........................................166–168, 172, 173 Fibroblasts ......................................................... 50, 84–89, 92, 93, 115, 116, 153, 167, 174, 180, 201–209 Flow cytometry ....................................................... 76, 78, 87–91, 93, 94, 125, 127, 129, 130, 134, 138, 197, 204, 207, 209 Fluorescent protein ..........................................12, 93, 177

G

L Lung MPCs .......................................................... 125, 126

M

Chrissa Kioussi (ed.), Stem Cells and Tissue Repair: Methods and Protocols, Methods in Molecular Biology, vol. 2155, https://doi.org/10.1007/978-1-0716-0655-1, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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STEM CELLS

214 Index

AND

TISSUE REPAIR: METHODS

AND

PROTOCOLS

O Osteoblasts ...................................................125, 201–209

P Pancreas ........................................................143, 193–199 Protein assay .................................................................. 177

R Regeneration ....................................................... 1, 24, 42, 43, 63–69, 71, 83, 99, 100, 107–112, 116, 126, 142, 184, 201, 202 Reprogramming .......................................................41–59, 86–89, 92, 93, 201–209

S Self-renewal .................................. 72, 142, 143, 159, 167 Silicone splints ............................................. 117, 120, 122 Skeletal muscles .................................................... 141–150 Skin .......................................................................... 33, 34, 103, 109, 110, 115–122, 128, 144, 168 Spermatogenesis .................................................. 151, 153, 160, 163, 165, 166, 177–179 Spermatogonial stem cells (SSC) ....................... 151–163, 165–169

Spermatogonial transplantation159, 160, 166, 167, 170, 173, 177–179 Spinal cord injuries.......................................................... 63 Stem cells ................................................ 1, 11–20, 23–37, 42, 43, 54, 59, 63–69, 71–79, 83–95, 99, 108, 115, 141, 151, 153, 165, 167, 183, 186–187, 201, 202 Stem cells from human exfoliated deciduous teeth (SHED).............................................................. 108

T Teeth ............................................. 99–105, 108, 110, 116 Testis151, 153–155, 158, 160, 161, 163, 166, 172, 178, 179 3D imaging.................................................. 193, 194, 199 Transdifferentiation............................................ 46, 48, 58

V Vascular stem cell ..................................... v, 71–78, 83–94 Vascular wall .................................................71–78, 83–94 Vasculogenic zone ........................................................... 72

W Wound healing ..................................................... 115–120