Skin Stem Cells: Methods and Protocols [Hardcover ed.] 1493988697, 9781493988693

Table of contents :
Front Matter ....Pages i-xv
In Vivo Genetic Alteration and Lineage Tracing of Single Stem Cells by Live Imaging (Olivia Farrelly, Paola Kuri, Panteleimon Rompolas)....Pages 1-14
Ex Vivo Imaging and Genetic Manipulation of Mouse Hair Follicle Bulge Stem Cells (Daniel Haensel, Melissa A. McNeil, Xing Dai)....Pages 15-29
Quantitative Detection of Low-Abundance Transcripts at Single-Cell Level in Human Epidermal Keratinocytes by Digital Droplet Reverse Transcription-Polymerase Chain Reaction (Frédéric Auvré, Julien Coutier, Michèle T. Martin, Nicolas O. Fortunel)....Pages 31-41
Qualitatively Monitoring Binding and Expression of the Transcription Factors Sp1 and NFI as a Useful Tool to Evaluate the Quality of Primary Cultured Epithelial Stem Cells in Tissue Reconstruction (Gaëtan Le-Bel, Sergio Cortez Ghio, Danielle Larouche, Lucie Germain, Sylvain L. Guérin)....Pages 43-73
Ribonucleoproteins Mediated Efficient In Vivo Gene Editing in Skin Stem Cells (Wenbo Wu, Ting Chen)....Pages 75-86
Isolation and Characterization of Cutaneous Epithelial Stem Cells (Stephanie R. Gillespie, David M. Owens)....Pages 87-99
Interfollicular Epidermal Stem Cells: Boosting and Rescuing from Adult Skin (Mariana T. Cerqueira, Ana M. Frias, Rui L. Reis, Alexandra P. Marques)....Pages 101-110
Whole-Mount Immunofluorescent Staining Coupled to Multicolor Lineage Tracing Model for Analyzing the Spatiotemporal Organization of Epidermal Stem Cells (Edwige Roy, Kiarash Khosrotehrani)....Pages 111-118
Isolation and Enrichment of Newborn and Adult Skin Stem Cells of the Interfollicular Epidermis (Stefano Sol, Dario Antonini, Caterina Missero)....Pages 119-132
Isolation and Cultivation of Epidermal (Stem) Cells (Xusheng Wang, Shiyang Dong, Yaojiong Wu)....Pages 133-138
One-Step Simple Isolation Method to Obtain Both Epidermal and Dermal Stem Cells from Human Skin Specimen (Hua Qian, Xue Leng, Jie Wen, Qian Zhou, Xin Xu, Xunwei Wu)....Pages 139-148
Isolation and Cultivation of Skin-Derived Precursors (Xiaoxiao Wang, Shiyang Dong, Yaojiong Wu)....Pages 149-152
Magnetic-Based Cell Isolation Technique for the Selection of Stem Cells (Petek Korkusuz, Sevil Köse, Nilgün Yersal, Selin Önen)....Pages 153-163
Isolation of Human Skin Epidermal Stem Cells Based on the Expression of Endothelial Protein C Receptor (Meilang Xue, Suat Dervish, Christopher J. Jackson)....Pages 165-174
Decellularized bSIS-ECM as a Regenerative Biomaterial for Skin Wound Repair (Mahmut Parmaksiz, Ayşe Eser Elçin, Yaşar Murat Elçin)....Pages 175-185
Protocols for Full Thickness Skin Wound Repair Using Prevascularized Human Mesenchymal Stem Cell Sheet (Lei Chen, Daniel Radke, Shaohai Qi, Feng Zhao)....Pages 187-200
Cultivation of Adipose-Derived Stromal Cells on Intact Amniotic Membrane-Based Scaffold for Skin Tissue Engineering (Ehsan Taghiabadi, Bahareh Beiki, Nasser Aghdami, Amir Bajouri)....Pages 201-210
Amniotic Membrane Seeded Fetal Fibroblasts as Skin Substitute for Wound Regeneration (Ehsan Taghiabadi, Bahareh Beiki, Nasser Aghdami, Amir Bajouri)....Pages 211-219
Skin Wound Healing: Refractory Wounds and Novel Solutions (Gabriel M. Virador, Lola de Marcos, Victoria M. Virador)....Pages 221-241
Isolation, Culture, and Motility Measurements of Epidermal Melanocytes from GFP-Expressing Reporter Mice (Lina Dagnino, Melissa Crawford)....Pages 243-256
Melanoblasts as Multipotent Cells in Murine Skin (Tsutomu Motohashi, Takahiro Kunisada)....Pages 257-266
Regeneration of Mouse Skin Melanocyte Stem Cells In Vivo and In Vitro (Ke Yang, Weiming Qiu, Pei-Rong Gu, Mingxing Lei)....Pages 267-284
Interactions Between Epidermal Keratinocytes, Dendritic Epidermal T-Cells, and Hair Follicle Stem Cells (Krithika Badarinath, Abhik Dutta, Akshay Hegde, Neha Pincha, Rupali Gund, Colin Jamora)....Pages 285-297
Isolating Immune Cells from Mouse Embryonic Skin (Ambika S. Kurbet, Srikala Raghavan)....Pages 299-305
Direct Conversion of Mouse Embryonic Fibroblasts into Neural Crest Cells (Tsutomu Motohashi, Takahiro Kunisada)....Pages 307-321
High-Titer Production of HIV-Based Lentiviral Vectors in Roller Bottles for Gene and Cell Therapy (Hazal Banu Olgun, Hale M. Tasyurek, Ahter Dilsad Sanlioglu, Salih Sanlioglu)....Pages 323-345
High-Grade Purification of Third-Generation HIV-Based Lentiviral Vectors by Anion Exchange Chromatography for Experimental Gene and Stem Cell Therapy Applications (Hazal Banu Olgun, Hale M. Tasyurek, Ahter D. Sanlioglu, Salih Sanlioglu)....Pages 347-365
Full-Thickness Human Skin Equivalent Models of Atopic Dermatitis (Gopu Sriram, Paul Lorenz Bigliardi, Mei Bigliardi-Qi)....Pages 367-383
Human Hair Follicle Associated-Pluripotent (hHAP) Stem Cells Differentiate to Cardiac Muscle Cells (Robert M. Hoffman)....Pages 385-392
Isolation of Normal Fibroblasts and Their Cancer-Associated Counterparts (CAFs) for Biomedical Research (BarboraBarbora Dvořánková, Lukáš Lacina, Karel Smetana Jr.)....Pages 393-406
Isolation of Cancer Stem Cells from Squamous Cell Carcinoma (Silvia Fontenete, Mirna Perez-Moreno)....Pages 407-414
Identification of Human Cutaneous Squamous Cell Carcinoma Cancer Stem Cells (Carlotta Olivero, Huw Morgan, Girish K. Patel)....Pages 415-433
Identification of Human Cutaneous Basal Cell Carcinoma Cancer Stem Cells (Huw Morgan, Carlotta Olivero, Girish K. Patel)....Pages 435-450
Back Matter ....Pages 451-463

Citation preview

Methods in Molecular Biology 1879

Kursad Turksen Editor

Skin Stem Cells 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, AL10 9AB, UK

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

Skin Stem Cells Methods and Protocols Second Edition

Edited by

Kursad Turksen Ottawa, ON, Canada

Editor Kursad Turksen Ottawa, ON, Canada

ISSN 1064-3745 ISSN 1940-6029 (electronic) ISBN 978-1-4939-8869-3 ISBN 978-1-4939-8870-9 (eBook) https://doi.org/10.1007/978-1-4939-8870-9 Library of Congress Control Number: 2018965200 © Springer Science+Business Media, LLC, part of Springer Nature 2019 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, express 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 Press imprint is published by the registered company Springer Science+Business Media, LLC part of Springer Nature. The registered company address is: 233 Spring Street, New York, NY 10013, U.S.A.

Preface Skin functions as the first defense against physical, chemical, and biological insults. In response to injuries resulting from such injuries, stem cells of the epidermal and dermal lineages need to activate a robust response to repair and maintain skin function. Over the last decade or so, efforts to understand the mechanisms underlying stem cell-derived repair have led to the isolation of numerous stem cell-like subpopulations from the epidermis and dermis. In this second edition of Skin Stem Cells: Methods and Protocols, I sought updated and new contributions from investigators who are pushing the field forward. It is my hope that this collection of protocols will be valuable both to people already in the field and to those now wishing to embark on studies of skin stem cells. I thank Patrick J. Marton, Executive Editor, Springer Protocols, for being available and supportive of the project from the beginning. It has been wonderful to work with David C. Casey, Editor of the Methods in Molecular Biology series. He is always there to provide support and ensure that there are no missing details that I may have overlooked. Thank you, David. A special thank you goes to Sarumathi Hemachandrane, Anand Ventakachalam, and the rest of the production group for making production of the volume seamless. Finally, I am very grateful to all the investigators who contributed to this volume. Their generosity and willingness to share often very hard-won approaches have made this volume possible. Ottawa, ON, Canada

Kursad Turksen

v

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

In Vivo Genetic Alteration and Lineage Tracing of Single Stem Cells by Live Imaging. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Olivia Farrelly, Paola Kuri, and Panteleimon Rompolas Ex Vivo Imaging and Genetic Manipulation of Mouse Hair Follicle Bulge Stem Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Daniel Haensel, Melissa A. McNeil, and Xing Dai Quantitative Detection of Low-Abundance Transcripts at Single-Cell Level in Human Epidermal Keratinocytes by Digital Droplet Reverse Transcription-Polymerase Chain Reaction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Fre´de´ric Auvre´, Julien Coutier, Miche`le T. Martin, and Nicolas O. Fortunel Qualitatively Monitoring Binding and Expression of the Transcription Factors Sp1 and NFI as a Useful Tool to Evaluate the Quality of Primary Cultured Epithelial Stem Cells in Tissue Reconstruction . . . . . . . . . . . . . . . . . . . . . . . . . Gae¨tan Le-Bel, Sergio Cortez Ghio, Danielle Larouche, Lucie Germain, and Sylvain L. Gue´rin Ribonucleoproteins Mediated Efficient In Vivo Gene Editing in Skin Stem Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Wenbo Wu and Ting Chen Isolation and Characterization of Cutaneous Epithelial Stem Cells . . . . . . . . . . . . . . . . Stephanie R. Gillespie and David M. Owens Interfollicular Epidermal Stem Cells: Boosting and Rescuing from Adult Skin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Mariana T. Cerqueira, Ana M. Frias, Rui L. Reis, and Alexandra P. Marques Whole-Mount Immunofluorescent Staining Coupled to Multicolor Lineage Tracing Model for Analyzing the Spatiotemporal Organization of Epidermal Stem Cells. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Edwige Roy and Kiarash Khosrotehrani Isolation and Enrichment of Newborn and Adult Skin Stem Cells of the Interfollicular Epidermis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Stefano Sol, Dario Antonini, and Caterina Missero Isolation and Cultivation of Epidermal (Stem) Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Xusheng Wang, Shiyang Dong, and Yaojiong Wu One-Step Simple Isolation Method to Obtain Both Epidermal and Dermal Stem Cells from Human Skin Specimen. . . . . . . . . . . . . . . . . . . . . . . . . . . . . Hua Qian, Xue Leng, Jie Wen, Qian Zhou, Xin Xu, and Xunwei Wu Isolation and Cultivation of Skin-Derived Precursors . . . . . . . . . . . . . . . . . . . . . . . . . . . . Xiaoxiao Wang, Shiyang Dong, and Yaojiong Wu

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15

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43

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Magnetic-Based Cell Isolation Technique for the Selection of Stem Cells . . . . . . . . . . ¨ nen ¨ n Yersal, and Selin O Petek Korkusuz, Sevil Ko¨se, Nilgu Isolation of Human Skin Epidermal Stem Cells Based on the Expression of Endothelial Protein C Receptor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Meilang Xue, Suat Dervish, and Christopher J. Jackson Decellularized bSIS-ECM as a Regenerative Biomaterial for Skin Wound Repair . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Mahmut Parmaksiz, Ays¸e Eser Elc¸in, and Yas¸ar Murat Elc¸in Protocols for Full Thickness Skin Wound Repair Using Prevascularized Human Mesenchymal Stem Cell Sheet . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Lei Chen, Daniel Radke, Shaohai Qi, and Feng Zhao Cultivation of Adipose-Derived Stromal Cells on Intact Amniotic Membrane-Based Scaffold for Skin Tissue Engineering . . . . . . . . . . . . . . . . . . . . . . . . . . Ehsan Taghiabadi, Bahareh Beiki, Nasser Aghdami, and Amir Bajouri Amniotic Membrane Seeded Fetal Fibroblasts as Skin Substitute for Wound Regeneration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ehsan Taghiabadi, Bahareh Beiki, Nasser Aghdami, and Amir Bajouri Skin Wound Healing: Refractory Wounds and Novel Solutions . . . . . . . . . . . . . . . . . . . Gabriel M. Virador, Lola de Marcos, and Victoria M. Virador Isolation, Culture, and Motility Measurements of Epidermal Melanocytes from GFP-Expressing Reporter Mice . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Lina Dagnino and Melissa Crawford Melanoblasts as Multipotent Cells in Murine Skin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Tsutomu Motohashi and Takahiro Kunisada Regeneration of Mouse Skin Melanocyte Stem Cells In Vivo and In Vitro. . . . . . . . . . Ke Yang, Weiming Qiu, Pei-Rong Gu, and Mingxing Lei Interactions Between Epidermal Keratinocytes, Dendritic Epidermal T-Cells, and Hair Follicle Stem Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Krithika Badarinath, Abhik Dutta, Akshay Hegde, Neha Pincha, Rupali Gund, and Colin Jamora Isolating Immune Cells from Mouse Embryonic Skin. . . . . . . . . . . . . . . . . . . . . . . . . . . . Ambika S. Kurbet and Srikala Raghavan Direct Conversion of Mouse Embryonic Fibroblasts into Neural Crest Cells. . . . . . . . Tsutomu Motohashi and Takahiro Kunisada High-Titer Production of HIV-Based Lentiviral Vectors in Roller Bottles for Gene and Cell Therapy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Hazal Banu Olgun, Hale M. Tasyurek, Ahter Dilsad Sanlioglu, and Salih Sanlioglu High-Grade Purification of Third-Generation HIV-Based Lentiviral Vectors by Anion Exchange Chromatography for Experimental Gene and Stem Cell Therapy Applications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Hazal Banu Olgun, Hale M. Tasyurek, Ahter D. Sanlioglu, and Salih Sanlioglu

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243 257 267

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Contents

ix

Full-Thickness Human Skin Equivalent Models of Atopic Dermatitis . . . . . . . . . . . . . . Gopu Sriram, Paul Lorenz Bigliardi, and Mei Bigliardi-Qi Human Hair Follicle Associated-Pluripotent (hHAP) Stem Cells Differentiate to Cardiac Muscle Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Robert M. Hoffman Isolation of Normal Fibroblasts and Their Cancer-Associated Counterparts (CAFs) for Biomedical Research . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Barbora Dvorˇa´nkova´, Luka´ˇs Lacina, and Karel Smetana Jr. Isolation of Cancer Stem Cells from Squamous Cell Carcinoma. . . . . . . . . . . . . . . . . . . Silvia Fontenete and Mirna Perez-Moreno Identification of Human Cutaneous Squamous Cell Carcinoma Cancer Stem Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Carlotta Olivero, Huw Morgan, and Girish K. Patel Identification of Human Cutaneous Basal Cell Carcinoma Cancer Stem Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Huw Morgan, Carlotta Olivero, and Girish K. Patel

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

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Contributors NASSER AGHDAMI  Department of Regenerative Biomedicine, Cell Science Research Center, Royan Institute for Stem Cell Biology and Technology, ACECR, Tehran, Iran DARIO ANTONINI  Department of Biology, University of Naples Federico II, Napoli, Italy; CEINGE Biotecnologie Avanzate (Center for Genetic Engineering), Napoli, Italy FRE´DE´RIC AUVRE´  Laboratoire de Ge´nomique et Radiobiologie de la Ke´ratinopoı¨e`se, CEA/ DRF/IBFJ/IRCM, Evry, France; INSERM U967, Fontenay-aux-Roses, France; Universite´ Paris-Diderot, Paris, France; Universite´ Paris-Saclay, Paris, France KRITHIKA BADARINATH  IFOM-inStem Joint Research Laboratory, Centre for Inflammation and Tissue Homeostasis, Institute for Stem Cell Biology and Regenerative Medicine, Bangalore, Karnataka, India; National Centre for Biological Sciences, Bangalore, Karnataka, India AMIR BAJOURI  Department of Regenerative Biomedicine, Cell Science Research Center, Royan Institute for Stem Cell Biology and Technology, ACECR, Tehran, Iran BAHAREH BEIKI  Skin and Stem Cell Research Center, Tehran University of Medical Sciences, Tehran, Iran PAUL LORENZ BIGLIARDI  Department of Dermatology, University of Minnesota, Minneapolis, MN, USA MEI BIGLIARDI-QI  Department of Dermatology, University of Minnesota, Minneapolis, MN, USA MARIANA T. CERQUEIRA  3B’s Research Group—Biomaterials, Biodegradable and Biomimetics, Avepark—Parque de Cieˆncia e Tecnologia, Zona Industrial da Gandra, Guimara˜es, Portugal; ICVS/3B’s—, PT Government Associate Laboratory, Braga, Portugal LEI CHEN  Department of Burns, First Affiliated Hospital of Sun Yat-sen University, Guangzhou, China TING CHEN  National Institute of Biological Sciences, Beijing, China JULIEN COUTIER  Laboratoire de Ge´nomique et Radiobiologie de la Ke´ratinopoı¨e`se, CEA/ DRF/IBFJ/IRCM, Evry, France; INSERM U967, Fontenay-aux-Roses, France; Universite´ Paris-Diderot, Paris, France; Universite´ Paris-Saclay, Paris, France MELISSA CRAWFORD  Department of Physiology and Pharmacology, Child Health Research Institute and Lawson Health Research Institute, University of Western Ontario, London, ON, Canada LINA DAGNINO  Department of Physiology and Pharmacology, Child Health Research Institute and Lawson Health Research Institute, University of Western Ontario, London, ON, Canada; Department of Physiology and Pharmacology, University of Western Ontario, London, ON, Canada XING DAI  Department of Biological Chemistry, School of Medicine, University of California, Irvine, CA, USA LOLA DE MARCOS  University of Navarra, Pamplona, Navarra, Spain SUAT DERVISH  The Westmead Institute, The University of Sydney, Westmead, NSW, Australia SHIYANG DONG  Graduate School at Shenzhen, Tsinghua University, Shenzhen, China

xi

xii

Contributors

ABHIK DUTTA  IFOM-inStem Joint Research Laboratory, Centre for Inflammation and Tissue Homeostasis, Institute for Stem Cell Biology and Regenerative Medicine, Bangalore, Karnataka, India BARBORA DVORˇA´NKOVA´  First Faculty of Medicine, Institute of Anatomy and BIOCEV, Charles University, Prague, Czech Republic AYS¸E ESER ELC¸IN  Tissue Engineering, Biomaterials and Nanobiotechnology Laboratory, Ankara University, Faculty of Science and Stem Cell Institute, Ankara, Turkey YAS¸AR MURAT ELC¸IN  Tissue Engineering, Biomaterials and Nanobiotechnology Laboratory, Ankara University, Faculty of Science, Ankara, Turkey; Biovalda Health Technologies, Inc., Ankara, Turkey OLIVIA FARRELLY  Department of Dermatology, Institute for Regenerative Medicine, University of Pennsylvania Perelman School of Medicine, Philadelphia, PA, USA SILVIA FONTENETE  Section for Cell Biology and Physiology, Department of Biology, Faculty of Science, University of Copenhagen, Copenhagen Ø, Denmark NICOLAS O. FORTUNEL  Laboratoire de Ge´nomique et Radiobiologie de la Ke´ratinopoı¨e`se, CEA/DRF/IBFJ/IRCM,, Evry, France; INSERM U967, Fontenay-aux-Roses, France; Universite´ Paris-Diderot, Paris, France; Universite´ Paris-Saclay, Paris, France ANA M. FRIAS  3B’s Research Group—Biomaterials, Biodegradable and Biomimetics, Avepark—Parque de Cieˆncia e Tecnologia, Zona Industrial da Gandra, Guimara˜es, Portugal; ICVS/3B’s—PT Government Associate Laboratory, Braga, Portugal LUCIE GERMAIN  Centre Universitaire d’Ophtalmologie—Recherche (CUO Recherche), CHU de Que´bec, Que´bec, QC, Canada; Centre de recherche en organoge´ne`se expe´rimentale de l’Universite´ Laval/LOEX, CHU de Que´bec—Universite´ Laval Research Center, Que´bec, QC, Canada; Department of Surgery, Faculty of Medicine, Universite´ Laval, Que´bec, QC, Canada; Department of Ophthalmology, Faculty of Medicine, Universite´ Laval, Que´bec, QC, Canada SERGIO CORTEZ GHIO  Centre de recherche en organoge´ne`se expe´rimentale de l’Universite´ Laval/LOEX, CHU de Que´bec—Universite´ Laval Research Center, Que´bec, QC, Canada; Department of Surgery, Faculty of Medicine, Universite´ Laval, Que´bec, QC, Canada STEPHANIE R. GILLESPIE  Department of Dermatology, College of Physicians & Surgeons, Columbia University Medical Center, New York, NY, USA SYLVAIN L. GUE´RIN  Centre Universitaire d’Ophtalmologie—Recherche (CUO Recherche), CHU de Que´bec, Que´bec, QC, Canada; Department of Ophthalmology, Faculty of Medicine, Universite´ Laval, Que´bec, QC, Canada RUPALI GUND  IFOM-inStem Joint Research Laboratory, Centre for Inflammation and Tissue Homeostasis, Institute for Stem Cell Biology and Regenerative Medicine, Bangalore, Karnataka, India; Department of Dermatology, Columbia University, New York, NY, USA PEI-RONG GU  Integrative Stem Cell Center, China Medical University Hospital, China Medical University, Taichung, Taiwan DANIEL HAENSEL  Department of Biological Chemistry, School of Medicine, University of California, Irvine, CA, USA AKSHAY HEGDE  IFOM-inStem Joint Research Laboratory, Centre for Inflammation and Tissue Homeostasis, Institute for Stem Cell Biology and Regenerative Medicine, Bangalore, Karnataka, India; Shanmugha Arts, Science, Technology and Research Academy (SASTRA) University, Thanjavur, Tamil Nadu, India ROBERT M. HOFFMAN  AntiCancer Inc., San Diego, CA, USA; Department of Surgery, University of California, San Diego, CA, USA

Contributors

xiii

CHRISTOPHER J. JACKSON  Sutton Arthritis Research Laboratory, Kolling Institute of Medical Research, The University of Sydney at Royal North Shore Hospital, St Leonards, NSW, Australia COLIN JAMORA  IFOM-inStem Joint Research Laboratory, Centre for Inflammation and Tissue Homeostasis, Institute for Stem Cell Biology and Regenerative Medicine, Bangalore, Karnataka, India KIARASH KHOSROTEHRANI  The University of Queensland, UQ Diamantina Institute, Translational Research Institute, Brisbane, QLD, Australia PETEK KORKUSUZ  Hacettepe University, Faculty of Medicine, Department of Histology and Embryology, Ankara, Turkey SEVIL KO¨SE  Atilim University, Faculty of Health Sciences, Department of Nutrition and Dietetics, Ankara, Turkey TAKAHIRO KUNISADA  Department of Tissue and Organ Development, Regeneration and Advanced Medical Science, Gifu University Graduate School of Medicine, Gifu, Japan AMBIKA S. KURBET  SASTRA University, Thanjavur, Tamil Nadu, India; Institute for Stem Cell Biology and Regenerative Medicine, Bangalore, India PAOLA KURI  Department of Dermatology, Institute for Regenerative Medicine, University of Pennsylvania Perelman School of Medicine, Philadelphia, PA, USA LUKA´Sˇ LACINA  First Faculty of Medicine, Institute of Anatomy and BIOCEV, Charles University, Prague, Czech Republic DANIELLE LAROUCHE  Centre de recherche en organoge´ne`se expe´rimentale de l’Universite´ Laval/LOEX, CHU de Que´bec—Universite´ Laval Research Center, Que´bec, QC, Canada; Department of Surgery, Faculty of Medicine, Universite´ Laval, Que´bec, QC, Canada GAE¨TAN LE-BEL  Centre Universitaire d’Ophtalmologie—Recherche (CUO Recherche), CHU de Que´bec, Que´bec, QC, Canada; Centre de recherche en organoge´ne`se expe´rimentale de l’Universite´ Laval/LOEX, CHU de Que´bec—Universite´ Laval Research Center, Que´bec, QC, Canada; Department of Surgery, Faculty of Medicine, Universite´ Laval, Que´bec, QC, Canada; Department of Ophthalmology, Faculty of Medicine, Universite´ Laval, Que´bec, QC, Canada MINGXING LEI  Integrative Stem Cell Center, China Medical University Hospital, China Medical University, Taichung, Taiwan; Institute of New Drug Development, College of Biopharmaceutical and Food Sciences, China Medical University, Taichung, Taiwan XUE LENG  Laboratory for Tissue Engineering and Regeneration, and Shandong Provincial Key Laboratory of Oral Tissue Regeneration, School of Stomatology, Shandong University, Jinan, Shandong, China ALEXANDRA P. MARQUES  3B’s Research Group—Biomaterials, Biodegradable and Biomimetics, Avepark—Parque de Cieˆncia e Tecnologia, Zona Industrial da Gandra, Guimara˜es, Portugal; ICVS/3B’s—, PT Government Associate Laboratory, Braga, Portugal; The Discoveries Centre for Regenerative and Precision Medicine, Headquarters at University of Minho, Guimara˜es, Portugal MICHE`LE T. MARTIN  Laboratoire de Ge´nomique et Radiobiologie de la Ke´ratinopoı¨e`se, CEA/DRF/IBFJ/IRCM, Evry, France; INSERM U967, Fontenay-aux-Roses, France; Universite´ Paris-Diderot, Paris, France; Universite´ Paris-Saclay, Paris, France MELISSA A. MCNEIL  Department of Biological Chemistry, School of Medicine, University of California, Irvine, CA, USA CATERINA MISSERO  Department of Biology, University of Naples Federico II, Napoli, Italy; CEINGE Biotecnologie Avanzate (Center for Genetic Engineering), Napoli, Italy

xiv

Contributors

HUW MORGAN  European Cancer Stem Cell Research Institute, Cardiff University, Cardiff, UK TSUTOMU MOTOHASHI  Department of Tissue and Organ Development, Regeneration and Advanced Medical Science, Gifu University Graduate School of Medicine, Gifu, Japan HAZAL BANU OLGUN  Human Gene and Cell Therapy Center of Akdeniz University Hospitals and Clinics, Antalya, Turkey CARLOTTA OLIVERO  European Cancer Stem Cell Research Institute, Cardiff University, Cardiff, UK ¨ NEN  Hacettepe University, Institute of Health Sciences, Department of Stem Cell SELIN O Sciences, Ankara, Turkey DAVID M. OWENS  Department of Dermatology, College of Physicians & Surgeons, Columbia University Medical Center, New York, NY, USA; Department of Pathology and Cell Biology, College of Physicians & Surgeons, Columbia University Medical Center, New York, NY, USA; Russ Berrie Medical Science Pavilion, New York, NY, USA MAHMUT PARMAKSIZ  Tissue Engineering, Biomaterials and Nanobiotechnology Laboratory, Ankara University, Faculty of Science and Stem Cell Institute, Ankara, Turkey GIRISH K. PATEL  European Cancer Stem Cell Research Institute, Cardiff University, Cardiff, UK MIRNA PEREZ-MORENO  Section for Cell Biology and Physiology, Department of Biology, Faculty of Science, University of Copenhagen, Copenhagen Ø, Denmark NEHA PINCHA  IFOM-inStem Joint Research Laboratory, Centre for Inflammation and Tissue Homeostasis, Institute for Stem Cell Biology and Regenerative Medicine, Bangalore, Karnataka, India; Manipal Academy of Higher Education, Manipal, Karnataka, India HUA QIAN  Laboratory for Tissue Engineering and Regeneration, and Shandong Provincial Key Laboratory of Oral Tissue Regeneration, School of Stomatology, Shandong University, Jinan, Shandong, China SHAOHAI QI  Department of Burns, First Affiliated Hospital of Sun Yat-sen University, Guangzhou, China WEIMING QIU  Department of Dermatology, Wuhan General Hospital of Chinese People’s Liberation Army, Wuhan, China DANIEL RADKE  Department of Biomedical Engineering, Michigan Technological University, Houghton, MI, USA SRIKALA RAGHAVAN  Institute for Stem Cell Biology and Regenerative Medicine, Bangalore, India RUI L. REIS  3B’s Research Group—Biomaterials, Biodegradable and Biomimetics, Avepark—Parque de Cieˆncia e Tecnologia, Zona Industrial da Gandra, Guimara˜es, Portugal; ICVS/3B’s—, PT Government Associate Laboratory, Braga, Portugal; The Discoveries Centre for Regenerative and Precision Medicine, Headquarters at University of Minho, Guimara˜es, Portugal PANTELEIMON ROMPOLAS  Department of Dermatology, Institute for Regenerative Medicine, University of Pennsylvania Perelman School of Medicine, Philadelphia, PA, USA EDWIGE ROY  The University of Queensland, UQ Diamantina Institute, Translational Research Institute, Brisbane, QLD, Australia AHTER DILSAD SANLIOGLU  Human Gene and Cell Therapy Center of Akdeniz University Hospitals and Clinics, Antalya, Turkey SALIH SANLIOGLU  Human Gene and Cell Therapy Center of Akdeniz University Hospitals and Clinics, Antalya, Turkey

Contributors

xv

KAREL SMETANA JR  First Faculty of Medicine, Institute of Anatomy and BIOCEV, Charles University, Prague, Czech Republic STEFANO SOL  Department of Biology, University of Naples Federico II, Napoli, Italy; CEINGE Biotecnologie Avanzate (Center for Genetic Engineering), Napoli, Italy GOPU SRIRAM  Faculty of Dentistry, National University of Singapore, Singapore, Singapore EHSAN TAGHIABADI  Department of Regenerative Biomedicine, Cell Science Research Center, Royan Institute for Stem Cell Biology and Technology, ACECR, Tehran, Iran; Skin and Stem Cell Research Center, Tehran University of Medical Sciences, Tehran, Iran HALE M. TASYUREK  Human Gene and Cell Therapy Center of Akdeniz University Hospitals and Clinics, Antalya, Turkey GABRIEL M. VIRADOR  Biology Department, Montgomery College, Rockville, MD, USA; University of Navarra, Pamplona, Navarra, Spain VICTORIA M. VIRADOR  Biology Department, Montgomery College, Rockville, MD, USA; Virador and Associates, Bethesda, MD, USA XIAOXIAO WANG  Graduate School at Shenzhen, Tsinghua University, Shenzhen, China XUSHENG WANG  Graduate School at Shenzhen, Tsinghua University, Shenzhen, China JIE WEN  Laboratory for Tissue Engineering and Regeneration, and Shandong Provincial Key Laboratory of Oral Tissue Regeneration, School of Stomatology, Shandong University, Jinan, Shandong, China WENBO WU  Peking University-Tsinghua University-National Institute of Biological Sciences Joint Graduate Program, School of Life Sciences, Peking University, Beijing, China; National Institute of Biological Sciences, Beijing, China XUNWEI WU  Laboratory for Tissue Engineering and Regeneration, and Shandong Provincial Key Laboratory of Oral Tissue Regeneration, School of Stomatology, Shandong University, Jinan, Shandong, China YAOJIONG WU  Graduate School at Shenzhen, Tsinghua University, Shenzhen, China MEILANG XUE  Sutton Arthritis Research Laboratory, Kolling Institute of Medical Research, The University of Sydney at Royal North Shore Hospital, St Leonards, NSW, Australia XIN XU  Laboratory for Tissue Engineering and Regeneration, and Shandong Provincial Key Laboratory of Oral Tissue Regeneration, School of Stomatology, Shandong University, Jinan, Shandong, China KE YANG  Department of Pediatric Research Institute, Chongqing Engineering Research Center of Stem Cell Therapy, Children’s Hospital of Chongqing Medical University, and Key Laboratory of Child Development and Disorders of Ministry of Education, Chongqing, China NILGU¨N YERSAL  Hacettepe University, Faculty of Medicine, Department of Histology and Embryology, Ankara, Turkey FENG ZHAO  Department of Biomedical Engineering, Michigan Technological University, Houghton, MI, USA QIAN ZHOU  Laboratory for Tissue Engineering and Regeneration, and Shandong Provincial Key Laboratory of Oral Tissue Regeneration, School of Stomatology, Shandong University, Jinan, Shandong, China

Methods in Molecular Biology (2019) 1879: 1–14 DOI 10.1007/7651_2018_172 © Springer Science+Business Media New York 2018 Published online: 22 September 2018

In Vivo Genetic Alteration and Lineage Tracing of Single Stem Cells by Live Imaging Olivia Farrelly, Paola Kuri, and Panteleimon Rompolas Abstract Studies characterizing stem cell lineages in different organs aim to understand which cells particular progenitors can give rise to and how this process is controlled. Because the skin contains several resident stem cell populations and undergoes constant turnover, it is an ideal tissue in which to study this phenomenon. Furthermore, with the advent of two-photon microscopy techniques in combination with genetic tools for cell labeling, this question can be studied non-invasively by using live imaging. In this chapter, we describe an experimental approach that takes this technique one step further. We combine the Cre and Tet inducible genetic systems for single clone labeling and genetic manipulation in a specific stem cell population in the skin by using known drivers. Our system involves the use of gain- and loss-of-function alleles activated only in a differentially labeled population to distinguish single clones. The same region within a tissue is imaged repeatedly to document the fate and interactions of single clones with and without genetic modifications in the long term. Implementing this lineage tracing approach while documenting changes in cell behavior brought about by the genetic alterations allows both aspects to be linked. Because of the inherent flexibility of the approach, we expect it to have broad applications in studying stem cell function not only in the skin, but also in other tissues amenable to live imaging. Keywords Clonal dynamics, Inducible genetic mouse tools, Lineage tracing, Live imaging, Skin, Stem cells, Two-photon laser scanning fluorescent microscopy

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Introduction

1.1 In Vivo Lineage Tracing in the Skin

In vivo lineage tracing has been the gold standard for identifying and studying stem cells in various organs. In a classical lineage tracing experiment, cells are labeled to allow the fate of their descendants to be followed through time [1]. This experimental setup has been widely used to characterize tissue-specific resident stem cell populations and their subsequent progeny. The visualization of stem cells through live imaging is a major step forward from static histological analysis and has significantly contributed to our understanding of stem cell behavior in their native in vivo environment. Studies carried out at a population-level provide information on the general aspects of stem cell behavior. However, by averaging out individual behaviors, they bypass the opportunity to shed light on the inherent heterogeneity of the

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stem cell population [2]. Therefore, gaining a deeper understanding of the underlying mechanisms that regulate individual stem cell activity requires tools to document behavior at the individual level. Observing how the behavior of individual stem cells is altered when a particular gene is turned on or off compared to that of their neighbors allows a causative relationship between specific genes and cellular activity to be established. Therefore, the capacity to both trace and genetically manipulate single cells over time ultimately contributes to dissecting the underlying molecular heterogeneity that characterizes some stem cell populations. Furthermore, cellular interactions between cells with different genetic compositions can be studied by differentially labeling single genetically altered and “normal” cells within the same tissue. Experiments of this nature would also be useful to uncover the genetic drivers of aberrant stem cell activity in the context of wound healing and disease. The skin is a well-studied and readily accessible organ that continuously self-regenerates, thanks to the contribution of several resident stem cell populations located within self-contained compartments. These include the interfollicular epidermis (IFE), hair follicles (HF), and sebaceous and sweat glands [3]. As in other adult epithelia, stem and progenitor cells in the skin constantly divide to replace the cells that are naturally lost through terminal differentiation and shedding. This process is crucial for the maintenance of organ homeostasis [4]. Due to the fast turnover of cells in the skin, conventional histological techniques are insufficient to provide a comprehensive understanding of complex mechanisms regulating stem cell activity [5]. Genomic tools provide a very detailed snapshot on the molecular heterogeneity in a cell population, but lack the temporal and spatial information required to put that information into the context of the three-dimensional tissue [6], a technical challenge that the field is currently tackling [7]. The combination of live imaging and powerful mouse genetic tools provides a unique opportunity for real-time single cell tracking in the adult skin. The approach has already provided unparalleled insight into the proliferation and differentiation dynamics of stem cells and their progeny [8, 9]. These genetic systems rely on the use of different promoters to drive the expression of a Cre recombinase in a specific stem cell population to activate expression of a fluorescent reporter. Labeled cells can be visualized in vivo in a noninvasive manner by using two-photon laser scanning fluorescent microscopy, which minimizes phototoxicity and photobleaching while increasing tissue imaging depth [10, 11]. Additionally, re-visiting the same area in multiple imaging sessions enables the direct tracking of progenitor and descendant cell fate over time. Studies that address how genetic changes impact stem cells on a population level lack the spatiotemporal resolution to distinguish variable cellular responses of singe cells. Because stem cell

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populations are heterogeneous, gaining a molecular understanding of the underlying mechanisms that regulate stem cell activity at the single cell level requires tools to genetically manipulate and monitor individual cells. In this chapter, we describe how live imaging can be combined with the use of two inducible systems, the Cre and Tet systems, to lineage trace cells whose gene expression is specifically altered. The method involves using the spatiotemporally regulated expression of Cre to generate single labeled clones within specific stem cell populations, while Tet drives the expression of gain- or loss-of-function alleles to genetically manipulate specific genes and signaling pathways in the labeled clones. In the example below, we propose a system in which expression of a single gene is modulated in fluorescently labeled skin stem cells whose long-term behavior can be recorded non-invasively at high resolution by two-photon microscopy. The activity of these cells is then compared to that of “normal” neighboring cells, which are also clonally marked but with a different fluorescent reporter. We use this design as proof of principle for a genetic system that can be expanded to modulate a wide assortment of signaling pathways and characterize their importance in stem cell dynamics in vivo. 1.2 Combining Cre-Mediated Recombination with TetracyclineControlled Transcriptional Activation

The P1 bacteriophage Cre-recombinase system efficiently mediates the recombination of palindromic LoxP recognition sites, resulting in the excision of the flanked DNA segment [12]. Cre activity is spatiotemporally controlled by using a cell population-specific promoter to control the expression of an inducible cre allele (CreER) in which the protein is fused to an estrogen receptor (ER) that sequesters it in the cytoplasm [13]. The chimeric CreER will translocate to the nucleus only in the presence of an ER ligand, most commonly, the estrogen analog tamoxifen (see Note 1). Importantly, the number of cells that undergo Cre-mediated recombination increases with Tamoxifen treatment in a dose-dependent manner. Therefore, as a general rule, it is necessary to titrate the drug dosage to confine Cre-mediated recombination to only few, spatially dispersed clones that are easily identifiable. The exact dose of tamoxifen must be determined to account for variations in promoter activity, as well as tissue variations that may affect Cre-recombinase expression and activation to achieve clonal labeling. It is also necessary for mice to carry a Cre reporter allele that tracks recombinase activity for normal cells to be identified and tracked at the individual level. Tetracycline(Tet)-inducible systems, derived from the E. coli tetracycline resistance operon, activate or repress genes under the control of the Tet Operon (tetO) sequence [14, 15]. Depending on whether the TetOFF or TetON system is used, two different transactivator fusion alleles can be used to drive expression of genes downstream of TetO sequences, the tTA (tetracycline transactivator protein) and rtTA (reverse tTA), respectively. Each has an opposite requirement of tetracycline or doxycycline, tetracycline’s

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commonly used analog. In the TetOFF system, doxycycline prevents tTA from binding to TetO sequences. Conversely, in the TetON system, doxycycline induces the binding of rtTA to TetO sequences and downstream gene expression is subsequently activated. Therefore, unlike the Cre-mediated recombination effects, which are irreversible, the activity of Tet-inducible systems can be modulated throughout the experiment by the administration or removal of doxycycline. To label and monitor the behavior of normal and genetically altered cells within the same tissue we combine the two inducible systems to lineage trace the behavior of normal and genetically altered cells within the same tissue. In the method outlined in this chapter, we use the LoxP-STOP-LoxP (LSL) allele, in which a STOP cassette is placed between a ubiquitous promoter and the Tet transactivator of choice. This allele links the two inducible systems because Cre recombination is required for the excision of the STOP sequence to allow the expression of the tTA. Therefore, the activation of TetO-driven alleles is restricted to the cells in which Cre-recombination is activated. This system can be combined with any available Cre transgenic line and TetO alleles, enabling a wide range of manipulation possibilities. Similar genetic approaches have previously been used for the activation in non-sensory regions of the ear [16], and in the skin, to induce YAP expression [17] and drive oncogene expression [18]. The novelty of our approach relies on the simultaneous differential fluorescent labeling of single cells and its combined use with two-photon microscopy to perform lineage tracing studies in live mice.

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Materials

2.1 Choice of Mouse Lines

Several inducible CreER reporter lines that target specific stem cell populations in the skin and other tissues are available from public vendors such as the Jackson Laboratory. The driver lines listed in Table 1 are three examples of CreER reporter lines that allow clonal labeling in unique stem cell populations in the epidermis (K14-CreER), isthmus/sebaceous gland (Lgr6-CreER), and hair follicle (Lgr5-CreER). Although there are several CreER reporter lines that can be used for lineage tracing, the presence of the mTom-LSL-mGFP allele ensures that cell membranes will be ubiquitously labeled by tdTomato (mT) expression. Because membrane labels can be used to visualize cellular morphology and tissue architecture, expression of tdTomato is necessary to determine spatial location of the clones within the epidermis. In cells where Cre recombinase is activated EGFP (mG) will replace tdTomato at the membrane to generate single “normal” clones (see Fig. 1a). Cre-recombinase will also turn on expression of the tTA in single cells. The Tet transactivator drives expression of both a reporter

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Table 1 Useful publicly available transgenic mouse lines Allele

Source

Reference

LSL-tTA

JAX Stock no. 008600

[19]

mTom-LSL-mGFP

JAX Stock no. 007676

[20]

TetO-H2BGFP

JAX Stock no. 005104

[21]

K14-CreERa

JAX Stock no. 005107

[22]

Lgr5-CreERa

JAX Stock no. 008875

[23]

Lgr6-CreERa

JAX Stock no. 016934

[24]

TetO-GeneofInterest

a

Available Cre-reporters that can be used to target different stem cell populations in the skin

TetO allele and the gain- or loss-of-function TetO allele of choice, thus labeling single altered clones. A model experimental timeline is shown in Fig. 1b (see Note 2). 2.2 Systemic Induction by Intraperitoneal Injection of Tamoxifen

1. Tamoxifen (Sigma). 2. Corn oil. 3. Scale. 4. 27 G Insulin syringe (BD). Preparation of Tamoxifen 1. Dissolve tamoxifen in corn oil to a final concentration of 20 mg/ml by incubating in a 37
C water bath. Vortex the solution frequently over the course of several hours. Tamoxifen is classified as carcinogenic and toxic, so always wear gloves when handling it. After tamoxifen is in solution, wrap with foil and store in the dark at 20
C.

2.3 Temporal Regulation of Transgene Expression by Doxycycline

1. Doxycycline (Sigma). 2. Drinking water. 3. Amber water bottle. Preparation of Doxycycline 1. Dissolve doxycycline in drinking water to a final concentration of 1 mg/ml by vortexing the solution for several minutes. This solution can be wrapped in foil and frozen in 5 ml aliquots at 20
C.

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A 1) Stem cell-specific promoter

CreER

CreER + Tamoxifen

3) Rosa26

Stop

loxP

2) Rosa26

tTA

Rosa26

mTomato

loxP

loxP

Rosa26

tTA loxP

mGFP loxP

Normal cell

mGFP loxP Normal clone

tTA

4) TetO

Gof/Lof

5) TetO

H2BGFP

+ Doxycycline Gof/Lof clone

B Tamoxifen

Day 0

Imaging re-visits

Fig. 1 Genetic system and experimental timeline. To generate single genetically altered and normal clones, five alleles are used in our system. A stem cell-specific driver to control expression of an inducible Cre recombinase (1). Addition of tamoxifen will enable the Cre-mediated excision of loxP-flanked sequences in two alleles: a reporter allele (2), in this case mT/mG, in which membrane-localized EGFP (mG) replaces the excised membrane-localized tdTomato (mT) in single clones, and the Tet transactivator allele (3), in this case LSL-tTA, where excision of a STOP cassette allows tTA expression. Expression of tTA in single clones will then enable activation of the TetO alleles: A gain- or loss-of-function (Gof/Lof) allele to alter a gene/pathway of interest (4) and a second reporter allele (5), in this case TetO-H2BGFP used to label the single “Lof/Gof clones” with nuclear EGFP. In this example, where the TetOFF system is used, doxycycline treatment will prevent binding of tTA to the TetO sequences, thus silencing TetO-driven gene expression (a). Before tamoxifen treatment, all cells are uniformly labeled with membrane tdTomato (“normal cells”). After induction of Cre-mediated recombination, on the first imaging session taking place at “Day 0”, single clones labeled either with membrane EGFP instead of red (“normal clones”) or carrying a nuclear EGFP (“Gof/Lof clones”) are identified. Subsequent imaging sessions allow live tracking of clonal dynamics through time. The frequency of imaging re-visits should be empirically determined according to experimental needs (b) 2.4 Two-Photon Imaging

Mouse Preparation 1. Fluorescent reporter mice containing all alleles (see Table 1).

2. PBS, pH 7.2. 3. Depilatory lotion (Nair).

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4. Tattoo ink (Optional; Dr. Ph. Martin’s Black Star India Ink). 5. LubriFresh P.M. sterile eye ointment (MAJOR). 6. Ketamine (Midwest Veterinary Supply). 7. Xylazine (Midwest Veterinary Supply). 8. 10 ml vial (GREER). 9. 30 G Insulin syringe for tattooing (Optional; BD). 10. 27 G Insulin syringe for IP injections (BD). 11. Scale. 12. Small, electric hair clipper (Wahl). 13. Cotton swabs. 14. Adhesive tape. 15. Wall-mounted anesthesia machine (VetEquip). 16. Isoflurane vaporizer (VetEquip). 17. Anesthesia breathing circuit and nose cone (Braintree Scientific). 18. O2 gas flow regulator for E-cylinders (VetEquip). 19. O2 tank (E-cylinder; Airgas). Imaging Equipment 1. Olympus FV1200MPE (Olympus) or equivalent multiphoton microscope with upright objective lens and motorized stage.

2. Chameleon Vision II (Coherent) or equivalent Ti:sapphire tunable laser. 3. 25 objective (Olympus, XLPLN25XWMP2) or equivalent infrared optimized objective of desired magnification. 4. Custom mounting stage with custom-made mounting spatulas [10] and micromanipulators (Edmund Optics, cat. nos. NT03682  2 and NT36-347  2). 5. Heating pad (CryoLogic, BioTerm SmartStage no. SS12). 6. Glass coverslip (22 mm; Electron Microscopy Sciences). 7. Super glue (Loctite). 8. Bull’s-eye level (McMaster-Carr). 9. Thumb screw (ProTanium). 10. Vacuum grease (Dow Corning). 11. Purified water (Milli-Q, Millipore). Preparation of Ketamine/Xylazine Solution 1. Combine 15 mg/ml ketamine (15 mg/ml) and xylazine (1 mg/ml) in PBS. Store solution in a sterile 10 ml vial at room temperature for up to 30 days.

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Image Analysis

1. Fiji/ImageJ.

Methods

3.1 Systemic Induction by Intraperitoneal Injection of Tamoxifen

Cre activation is induced with a single intraperitoneal (IP) injection of tamoxifen (1 μg/g in corn oil) at ~postnatal day 50 (P50) to clonally label single epidermal cells. 1. Weigh the mouse to determine injection dose. For full induction, use approximately 75 mg tamoxifen/kg body weight. Be sure to titrate the dose of tamoxifen beforehand to determine the optimal dose for clonal induction using a particular Cre reporter. 2. Administer tamoxifen via IP injection using an institution approved animal injection protocol. 3. Tamoxifen remains in the mouse’s system for ~3 days, and recombination events can occur during this time. Therefore, the first imaging time-point should be at least 3 days after tamoxifen injection.

3.2 Temporal Regulation of Transgene Expression by Doxycycline

After the addition of tamoxifen, cells that express tTA following recombination will also start to express H2BGFP (and any other transgenes under the control of the TetO element). To reverse the expression of all TetO alleles, the binding of tTA can be inhibited by treatment with doxycycline. This would lead to a reversal of both the genetic modifications and reporter expression, returning “Gof/ Lof” clones to a “normal” state (see Note 3) Therefore, when TetO inhibition is desired, administer doxycycline (1 mg/ml in drinking water) at specified times. 1. Add the drinking water with dissolved doxycycline to an amber water bottle and place in the mouse cage at selected timepoints.

3.3 Two-Photon Imaging

3.3.1 Preparing Mouse for Imaging

On the fourth day after tamoxifen induction, the first time-point in the time-course clonal analysis is collected by two-photon microscopy. 1. Weigh the mouse to determine injection dose of the ketamine (100 mg/kg)/xylazine (10 mg/kg) solution. Administer tamoxifen via IP injection with a 27 G insulin syringe using an institution approved animal injection protocol. 2. After the mouse is fully sedated, place the mouse on heating pad (see Note 4) on the stage. Apply eye ointment to keep eyes lubricated throughout imaging session and prevent damage. Next, use the electric clippers, appropriate for veterinary use in

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small animals, to shave the hair covering the area of skin to be imaged. 3. Apply depilatory cream to the shaved area using a cotton swab. Let the cream sit for 2 min, then use cotton swabs soaked in PBS to gently remove the cream. 4. Optional: Mark the area of the skin that is being imaged for later re-visiting by creating a small punctate tattoo. Dip a 30-G needle into an ink solution and quickly poke the skin of the anesthetized mouse. The pigment should be visible macroscopically. 3.3.2 Mounting Mouse on Imaging Stage

1. Screw the skin-mounting spatula onto the micromanipulator and make sure it is level using a bull’s-eye level. 2. Place the mouse’s skin onto the spatula and ensure that the skin is completely flat. 3. Screw the coverslip spatula onto the second micromanipulator and make sure it is level using a bull’s-eye level. Lower the coverslip spatula using the micromanipulator towards the mounted skin. 4. Place a drop of super glue onto the edge of a round coverslip and gently press it onto the tip of the coverslip spatula to secure. After the super glue has dried, use the micromanipulator to lower the coverslip until it lightly presses down on the area of the skin to be imaged (see Fig. 2b).

3.3.3 Preparing Stage for Imaging

1. Move the stage from the preparation area to the imaging platform. Immediately plug in the outlet of the heating pad to the power supply. 2. Position the nose cone around the snout of the mouse and secure the connected tubing to the stage using a piece of adhesive tape (see Fig. 2a). 3. Adjust the oxygen tank and vaporizer outlet to deliver 1 L/min O2 and 1% isoflurane to the mouse throughout the imaging session. 4. If using a 25 water-immersion lens, deposit a drop of purified water to create a meniscus on the top of the coverslip. 5. Use the appropriate filters to view the fluorophores used on the mouse. Adjust the X-, Y-plane and Z-axis controllers to focus on the area of interest.

3.3.4 Two-Photon Image Acquisition

1. Turn off all the lights in the room (see Note 5). 2. Use an Olympus FV1200MPE multiphoton microscope equipped with a Chameleon Vision II laser (or equivalent microscope and laser). Set up to image with a 25 objective or an equivalent objective.

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Fig. 2 Mouse mounting. An anesthetized mouse on stage of confocal microscope. Anesthetic is delivered through nose cone (a). For ear mounting two spatulas are used, one to support the tissue form below (skin mounting spatula) and a second one to which the coverslip is glued placed on top (coverslip spatula). The ear is held firmly in place between the both spatulas (b). High magnification image of the ear following clonal induction. Green channel is shown (c). High magnification image of ear epidermis after clonal induction. Membrane EGFP (“normal clones”) and nuclear EGFP (“Gof/Lof clones”) are easily identifiable among red membrane-labeled cells (d)

3. Tune the laser to a wavelength between 900 and 1000 nm. This is suitable to visualize both GFP and Tomato. However, 940 nm is optimal for green-yellow fluorophores (e.g., GFP and YFP) and 1040 nm is optimal for orange-red fluorophores (e.g., RFP, Tomato and mCherry). 4. Set the upper and lower limits of the Z stack. Normally, the epidermis is set as the start-point and the dermis just below the tip of the hair follicles are set as the endpoint. Set the step size between 1 and 3 μm. 5. Adjust the power increment to clearly visualize the cells of interest. 6. Acquire image stacks. 3.3.5 Performing Re-Visits

1. Prepare the mouse as detailed above to re-visit the same area of the skin in the same mouse. 2. Use the tattoo and clusters of hair follicles as landmarks to find the same area. 3. Repeat steps from Sect. 3.3 to set up and image the mouse.

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Fig. 3 Image analysis. Example data set of a K14-CreER::mTom-LSL-mGFP::LSL-tTA::TetO-H2BGFP mouse acquired daily over 4 days. Overlay of green and red channels of entire imaged region of the skin at Day 0, basal layer is shown (a). Single labeled clones (membrane localized EGFP or nuclear EGFP) are distinguishable within the uniformly fluorescently labeled tissue (membrane tdTomato), with the hair follicle landmarks visible using the latter. Therefore, cells will be either “normal” (red membrane), “normal clones” (green membrane) or genetically modified (green nuclei with red membrane) (b). A region of interest is chosen for in-depth analysis of cell fate (a0 ). To determine the fate of single labeled clones, basal progenitor cells are looked at individually, along the entire Z stack on Day 0 (a00 ). For the chosen example, a single basal progenitor gives rise to four cells throughout the image period. Two of those can be seen in the suprabasal layer, indicating commitment to differentiation. The upward movement of cells can be more easily visualized by using orthogonal views of the acquired Z stack (c). Overall, in these panels, three division and two differentiation events are documented. A possible lineage tree describing the division events (d). An accurate lineage tree cannot be determined with this experimental setup alone, but a label-dilution assay compatible with our system would enable this analysis (see Note 3). Scale bar 25 μm unless otherwise indicated

3.4

Image Analysis

After image acquisition, cell fate of labeled clones is manually determined. Raw data from the imaging re-visits is observed to document cell division and differentiation events. The first by quantifying the number of cells that a single clone identified at Day 0 gave rise to and the second by determining how many of those clones underwent terminal differentiation. Because the normal and Gof/Lof clones are uniquely labeled, the consequences of the genetic alteration(s) can be inferred from differences in the behavior of clones belonging to each population (see Fig. 3). 1. After image acquisition, use Fiji/ImageJ or a comparable image analysis software for clonal analysis. After importing the raw image files into the software, choose regions of interest

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within the first acquired time-point and select clones for analysis. 2. For each re-visit, document the fate of identified clones by quantifying each cell’s division events. The identity of the daughter cells can be determined by their location in the Zaxis because stem and progenitor cells will remain in the basal layer, while differentiating cells will migrate upwards. 3. Document these changes manually by comparing changes in cell number and location between time-points. A high resolution in the Z-axis and a tissue imaged as flat as possible will facilitate analysis.

4

Notes 1. It is well documented that unwanted Cre activity can occur even without treatment with tamoxifen, known as “leakiness” [25]. This disadvantage must be considered when analyzing the data. If, at Day 0, clusters of clones instead of single ones are observed, it is likely that the recombination event precedes tamoxifen induction. These cells should be disregarded from further analysis. 2. Mice used for live imaging should be bred into an albino background because pigment absorbs intensely in the infrared spectrum, which may be detrimental to the health of the tissue following prolonged exposure. Additionally, pigments produce strong auto-fluorescence that interferes with signal detection. 3. Administration of doxycycline will inhibit expression of all TetO alleles, including the fluorescent nuclear label. Therefore, after doxycycline administration, every round of cell division will dilute the nuclear label by half, which can be taken advantage of to carry out a label-dilution experiment [2, 9]. In this assay, the origin of the clones is determined based on the change in fluorescence level after every division. Quiescent cells retain their original level of fluorescence, whereas the levels of proliferative cells will fade over time. 4. Because mice are unable to regulate body temperature when under anesthesia, special care must be taken to prevent hypothermia during the experiment. Ensure that the anesthetized mice are place on a heating pad at all times. 5. A lightproof curtain can be placed around the microscope to minimize light leakage.

In Vivo Genetic Alteration and Lineage Tracing of Single Stem Cells by Live. . .

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in mammalian cells using a ligand-dependent chimeric Cre recombinase. Proc Natl Acad Sci U S A 92:6991–6995 14. Gossen M, Bujard H (1992) Tight control of gene expression in mammalian cells by tetracycline-responsive promoters. Proc Natl Acad Sci U S A 89:5547–5551 15. Gossen M, Freundlieb S, Bender G, Mu¨ller G, Hillen W, Bujard H (1995) Transcriptional activation by tetracyclines in mammalian cells. Science 268:1766–1769 16. Pan W, Jin Y, Stanger B, Kiernan AE (2010) Notch signaling is required for the generation of hair cells and supporting cells in the mammalian inner ear. Proc Natl Acad Sci 107:15798–15803. https://doi.org/10. 1073/pnas.1003089107 17. Schlegelmilch K, Mohseni M, Kirak O, Pruszak J, Rodriguez JR, Zhou D, Kreger BT, Vasioukhin V, Avruch J, Brummelkamp TR, Camargo FD (2011) Yap1 acts downstream of α-catenin to control epidermal proliferation. Cell 144:782–795. https://doi.org/10. 1016/j.cell.2011.02.031 18. Grachtchouk M, Pero J, Yang SH, Ermilov AN, Michael LE, Wang A, Wilbert D, Patel RM, Ferris J, Diener J, Allen M, Lim S, Syu L-J, Verhaegen M, Dlugosz AA (2011) Basal cell carcinomas in mice arise from hair follicle stem cells and multiple epithelial progenitor populations. J Clin Invest 121:1768–1781. https://doi.org/10.1172/JCI46307 19. Wang L, Sharma K, Deng H-X, Siddique T, Grisotti G, Liu E, Roos RP (2008) Restricted expression of mutant SOD1 in spinal motor neurons and interneurons induces motor neuron pathology. Neurobiol Dis 29:400–408. https://doi.org/10.1016/j.nbd.2007.10.004 20. Muzumdar MD, Tasic B, Miyamichi K, Li L, Luo L (2007) A global double-fluorescent Cre reporter mouse. Genesis 45:593–605. https:// doi.org/10.1002/dvg.20335 21. Tumbar T (2004) Defining the epithelial stem cell niche in skin. Science 303:359–363. https://doi.org/10.1126/science.1092436 22. Vasioukhin V, Degenstein L, Wise B, Fuchs E (1999) The magical touch: genome targeting in epidermal stem cells induced by tamoxifen application to mouse skin. Proc Natl Acad Sci U S A 96:8551–8556 23. Barker N, van Es JH, Kuipers J, Kujala P, van den Born M, Cozijnsen M, Haegebarth A, Korving J, Begthel H, Peters PJ, Clevers H (2007) Identification of stem cells in small

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Methods in Molecular Biology (2019) 1879: 15–29 DOI 10.1007/7651_2018_136 © Springer Science+Business Media New York 2018 Published online: 25 February 2018

Ex Vivo Imaging and Genetic Manipulation of Mouse Hair Follicle Bulge Stem Cells Daniel Haensel, Melissa A. McNeil, and Xing Dai Abstract Stem cells that reside in the bulge of adult mouse hair follicles are a leading model of tissue stem cell research. Ex vivo culturing, molecular and cell biological characterizations, as well as genetic manipulation of fluorescence-activated cell sorting-isolated bulge stem cells offer a useful experimental pipeline to complement in vivo studies. Here we describe detailed methods for culturing, immunostaining, live cell imaging, and adenoviral infection of bulge stem cells for downstream applications such as in vitro clonal and in vivo patch assays. Keywords Adenovirus, Bulge stem cells, Clonal assay, Hair follicle, Immunofluorescence, Live cell imaging, Patch assay

1

Introduction The hair follicle is a skin appendage that undergoes dramatic remodeling during the postnatal hair cycle, which includes a growth phase (anagen), a destruction phase (catagen), and a resting phase (telogen) [1]. Fueling the hair cycle are the hair follicle stem cells (HFSCs) that reside in the bulge, a part of the outer root sheath that bulges out from the base of the non-cycling portion of the hair follicle, and their immediate progeny in the secondary hair germ (HG) [2, 3]. Transition from telogen into anagen occurs when the HG cells receive signals from the adjacent dermal papilla and begin to proliferate and differentiate into multiple cell types of the regenerative portion of the hair follicle, including the inner root sheath and the hair matrix that later produces the actual hair shaft [2]. The bulge HFSCs are generally quiescent but become proliferative during early anagen and differentiate into the outer root sheath that encases the inner parts of the growing hair follicle [2]. In catagen, most of the hair follicle cells that are located beneath the bulge undergo apoptosis, leaving a structure called strand which retracts up to the base of the bulge [1]. Some of the outer root sheath cells escape apoptosis and are incorporated into the base of the non-cycling portion of the hair follicle where they form a new

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bulge and a new HG, which are HFSCs for the next hair cycle [2]. Understanding the function of HFSCs and the molecular regulation of their quiescence, activation, and differentiation represents an important direction in adult stem cell biology and may have implications on how we prevent and treat hair loss. Various tools and techniques have been generated and/or applied to study HFSCs. With the knowledge of unique markers of the various hair follicle cell types, immunofluorescence can be used to examine the presence, absence, or fate alterations of cells in the context of various genetic perturbations. HFSCs can be identified by immunofluorescence using classical HFSC markers, such as CD34, or with pulse chase methods to identify HFSCs as labelretaining cells [2]. These analyses provide static snapshots of HFSCs, as skin must be frozen or fixed before sectioning and further analysis. More sophisticated imaging techniques now have abilities to track the migration, proliferation, and overall behavior of individual cells within the hair follicle in real time during various stages of regeneration [4, 5]. Coupled with fluorescent genetic labeling, these pioneering live imaging techniques uncover unprecedented information about stem cell dynamics in vivo. With flow cytometry and fluorescence-activated cell sorting (FACS), HFSCs can be quantified and isolated for downstream purposes such as gene expression analysis and in vitro assays [6, 7]. FACS-sorted HFSCs can be cultured and expanded on mitotically inactivated fibroblast feeders [6]. Cultured HFSCs can be functionally assessed for clonal growth potential and, with serial passaging, long-term self-renewal capability, as well as for regenerative capacity in host animals [6, 8, 9]. Here we describe methods for culturing and downstream analyses of sorted bulge HFSCs. Specifically, we expand on the existing clonal growth assay by incorporating live cell imaging to monitor the division events and movement tracks of individual cells within the growing bulge HFSC colonies. Furthermore, we provide protocols for growing bulge HFSCs on glass for immunofluorescence and for efficient adenovirus infection that minimally impacts their regenerative capacity in vivo.

2

Materials

2.1 J2-3T3 Fibroblast Culture, Mitotic Inactivation, and Feeder Layer Preparation

1. J2-3T3 fibroblasts. 2. F media (see [6] for detailed instructions). 3. Mitomycin C (Fisher Scientific, Cat. No. BP2531-2). 4. 0.22-μm filter (Millex, Cat. No. SLGV033RS). 5. 10-mL syringe (BD, Cat. No. 309604). 6. 1 PBS. 7. 15-mL conical vials.

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2.2 Plating and Culture of Primary HFSCs

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1. Tissue culture dishes with mitotically inactivated J2-3T3 fibroblast layer (requires reagents from Sect. 2.1). 2. E media (see [6] for detailed instructions). 3. 1 PBS. 4. Versene: (a) 200 mL 10 PBS. (b) 0.4 g EDTA disodium salt (Sigma, Cat. No. E-6511). (c) Bring up to 2 L with deionized H2O. (d) Autoclave. (e) 8 mL sterile 25% glucose solution in deionized H2O. 5. 0.1% trypsin (Sigma, Cat. No. T4799). 6. 15-mL conical vials.

2.3

Clonal Assay

1. 6-well tissue culture plate (Falcon, Cat. No. 353046) or 60-mm gridded plates (Corning, Cat. No. 430166). 2. Tissue culture dishes with mitotically inactivated J2-3T3 fibroblast layer in F media. 3. E media. 4. 0.5% crystal violet (Sigma, Cat. No. HT90132-1L) in a 1:1 methanol/H2O solution.

2.4

Live Cell Imaging

1. Keyence BZ-X700 microscope (or equivalent microscope capable of live cell imaging). 2. 6-well tissue culture plate (Falcon, Cat. No. 353046). 3. Mitotically inactive 3T3 fibroblast feeders. 4. F media. 5. E media. 6. Freshly sorted or passaged HFSCs.

2.5 Immunofluorescence

1. Collagen I solution at a concentration of 25 μg/mL in 0.02 M acetic acid (Sigma, Cat. No. C9791). 2. Glass coverslips (Fisher Scientific, Cat. No. 12-546). 3. Optional: 0.22-μm low protein-binding filter and 10-mL syringe. 4. F media. 5. E media. 6. 1 PBS. 7. 4% paraformaldehyde, made from powder (MP Biomedicals, Cat. No. 150146) in 1 PBS. 8. 1 PBS with 0.1% Triton X-100 (Sigma, Cat. No. T9284).

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9. 20% normal goat serum (NGS)-gelatin solution (20 mL): (a) 4 mL NGS (Invitrogen, Cat. No. 16210-064). (b) 200 μL 10% Triton X-100 (Sigma, Cat. No. T9284). (c) 2 mL 10 PBS. (d) 200 μL 2% NaN3 (Alfa Products, Cat. No. 50101). (e) 200 μL 5% Tween 20 (Fisher Scientific, Cat. No. EC500018-3). (f) 200 μL 1% gelatin (Sigma, Cat. No. G-1890). (g) 13.2 mL distilled H2O. l

Mix and heat inactivate in 55  C water bath for 30 min. Store at 4  C for about 2 weeks.

10. 10% NGS-gelatin solution (10 mL): (a) 5 mL 20% NGS-gelatin solution (from step 9). (b) 50 μL 10% Triton X-100 (Sigma, Cat. No. T9284). (c) 500 μL 10 PBS. (d) 50 μL 2% NaN3 (Alfa Products, Cat. No. 50101). (e) 50 μL 5% Tween 20 (Fisher Scientific, Cat. No. EC500018-3). (f) 50 μL 1% gelatin (Sigma, Cat. No. G-1890). (g) 4.3 mL distilled H2O. 11. 2% NGS-gelatin solution: (a) 1 mL 20% NGS-gelatin (from step 9). (b) 200 μL 10% Triton X-100 (Sigma, Cat. No. T9284). (c) 900 μL 10 PBS. (d) 200 μL 2% NaN3 (Alfa Products, Cat. No. 50101). (e) 200 μL 5% Tween 20 (Fisher Scientific, Cat. No. EC500018-3). (f) 200 μL 1% gelatin (Sigma, Cat. No. G-1890). (g) 7.75 mL distilled H2O. 12. Primary antibodies of interest. 13. Appropriate secondary antibodies. 14. DAPI (Life, Cat. No. D1306). 15. Vectashield (Vector, Cat. No. H1000). 2.6 Adenoviral Infection of HFSCs

1. 6-well tissue culture dishes (Falcon, Cat. No. 353046) with mitotically inactivated J2-3T3 fibroblast layer. 2. 1 PBS. 3. E media. 4. Versene.

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5. 0.1% trypsin (Sigma, Cat. No. T4799). 6. 15-mL conical vial. 7. IRES-GFP adenovirus (Vector Biolabs, Cat. No. 1761). 2.7 Patch Reconstitution Assay with Virally Infected HFSCs

3

1. Detailed lists of required reagents for the (1) isolation of neonatal dermal cells and (2) patch reconstitution assay can be found in [7]. 2. Reagents from Sect. 2.6 are required for generating virally infected HFSCs.

Methods

3.1 J2-3T3 Fibroblast Culture, Mitotic Inactivation, and Feeder Layer Preparation

Culturing healthy fibroblasts and generation of a mitotically inactive feeder layer are a critical component of HFSC culture. 1. For J2-3T3 growth and propagation, see the detailed protocol in [6]. 2. Allow cells to reach 100% confluency. 3. Prepare the mitomycin C solution. (a) Add 5 mL 1 PBS to 2 mg vial of mitomycin C and mix (0.4 mg/mL). (b) Filter sterilize with 0.22-μm filter. 4. Combine 12 mL F media with 240 μL of filter sterilized 0.4 mg/mL mitomycin C, and add to a confluent 100-mm plate of J2-3T3 fibroblasts (see Note 1). 5. Swirl gently to mix and incubate at 37  C in incubator with 5% CO2 for 2 h. 6. Aspirate media and rinse cells five times with 2 mL 1 PBS (see Note 2). 7. Add 2 mL 0.1% trypsin and incubate until cells start to lift off the plate (about 10 min). 8. Gently remove cells with P1000. 9. Add 8 mL F media to inactivate trypsin and transfer cell suspension to a 15-mL conical vial and centrifuge to pellet at 1000 RPM for 5 min. 10. Aspirate the supernatant and resuspend the cell pellet in 5 mL F media and count cells. 11. Seed appropriate number of mitomycin C-treated J2-3T3 fibroblasts in F media based on plate size. Use Table 1 for guide. 12. Swirl plate gently to evenly distribute feeders (see Note 3).

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Table 1 Desired number of mitotically inactive J2-3T3 fibroblasts to plate depending on plate size Plate type

J2-3T3 Fibroblast number

100 mm

1,000,000

60 mm

~360,000

35 mm

~170,000

6 Well

~170,000

13. Culture at 37  C in incubator with 5% CO2 for 2 days to allow fibroblasts to attach fully and spread to cover the entire dish before adding HFSCs (see Note 4). 14. Fibroblast feeder plates can be cultured for up to 1 week before adding HFSCs. F media should be changed every 3 days until usage. 3.2 Plating Freshly Isolated HFSCs

To plate HFSCs, a media switch from F media to E media must first occur. 1. Epidermal single cell suspension and sorting of HFSCs can be done using the detailed protocol in [6]. 2. Prepare mitomycin C-treated J2-3T3 feeder layer by removing F media and then rinsing twice with 2 mL 1 PBS. 3. Add appropriate amount of E media to plate or well. 4. After HFSCs are obtained by FACS, count cells and add appropriate number of cells to each plate or well. 5. Swirl plate to mix. 6. Culture HFSCs at 35  C in incubator with 5% CO2 (see Note 5). 7. Replace E media every 3 days. 8. Colonies should become visible after ~7 days and then rapidly expand (see Note 6).

3.3

Passaging HFSCs

After ~2 weeks of culture and significant colony growth, HFSCs can be passaged onto a new mitomycin C-treated J2-3T3 fibroblast feeder layer. 1. Aspirate media. Add 2 mL versene and let sit for 2 min. 2. Vigorously pipette the versene with a P1000 to spray off all fibroblasts (see Note 7). 3. Aspirate versene, wash plate with 5 mL E media to remove residual fibroblasts, and then aspirate.

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4. Add 2 mL 0.1% trypsin (no EDTA) and let incubate until cells start to lift up (about 10 min). 5. Add 8 mL E media to inactivate trypsin and spray to remove residual cells. 6. Put cells in a 15-mL conical vial and centrifuge to pellet at 1000 RPM for 5 min. 7. Aspirate media, resuspend pellet in 5 mL E media, and count cells. 8. Prepare mitomycin C-treated J2-3T3 feeder layer by removing F media and then rinsing twice with 2 mL 1 PBS. 9. Add appropriate amount of E media to plate or well. 10. Add appropriate number of HFSCs from step 7 onto mitomycin C-treated J2-3T3 feeder layer. 11. Culture HFSCs at 35  C in incubator with 5% CO2 (see Note 5). 12. Replace E media every 3 days. 13. Colonies should become visible after ~4 days and then rapidly expand (see Note 8). Clonal Assay

Clonal assays involve plating a low number of cells (1000 cells/ cm2) such that each colony is generated by a single cell. There is flexibility in terms of what plate size to use. Generally use of a 6-well plate is recommended for ease of generating technical replicates. Gridded plates are useful for tracking the same colony over time and are also ideal for live cell imaging (Sect. 3.5).

3.4.1 Clonal Growth Assay

1. Generate mitomycin C-treated J2-3T3 fibroblast feeder layer as described in Sect. 3.1 using 6-well plates.

3.4

2. Prepare mitomycin C-treated J2-3T3 fibroblast feeder layer for HFSC as described in Sect. 3.2. 3. Add 1000 cells/cm2 of HFSCs (~9,500 cells for 6-well plate) to appropriate volume of E media (2–3 mL for 6-well plate) in 15-mL conical vial, gently mix, and then add to well (see Note 9). 4. Swirl plate to mix. 5. Culture HFSCs at 35  C in incubator with 5% CO2 (see Note 5). 6. Replace E media every 3 days. 7. Culture HFSCs for 2 weeks. 8. After 2 weeks of culture, aspirate media. Add 2 mL versene and let sit for 2 min. 9. Vigorously pipette the versene with a P1000 to spray off all fibroblasts (see Note 7). 10. Aspirate versene, wash plate with 2 mL 1 PBS to remove residual fibroblasts, and then aspirate.

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11. Add 2 mL 0.5% crystal violet staining solution in a 1:1 solution of water:methanol and incubate for 30 min. 12. Remove staining solution and then rinse with deionized water until water goes clear. 13. Allow plates to dry and then image and count/measure colonies. 3.4.2 Tracking Individual Colonies Using Gridded Plates

1. Prepare mitomycin C-treated J2-3T3 fibroblast feeder layer using 60-mm gridded plates. 2. Add 1000 cells/cm2 of HFSCs (~21,000 cells per 60-mm gridded plate) to appropriate volume of E media (3 mL per 60-mm gridded plate) in 15-mL conical vial, gently mix, and then add to pre-prepared feeder plate (see Note 9). 3. Swirl plate to mix. 4. Culture HFSCs at 35  C in incubator with 5% CO2 (see Note 5). 5. Replace E media every 3 days. 6. After 7 days of culture, go through grids and look for colonies to track, and note colony locations. 7. Image individual colonies every 24 h (Fig. 1). 8. End-point clonal analysis can also be done by completing steps 8–13 in Sect. 3.4.1.

Fig. 1 Growth of a single HFSC colony over time. Note rapid expansion of colony size during the indicated time frame. Arrow indicates location of a grid on plate

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3.5

Live Cell Imaging

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Live cell imaging of HFSC colonies enables analysis of the migratory and proliferative behaviors of cells within the expanding colonies. We use the Keyence BZ-X700 live imaging system, which can utilize multiple tissue culture plate types. 1. Prepare cells for imaging by completing steps 1–6 in Sect. 3.4.2. 2. Consider starting live imaging at 7 days after plating of HFSCs as colonies should be visible at this time (see Note 10). 3. Adjust microscope setup to maintain temperature and CO2 levels during live imaging. 4. Image each colony at 10 magnification every 15 min or less (see Note 11) for an 18-h duration. 5. Analysis of individual cells within colonies can be done using the “Manual Tracking” plugin in the ImageJ software from FIJI. (a) Carefully track individual cells through each image frame during the 18-h period to generate cell movement tracks with coordinates. (b) Set the original (X,Y) coordinate to the origin (0, 0), adjust to be in microns based on microscope’s field of view, and then graph. (c) Consider calculating directionality and velocity of migration. 6. If desired, individual cell divisions can be counted by manually going through each frame.

3.6 Immunofluorescence

Immunofluorescence of HFSCs presents a challenge due to the technical complications of growing HFSCs on glass to allow for optimal colony growth, morphology, and imaging. We have assessed several commonly used glass-coating procedures, such as using poly-L-lysine, collagen I or IV, laminin, or fibronectin to coat glass coverslips before culturing, and found them all to yield less than ideal growth and morphology. Ultimately, co-culture using coverslips pre-seeded with a fibroblast feeder layer is the best at preserving HFSC viability and morphology for immunofluorescent imaging. 1. Generate working collagen solution (25 μg/mL in 0.02 M acetic acid) as per manufacturer’s instructions. Filter through a low protein-binding filter to sterilize. 2. Cover surface of glass coverslips or glass chamber slides with working collagen solution for 1 h at room temperature in tissue culture hood. 3. Remove collagen and rinse three times with 1 PBS. 4. Coated coverslips/chamber slides can be store at 4  C for up to a month if not immediately used.

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5. Prepare mitomycin C-treated J2-3T3 fibroblast feeder layer as described in Sect. 3.2 using either chamber slides or glass coverslips. If using glass coverslips, place coverslips in 6-well plates, and add appropriate number of mitomycin C-treated J2-3T3 fibroblasts. 6. Add the needed volume of E media along with the desired number of HFSCs in 15-mL conical vial, gently mix, and then add to plate (see Note 9 regarding media volume and cell density). 7. Culture HFSCs at 35  C in incubator with 5% CO2 (see Note 5). 8. Replace E media every 3 days. 9. Colonies become visible in approximately 7 days and fixation for immunofluorescence can be done when colonies reach the desired size. 10. Gently wash cells with 1 PBS, and then fix cells for 15 min in ice cold 4% paraformaldehyde. 11. Wash for 10 min with 1 PBS, and then twice in 1 PBS containing 0.1% Triton X-100 for 10 min each. 12. Block with 20% NGS-gelatin solution for 30 min at room temperature to overnight at 4  C. 13. Incubate with primary antibody diluted in 10% (or lower, depending on the specific antibody used) NGS-gelatin solution overnight at 4  C. Use of an anti-keratin 14 (K14) antibody helps to distinguish HFSC colonies from the fibroblast feeders (Fig. 2).

Fig. 2 Immunofluorescent staining for K14 expression in cultured HFSCs. DAPI stains the nuclei

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14. Wash three times with 1 PBS for 10 min at room temperature, and then wash with 2% NGS-gelatin solution for 15 min at room temperature. 15. Incubate with secondary antibody diluted in 2% NGS-gelatin solution for 1 h at room temperature in the dark. 16. Wash three times with 1 PBS containing 0.1% Triton X-100 for 10 min at room temperature in the dark. 17. Stain with DAPI at a final concentration of 2 μg/mL in 1 PBS for 10–20 min in the dark. 18. Wash three times with 1 PBS for 5 min each. 19. Mount slides with Vectashield and seal edges with nail polish. 20. Image colonies using epifluorescence or confocal microscopy. Note that the HFSCs often push the fibroblast feeders away as the colonies expand, resulting in a single layer of cells. However, the HFSC colonies may begin to stratify if cultured for too long. 3.7 Adenoviral Infection of HFSCs

Viral infection of HFSCs enables manipulation of gene expression and functional characterization. We found it necessary to temporarily remove the mitomycin C-treated J2-3T3 fibroblast feeders in order to maximize the viral infection efficiency of HFSCs. In our experiments, we utilized commercially available adenoviruses that express GFP so infection efficiency can be estimated based on GFP expression (Fig. 3). 1. Prepare mitomycin C-treated J2-3T3 fibroblast feeder layer as described in Sect. 3.2. 2. Add 1000 cells/cm2 of HFSCs (~9500 cells for 6-well plate) to appropriate volume of E media (2–3 mL for 6-well plate) in 15-mL conical vial, gently mix, and then add to well (see Note 9). 3. Swirl plate to mix. 4. Culture HFSCs at 35  C in incubator with 5% CO2 (see Note 5). 5. Replace E media every 3 days. 6. Culture HFSCs for 2 weeks so there are enough cells to infect. 7. Harvest HFSCs in a single well by following steps 1–7 in Sect. 3.3, and use for counting the number of cells (see Note 12). 8. To the remaining wells, add 2 mL versene and let sit for 2 min. 9. Vigorously pipette the versene solution with a P1000 to spray off all fibroblasts (see Note 7). 10. Aspirate versene, wash plate with 2 mL E media to remove residual fibroblasts, and then aspirate. 11. Mix appropriate amount of viruses based on cell count in step 7 to achieve multiplicity of infection (MOI) of 50 to 1 mL E media in a 1.5-mL Eppendorf tube and then carefully add to a well.

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48 h

24 h

72 h

B

% GFP+

100 90 80 70 60 50 40 30 20 10 0

1 2 3 Days After Infection

Fig. 3 GFP expression as a measure for efficiency of adenovirus infection in HFSCs. GFP protein was visualized by (a) fluorescence microscopy and (b) flow cytometry

12. Incubate cells with viruses overnight at 35  C in incubator with 5% CO2. 13. The next day, check cells under the microscope for GFP expression (Fig. 3). 14. Prepare mitomycin C-treated J2-3T3 fibroblast feeder layer as described in Sect. 3.2. 15. Remove the virus-containing media from HFSCs and then briefly rinse with 2 mL 1 PBS. 16. Add 1 mL 0.1% trypsin (no EDTA) and let incubate until cells start to lift up (about 10 min). 17. Add 4 mL E media to inactivate trypsin and spray to remove residual cells. 18. Put cells in a 15-mL conical vial and centrifuge at 1000 RPM for 5 min to pellet cells. 19. Aspirate media, resuspend pellet in 2 mL E media, and count cells.

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20. Add 1000 cells/cm2 of HFSCs (~9,500 cells for 6-well plate) to appropriate volume of E media (2–3 mL for 6-well plate) in 15-mL conical vial, gently mix, and then add to well (see Notes 9 and 13). 21. Culture HFSCs at 35  C in incubator with 5% CO2 (see Note 5). 22. Replace E media every 3 days. 23. Colonies should begin to be visible after ~4 days and then rapidly expand (see Note 8). 24. Proceed with downstream applications. 3.8 Patch Reconstitution Assay with Virally Infected HFSCs

The regenerative capacity of virally infected HFSCs can be assessed using a well-established “patch” assay [7], where HFSCs can be combined with newborn dermal cells and then subcutaneously injected into the backs of immunocompromised Nu/J mice. After 2 weeks, hair follicles form at the site of injection (Fig. 4). 1. Generate virally infected HFSCs by following steps in Sect. 3.7. Cells should have been growing for 2 weeks and ready for use once step 2 below is completed. 2. Isolate newborn primary dermal cells using the protocol from [7]. 3. Harvest HFSCs by following steps 1–7 in Sect. 3.3. 4. Combine 100,000 infected HFSCs with 500,000 newborn dermal cells in 1.5-mL Eppendorf tube and centrifuge at 1000 RPM for 5 min to pellet the cells. 5. As dermal-only negative control, add 500,000 newborn dermal cells to 1.5-mL Eppendorf tubes, and centrifuge to pellet at 1000 RPM for 5 min (see Note 14). 6. Very carefully remove the media from the vials as to not dislodge the cell pellets.

Fig. 4 Result of a “patch” assay using newborn keratinocytes (left) and infected HFSCs (right)

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7. Add 50 μL of E media to each pellet and then place the cells on ice and bring to the appropriate mouse procedure room. 8. Follow approved mouse protocols to sedate Nu/J mice, and sterilize the outer skin surface surrounding the desired injection site. 9. Carefully resuspend the cells in each 1.5-mL Eppendorf tube, and then draw cell suspension into syringe with a 25G needle. 10. Carefully inject the cell suspensions into the backs of the Nu/J mice (see [7] for detailed injection instructions). 11. Return mice to their cages and follow approved post-operation procedures. 12. Sacrifice mice 2 weeks later to examine hair follicles at each of the injection sites.

4

Notes 1. It is critical to not use over-confluent J2-3T3 fibroblasts. 2. If not used immediately, add 10 mL F media to mitomycin C-treated J2-3T3 fibroblasts. Cells can be kept in the incubator for a week before use. 3. Even distribution is critical for optimal HFSC growth. 4. It generally takes ~2 days for mitomycin C-treated J2-3T3 fibroblasts to attach and spread over the entire dish. It is not recommended that HFSCs be added until the tissue culture plastic is fully covered. 5. After initial seeding of HFSCs, do not move plate within the first 72 h to allow the cells to attach to the feeders. 6. HFSCs grow as colonies. Colonies derived from HFSCs isolated from p49 mice (when their hair follicles are in telogen) become visible within a week and will begin to expand very rapidly after coming out of quiescence. 7. Feeders are easily removed and HFSCs will remain attached. HFSCs will begin to detach if treated for extended amount of time. 8. Passaged HFSCs form colonies faster than freshly sorted HFSCs. 9. Adding cells to the full volume of media in a conical vial followed by mixing facilitates even distribution of HFSCs in the well. 10. It is recommended that imaging be performed between 7 and 9 days. Generally, beyond 10 days after plating, the HFSC colonies become too large for an entire colony to be captured in a single field of view.

Experimentation with Hair Follicle Stem Cells

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11. Imaging every 15 min or less is recommended to capture the dynamic cellular changes, as HFSCs expand fairly quickly at this time. 12. An extra well is used for counting cells before infection to calculate how much viruses to add to achieve an ideal MOI. 13. This plating density is good for a clonal assay. Different plating density may be desired per specific downstream assay. 14. A dermal-only control is necessary to gauge how much the dermal cells might be contaminated with newborn epidermal cells, which are regeneration-competent. References 1. Alonso L, Fuchs E (2006) The hair cycle. J Cell Sci 119(Pt 3):391–393 2. Hsu YC, Li L, Fuchs E (2014) Emerging interactions between skin stem cells and their niches. Nat Med 20(8):847–856 3. Ito M, Kizawa K, Hamada K, Cotsarelis G (2004) Hair follicle stem cells in the lower bulge form the secondary germ, a biochemically distinct but functionally equivalent progenitor cell population, at the termination of catagen. Differentiation 72(9-10):548–557 4. Rompolas P, Mesa KR, Greco V (2013) Spatial organization within a niche as a determinant of stem-cell fate. Nature 502(7472):513–518 5. Rompolas P et al (2012) Live imaging of stem cell and progeny behaviour in physiological hairfollicle regeneration. Nature 487(7408):496–499

6. Nowak JA, Fuchs E (2009) Isolation and culture of epithelial stem cells. Methods Mol Biol 482:215–232 7. Zheng Y, Hsieh JC, Escandon J, Cotsarelis G (2016) Isolation of mouse hair follicle bulge stem cells and their functional analysis in a reconstitution assay. Methods Mol Biol 1453:57–69 8. Blanpain C, Lowry WE, Geoghegan A, Polak L, Fuchs E (2004) Self-renewal, multipotency, and the existence of two cell populations within an epithelial stem cell niche. Cell 118(5):635–648 9. Adam RC et al (2015) Pioneer factors govern super-enhancer dynamics in stem cell plasticity and lineage choice. Nature 521(7552):366–370

Methods in Molecular Biology (2019) 1879: 31–41 DOI 10.1007/7651_2018_149 © Springer Science+Business Media New York 2018 Published online: 08 May 2018

Quantitative Detection of Low-Abundance Transcripts at Single-Cell Level in Human Epidermal Keratinocytes by Digital Droplet Reverse Transcription-Polymerase Chain Reaction Fre´de´ric Auvre´, Julien Coutier, Miche`le T. Martin, and Nicolas O. Fortunel Abstract Genetic and epigenetic characterization of the large cellular diversity observed within tissues is essential to understanding the molecular networks that ensure the regulation of homeostasis, repair, and regeneration, but also pathophysiological processes. Skin is composed of multiple cell lineages and is therefore fully concerned by this complexity. Even within one particular lineage, such as epidermal keratinocytes, different immaturity statuses or differentiation stages are represented, which are still incompletely characterized. Accordingly, there is presently great demand for methods and technologies enabling molecular investigation at single-cell level. Also, most current methods used to analyze gene expression at RNA level, such as RT-qPCR, do not directly provide quantitative data, but rather comparative ratios between two conditions. A second important need in skin biology is thus to determine the number of RNA molecules in a given cell sample. Here, we describe a workflow that we have set up to meet these specific needs, by means of transcript quantification in cellular micro-samples using flow cytometry sorting and reverse transcriptiondigital droplet polymerase chain reaction. As a proof-of-principle, the workflow was tested for the detection of transcription factor transcripts expressed at low levels in keratinocyte precursor cells. A linear correlation was found between quantification values and keratinocyte input numbers in a low quantity range from 40 cells to 1 cell. Interpretable signals were repeatedly obtained from single-cell samples corresponding to estimated expression levels as low as 10–20 transcript copies per keratinocyte or less. The present workflow may have broad applications for the detection and quantification of low-abundance nucleic acid species in single cells, opening up perspectives for the study of cell-to-cell genetic and molecular heterogeneity. Interestingly, the process described here does not require internal references such as house-keeping gene expression, as it is initiated with defined cell numbers, precisely sorted by flow cytometry. Keywords ddPCR, Keratinocytes, Low-abundance nucleic acids, Single-cell level, Transcript quantification

1

Introduction Wide cellular diversity is found within all tissues. It is required to control homeostasis, repair, and regeneration, and can be disturbed by pathological dysregulations. Skin is a perfect illustration of this diversity, being composed of multiple cell lineages, each of which comprises various cell subpopulations with distinct cellular and

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molecular properties. Even within a single cell lineage, such as keratinocytes constituting the epithelial part of the epidermis, different immaturity statuses and differentiation stages can be detected and separated, including diverse subpopulations of stemand progenitor-cells [1–6]. Further cellular and molecular studies remain needed for full characterization. In previous issues of Methods Mol Biol, our group presented an experimental model enabling functional investigation of keratinocyte stem- and progenitor-cells at single-cell level, based on multi-parallel clonal micro-cultures [7] and a dye-tracking approach to monitor the cultured keratinocyte cycle [8]. The present study describes a method suitable for detection and quantification of transcripts in samples with low cell numbers, down to single-cell level, using flow cytometry sorting and reverse transcription-digital droplet polymerase chain reaction (RT-ddPCR). The goal was to set up a method suitable for acquisition of quantitative data on transcript expression in normal and pathological samples of human epidermal cells, and sensitive enough to enable the study of transcripts expressed at low levels in single cells. Briefly, digital PCR is a refinement of classical PCR methods that allows absolute quantification of nucleic acids, including cDNA (RT-dPCR). The principle of digital PCR consists in fractionation of the sample reaction volume into thousands of smaller reactions, so that individual nucleic acid targets are partitioned in separate micro-volumes. In digital droplet PCR (ddPCR) technology, sample fractionation is based on water-oil emulsion. Each PCR reaction volume is fractionated into nanoliter-size samples which are encapsulated into oil droplets. PCR amplification is then performed similarly to classical TaqMan assay, giving rise to a mix of positive fluorescent droplets (presence of the PCR target sequence) together with negative non-fluorescent droplets (absence of the PCR target sequence). The fraction of fluorescent droplets is determined on binary readout of the numbers of positive and negative droplets per sample. Interpretation of ddPCR signals is based on the fact that the distribution of target molecules within the droplet ensemble follows a Poisson distribution. The correspondence between the proportion of fluorescent droplets and absolute quantity of copies of the target sequence in the input sample can therefore be determined according to this probability distribution. Readout of positive droplets thus provides a quantitative estimate of specific expressed sequences (see [9, 10]). We expect RT-ddPCR to provide a useful tool for different areas of fundamental, translational, and clinical research on skin. The wide range of applications may include detection and quantification of low-abundance messenger RNAs (mRNAs) and mRNA variants, and also analysis of diverse epigenetic molecular species such as long noncoding RNAs (LncRNAs) [11] and micro-RNAs (miRNAs) [12], the functions of which in normal skin

Quantitative Detection of Low-Abundance Transcripts at Single-Cell Level in. . .

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morphogenesis, homeostasis, aging, and regenerative capacity are far from being fully understood. In particular, analysis of low-abundance nucleic species is of great importance for further exploration of the stemness concept [13], notably in skin. In the domain of pathophysiology, RT-ddPCR may be applied to characterize mRNA expression dysregulation. It is also being investigated for the detection of pathogens, including viral genomes [14]. RT-ddPCR applications are exemplified in the present study by detection and quantification of two transcripts from transcription factor genes which are studied by our group for their role in the regulation of the cell-fate decision in human keratinocyte precursor cells, notably in the maintenance of their immature character: MXD4 and KLF4 transcripts, which respectively encode MAD4 and KLF4 proteins ([15]; for a review, see [16]). Transcription factors are often characterized by low transcript and protein expression levels in normal cells. Like low-abundance nucleic acid species in general, their study thus depends on the availability of sensitive quantification methods, especially in the perspective of working at single-cell level. Finally, considering the current demand for validation of wide genomic and epigenomic data based on massive next-generation sequencing, which is in continuous expansion, the workflow described in this chapter may constitute an efficient tool for the qualification of samples and technical processes. In particular, it may contribute to the development and validation of genomewide analyses applied in cell micro-samples, including at singlecell level. Notably, the biological significance of global profiling approaches such as transcriptome analysis by RNA-sequencing applied on cell micro-samples depends on careful checking to ensure that low-level expressions are correctly detected and not excluded from the analysis pipeline.

2 2.1

Materials Cellular Material

The cells used in the example of RT-ddPCR analysis shown here were cultured human epidermal keratinocyte precursor cells, defined as the progeny of the basal stem- and progenitor-cell fraction obtained by the previously described selection method based on rapid adhesion on collagen [17]. The cells were used at passage1 stage after primary culture. The example, presented as a proof-ofprinciple, is nonrestrictive, and the workflow can be directly applied to the study of the different cell types in any tissue. In skin, suitable cellular material non-exhaustively includes the various immature or differentiated keratinocyte subpopulations and dermal fibroblast subtypes [18].

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Equipment

2.2.1 Flow Cytometry

Cell sorter: MoFlo Astrios EQ (Beckman-Coulter Life Sciences), allowing 6-way sorting and deposition of defined cell numbers, including single cells. Excitation sources (lasers). Wavelength/power: 355 nm/100 mW; 405 nm/55 mW; 488 nm/200 Mw; 561 nm/200 mW; and 642 nm/100 mW. Biological safety cabinet, Class II Type A2, SteriGARD (Baker). Summit™ software system v6.3 (Cytomation, Inc.).

2.2.2 Microplate Sealing

PX1™ PCR plate sealer (Bio-Rad; cat. no. 1814000).

2.2.3 Droplet Generation

QX200 droplet generator (Bio-Rad; cat. no. 1864002).

2.2.4 Sample Lysis, cDNA Synthesis, and PCR Amplification

C1000 Touch thermal cycler (Bio-Rad).

2.2.5 Readout

QX200™ droplet reader (Bio-Rad; cat. no. 1864003). QuantaSoft™ software, regulatory edition (Bio-Rad; no. 1864011).

2.3 Reagents and Products 2.3.1 Flow Cytometry Cell Sorting

cat.

Flow cytometry tubes, 5 ml polypropylene round-bottom (Falcon; cat. no. 352063). BSA (Sigma; cat. no. A9418). DPBS-2% BSA filtered through a 0.22 μm filter and stored as frozen aliquots at 20  C. Hard-shell® 96-well PCR plates, clear well, white shell (Bio-Rad; cat. no. HSP9601).

2.3.2 Cell Lysis

Tris-EDTA buffer (10 mM Tris–HCl pH 8.0, 0.1 mM EDTA) (Sigma-Aldrich; cat. no. T9285). SingleShot™ cell lysis buffer (Bio-Rad; cat. no. 172-5080). Proteinase K solution (Bio-Rad; cat. no. 172-5080). DNAse solution (Bio-Rad; cat. no. 172-5080). Microseal® ‘B’ PCR plate sealing film (Bio-Rad; cat. no. MSB1001).

2.3.3 Reverse Transcription

Distilled water, DNase/RNase-free (Invitrogen; cat. no. 0977035). SuperScript™ VILO™ cDNA synthesis [reverse-transcriptase 5 RT buffer and 10 VILO SuperScript enzyme] (Invitrogen; cat. no. 11754050). Microseal® ‘B’ PCR plate sealing film (Bio-Rad; cat. no. MSB1001).

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2.3.4 Polymerase Chain Reaction

ddPCR™ supermix for probes (No dUTP) (Bio-Rad; cat. no. 1863024). ddPCR™ 96-well plates (Bio-Rad; cat. no. 12001925). PCR plate heat seal, foil, pierceable (Bio-Rad; cat. no. 1814040).

2.3.5 Droplet Generation

DG8 cartridges and gasket (Bio-Rad; cat. no. 1864007). Droplet generation oil (Bio-Rad; cat. no. 1863005).

2.3.6 Signal Readout

ddPCR droplet reader oil (Bio-Rad; cat. no. 1863004).

3

Methods

3.1 Description of the Workflow

As described in detail in Subheading 3 below, the workflow that we have set up for transcript quantification in keratinocyte microsamples includes the following experimental steps, schematized in Fig. 1: 1. Automated deposition of defined numbers of cells in microwells using flow cytometry. 2. Cell lysis and RNA reverse transcription. 3. Sample preparation for polymerase chain reaction (PCR mixture). 4. Droplet generation (sample partitioning through water-oil emulsion). 5. Polymerase chain reaction (generation of fluorescent droplets). 6. Readout (quantification of positive and negative droplets). In the example of a RT-ddPCR experiment presented here, the feasibility of MXD4 and KLF4 transcript detection was investigated in samples with decreasing quantities of input cellular material. – For KLF4 transcript amplification, primers were selected in exons 2 and 3 of the human KLF4 gene (Thermo-Fisher Scientific; primers ref. HS00358836_m1; fluorescent probe FAM-MGB), producing a 110 base-length amplicon. – For MXD4 transcript amplification, primers were selected in exons 3 and 4 of the human MXD4 gene (Thermo-Fisher Scientific; primers ref. HS01557630_m1; fluorescent probe FAM-MGB), producing a 60 base-length amplicon. The tested keratinocyte quantities comprised 40 cells, 20 cells, 5 cells, and 1 single cell. For each cell number, 20–47 replicates were analyzed. The four micro-sample sizes gave rise to interpretable signals for both KLF4 and MXD4 transcripts, as shown in Fig. 2. Moreover, a proportionality between transcript quantification values and input keratinocyte numbers was found, indicating that the workflow was functional in this low cell-quantity range,

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a

b Deposition of defined cell numbers

Sample partitioning and target amplification

Flow cytometry

Lysis RT

Water-oil emulsion

ddPCR Target sequence Non-target sequences

Micro-samples (single cells)

c

d Binary readout

Fluorescent signal

Fluorescence

Signal analysis in individual droplets Positive

Threshold Negative Analyzed droplets

Fig. 1 Schematic representation of the RT-ddPCR workflow. (a) Generation of micro-samples by flow cytometry cell-sorting and automated deposition. Defined cell numbers are deposited in multi-well plates containing the lysis solution. This input cellular material can be single cells. (b) Cell lysis, reverse transcription, sample partitioning, and ddPCR target amplification. Steps from cell lysis to cDNA synthesis and preparation of the amplification mixture are performed in the same well to avoid material loss. Sample partitioning is then achieved based on the principle of water-oil emulsion. The ddPCR reaction is then performed. (c) ddPCR amplification gives rise to two droplet populations: droplets that initially contained the target become fluorescent after PCR amplification, whereas droplets that did not contain the target remain non-fluorescent. Droplets are analyzed individually for fluorescence signal. (d) The ddPCR signal is analyzed on binary readout. The absolute number of molecular targets present in initial samples can be determined from the proportion of positive and negative droplets based on Poisson distribution

including single-cell analysis. The method repeatedly enabled detection of KLF4 and MXD4 transcripts in single keratinocytes, and estimated their expression levels in the range of 9.5  10.1 and 10.2  6.1 copies per cell, respectively. Characterization of this heterogeneity is critical for further understanding the intrinsic properties that can distinguish different individual cells. In

Quantitative Detection of Low-Abundance Transcripts at Single-Cell Level in. . .

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Fig. 2 Detection and quantification of KLF4 and MXD4 transcripts in micro-samples of human epidermal keratinocyte precursor cells by RT-ddPCR. Input keratinocyte quantities were 40, 20, and 5 cells, or 1 cell. In the example shown, each input cell quantity was analyzed in 20–47 replicate RT-ddPCR points. Dots represent RT-ddPCR transcript quantification values obtained with equivalent replicate cell samples. (a, b) A linear correlation between quantification values and input keratinocyte numbers was found for the two transcripts (see linear scale representations). Correlation coefficients (R2) were 0.9778 and 0.9781 for KLF4 and MXD4 transcripts, respectively. Moreover, the possibility for transcript quantification in single cells was validated, providing estimates of KLF4 and MXD4 transcript numbers in the range of 9.5  10.1 and 10.2  6.1 copies per individual keratinocyte precursor cell, respectively (see log-scale representations). Dots correspond to ddPCR analyses of replicate biological samples from three independent experiments. Means  SD are indicated

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summary, this methodological set-up validates the feasibility of detection and quantification of low-abundance transcripts in single human keratinocytes. Interestingly, the present process does not require internal references such as house-keeping gene expression, which frequently constitutes a difficult issue in RT-PCR. Indeed, the workflow described here is initiated with defined cell numbers, which is precisely ensured by flow cytometry sorting. It is important to bear in mind that high-abundance nucleic species can saturate the system, even with single cells as input material, requiring sample dilution. For example, to test RT-ddPCR detection of the strongly expressed 18S rRNA in single keratinocytes, a 1/400 dilution of cDNA samples was used, allowing detection in the range of 10,000–15,000 copies per cell. Accordingly, when quantification is the objective, we recommend checking that the cellular abundance of the specific targets matches the quantitative detection range of the method, by systematically establishing a curve of cell quantity versus signal. 3.2 Automated Deposition of Defined Numbers of Cells in Micro-Wells Using Flow Cytometry

1. Prepare a volume of lysis solution corresponding to 5 μl/sample: mix 4.2 μl Tris-EDTA buffer, 0.64 μl singleshot cell lysis buffer, 0.08 μl proteinase K solution, and 0.08 μl DNAse solution (see Note 1). 2. Distribute the lysis solution in a 96-well PCR plate. 3. Suspend cell samples in PBS-2% BSA, in flow cytometry tubes (see Note 2). 4. Set the MoFlo at a low sheath pressure (20 psi), use a 100 μm nozzle, and adjust drop drive frequency at 40 kHz, voltage at 55 V. 5. Set sorting gates according to defined morphological and phenotypic characteristics, and for cell doublet exclusion (see Note 3). 6. Process for deposition of defined numbers of cells into microwells (see Notes 4 and 5). 7. Seal the micro-well plate. 8. Centrifuge the plate for 1 min at 500  g.

3.3 Cell Lysis and RNA Reverse Transcription

1. Process for cell lysis using the following thermocycler program: 10 min at 25  C, then 5 min at 75  C, and at 7  C until sample storage or further processing (see Note 6). 2. Prepare a volume of reverse transcription mix corresponding to 6 μl/sample: mix 3 μl DNase/RNase-free H2O, 2 μl 5 RT buffer, and 1 μl 10 VILO SuperScript enzyme (see Notes 1 and 7). 3. Remove the sealing film from the plate. 4. Add 6 μl RT mixture to each 5 μl cell lysate, and homogenize by gentle pipetting.

Quantitative Detection of Low-Abundance Transcripts at Single-Cell Level in. . .

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5. Seal the micro-well plate. 6. Process for reverse transcription (cDNA synthesis) using the following thermocycler program: 10 min at 25  C, followed by 60 min at 42  C, 5 min at 85  C, and then 7  C until next step. 3.4 Sample Preparation for Polymerase Chain Reaction (ddPCR Mixture)

1. Prepare a volume of ddPCR mixture corresponding to 11 μl/ sample: mix 10 μl of ddPCR supermix for probes and 1 μl probes (see Notes 1 and 8).

3.5 Droplet Generation (Sample Partitioning Through Water-Oil Emulsion)

1. Carefully transfer the samples (20 μl) to the droplet generation cartridge (see Note 9).

2. Remove the sealing film from the plate. 3. Add 11 μl ddPCR mixture to each 11 μl cDNA sample, and homogenize by gentle pipetting.

2. Fill cartridge wells with 70 μl droplet generation oil. 3. Cover the wells with the rubber seal (see Note 10). 4. Process for water-oil emulsion using the droplet generator.

3.6 Polymerase Chain Reaction (Generation of Fluorescent Droplets)

1. Carefully transfer the samples (40 μl) to a new 96-well plate (see Note 11).

3.7 Readout (Quantification of Positive and Negative Droplets)

1. Place the plate in the droplet reader without removing the sealing film.

4

2. Seal the micro-well plate using a pierceable film. 3. Process polymerase chain reaction (ddPCR) using the following thermocycler program: 10 min at 95  C, followed by 40 cycles consisting of 30 s at 94  C and 1 min at 60  C, and then 10 min at 98  C, and finally 7  C until next step (see Note 12).

2. Quantify positive and negative droplets. 3. Process for data analysis and interpretation using the QuantaSoft™ software (see Note 13).

Notes 1. Prepare more volume than strictly needed, due to loss during pipetting, which can be estimated as 20%. 2. Filtering the cell suspension is recommended to strictly avoid aggregates. 3. Use the area-versus-peak ratio on a fluorescence parameter for doublet discrimination. 4. We suggest analyzing samples in at least 4–8 replicates.

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5. Check the accuracy of cell deposition by visual microscopic observation, as previously described [6, 7]. 6. At this stage, samples can be stored frozen at

80  C.

7. We recommend the use of high fidelity enzymes to ensure high accuracy and sensitivity. 8. cDNA amplification primers are preferentially selected in different exons of the gene. The fluorescent probes that are used to generate the PCR signal are available with different wavelengths (TaqMan principle, the presented example, FAM-MGB, Thermo-Fisher Scientific). 9. Strictly avoid making bubbles. 10. Check that the rubber seal is correctly adjusted. 11. Pipet very gently, as droplets are fragile and can be easily destroyed. 12. Set the thermocycler so that lip temperature is maintained at 105  C, and temperature changes are operated at a low speed of 2  C/s (gradient), to maintain droplet integrity. 13. Molecular target distribution within droplets after sample partitioning follows a Poisson distribution. Accordingly, the absolute number of molecular targets present in initial samples can be determined from the proportion of positive and negative droplets, based on Poisson distribution [9, 10].

Acknowledgments We thank Genopole® (Evry, France), and particularly Julien Picot, who provided support for equipment and infrastructure. This work was supported by grants from: CEA and INSERM (UMR967) and the De´le´gation Ge´ne´rale de l’Armement (DGA) grants; FUI-AAP13 and the Conseil Ge´ne´ral de l’Essonne within the STEMSAFE grant; and EURATOM (RISK-IR, FP7, grant 323267). Julien Coutier received a CEA-DGA thesis fellowship grant. References 1. Li A, Simmons PJ, Kaur P (1998) Identification and isolation of candidate human keratinocyte stem cells based on cell surface phenotype. Proc Natl Acad Sci U S A 95:3902–3907 2. Fortunel NO, Hatzfeld JA, Rosemary PA, Ferraris C, Monier MN, Haydont V, Longuet J, Brethon B, Lim B, Castiel I, Schmidt R, Hatzfeld A (2003) Long-term expansion of human functional epidermal precursor cells: promotion of extensive

amplification by low TGF-beta1 concentrations. J Cell Sci 116:4043–4052 3. Larderet G, Fortunel NO, Vaigot P, Cegalerba M, Malte`re P, Zobiri O, Gidrol X, Waksman G, Martin MT (2006) Human side population keratinocytes exhibit long-term proliferative potential and a specific gene expression profile and can form a pluristratified epidermis. Stem Cells 24:965–974 4. Rachidi W, Harfourche G, Lemaitre G, Amiot F, Vaigot P, Martin MT (2007) Sensing

Quantitative Detection of Low-Abundance Transcripts at Single-Cell Level in. . . radiosensitivity of human epidermal stem cells. Radiother Oncol 83:267–276 5. Harfouche G, Vaigot P, Rachidi W, Rigaud O, Moratille S, Marie M, Lemaitre G, Fortunel NO, Martin MT (2010) Fibroblast growth factor type 2 signaling is critical for DNA repair in human keratinocyte stem cells. Stem Cells 28:1639–1648 6. Fortunel NO, Cadio E, Vaigot P, Chadli L, Moratille S, Bouet S, Rome´o PH, Martin MT (2010) Exploration of the functional hierarchy of the basal layer of human epidermis at the single-cell level using parallel clonal microcultures of keratinocytes. Exp Dermatol 19:387–392 7. Fortunel NO, Vaigot P, Cadio E, Martin MT (2010) Functional investigations of keratinocyte stem cells and progenitors at a single-cell level using multiparallel clonal microcultures. Methods Mol Biol 585:13–23 8. Chadli L, Cadio E, Vaigot P, Martin MT, Fortunel NO (2013) Monitoring the cycling activity of cultured human keratinocytes using a CFSE-based dye tracking approach. Methods Mol Biol 989:83–97 9. Hindson BJ, Ness KD, Masquelier DA, Belgrader P, Heredia NJ, Makarewicz AJ, Bright IJ, Lucero MY, Hiddessen AL, Legler TC, Kitano TK, Hodel MR, Petersen JF, Wyatt PW, Steenblock ER, Shah PH, Bousse LJ, Troup CB, Mellen JC, Wittmann DK, Erndt NG, Cauley TH, Koehler RT, So AP, Dube S, Rose KA, Montesclaros L, Wang S, Stumbo DP, Hodges SP, Romine S, Milanovich FP, White HE, Regan JF, Karlin-Neumann GA, Hindson CM, Saxonov S, Colston BW (2011) High-throughput droplet digital PCR system for absolute quantitation of DNA copy number. Anal Chem 83:8604–8610 10. Pinheiro LB, Coleman VA, Hindson CM, Herrmann J, Hindson BJ, Bhat S, Emslie KR (2012) Evaluation of a droplet digital polymerase chain reaction format for DNA copy number quantification. Anal Chem 84:1003–1011 11. Kretz M, Siprashvili Z, Chu C, Webster DE, Zehnder A, Qu K, Lee CS, Flockhart RJ, Groff AF, Chow J, Johnston D, Kim GE, Spitale RC, Flynn RA, Zheng GX, Aiyer S, Raj A, Rinn JL,

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Chang HY, Khavari PA (2013) Control of somatic tissue differentiation by the long non-coding RNA TINCR. Nature 493:231–235 12. Nagosa S, Leesch F, Putin D, Bhattacharya S, Altshuler A, Serror L, Amitai-Lange A, Nasser W, Aberdam E, Rouleau M, Tattikota SG, Poy MN, Aberdam D, Shalom-Feuerstein R (2017) microRNA-184 induces a commitment switch to epidermal differentiation. Stem Cell Reports 9:1991–2004 13. Fortunel NO, Otu HH, Ng HH, Chen J, Mu X, Chevassut T, Li X, Joseph M, Bailey C, Hatzfeld JA, Hatzfeld A, Usta F, Vega VB, Long PM, Libermann TA, Lim B (2003) Comment on “‘Stemness’: transcriptional profiling of embryonic and adult stem cells” and “a stem cell molecular signature”. Science 302:393 14. Arvia R, Sollai M, Pierucci F, Urso C, Massi D, Zakrzewska K (2017) Droplet digital PCR (ddPCR) vs quantitative real-time PCR (qPCR) approach for detection and quantification of Merkel cell polyomavirus (MCPyV) DNA in formalin fixed paraffin embedded (FFPE) cutaneous biopsies. J Virol Methods 246:15–20 15. Hurlin PJ, Que´va C, Koskinen PJ, Steingrı´msson E, Ayer DE, Copeland NG, Jenkins NA, Eisenman RN (1995) Mad3 and Mad4: novel Max-interacting transcriptional repressors that suppress c-myc dependent transformation and are expressed during neural and epidermal differentiation. EMBO J 14:5646–5659 16. Sur I (2009) Kru¨ppel-like factors 4 and 5: unity in diversity. Curr Genomics 10:594–603 17. Fortunel NO, Chadli L, Bourreau E, Cadio E, Vaigot P, Marie M, Deshayes N, RathmanJosserand M, Leclaire J, Martin MT (2011) Cellular adhesion on collagen: a simple method to select human basal keratinocytes which preserves their high growth capacity. Eur J Dermatol 21(Suppl 2):12–20 18. Mine S, Fortunel NO, Pageon H, Asselineau D (2008) Aging alters functionally human dermal papillary fibroblasts but not reticular fibroblasts: a new view of skin morphogenesis and aging. PLoS One 3:e4066

Methods in Molecular Biology (2019) 1879: 43–73 DOI 10.1007/7651_2018_153 © Springer Science+Business Media New York 2018 Published online: 27 May 2018

Qualitatively Monitoring Binding and Expression of the Transcription Factors Sp1 and NFI as a Useful Tool to Evaluate the Quality of Primary Cultured Epithelial Stem Cells in Tissue Reconstruction Gae¨tan Le-Bel, Sergio Cortez Ghio, Danielle Larouche, Lucie Germain, and Sylvain L. Gue´rin Abstract Electrophoretic mobility shift assays and Western blots are simple, efficient, and rapid methods to study DNA–protein interactions and protein expression, respectively. Primary cultures and subcultures of epithelial cells are widely used for the production of tissue-engineered substitutes and are gaining popularity as a model for gene expression studies. The preservation of stem cells through the culture process is essential to produce high quality substitutes. However, the increase in the number of cell passages is associated with a decrease in their ability to proliferate until senescence is reached. This process is likely to be mediated by the altered expression of nuclear-located transcription factors such as Sp1 and NFI, whose expression has been documented to be required for cell adhesion, migration, and differentiation. In some of our recent studies, we observed a correlation between reconstructed tissues exhibiting poor histological and structural characteristics and a low expression of Sp1 in their constituting epithelial cells. Therefore, monitoring both the expression and DNA binding of these transcription factors in human skin and corneal epithelial cells is a useful tool for characterizing the quality of primary cultured epithelial cells. Keywords Cornea, Corneal epithelial cells, EMSA, Epidermis, Gel mobility shift assay, NFI, Sp1, Stem cells, Tissue engineering, Transcription factor, Western blot

Abbreviations AEBSF anhCEC anhK APS ATP BCA ccDME-Ham ckDME-Ham DMEM DMSO DNA DTT

4-(2-Aminoethyl)benzenesulfonyl fluoride Adult normal human corneal epithelial cells Adult normal human keratinocytes Ammonium persulfate Adenosine triphosphate Bicinchoninic acid Complete corneal epithelial cell culture medium Complete keratinocyte culture medium Dulbecco’s modified Eagle’s medium Dimethyl sulfoxide Deoxyribonucleic acid Dithiothreitol

43

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ECL EDTA EGF EMSA Ham HEPES i3T3 iHFL MMP9 NE NFI nhCEC nhK nnhCEC nnhK PAGE PBS PMSF PNK poly(dI:dC) PVDF SCC SDS Sp1 STE TBS TBS-T tDMEM TE TEMED TM

1

Enhanced chemiluminescence Ethylenediaminetetraacetic acid Epidermal growth factor Electrophoretic mobility shift assay Ham’s medium 4-(2-Hydroxyethyl)-1-piperazineethanesulfonic acid Irradiated Swiss 3T3 Irradiated human feed layer Matrix metallopeptidase 9 Nuclear extract Nuclear factor I Normal human corneal epithelial cell Normal human keratinocytes Newborn normal human corneal epithelial cells Newborn normal human keratinocytes Polyacrylamide gel electrophoresis Phosphate buffered saline Phenylmethylsulfonyl fluoride T4 polynucleotide kinase Poly(deoxyinosinic-deoxycytidylic) acid sodium salt Polyvinylidene fluoride Supershifted complexes Sodium dodecyl sulfate Specificity protein 1 Sodium Tris–HCl EDTA Tris Buffered Saline Tris Buffered Saline tween Tissue transport medium Tris–HCl EDTA Tetramethylethylenediamine Melting temperature

Introduction When developing bioengineered tissues for transplantation, the preservation of epithelial stem cells is of the utmost importance to warrant graft survival and long-term regeneration of the epithelium. Consequently, it has become particularly important to develop molecular tools to validate cell culture methods currently used for stem cell isolation, expansion, and preservation. The use of a fibroblast feeder layer, as described by Rheinwald and Green, provides a solid basis to achieve success in culturing human epithelial cells for clinical applications [1]. Although the absence of allogeneic feeder cell layer would be desirable during the production of autologous tissues for grafting purposes, the presence of fibroblasts and/or their secreted products greatly improves the morphological characteristics and proliferative potential of epithelial cells [2–5]. We

Quality of Primary Cultured Epithelial Stem Cells

45

have previously shown that in cultures of human skin keratinocytes, feeder layers of irradiated Swiss 3T3 (i3T3) fibroblasts or irradiated human fibroblast (iHFL) delay keratinocyte terminal differentiation by preventing extinction of the ubiquitous transcription factors Sp1 and NFI [6, 7]. Furthermore, the progressive loss of Sp1 and NFI expression during the last passages of cultured human skin keratinocytes or corneal epithelial cells also correlates with their growth arrest and terminal differentiation [6–9]. Monitoring both the expression and DNA binding properties of Sp1 and NFI is therefore useful as a tool in order to assess the quality of the epithelial cell cultures that are used for the production of engineered tissues. This chapter details the methods for harvesting and processing nuclear protein extracts of epithelial cells that have been cultured on an irradiated iHFL feeder layer as described (for a detailed protocol, see [10]), as well as the further analysis of Sp1 and NFI by Western blotting and electrophoretic mobility shift assay (EMSA).

2 2.1

Materials Cell Culture

2.1.1 Base Medium

2.1.2 Sera

DME-Ham. Combine three parts Dulbecco’s modified Eagle’s medium (DMEM; cat. no. 12800-082, Gibco) with one part Ham’s medium (cat. no. 21700-075, Life Technologies) in apyrogenic ultrapure water. Add 3.07 g/L of NaHCO3 (36.54 mM; cat. no. 3509-05, J.T. Baker), 24.3 mg/L of adenine (0.18 mM; cat. no. 0183-50G, Amresco) solubilized in 312.5 μL/L of 2 N HCL (2 M, cat. no. 0336-03, J.T. Baker). Adjust pH to 7.1. Sterilize by filtration through a 0.22 μm low binding disposable filter (see Subheading 2.1.7, item 1) and store in the dark at 4
C. 1. Fetal calf serum (cat. no. SH30070, HyClone). 2. Fetal clone II serum (cat. no. SH30066, HyClone). Thaw sera at 4
C or in cold water (see Subheading 4; Note 1). Gently swirl it to resuspend its components. Inactivate in a 56
C water bath for 30 min. To avoid repeated thawing and freezing cycles, distribute in single use aliquots. Store at 20
C, or at 80
C for long-term storage.

2.1.3 Additives

1. Insulin (cat. no. 91077C, SAFC Bioscience). Dissolve at 5 mg/ mL in 5 mM HCl. Yields a 1000 stock solution. Can be stored at 4
C. 2. Epidermal growth factor (EGF; cat. no. 236-GMP, R&D Systems). Dissolve at 200 μg/mL in 10 mM HCl. Dilute (1:20) the solution with DME-Ham (see Subheading 2.1.1) containing 10% v/v Fetal clone II serum (see Subheading 2.1.2, item 2). Yields a 1000 stock solution.

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3. Hydrocortisone (DIN. 872520, Teva). Dissolve at 5 mg/mL in 95% ethanol. Dilute (1:25) the solution with DME-Ham (see Subheading 2.1.1). Yields a 500 stock solution. 4. Penicillin G/Gentamicin (DIN. 02220296, Fresenius Kabi, and cat. no. GE152, Galenova, respectively). Dissolve penicillin G at 50000 IU/mL and active gentamicin at 12.5 mg/mL in apyrogenic ultrapure water. Yields a 500 stock solution. 5. Isoproterenol hydrochloride (DIN. 00897639, Sandoz). Store at 4
C. Vials are at 0.2 mg/mL and are single use. Leftover isoproterenol should not be stored. 1000 stock solution. 6. Fungizone (DIN. 00029149, Bristol Meyers Squibb Canada). Dissolve amphotericin B at 0.25 mg/mL in apyrogenic ultrapure water. Yields a 500 stock solution. For all additives, except isoproterenol (see Subheading 2.1.3, item 5), sterilize by filtration through a 0.22 μm low binding disposable filter (see Subheading 2.1.7, item 1). To avoid repeated thawing and freezing cycles, distribute in single use aliquots. Store at 80
C. 2.1.4 Complete Media

1. Tissue transport medium (tDMEM). Add 10% (v/v) of fetal calf serum (see Subheading 2.1.2, item 1) to high (4.5 g/L) glucose DMEM (containing sodium pyruvate and L-glutamine; cat. no. 10-013, Mediatech). Then add penicillin G/gentamicin and fungizone (dilute to 1; see Subheading 2.1.3, items 4 and 6). Store in the dark at 20 to 80
C for up to 6 months or at 4
C for up to 10 days. 2. Complete keratinocyte culture medium (ckDME-Ham). First, add 5% (v/v) of fetal clone II serum (see Subheading 2.1.2, item 2). Then, add insulin (dilute to 1; see Subheadings 2.1.3, items 1 and 4; Note 2) to DME-Ham (see Subheading 2.1.1), 1.06 mL/L of isoproterenol (see Subheading 2.1.3, item 5), epidermal growth factor, hydrocortisone, and penicillin G/gentamicin (dilute to 1; see Subheading 2.1.3, items 2–4). Store in the dark at 4
C for up to 10 days. 3. Complete corneal epithelial cell culture medium (ccDMEHam). First, add 5% (v/v) of fetal clone II serum (see Subheading 2.1.2, item 2). Then, add insulin (dilute to 1; see Subheadings 2.1.3, items 1 and 4; Note 2) to DME-Ham (see Subheading 2.1.1), 1.06 mL/L of isoproterenol (see Subheading 2.1.3, item 5), epidermal growth factor, hydrocortisone, and penicillin G/gentamicin (dilute to 1; see Subheading 2.1.3, items 2–4). Store in the dark at 4
C for up to 10 days. 4. Cryopreservation medium for normal human keratinocytes. Add dimethyl sulfoxide (DMSO; cat. no. D2438, Sigma Aldrich) in fetal calf serum, at 10% (v/v), (see Subheading 2.1.2, item 1). Keep on ice or store at 4
C and use within the day (see Subheading 4; Note 3).

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5. Cryopreservation medium for normal human corneal epithelial cell. Add dimethyl sulfoxide (DMSO; cat. no. D2438, Sigma Aldrich) to fetal clone II serum, at 10% (v/v) (see Subheading 2.1.2, item 2). Keep on ice or store at 4
C and use within the day (see Subheading 4; Note 3). All frozen components can be thawed at 4
C (see Subheading 4; Note 1). 2.1.5 Other Solutions

1. Phosphate buffered saline (PBS). Dissolve NaCl (cat. no. 3627-01, J.T. Baker) at 127 mM, KCl (cat. no. 3046-01, J.T. Baker) at 2.7 mM, Na2HPO4 (cat. no. 3827-01, J.T. Baker) at 6.5 mM and KH2PO4 (cat. no. 3248-01, J.T. Baker) at 1.5 mM in apyrogenic ultrapure water. Verify pH is between 7.35 and 7.45. Yields a 10 stock solution. Store at room temperature. 2. PBS—Penicillin G/Gentamicin/Fungizone (PBS-P/G/F). Dilute 10 PBS (see Subheading 2.1.5, item 1) to 1 with apyrogenic ultrapure water. Sterilize by filtration through a 0.22 μm low binding disposable filter (see Subheading 2.1.7, item 1). Add Penicillin G/Gentamicin and Fungizone (dilute to 1; see Subheading 2.1.3, items 4 and 6). Store at 4
C. 3. HEPES/KCl/NaCl. Dissolve 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) at 0.1 M (cat. no. 7745-14, Malinckrodt Chemicals), KCl (cat. no. 3046-01, J.T. Baker) at 67 mM, and NaCl (cat. no. 3627-01, J.T. Baker) at 1.42 M in apyrogenic ultrapure water. Adjust pH to 7.3. Yields a 10 stock solution. Store in the dark at 4
C (see Subheading 4; Note 4). 4. HEPES/KCl/NaCl—CaCl2. Dilute 10 HEPES/KCl/NaCl to 1 (see Subheading 2.1.5, item 3) with apyrogenic ultrapure water. Add CaCl2 to 1 mM (cat. no. 1336-01, J.T. Baker). Adjust pH to 7.45. Store in the dark at 4
C. 5. Thermolysin (cat. no. T7902, Sigma-Aldrich). Dissolve at 500 μg/mL in HEPES/KCl/NaCl—CaCl2 (see Subheading 2.1.5, item 4). Sterilize by filtration through a 0.22 μm low binding disposable filter (see Subheading 2.1.7, item 1). Store at 4
C and use within the day. 6. Dispase II (cat. no. 4942078001, Sigma-Aldrich). Dissolve at 2.5 mg/mL in HEPES/KCl/NaCl—CaCl2 (see Subheading 2.1.5, item 4). Adjust pH to 7.4. Sterilize by filtration through a 0.22 μm low binding disposable filter (see Subheading 2.1.7, item 1). Store at 4
C and use within the hour. 7. Trypsin/EDTA. Dissolve trypsin 1-250 (cat. no. 27250-018, Gibco) at 0.05% w/v, EDTA (cat. no. 8995-01, J.T. Baker) at 0.01% w/v and D-glucose (cat. no. 1920-05, J.T. Baker) at 2.8 mM in 1 PBS (see Subheading 2.1.5, item 1). Add 100,000 IU/L of penicillin G (Novopharm), 25 mg/L active

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gentamicin (cat. no. GE152, Galenova), and 0.00075% (v/v) pre-sterile filtered 0.1% phenol red-water solution (cat. no. T254-03, J.T. Baker). Adjust pH to 7.45. Sterilize by filtration through a 0.22 μm low binding disposable filter (see Subheading 2.1.7, item 1). To avoid repeated thawing and freezing cycles, distribute in single use aliquots. Store at 20 to 80
C. 2.1.6 Tissues and Cells

1. Normal human keratinocytes (nhK). Adult normal human keratinocytes (anhK) or newborn normal human keratinocytes (nnhK) are isolated from surgically removed, 3–6 cm skin and foreskin samples, respectively. 2. Normal human corneal epithelial cells (nhCECs) are isolated from the corneas of human postmortem donors unsuitable for transplantation (Banque nationale d’yeux du CHU de Que´bec, Quebec, Canada). 3. Irradiated human fibroblast layer (iHFL). Fibroblasts extracted from normal skin samples are seeded at 6000 cells/cm2 in a 75 cm2 culture flask (Tissue culture flask, cat. no. 35310, BD Falcon) in 15 mL of ckDME-Ham or ccDME-Ham (see Subheading 2.1.4, items 2 and 3). Incubate in 8% CO2, 100% humidity atmosphere at 37
C. After 4 days (80% confluence), irradiate at 6000 rads with a Gammacell irradiator (60Co source) (see Subheading 4; Note 5). Then, freeze the cells (see Subheading 3.1.6) or seeded at 6000 cells/cm2 in a 75 cm2 culture flask in 15 mL of ckDME-Ham or ccDME-Ham (see Subheading 2.1.4, items 2 and 3). Medium must be changed every 7 days. iHFL cultures are incubated in an 8% CO2 and 100% humidity atmosphere at 37
C. iHLF cultures must be used within 30 days.

2.1.7 Labware

1. For volumes inferior to 100 mL: 0.22 μm low binding disposable filter (cat. no. SLGV033RS, EMD Millipore). For volumes superior to 100 mL: filtration unit mounted with a 47 mm diameter and 0.22 μm filter set (cat. no. 28199-324, VWR, and cat. no. GVWP04700, EMD Millipore, respectively). 2. Sterile containers (cat. no. NCS902-10, Starplex). 3. 15 and 50 mL tubes (cat. no. 352095 and cat. no. 352070, Corning). 4. 35  40 mm and 100  15 mm cell culture Petri dishes (cat. no. 321008, and cat. no. 353003, Corning). 5. Dissecting curved forceps (cat. no. 08-953F, Fisher Scientific). 6. Size 4 scalpel (cat. no. 08-917-5, Fisher), size 22 blade (cat. no. SMI 1/73-0422, Southmedic, respectively). 7. Trypsination unit, Celstir® 50 mL suspension culture flask (cat. no. 356875, Wheaten).

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8. Parafilm® M (cat. no. 13-374-16, Fisher). 9. 25 or 75 cm2 tissue culture flasks (cat. no. 353109, Corning and cat. no. 353110, Corning, respectively). 10. Sterile cryogenic vials (cat. no. 5000-0020, Nalgene). 11. Freezing container (cat. no. 15-350-50, Nalgene). 12. Sterile 7 cm  7 cm gauze (cat. no. A3120, AMD-Ritmed). 13. Dissecting curved scissor (cat. no. E3220, Storz). 14. 8 mm diameter trephine (cat. no. ACKP825, Medline). 15. Dissecting stereomicroscope (cat. no. 495015-0021-000, Zeizz). 2.2 Electrophoretic Mobility Shift Assay 2.2.1 Reagents

1. [γ32P] dATP (cat. no. NEG502Z250UC, Perkin Elmer) (see Subheading 4; Note 6). 2. 50 ng of a synthetic double-stranded oligonucleotide (refer to Table 1) bearing the high affinity binding site for the transcription factor Sp1 (see Subheading 4; Note 7). 3. 50 ng of a synthetic double-stranded oligonucleotide (refer to Table 1) bearing the high affinity binding site for the transcription factor NFI (see Subheading 4; Note 7). 4. T4 polynucleotide kinase (PNK) and 10 kinase buffer (cat. no. M0201S and cat. no. B0201S, BioLabs). 5. Siliconized glass wool. Glass wool (cat. no. 3950 9989, Corning) can be siliconized by soaking them in a 5% solution of Sigmacote (cat. no. SL2-100ML, Sigma Aldrich) in heptan (cat. no. 246654, Sigma-Aldrich). As the organic solvent evaporates, the Sigmacote is deposited on the glass wool fibers and baked for 2 h at 180
C before use. Store at room temperature. 6. Extract (crude or enriched) containing cell or tissue nuclear proteins (see Subheading 3.2.1) (see Subheading 4; Note 8). 7. BCA Protein Assay Kit (cat. no. 23225, Thermo Fisher Scientific) (see Subheading 4; Note 9).

Table 1 Sequence of oligonucleotides bearing the high affinity binding sites for the transcription factors Sp1 and NFI

Oligonucleotide

Top strand (50 –30 ) Bottom strand (50 –30 )

Sp1

GATCATATCTGCGGGGCGGGGCAGACACAG GATCCTGTGTCTGCCCCGCCCCGCAGATAT

NFI

TTATTTTGGATTGAAGCCAATATGAG CTCATATTGGCTTCAATCCAAAATAA

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2.2.2 Solutions

1. Three nuclear extract (NE) solutions. For NE1 dissolve NaCl at 0.14 M and Tris–HCl pH 7.5 (cat.no. 600-126-CG, Wisent Bioproducts) at 10 mM in apyrogenic ultrapure water. For NE2 dissolve KCl at 25 mM, Mg(CH3COO)2 (cat. no. M5661-50G, Sigma-Aldrich) at 2 mM, dithiothreitol (DTT) at 1 mM (cat. no. 10708984001, Sigma-Aldrich), and Tris–HCl pH 7.5 at 10 mM in apyrogenic ultrapure water. For NE3 dissolve KCl at 0.35 mM, Mg(CH3COO)2 at 2 mM, Tris–HCl pH 7.5 at 10 mM, and DTT at 1 mM in apyrogenic ultrapure water. For all three nuclear extract solutions, filter through a 0.22 μm low binding disposable filter (see Subheading 2.1.7, item 1) and store at 4
C. 2. A solution of protease inhibitor cocktail (cat. no. 11836153001, Sigma-Aldrich). Dissolve one tablet in 1 mL of apyrogenic ultrapure water to obtain a 25 solution. Store at 20
C (see Subheading 4; Note 10). 3. A dialysis buffer: Dissolve KCl at 50 mM, MgCl2 (cat.no. M33-500, Fisher Scientific) at 1 mM, K3PO4 pH 7.4 (cat. no. PP-0227-500G, Laboratoire MAT) at 20 mM K3PO4, β-mercaptoethanol (cat. no. 190242, ICN Biochemical) at 1 mM, 20% (v/v) glycerol (cat. no. 5350-1-40, Caledon Laboratory Chemical), 1 mM DTT and phenylmethylsulfonyl fluoride (PMSF) at 0.5 mM (cat. no. P7626-5G, Sigma-Aldrich) in apyrogenic ultrapure water. Store at 4
C and add DTT and PMSF just before use (see Subheading 4; Note 11). 4. TE pH 8. Dissolve Tris–HCl pH 8 at 10 mM and ethylenediaminetetraacetic acid (EDTA) pH 8 at 1 mM (cat. no. BD9232, VWR) in apyrogenic ultrapure water. Store at room temperature. 5. STE solution. Dissolve Tris–HCl pH 8 at 10 mM, ethylenediaminetetraacetic acid (EDTA) pH 8 at 1 mM, and NaCl at 100 mM in apyrogenic ultrapure water. Store at room temperature. 6. Sephadex G-50 Solution. Slowly add 30 g of Sephadex G-50 (cat. no. 17-0060-01, Sigma-Aldrich) to 250 mL of TE pH 8 (see Subheading 2.2.2, item 4). Make sure the powder is well dispersed. Incubate overnight at room temperature. Decant the supernatant and replace with an equal volume of TE pH 8. Store at 4
C in a screw-capped bottle. 7. A stock solution of 40% (w/v) acrylamide prepared in a 39:1 (w/w) ratio of acrylamide and N0 ,N0 -methylene bis-acrylamide (cat. no. AB1032 and BB0025, Bio Basic) in apyrogenic ultrapure water. Store in the dark at room temperature (see Subheading 4; Note 12). 8. Ammonium persulfate (APS) 10% (cat. no. A3678-25G, Sigma-Aldrich) in apyrogenic ultrapure water. Store to 4
C and use within 6 months.

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9. Tetramethylethylenediamine (TEMED) (cat. no. 161-0800, Bio-Rad). 10. A stock solution of 5 Tris-glycine. Dissolve Tris base (cat. no. BP152-5, Fisher Scientific) at 250 mM, EDTA at 12.5 mM and glycine (cat. no. GB0235, BioBasic Canada) at 2 M in apyrogenic ultrapure water. Store in the dark at room temperature (see Subheading 4; Note 13). 11. A stock solution of 2 binding buffer. Dissolve HEPES pH 7.9 at 20 mM, glycerol at 20%, EDTA at 0.2 mM EDTA, and PMSF at 0.5 mM in apyrogenic ultrapure water. Distribute in single use aliquots (250 μL) and store at 20
C. 12. A stock solution of 6 loading buffer. Dissolve bromophenol blue (cat. no. 11439-5G, Sigma-Aldrich) at 0.25%, xylene cyanol (cat. no. BP565-10, Fisher Scientific) at 0.25%, and sucrose (cat. no. SU-1000-SKG, Laboratoire MAT) at 40% in apyrogenic ultrapure water. Store at room temperature. 13. A stock solution of 1 μg/μL poly(deoxyinosinicdeoxycytidylic) acid sodium salt (poly(dI:dC)) (cat. no. P4929-10UN, Sigma-Aldrich). Distribute in single use aliquots (50 μL) and store at 20
C. 2.2.3 Labware

1. Homogenizer potter, size: 5 mL (cat. no. 02-911-525, Fisher Scientific). 2. Beckman Optima TLX, Optima TL, or TL-100 Ultracentrifuge with rotor TLA-120.2 (10  2.0 mL, 8  34 mm). 3. Dialysis molecular porous membrane. Dialysis Tubing, 3.5 K MWCO, is composed of regenerated-cellulose dialysis tubing and supplied in 8 in. (20 cm) sticks 16 mm circular internal diameter (I.D.). The hydrated Dialysis Tubing holds 2–10 mL of sample per centimeter of length (cat. no. 88242, Thermo Fisher Scientific). 4. Ependorff tubes (cat. no. T9661-1000EA, Sigma-Aldrich). 5. 15 mL tubes (cat. no. 352095, Corning). 6. Syringes 1 mL (cat. no. 14-817-25, Fisher Scientific). 7. Aqueous counting scintillant (cat. no. 88247501, VWR). 8. Scintillation vials (cat. no. 73.662.500, Sarstedt). 9. Scintillation beta counter (LS 6000SC, Beckman). 10. Standard vertical electrophoresis apparatus for polyacrylamide gels, a gel length of 15 cm is adequate (see Subheading 4; Note 14). 11. Whatman chromatographic paper (cat. no. 05-714-4, Fisher Scientific) and plastic wrap.

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12. Autoradiography cassettes (electrophoresis systems, FBXC 810, Fisher Scientific). 13. Scientific imaging films (cat. no. 28906839, GE Healthcare). 14. Spatula (cat. no. S50789A, Fisher Scientific). 15. Gel dryer (cat. no. 1651745, Bio Rad). 2.3 Western Blot Analyses 2.3.1 Reagents

1. Nuclear extract (see Subheading 3.2.1). 2. Prestained molecular weight markers (cat. no. 26612, Thermo Fisher Scientific). 3. Polyvinylidene fluoride (PVDF) membrane 0.45 μm (cat. no. 162-0264, Bio-Rad) and Whatman chromatographic paper (cat. no. 05-714-4, Fisher Scientific). 4. Anti-Sp1 rabbit polyclonal IgG Sp1 (clone PEP2) (cat. no. Sc-59, Santa Cruz Biotechnology) at 1/250. 5. Anti-NFI rabbit polyclonal IgG NFI (H-300X) no. Sc-5567, Santa Cruz Biotechnology) at 1/2000.

(cat.

6. Peroxidase-conjugated AffiniPure goat Anti-Rabbit IgG (H +L) (cat. no. 111-036-003, Jackson ImmunoResearch Laboratories Inc.) at 1/2500. 7. ECL Plus Western blotting detection system (cat. no. 80196, Thermo Fisher Scientific). 2.3.2 Solutions

1. 2 SDS PAGE sample buffer. Dissolve Tris–HCl pH 6.8 at 0.5 M, DTT at 400 mM, SDS (sodium dodecyl sulfate) (cat. no. SE-0099-500G, Laboratoire MAT) at 8%, bromophenol blue at 0.4%, and glycerol at 40% in apyrogenic ultrapure water. Store at room temperature. 2. A stock solution of 30% (w/v) acrylamide prepared in a 29:1 (w/w) ratio of acrylamide and N0 ,N0 -methylene bis-acrylamide in apyrogenic ultrapure water. Store in the dark at room temperature (see Subheading 4; Note 12). 3. Running gel buffer. Dissolve Tris-base at 1.5 M in apyrogenic ultrapure water and adjust pH to 8.8. Store at 4
C. 4. Stacking gel buffer. Dissolve Tris–base at 0.5 M in apyrogenic ultrapure water and adjust pH to 6.8. Store at 4
C. 5. Ammonium persulfate (APS) 10% (cat. no. A3678-25G, Sigma-Aldrich) in apyrogenic ultrapure water. Store at 4
C and use within 6 months. 6. 10 SDS PAGE running buffer. For 1 L, dissolve 10 g SDS, 30.3 g Tris-base, and 144.1 g glycine in 800 mL of apyrogenic ultrapure water. Adjust volume to 1 L and store at room temperature.

Quality of Primary Cultured Epithelial Stem Cells

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7. 10 transfer buffer. For 1 L, dissolve 30.3 g Tris-base and 140.6 g glycine in 800 mL of apyrogenic ultrapure water. Adjust volume to 1 L and store at room temperature. 8. Methanol (cat. no. A412P-4, Fisher Scientific). 9. Tetramethylethylenediamine (TEMED; cat. no. 161-0800, Bio-Rad). 10. 10% SDS (cat. no. SE-0099-500G, Laboratoire MAT) solution in apyrogenic ultrapure water. Store at room temperature. 11. Butanol (cat. no. A399S-4, Fisher Scientific). 12. 10 Tris Buffered Saline (TBS). Dissolve Tris-base at 0.1 M and NaCl at 1.5 M in apyrogenic ultrapure water. Adjust pH to 8 and store at room temperature. 13. 1 Tris Buffered Saline Tween (TBS-T). Dilute (1:10) 10 TBS (see Subheading 2.3.2, item 12) with apyrogenic ultrapure water and add Tween 20 (cat. no. P1379-1L, SigmaAldrich) at 0.1% (v/v). Store at room temperature. 14. Blocking solution. Dissolve milk powder at 5% (w/v) in 1 TBS-T (see Subheading 2.3.2, item 13). 2.3.3 Labware

1. Mini-protean cell with no. 1658033, Bio-Rad).

mini

Trans-Blot

module

(cat.

2. Autoradiography cassettes (electrophoresis systems, FBXC 810, Fisher Scientific). 3. Scientific imaging films (cat. no. 28906839, GE Healthcare). 4. Acetate sheets (cat. no. S25218A, Fisher Scientific).

3 3.1

Methods Cell Culture

3.1.1 Tissue Sampling and Transport

Ethical approval and informed consent must be obtained for each human tissue. 1. Immediately following aseptic (see Subheading 4; Note 15) surgical removal, put the skin sample (see Subheading 2.1.6, item 1) into a sterile container (see Subheading 2.1.7, item 2) filled with cold (4
C) tDMEM (see Subheading 2.1.4, item 1). 2. Shortly after donor’s death, eyes or only the corneas (see Subheading 2.1.6, item 2) are extracted and placed in a sterile container (see Subheading 2.1.7, item 2) by qualified staff. 3. Samples must be kept on ice and transported to a cell culture facility without delay. Manipulations must be performed under a sterile laminar flow hood cabinet.

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3.1.2 Isolation of Normal Human Skin Keratinocytes

1. Wash the skin sample (see Subheadings 2.1.6, item 1 and 3.1) by transferring it into a 50 mL tube (see Subheading 2.1.7, item 3) containing 30 mL PBS-P/G/F (see Subheading 2.1.5, item 2) and agitating vigorously. With sterile forceps, transfer the sample into another 50 mL tube containing 30–45 mL PBS-P/ G/F. Repeat eight times for a total of ten washes. 2. Use curved forceps (see Subheading 2.1.7, item 5) to spread out the skin sample (epidermis facing top) in a 100 mm Petri dish (see Subheading 2.1.7, item 4). 3. Use a scalpel with a size 22 blade (see Subheading 2.1.7, item 6) to cut the sample into as many 3 mm  100 mm strips as necessary. 4. Add 10 mL of cold thermolysin (see Subheading 2.1.5, item 5) into the Petri dish containing the small skin strips (see Subheading 4; Note 16). 5. Seal the Petri dish with Parafilm® M (see Subheading 2.1.7, item 8) and incubate overnight at 4
C. 6. Use two curved forceps to carefully separate the epidermis from the dermis (delicately pull hair if present to collect follicle keratinocytes). To dissociate nhKs, put the epidermal strips (and hair) in a Celstir® suspension culture flask (see Subheading 2.1.7, item 7) containing 20 mL warm (37
C) trypsin/EDTA (see Subheading 2.1.5, item 7). 7. Incubate under agitation for 15–30 min at 37
C. 8. Add 10 mL of ckDME-Ham into the flask. Transfer the 20 mL nhK suspension into a 50 mL tube containing 10 mL of ckDME-Ham. Rinse the culture suspension flask with 10 mL ckDME-Ham (see Subheading 2.1.4, item 2) and add it to the 50 mL tube. 9. Use an automated cell counter or a hematimeter to count nhKs. Count is expected to be approximately 3–4  106 nhK/cm2 skin sample and cell size should be between 9 and 15 μm. 10. Use trypan blue staining and a hematimeter to estimate cell viability. Cell viability is expected to be greater than 80%. 11. Centrifuge (300  g) the nhK suspension for 10 min at room temperature. 12. Remove the supernatant and resuspend nhKs at the desired concentration in ckDME-Ham. 13. Seed nhKs at no less than 100,000/25 cm2 (see Subheading 4; Note 17) into a culture flask (see Subheading 2.1.7, item 9) pre-seeded with iHFL (see Subheading 2.1.6, item 3). Seed directly into the culture medium. Total medium volume should not exceed 7 mL/25 cm2. If iHFL medium change

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was scheduled the same day nhK are seeded (7 days since last iHFL medium change), replace half the iHFL conditioned medium with warm (37
C) ckDME-Ham. 14. When the nhKs reach 80% confluence, subculture (see Subheading 3.1.5) or freeze (see Subheading 3.1.6) cells. 3.1.3 Isolation of Normal Human Corneal Epithelial Cells

1. Human corneal epithelial cells are obtained from cornea of postmortem human donors that are unsuitable for transplantation. The eye sample is brought to the Banque nationale d’yeux du CHU de Que´bec to be evaluated (see Subheading 4; Note 18). 2. Wash the eye specimen in a 50 mL centrifuge tube (see Subheading 2.1.7, item 3) containing 30 mL PBS-P/G/F (see Subheading 2.1.5, item 2). Agitate gently 1–2 min. With sterile curved forceps (see Subheading 2.1.7, item 5), transfer the eye specimen in another tube filled with PBS-P/G/F. Repeat this step three times. 3. Place the eye specimen into a 100 mm Petri dish (see Subheading 2.1.7, item 4). 4. Surround the eye specimen with a folded sterile gauze. This helps in holding the eye without having to touch it. 5. With the size 22 scalpel blade (see Subheading 2.1.7, item 6), make a small opening of 2–3 mm in the sclera. 6. With curved scissors (see Subheading 2.1.7, item 13), cut-out the cornea to obtain only the limbus and the central cornea. Avoid leaving the sclera to eliminate conjunctival epithelial cell contamination. 7. With two curved forceps, peel-off the iris. Do this step while holding the cornea in the air to avoid any damage to the epithelium during the procedure. 8. Place the limbal ring into a 35 mm tissue culture Petri dish (see Subheading 2.1.7, item 4). The limbal ring is obtained by separating the limbus from the central cornea with an 8 mm diameter trephine (see Subheading 2.1.7, item 14). Position the epithelium upward and add 5 mL of cold (4
C) dispase II (see Subheading 2.1.5, item 6). Seal the Petri dish with parafilm (see Subheading 2.1.7, item 8). 9. Incubate overnight at 4
C. 10. With two curved forceps, mechanically separate the epithelium from the stroma under a dissecting microscope (see Subheading 2.1.5, item 15). To dissociate nhCEC, put the epithelium in a Celstir® suspension culture flask (see Subheading 2.1.7, item 7) containing 10 mL warm (37
C) trypsin/EDTA (see Subheading 2.1.5, item 7).

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11. Incubate under agitation for 10 min at 37
C. 12. Add 10 mL of ccDME-Ham (see Subheading 2.1.4, item 3) into the flask. Transfer the 20 mL nhCEC suspension into a 50 mL tube containing 10 mL of ccDME-Ham. Rinse the culture suspension flask with 10 mL ccDME-Ham (see Subheading 2.1.4, item 3) and add it to the 50 mL tube. 13. Centrifuge (300  g) the nhCEC suspension for 10 min at room temperature. 14. Remove the supernatant and resuspend nhCECs at the desired concentration in 5 mL of warm (37
C) ccDME-Ham. 15. Seed nhCECs into a culture flask (see Subheading 2.1.7, item 9) pre-seeded with iHFL (see Subheading 2.1.6, item 3). Seed directly into the culture medium. Total medium volume should not exceed 7 mL/25 cm2. If iHFL medium change was scheduled the same day nhCEC are seeded (7 days since last iHFL medium change), replace half the iHFL conditioned medium with warm (37
C) ccDME-Ham (see Subheading 4; Note 17). 16. When the nhCECs reach 80% confluence, subculture (see Subheading 3.1.5) or freeze (see Subheading 3.1.6) cells. 3.1.4 Culture

1. Incubate nhK-iHFL or nhCEC-iHFL cocultures (see Subheadings 3.1.2, step 13 and 3.1.6, step 10) in an 8% CO2 and 100% humidity atmosphere at 37
C. 2. Change the culture medium three times a week, every 2–3 days. Remove the medium from the culture flask. Replace it with warm (37
C) ckDME-Ham or ccDME-Ham (see Subheading 2.1.4, items 2 and 3), depending on cell type. 3. Monitor cell confluence (see Subheading 4; Note 19) daily under a microscope. 4. When cells reach 75–95% confluence, subculture (see Subheading 3.1.5) or cryopreserve (see Subheading 3.1.6) them. Do not let cells reach 100% confluence.

3.1.5 Subculture (Passage)

1. Remove the culture medium (see Subheading 3.1.4). 2. Depending on culture flask size (see Subheading 2.1.7, item 9), swiftly rinse cells with either 1 or 2 mL (for either 25 or 75 cm2 culture flasks respectively) trypsin/EDTA (see Subheading 2.1.5, item 7). Remove it. 3. Depending on culture flask size (see Subheading 2.1.7, item 9), add either 3 or 8 mL (for either 25 or 75 cm2 culture flasks respectively) of trypsin/EDTA into the culture flask. 4. Incubate at 37
C until all cells are completely detached from the flask (verify cell detachment under a microscope). Time for

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complete detachment should be around 10 min. Do not incubate for more than 15 min (see Subheading 4; Note 20). 5. Depending on culture flask size, neutralize trypsin activity by adding either 3 or 8 mL of ckDME-Ham or ccDME-Ham (see Subheading 2.1.4, items 2 and 3) depending on cell type and swirl the cell suspension. 6. Vigorously pipette the cell suspension up and down at least ten times to ensure suspension homogeneity. 7. Transfer the cell suspension into a 50 mL tube (see Subheading 2.1.7, item 3). 8. Thoroughly rinse the culture flask with 4 mL ckDME-Ham or ccDME-Ham (see Subheading 2.1.4, items 2 and 3), depending on cell type. Transfer the suspension into the 50 mL tube. 9. Use an automated cell counter or a hematimeter to count cells. 10. Use trypan blue staining and a hematimeter to estimate cell viability. Cell viability is expected to be greater than 95%. 11. Centrifuge (300  g) the cell suspension for 10 min at room temperature. 12. Remove the supernatant and resuspend cells at the desired concentration in ckDME-Ham or ccDME-Ham (see Subheading 2.1.4, items 2 and 3), depending on cell type. 13. Seed cells at no less than 100,000/25 cm2 (see Subheading 4; Note 17) into a culture flask pre-seeded with iHFL (see Subheading 2.1.6, item 3). Seed directly into the culture medium. Total medium volume should not exceed 7 mL/25 cm2. If iHFL medium change was scheduled the same day cells are seeded (7 days since last iHFL medium change), replace half the iHFL conditioned medium with warm (37
C) ckDME-Ham or ccDME-Ham (see Subheading 2.1.4, items 2 and 3), depending on cellular type. 3.1.6 Cryopreservation

1. Fill a freezing container (see Subheading 2.1.7, item 11) with 100% isopropyl alcohol. Store at 4
C until cool. 2. Follow steps 1–11 from Subheading 3.1.5. 3. Remove the supernatant and resuspend cells at the desired concentration (max. 10  106/mL) in nhK or nhCEC cryopreservation medium (see Subheading 2.1.4, items 4 and 5), depending on cell type. Put the tube on ice. 4. Aliquot in cryogenic vials on ice (see Subheading 2.1.7, item 10). 5. Put the cryogenic vials in the freezing container (see Subheading 2.1.7, item 11). 6. Store the container overnight at 80
C. In these conditions, cell temperature should drop 1
C/min. 7. Store cryogenic vials in liquid nitrogen for long-term storage.

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3.1.7 Thawing

1. Put the cryogenic vial (see Subheading 3.1.6, step 7) in a 37
C water bath. Do not let the cell suspension thaw completely. A small ice pellet should remain. 2. Depending on cell type, add 0.5–1 mL of cold (4
C) ckDMEHam or ccDME-Ham (see Subheading 2.1.4, items 2 and 3) into the cryogenic vial. 3. As soon as the remaining ice has melted, transfer the content of the cryogenic vial into a 50 mL tube (see Subheading 2.1.7, item 3) containing 8–10 mL of cold (4
C) ckDME-Ham or ccDME-Ham (see Subheading 2.1.4, items 2 and 3), depending on cell type. 4. Centrifuge (300  g) the cell suspension for 10 min at room temperature. 5. Remove the supernatant and resuspend cells in 10 mL of warm (37
C) ckDME-Ham or ccDME-Ham (see Subheading 2.1.4, items 2 and 3), depending on cell type. 6. Use an automated cell counter or a hematimeter to count the cells. 7. Use trypan blue staining and a hematimeter to estimate cell viability. Cell viability is expected to be greater than 80%. 8. Centrifuge (300  g) the cell suspension for 10 min at room temperature. 9. Remove the supernatant and resuspend cells at the desired concentration in ckDME-Ham or ccDME-Ham, depending on cell type (see Subheading 4; Note 21). 10. Seed cells at no less than 100,000/25 cm2 (see Subheading 4; Notes 18 and 19) into a culture flask pre-seeded with iHFL (see Subheading 2.1.6, item 3). Seed directly into the culture medium. Total medium volume should not exceed 7 mL/25 cm2. If iHFL medium change was scheduled the same day cells are seeded (7 days since last iHFL medium change), replace half the iHFL conditioned medium with warm (37
C) ckDME-Ham or ccDME-Ham (see Subheading 2.1.4, items 2 and 3), depending on cell type.

3.2 Electrophoretic Mobility Shift Assay

Always wear gloves when preparing samples, reagents, and materials to prevent contamination with hands’ proteins.

3.2.1 Preparation of Crude Nuclear Extracts

1. Resuspend the cell pellet (10–50  106 of cells) in 5 mL NE1 in a 15 mL centrifuge tube (see Subheadings 2.2.2, item 1 and 2.2.3, item 5). Centrifuge at 900  g 5 min at 4
C; use a table centrifuge. 2. Remove the supernatant and resuspend the pelleted cells in 5 mL NE1 and centrifuge 5 min at 4
C.

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3. Remove the supernatant, resuspend cells in 5 mL NE2 (see Subheading 2.2.2, item 1), and centrifuge at 900  g 5 min at 4
C. 4. Remove the supernatant, resuspend cells in 2 mL NE2, and incubate 5 min on ice. 5. On ice, transfer the cellular suspension in the homogenizer potter (see Subheading 2.2.3, item 1) and push down and up the piston 20 times. 6. Centrifuge at 500  g 3 min at 4 supernatant.




C and remove the

7. Resuspend the pellet in 1 mL NE3 (see Subheading 2.2.2, item 1). Add 1/10 the cell pellet volume of KCL 4M and protease inhibitor cocktail (dilute to 1; see Subheading 2.2.2, item 2). 8. On ice, transfer the cell suspension in the homogenizer potter and push down and up the piston ten times. 9. With an ultracentrifuge, centrifuge in a 1.5 mL tube at maximal speed (16,000  g) 5 min at 4
C. The supernatant (nuclear extract) can be stored at 80
C before completion of the protocol. 10. If samples have been stored at 80
C, they must be thawed at 4
C. Transfer the supernatant in a conical tube suitable for the TL 100.2 rotor and centrifuge 2 h at 50,000 rpm (89,000  g) at 4
C (see Subheading 2.2.3, item 2). 11. Transfer the supernatant in a dialysis molecular porus membrane tubing (see Subheading 2.2.3, item 3), and close the tube with two universal closures prior to dialyzing 1 h against dialysis buffer (see Subheading 2.2.2, item 3) at 4
C. 12. Determine the total protein concentration using the BCA Protein Assay Kit (see Subheading 2.2.1, item 7). Evaluate by Coomassie blue staining the quality of the proteins from the crude nuclear extracts following their separation by gel electrophoresis (see example in Fig. 1a). 13. Distribute nuclear extract in 50–70 μL aliquot and store at 80
C. 3.2.2 Preparation of Purification Spun-Column

1. Plug the bottom of a 1 mL disposable syringe (see Subheading 2.2.3, item 6) with a small amount of siliconized glass wool (see Subheading 2.2.1, item 5). In the syringe, prepare a column (0.9-mL bed volume) of Sephadex G-50 solution (see Subheading 2.2.2, item 6). 2. Insert the spun-column into a 15 mL centrifuge tube (see Subheading 2.2.3, item 5) as shown in Fig. 5. Centrifuge at 1600  g for 75 s to pack down the Sephadex. Repeat until

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Fig. 1 Binding and expression of the transcription factor Sp1 by EMSA and Western blot. Crude nuclear extracts were prepared from nnhK grown at passage 3 (P3) to passage 12 (P12) with i3T3. (a) 10 μg nuclear proteins from each cell passage were separated by gel electrophoresis (SDS-PAGE) and then stained with Coomassie blue in order to ensure that no protein degradation was occurring from one sample to another. (b) A synthetic double-stranded oligonucleotide bearing the high-affinity binding site for the transcription factor Sp1 was 50 -end labeled and incubated with or without (P) crude nuclear extracts prepared from nnhK at each cell passage (P3–P12). The EMSA was conducted at 4
C on a 6% native polyacrylamide gel for 5 h at 120 V. The position corresponding to Sp1 and Sp3, as well as that of the free probe (U) is indicated. (c) Expression of Sp1 was monitored on the crude nuclear extracts (10 μg from each) used in panel (b) by Western blot using the Sp1 antibody. The results presented in this figure indicate clearly that Sp1 is expressed in human skin keratinocytes between cell passages, reaching high levels between passages P2–P4 and P9–P10

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the packed Sephadex reaches a column volume of approximately 0.9 mL. 3. Add 100 μL of STE (see Subheading 2.2.2, item 5) and centrifuge at 1600  g for 75 s to wash the column. Discard the flowthrough and repeat this step (see Subheading 4; Note 22). 4. Place the spun-column in a new 15 mL centrifuge tube containing an Eppendorf tube without a cap (see Subheading 2.2.3, item 4) in order to collect the labeled DNA (see Fig. 5). 5. The spun-column is ready for use. Do not store, use immediately. 3.2.3 Labeling and Purification of a DoubleStranded Sp1 and NFI Oligonucleotide

1. Anneal equal amounts of the complementary Sp1or NFI synthetic oligonucleotides (see Subheading 2.2.1, items 2 and 3; Table 1) (Example: for 1 μg/μL stock: 25 ng of 1 μg/μL of each strands in 50 μL total volume). Heat the resulting mix at 4
C over the specific melting temperature (TM) of the oligonucleotides (alternatively, you may heat the oligonucleotide mix at 90
C) for 90 s and let cool at room temperature. When double-stranded oligonucleotide reaches room temperature, store at 20
C prior to use. 2. Use 50 ng (2 μL of 25 ng/μL solution) of the Sp1 or NFI double-stranded oligonucleotide prepared (see Subheading 3.2.3, step 1) and label DNA with 5 μL T4 PNK buffer (see Subheading 2.2.1, item 4), 37 μL of apyrogenic ultrapure water, 1 μL of T4 PNK (see Subheading 2.2.1, item 4), and 5 μL of 10 μCi/mL [γ32P] dATP (see Subheading 2.2.1, item 1). Incubate for 45 min at 37
C and add 50 μL STE (see Subheading 2.2.2, item 5) to obtain a total volume of 100 μL. 3. Add the 100 μL Sp1 or NFI labeled probe in spun-column (see Subheading 3.2.2) and centrifuge at 1600  g for 75 s. Spuncolumn centrifugation separates the labeled probe (approximately 100 μL recovered in the Eppendorf tube) from the unincorporated [32P]dNTPs which remain in the column (Fig. 5). 4. Pour 5 mL of aqueous counting scintillant (see Subheading 2.2.3, item 7) in scintillation vials (see Subheading 2.2.3, item 8), add 1 μL of the labeled Sp1 or NFI probe, and count (number of decays detected by minute) using a scintillation beta counter (see Subheading 2.2.3, item 9) by Cerenkov counting. 5. Store labeled probe at 20
C in lead vial (see Subheading 4; Note 23).

3.2.4 Sp1-DNA and NFI-DNA Interaction in EMSA

1. Rigorously clean and dry the electrophoresis apparatus (see Subheading 2.2.3, item 10) and its accessories prior to use. Gel plates should be cleaned using any good quality

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commercial soap and then rinsed with 95% ethanol. One plate can be treated with a coat of Sigmacote (chlorinated organopolysiloxane in heptane) to facilitate gel removal from after running (see Subheading 2.2.1, item 5). 2. Prepare an 8% polyacrylamide gel as follows; mix 10 mL of 5 Tris-Glycine (see Subheading 2.2.2, item 10), 10 mL of 40% acrylamide (39:1) stock solution (see Subheading 2.2.2, item 7), and 30 mL of apyrogenic ultrapure water to obtain 50 mL. Add 360 μL of 10% APS (see Subheading 2.2.2, item 8) and 50 μL of TEMED (see Subheading 2.2.2, item 9). Carefully stir and pour the acrylamide solution between the plates. Place a 14-well comb and allow the gel to set for 30 min, then mount the gel in the electrophoresis tank and fill the chamber with 1 Tris-Glycine buffer (dilute 5 Tris-Glycine in apyrogenic ultrapure water). 3. Pre-run the gel at 120 V at 4
C until the current becomes stable (this usually takes 30 min on average). This pre-running step ensures that the gel will be at a constant temperature at the moment of sample loading. 4. When the gel is ready for loading, wash all wells and prepare samples as follows: for each sample, mix 12 μL of 2 binding buffer (see Subheading 2.2.2, item 11), 0.5 μL of 1 μg/μL (see Subheading 4; Note 24) poly(dI:dC) (see Subheading 2.2.2, item 13), and 0.6 μL of 2 M KCl (see Subheading 4; Note 25). Then add 100,000 cpm of labeled probe (see Subheading 3.2.3). When possible, pool together invariant components in one microcentrifuge tube and then redistribute equal amounts into different tubes accounting for the different experimental conditions. Finally, add 10 μg nuclear proteins (see Subheading 3.2.1) and apyrogenic ultrapure water to a final volume of 24 μL. Mix gently each tube and incubate at room temperature for 3 min. As a control, prepare a sample without nuclear extract and add 2 μL of 6 loading buffer (see Subheading 2.2.2, item 12). 5. Load samples by changing the pipet tip for each sample and run the gel at 120 V for 5h30 at 4
C (see Subheading 4; Note 26). 6. After the gel has run, disassemble the apparatus with a spatula (see Subheading 2.2.3, item 14) and remove one of the glass plates. Place a Whatman paper (see Subheading 2.2.3, item 11) over the gel and carefully lift the gel off the remaining plate. Make sure that the gel is well fixed on the Whatman paper before lifting the gel to avoid gel tearing. Place a plastic wrap (see Subheading 2.2.3, item 11) over the gel and dry at 80
C for 60 min (see Subheading 2.2.3, item 15).

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Fig. 2 Analysis of Sp1 binding in human corneal epithelial cells by EMSA analyses. A synthetic doublestranded oligonucleotide bearing the high-affinity binding site for Sp1 was 50 -end labeled and incubated with 5 μg crude nuclear proteins prepared from four distinct primary cultured newborn (nnhCEC) or adult (anhCEC) normal human corneal epithelial cells grown with i3T3. (a) nnhCEC6 grown from P3 to P20, (b) anhCEC17 grown from P3 to P10, (c) anhCEC61 grown from P3 to P8, and (d) anhCEC37 grown from P3 to P8. Formation of DNA–protein complexes was evaluated on a native 6% polyacrylamide gel. (e) The Sp1 probe was incubated either alone (P) or with 5 μg nuclear extract from nnhCEC6 grown at passage P3 (C). To validate the specificity for the formation of the Sp1/Sp3 DNA–protein complexes, competition analyses in EMSA were performed. Approximately 5 μg nuclear proteins from nnhCEC6 cells at passage P3 were incubated along with a 250-fold molar excess of an unlabeled double-stranded oligonucleotide bearing the high affinity binding site for Sp1 (250 Sp1) or in the presence of polyclonal antibodies directed against Sp1 and Sp3 [added individually (Sp1 Ab or Sp3 Ab) or in combination (Sp1/3 Abs)]. Formation of the Sp1 and Sp3 (Sp1/Sp3) complexes, as well as that of their corresponding supershifted complexes (SSC), was then monitored by EMSA on a native 6% polyacrylamide gel. U: free probe. The results presented in this figure indicate that as with nhKs, DNA binding of Sp1 varies greatly between cell passages in nhCECs. Furthermore, the cell cultures that sustain the highest and more prolonged expression/DNA binding of Sp1 from one passage to another (such as with nnhCEC6 cells) are also those that can sustain a higher number of cell passages in culture

7. Place an X-ray film (see Subheading 2.2.3, item 13) over the dried gel in an autoradiography cassette (see Subheading 2.2.3, item 12) and expose at 80
C overnight or less, depending on the signal intensity (see examples on Figs. 1b, 2, 3a, and 4).

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Fig. 3 Analysis of NFI DNA binding and expression by EMSA and Western blot. Crude nuclear extracts were prepared from anhCEC grown to 80% confluence with iHFL, with i3T3 or without feeder layer (). The cells were grown until seven passages. (a) A synthetic double-stranded oligonucleotide bearing the high-affinity binding site for the transcription factor NFI was 50 -end labeled and incubated with or without (P) crude nuclear extracts prepared from the anhCEC. The EMSA was conducted at 4
C on an 8% native polyacrylamide gel for 5 h 30 at 120 V. The position corresponding to NFI as well as that of the free probe (U) is indicated. (b) Expression of NFI was monitored in the crude nuclear extracts (10 μg from each) used in panel (a) by Western blot using the NFI antibody. The results presented in this figure show DNA binding and expression of NFI in anhCEC cultured on several passages with i3T3 and iHFL 3.3 Detection of the Sp1 and NFI Protein in Western Blot

Always wear gloves when preparing samples, reagents, and materials to prevent contamination with hands’ proteins.

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Fig. 4 Analysis of NFI binding in human cornea epithelial cells by EMSA analyses. Crude nuclear extracts were prepared from anhCEC grown to 80% confluence with an iHFL feeder layer. A synthetic double-stranded oligonucleotide bearing the highaffinity binding site for NFI was 50 -end labeled and incubated of crude nuclear extracts are prepared from anhCEC cultured up to passage P4. Formation of DNA–protein complexes was evaluated on a native 8% polyacrylamide gel. The NFI probe was incubated with 10 μg nuclear extract from P4 anhCEC (C) in the presence of a 25 or 250-fold molar excess of an unlabeled oligonucleotide bearing the high affinity binding site for NFI or Sp1. The same nuclear extract was incubated in the presence of polyclonal antibodies directed against either NFI (Ab NFI). Formation of the NFI complexes, as well as that of their corresponding supershifted complexes (SSC), was then monitored by EMSA. U: free probe. The results presented in this figure indicate that a single, diffuse DNA–protein complex typical of NFI binding to DNA was observed upon incubation of nuclear proteins from anhCEC with the NFI labeled probe. The binding specificity was further demonstrated by the disappearance of the NFI complex and the formation of a new supershifted complex (SSC) of low electrophoretic mobility upon addition of an antibody that recognizes all NFI isoforms 3.3.1 Sample Preparation

1. Typically use 10 μg of nuclear proteins (see Subheading 3.2.1) per lane. If necessary, dilute sample in apyrogenic ultrapure water to get the same volume for each sample (see Subheading 4; Note 27). For each sample, add the desired amount of proteins to a micro-centrifuge tube followed by 2 SDS PAGE sample buffer (see Subheading 2.3.2, item 1). 2. Incubate at 95–100
C for 3 min.

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3. Quick spin the samples. 4. Load immediately on the previously prepared gel (see Subheading 3.3.2). 3.3.2 Electrophoresis (Stacking and Running Gels) and Transfer

1. Place a short plate on top of the 1.5 mm spacer plate. 2. Slide the two plates into the casting frame, keeping the short plate facing front and make sure both plates are flush at the bottom on a level surface. 3. Lock the pressure cams to secure the glass plates. 4. Engage the spring-loaded lever and place the gel cassette assembly on the casting strand gasket. Make sure the horizontal ribs on the back of the casting frame are flush against the face of the casting stand and the glass plates are perpendicular to the level surface. 5. Prepare an 8% running gel as follows; mix 5 mL of running gel buffer (see Subheading 2.3.2, item 3), 5.3 mL of 30% acrylamide (39:1) stock solution (see Subheading 2.3.2, item 2), 0.2 mL of 10% SDS solution (see Subheading 2.3.2, item 10), and 9.3 mL of apyrogenic ultrapure water. Add 200 μL of 10% APS (see Subheading 2.3.2, item 5) and 20 μL of TEMED (see Subheading 2.3.2, item 9). The gel concentration may vary according to the separation that you need and the protein under study. For the detection of Sp1 and NFI, an 8% gel is recommended. 6. Take up 7.5 mL of running gel. Put the pipet at the edge of glass plate, hold at a 90
angle, and pour the 7.5 mL between the glass plates. 7. Add 500–1000 μL of butanol (see Subheading 2.3.2, item 11) saturated in apyrogenic ultrapure water. Let sit for 30–45 min. 8. When the gel is polymerized, invert the apparatus and empty the butanol. Blot with Whatman paper (see Subheading 2.3.1, item 3). 9. Prepare a staking gel as follows; mix 1.9 mL of staking gel buffer (see Subheading 2.3.2, item 4), 975 μL of 30% acrylamide (39:1) stock solution (see Subheading 2.3.2, item 2), 75 μL of 10% SDS solution (see Subheading 2.3.2, item 10), and 4.6 mL of apyrogenic ultrapure water. Add 75 μL of 10% APS (see Subheading 2.3.2, item 5) and 7.5 μL of TEMED (see Subheading 2.3.2, item 9). 10. Pour-in the stacking gel and place a comb into the gel. Let sit for 20–30 min. 11. Remove the gel cassette sandwich from the casting frame and place it into the electrode assembly with the short plate facing inward.

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12. Slide the gel cassette sandwich and electrode assembly into the clamping frame. 13. Press down the electrode assembly while closing the two cam levers of the clamping frame. 14. Lower samples into the mini tank and add 400 mL of 1 SDS PAGE running buffer. Dilute (1:10) 10 SDS PAGE running buffer (see Subheading 2.3.2, item 6) in apyrogenic ultrapure water. 15. Prepare and load the samples (see Subheading 3.3.1) and load 5 μL of prestained molecular weight markers (see Subheading 2.3.1, item 2) in the first well. 16. Run at 100 V for 1 h 30 min (depending on the extent of protein separation that you need). 17. Before the run is completed, get out a Pyrex dish and fill it with 1 transfer buffer. Dilute (1:10) 10 transfer buffer (see Subheading 2.3.2, item 7) and 20% methanol (see Subheading 2.3.2, item 8) in apyrogenic ultrapure water. 1 transfer buffer can be stored at 4
C before use. 18. Soak one PVDF membrane (see Subheading 2.3.1, item 3), four Whatman papers, and two fiber pads per gel for at least 5 min. PVDF must be activated in a methanol bath (see Subheading 2.3.2, item 8) prior to use. 19. Stop the electrophoresis and remove the gel. Separate plates and peel the gel off the plate into the 1 transfer buffer. 20. Open the gel cassette sandwich with the black side (or anode side) down and stack in this order: fiber pad, two Whatman papers, gel, PVDF membrane, two Whatman papers, and fiber pad. Make sure to roll out air bubbles with plastic pipette. 21. Close and put the twice gel cassette sandwich into the transfer chamber in the appropriate orientation (take care at the anode and cathode sides). Put in the frozen block supplied with the unit. Fill the transfer chamber with 1 transfer buffer. 22. Transfer at 120 V 1 h 30 min in the cold room (4
C). 23. Open each sandwich. Carefully remove membranes and put them in a clean container (pipet tip box for example) filled with 1 TBS. Dilute (1:10) 10 TBS (see Subheading 2.3.2, item 12) in apyrogenic ultrapure water. 3.3.3 Membrane Blocking and Antibody Incubations

1. Rinse each membrane twice in 1 TBS. Dilute (1:10) 10 TBS (see Subheading 2.3.2, item 13) in apyrogenic ultrapure water. 2. Block nonspecific antibody binding sites with 25 mL of blocking solution (see Subheading 2.3.2, item 14) for 1 h at room temperature.

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Syringe 1mL SephadexG-50

15 mL tube

Siliconized glass wool

Uncapped Eppendorf tube

Fig. 5 Spun-column schematic diagram

3. Primary antibody preparation. Dilute in blocking solution at the working dilution. Remove the blocking solution and add the primary antibody. For the anti-Sp1 see Subheading 2.3.1, item 4 and anti-NFI see Subheading 2.3.1, item 5. Incubate overnight at 4
C under agitation. 4. Wash each membrane in 1 TBS-T (see Subheading 2.3.2, item 13) three times for 5 min at room temperature under agitation. 5. Secondary antibody preparation. Dilute in blocking solution at the working dilution. Add secondary antibody (see Subheading 2.3.1, item 6). Incubate 3 h on shaker at room temperature under agitation. 6. Wash in 1 TBS-T three times for 5 min and twice for 5 min in distilled water at room temperature. 3.3.4 Detection of Sp1 and NFI Protein

1. Drip dry each membrane onto an acetate sheet (see Subheading 2.3.3, item 4) for 5 min with the protein side facing up. 2. Mix the ECL reagents (see Subheading 2.3.1, item 7) and pour onto the membrane. Incubate 3 min at room temperature. 3. Dump the solution off the membrane and cover the membrane with an acetate sheet. 4. In a darkroom, expose to film (see Subheading 2.3.3, item 3) for various periods of time (5, 15, 30, and 60 s or more, depending on the strength of signal) in an autoradiography cassette (see Subheading 2.3.3, item 2). A few different exposures will likely be needed. The band corresponding to Sp1 should be visible between 95 and 105 kDa (see example in Fig. 1c). The band corresponding to NFI should be visible

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between 35 and 50 kDa (see example in Fig. 3b). Actin expression may also be monitored by Western blot for normalization purpose.

4

Notes 1. Sera and additives can be thawed more rapidly at room temperature or in a 37
C water bath. However, do not refreeze. Rather, we recommend using immediately or dilute in culture medium at working dilution for further utilization. 2. Serum must be added first, followed by insulin. Insulin must be added with a new sterile plastic pipette. 3. DMSO is a toxic oxidative agent at temperatures above 10
C. Working with cells in contact with a solution containing DMSO must be done quickly and on ice. 4. HEPES may undergo degradation when exposed to light and might become toxic. 5. Fibroblasts can be cryopreserved in liquid nitrogen after irradiation (or preferred treatment). 6. [γ32P] dATP emits high-energy beta radiations. 60Co is also a radioactive material. Refer to the rules established by your local control radioactivity agency for handling and proper disposal of radioactive materials and waste. 7. The proteins Sp1 and NFI are ubiquitous transcription factors usually reported to activate gene transcription in response to physiological and pathological stimuli. They bind with high affinity to GC-rich motifs (see Table 1) and regulate the expression of a large number of genes involved in a variety of processes such as cell growth, apoptosis, differentiation, and immune responses. They are highly regulated by posttranslational modifications (phosphorylation, sumoylation, proteolytic cleavage, glycosylation, and acetylation) [11–13]. 8. When using crude nuclear extracts for detecting DNA–protein complexes in EMSA, their quality is very critical. Whenever possible, nuclei purification procedures using sucrose pads [14] are preferred to eliminate contamination by cytosolic proteins, which most often also contain substantial amounts of proteases. 9. BCA Protein Assay Kit is a two-component, high-precision, detergent-compatible assay reagent set to measure (A562nm) total protein concentration compared to a protein standard. BCA reagents provide accurate determination of protein concentration with most sample types encountered in protein research. Compared to most dye-binding methods, the BCA assay is much less affected by protein composition differences,

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providing greater protein-to-protein uniformity. However, unlike the Coomassie dye-binding methods, the universal peptide backbone also contributes to color formation, helping to minimize variability caused by protein compositional differences. 10. This protease inhibitor cocktail is a mixture of protease inhibitors with broad specificity for the inhibition of serine, cysteine, aspartic proteases, and aminopeptidase. It contains 4-(2-aminoethyl)benzenesulfonyl fluoride (AEBSF), pepstatinA, E-64, bestatin, leupeptin, and aprotinin. It contains no metal chelators. Use at 1:25 for cell lysates or tissue extracts. 11. Prepare 1 L of dialysis buffer solution to treat ten samples. 12. Acrylamide is a potent neurotoxic compound that is easily absorbed through skin. Wearing gloves and a mask to avoid direct contact with the skin or inhalation is therefore required when manipulating acrylamide dried or in solution. The acrylamide solution is light-sensitive and should be kept away from direct light. 13. The concentration of the polyacrylamide gel used in EMSA is primarily dictated by both the size of the labeled probe selected and the resolution of the DNA–protein complexes obtained. It can vary from 4% up to 12%. 14. Nearly all vertical electrophoresis apparatus can be used to perform EMSA analyses. Although gel electrophoresis is performed at room temperature in some EMSA protocols, we recommend 4
C. With some apparatus this can be easily achieved using a specially designed cooling unit. For apparatus not equipped with a cooling unit, simply run the gel in a cold room. 15. The skin sample should be disinfected with 0.5% v/v chlorhexidine gluconate in isopropyl alcohol. Using any other disinfectant may compromise cell viability. Thoroughly rinse off the disinfectant from the skin sample with cold tDMEM. Failing to eliminate the disinfectant from the system will also compromise cell viability. 16. Thermolysin allows for a better separation of the epidermis from the dermis, thus limiting fibroblast colonization in nhK cultures [15]. 17. The seeding density is determined by the operator. nhK or nhCEC densities are expected to double each day. Proliferation can vary from one population to another and decreases as passages advance. It is recommended to seed at no less than 100,000 cells/cm2 with an uncharacterized population. 95% confluence should be reached within 5–7 days for the first few passages.

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18. Protocols must be approved by the institution’s committee for the protection of human subjects. It is preferable to use corneas with as short as possible postmortem time to improve cultivation of nhCEC. 19. Here, we define confluence as the approximate percentage of the culture flask surface area occupied by epithelial cells. It is estimated under a microscope by assessing how much of a given field of vision is occupied by epithelial cells. Do not take the iHFL into account when evaluating confluence. Do not let more confluent areas differentiate for the sake of obtaining a higher mean confluence. However, note that trypsinizing cells under 75% confluence may result in count errors due to overrepresentation of iHFL. 20. For optimal efficiency, do not stack culture flasks on top of each other in the incubator. Temperature is typically higher on the flask surface directly in contact with the incubator shelf. 21. A double wash is preferable to completely remove DMSO from the suspension (see Subheading 4; Note 3). 22.

32

P has a half-life of 14.3 days. Make an account of the labeled probe every week to evaluate the residual cpm. Remake probe labeling when the probe reaches less than 50,000 cpm/μL.

23. If the spun-column is being used to change the buffer, the spun-column should be washed four times (step 3) with the desired buffer in order to equilibrate the Sephadex G-50. 24. Nonspecific DNA–protein interactions are usually prevented by the addition to the reaction mix of 1–5 μg of a nonspecific competitor DNA. Although this is clearly very effective when crude nuclear extracts are used, such high concentrations of nonspecific competitor DNA were found to compete even for specific DNA–protein complexes when enriched preparations of nuclear proteins are used in EMSA [16]. The more enriched the nuclear protein of interest, the lower the amount of nonspecific competitor required. For example, we routinely use 1–2 μg poly(dI:dC) with crude nuclear proteins, 250 ng when the nuclear extract is enriched on heparin-sepharose column, and no more than 25–50 ng with purified or recombinant proteins. 25. The signal strength of a shifted DNA–protein complex can be substantially increased by favoring the odds for the interaction between the protein of interest and its target sequence. This can easily be achieved with enriched preparations of nuclear proteins either by increasing the amount of the labeled probe selected, or by decreasing the concentration of poly(dI:dC), or both. Furthermore, the DNA binding ability of some nuclear proteins proved to be highly dependent on the amount of salt (usually provided by KCl) that is present in the reaction mix.

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Transcription factors such as NFI and Sp1 best interact with their respective target sequence in the presence of 100 mM and 150 mM KCl, respectively [17]. It is therefore useful to evaluate which KCl concentration best allow binding of nuclear proteins to a specific DNA target probe. 26. Formation of DNA–protein complexes is highly dependent on the voltage selected for their migration into the polyacrylamide gel [18]. We have found that reducing the migration time by running the EMSA at voltage lower than 120 V (usually corresponding to 10 mA for a single gel) on a 6% polyacrylamide gel rendered very unstable the formation of most DNA–protein complexes and consequently resulted in difficulty to detect them. 27. All sample volumes and total protein amounts must be the same.

Acknowledgments The authors would like to thank current and former members of the LOEX and CUO-Recherche laboratories who contributed to develop and improve the foregoing protocols. References 1. Rheinwald JG, Green H (1975) Serial cultivation of strains of human epidemal keratinocytes: the formation keratinizing colonies from single cells. Cell 6(3):331–343 2. Green H, Rheinwald JG, Sun TT (1977) Properties of an epithelial cell type in culture: the epidermal keratinocyte and its dependence on products of the fibroblast. Prog Clin Biol Res 17:493–500 3. McLoughlin CB (1961) The importance of mesenchymal factors in the differentiation of chick epidermis. J Embryol Exp Morphol 9:370–384 4. Melbye SW, Karasek MA (1973) Some characteristics of a factor stimulating skin epithelial cell growth in vitro. Exp Cell Res 79:279–286 5. Wessells NK (1964) Substrate and nutrient effects upon epidermal basal cell orientation and proliferation. Proc Natl Acad Sci U S A 52:252–259 6. Masson-Gadais B, Fugere C, Paquet C, Leclerc S, Lefort NR, Germain L, Guerin SL (2006) The feeder layer-mediated extended lifetime of cultured human skin keratinocytes is associated with altered levels of the transcription factors Sp1 and Sp3. J Cell Physiol 206:831–842

´ , Lavoie A, Larouche D, 7. Bisson F, Rochefort E Zaniolo K, Simard-Bisson C, Damour O, Auger FA, Gue´rin SL, Germain L (2013) Irradiated human dermal fibroblasts are as efficient as mouse fibroblasts as a feeder layer to improve human epidermal cell culture lifespan. Int J Mol Sci 14(3):4684–4704 8. Gaudreault M, Carrier P, Larouche K, Leclerc S, Giasson M, Germain L, Guerin SL (2003) Influence of Sp1/Sp3 expression on corneal epithelial cells proliferation and differentiation properties in reconstructed tissues. Invest Ophthalmol Vis Sci 44:1447–1457 9. Duval C, Gaudreault M, Vigneault F, TouzelDescheˆnes L, Rochette PJ, Masson-Gadais B, Germain L, Gue´rin SL (2012) Rescue of the transcription factors Sp1 and NFI in human skin keratinocytes through a feeder-layerdependent suppression of the proteasome activity. J Mol Biol 418:281–299 10. Larouche D, Paquet C, Fradette J, Carrier P, Auger FA, Germain L (2009) Regeneration of skin and cornea by tissue engineering. Methods Mol Biol 482:233–256 11. Li L, Davie JR (2010) The role of Sp1 and Sp3 in normal and cancer cell biology. Ann Anat 192:275–283

Quality of Primary Cultured Epithelial Stem Cells 12. Zhao C, Meng A (2005) Sp1-like transcription factors are regulators of embryonic development in vertebrates. Develop Growth Differ 47:201–211 13. Li L, He S, Sun JM, Davie JR (2004) Gene regulation by Sp1 and Sp3. Biochem Cell Biol 82:460–471 14. Leclerc S, Eskild W, Guerin SL (1997) The rat growth hormone and human cellular retinol binding protein 1 genes share homologous NF1-like binding sites that exert either positive or negative influences on gene expression in vitro. DNA Cell Biol 16:951–967 15. Germain L, Rouabhia M, Guignard R, Carrier L, Bouvard V, Auger FA (1993) Improvement of human keratinocyte isolation

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and culture using thermolysin. Burns 19(2):99–104 16. Roder K, Schweizer M (2001) Running-buffer composition influences DNA-protein and protein-protein complexes detected by electrophoretic mobility-shift assay (EMSA). Biotecnol Appl Biochem 33:209–214 17. Gaudreault M, Vigneault F, Leclerc S, Guerin SL (2007) Laminin reduces expression of the human alpha6 integrin subunit gene by altering the level of the transcription factors Sp1 and Sp3. Invest Ophthalmol Vis Sci 48:3490–3505 18. Vossen KM, Fried MG (1997) Sequestration stabilizes lac repressor-DNA complexes during gel electrophoresis. Anal Biochem 245:85–92

Methods in Molecular Biology (2019) 1879: 75–86 DOI 10.1007/7651_2018_115 © Springer Science+Business Media New York 2018 Published online: 14 November 2018

Ribonucleoproteins Mediated Efficient In Vivo Gene Editing in Skin Stem Cells Wenbo Wu and Ting Chen Abstract The clustered regularly interspaced, short palindromic repeats (CRISPR)-Cas9 system functions like an adaptive immune system in a variety of microbes and has recently been engineered as a powerful tool for manipulating genomic sequences in a huge variety of cell types. In mammals, CRISPR/Cas9 has the potential to bring curative therapies to patients with genetic diseases, although it remained unknown whether suitable in vivo methods for its use are feasible. It is now appreciated that the efficient delivery of these genome-editing tools into most tissue types, including skin, remains a major challenge. Here, we describe a detailed protocol for performing in vivo gene editing of genomic sequences in mouse skin stem cells using Cas9/sgRNAs ribonucleoproteins in combination with electrotransfer technology. We here present all of the required methods needed for the protocol, including molecular cloning, in vitro sgRNA expression and sgRNA purification, Cas9 protein purification, and in vivo delivery of cas9 ribonucleoproteins. This protocol provides a novel in vivo gene editing strategy using ribonucleoproteins for skin stem cells and can potentially be used as curative treatment for genetic diseases in skin and other somatic tissues. Keywords Cas9/sgRNA ribonucleoproteins, Electrotransfer, In vivo gene editing, Skin stem cells

1

Introduction

1.1 The CRISPR/Cas9 Genome-Editing Tool

The clustered regularly interspaced, short palindromic repeat (CRISPR)-associated (Cas) system is an RNA-guided DNA cleavage system initially identified in bacteria and archaea that functions like an adaptive immune system [1, 2]. RNA-guided DNA cleavage is an effective mechanism for the silencing of foreign genes in prokaryotes and has recently emerged as a powerful tool for genome engineering; it is also being explored as a potential therapeutic method to treat patients with genetic diseases [3–6]. CRISPR/Cas9-mediated somatic genome editing has been reported in multiple mice organs, including lung, liver, brain, pancreas, and muscle [7–14]. To date, most approaches have relied on virus-based delivery systems, which have drawbacks such as the potential integration of viral DNA into a host genome, off-target effects due to prolonged expression of genome-editing machinery, and the possible activation of virus-triggered host immune responses. Efficient alternative methods for the in vivo delivery of

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genome-editing machinery that can circumvent these critical problems are needed. Additionally, it can be anticipated that most therapeutic applications of CRISPR/Cas9 will require tissuespecific delivery. To date, achieving this has remained challenging for most tissues, including skin. 1.2 The Electrotransfer Technology

2

Electroporation refers to an increase in cell membrane permeability that results from the formation of aqueous pores in a lipid bilayer that are generated by the application of external pulsed electric fields [15]. Electroporation has been widely used in biomedical applications, including DNA vaccination, electrochemotherapy for local treatment in oncology, nonthermal irreversible electroporation as a minimally invasive surgical procedure to ablate tissue as well as in gene electrotransfer protocols for various cells and tissues. In animals, gene electrotransfer was first demonstrated in skin [16], then in liver, muscle, brain, kidney, testis, cartilage, arteries, prostate, and cornea [15, 17–21], albeit with transfection efficiencies that vary considerably among these different tissues. Alongside the development of gene editing tools, electroporation is increasingly being seen as a very important gene delivery strategy that can effectively circumvent the need for a virus-mediated delivery. There are multiple reports detailing the successful in vivo electroporation-based delivery of genome-editing machinery in mice organs, including in retina and brain [11, 22]. Here, we demonstrate the in vivo application of Cas9/sgRNA ribonucleoproteins and electroporation to mediate gene editing in the skin stem cells of postnatal mice [23]. In this protocol, we present a step-by-step protocol for editing the genomes of mouse skin stem cells using CRISPR/Cas9 in combination with electrotransfer technology.

Materials

2.1 Molecular Biology Reagents

1. Plasmid: pSpCas9(BB)-2A-GFP CRISPR/Cas9 (pX458) plasmid (Addgene, plasmid #48138) 2. DNA oligos (Standard, desalted) for cloning of chosen guides 3. PCR reagents: PrimeSTAR GXL DNA Polymerase (Takara) 4. Enzyme: BbsI, T4 ligase (NEB, New England Biolabs) 5. 10 T4 DNA ligase buffer 6. Ultrapure water (RNAse-/DNAse-free) 7. Escherichia coli competent cells (e.g., DH5α strain) 8. Luria–Bertani (LB) agar plates and liquid media containing 50 μg/mL 9. Ampicillin

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10. Gel DNA Recovery kit (Zymo Research) 11. EndoFree Maxi Plasmid kit (TIANGEN) 12. Standard DNA gel electrophoresis reagents 2.2 Verification of the Function of sgRNAs

1. TIAMamp Genomic DNA kit (TIANGEN) 2. PCR primers for amplification of genomic DNA used in the T7 endo I assay 3. Gel DNA Recovery kit (Zymo Research) 4. T7 endonuclease I 5. 0.25 M EDTA 6. 1.5% agarose gel

2.3 In Vitro sgRNA Transcription

1. T7-sequence containing forward primers and a universal reverse primer for generation of sgRNA templates 2. MEGAscript T7 Transcription kit (Thermo Fisher Scientific) 3. RNA purification reagents including phenol:chloroform:isoamyl alcohol (25:24:1, pH < 5), chloroform, and isopropanol 4. Reagents used in denaturing urea polyacrylamide gels, including urea, 10 TBE, 40% acrylamide (19 acryl:1 bis-acryl), and TEMED 5. Ethidium bromide 6. Standard heat block apparatus 7. Low range ssRNA ladder (NEB, #N0364S) 8. UV transilluminator

2.4 Recombinant Cas9 Protein Expression and Purification

1. E. coli BL21 cells 2. LB broth 3. Ampicillin stock solution (100 μg/mL) 4. Isopropyl β-D-thiogalactopyranoside (IPTG) 5. Shaking incubator 6. Sonication buffer [1 phosphate buffer, pH 7.8, 1.5 M NaCl, 10% glycerol, 10 mM imidazole, and 1 proteinase inhibitor cocktail (Roche)] 7. Sonicator 8. Ni Sepharose 6 Fast Flow (GE Healthcare) 9. Wash buffer (1 phosphate buffer, pH 7.8, 1.5 M NaCl, 10% glycerol, and 20 mM imidazole) 10. Elution buffer (1 phosphate buffer, pH 7.8, 1.5 M NaCl, 10% glycerol, and 250 mM imidazole) 11. Dialysis buffer A (20 mM Tris–HCl, pH 8.0, 150 mM KCl, 10% glycerol), dialysis buffer B (20 mM Tris–HCl, pH 8.0,

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150 mM KCl, 20% glycerol), and dialysis buffer C (20 mM Tris–HCl, pH 8.0, 150 mM KCl, and 50% glycerol) 12. Coomassie blue staining reagents 13. Gradient SDS-PAGE gel 14. PageRuler Prestained Protein Ladder (Thermo Fisher Scientific) 15. Neon Transfection System (Thermo Fisher Scientific) 16. Cas9 antibody (mAb) (Active Motif, #61577); and Donkey Anti-Mouse IgG antibody (Jackson ImmunoResearch) 2.5 Intradermal Injection and In Vivo Electroporation

1. Dynabeads (Invitrogen) 2. 29-gauge needle (BD) 3. 1 Dulbecco’s phosphate-buffered saline (DPBS) 4. Square wave pulse generator (BTX, model ECM 830) 5. 5-mm-diameter tweezertrodes (BTX, #45-0489)

3

Methods

3.1 Design and Generation of CRISPR/ Cas9 Genome-Editing Vectors

For the design and cloning of CRISPR/Cas9 vectors, we advise following the Nature Protocol or Springer Protocol from the research group of Dr. Feng Zhang Lab [24, 25]. In general, the sgRNAs can be designed by searching for “GG” or “CC” sequences near the sites of interest using the online CRISPR design tool of the Zhang Lab (http://crispr.mit.edu/). We use the pSpCas9(BB)-2A-GFP CRISPR/Cas9 plasmid (Addgene, plasmid #48138) and insert the specific sgRNA target sequence as described in their protocol.

3.2 Verification of the Function of sgRNAs

1. Genome DNA extraction Extract genomic DNA from transfected cells using a QuickExtract DNA Extraction kit following the manufacturer’s recommended protocol. 2. T7 endonuclease I assay The T7 endonuclease I assay is performed as described in the aforementioned Nature or Springer Protocols. Briefly, targeted genomic loci are amplified from genomic DNA using the specific primers with amplicon sizes between 600 and 1000 bp. The PCR reactions are conducted with a high-fidelity enzyme (e.g., PrimeSTAR GXL DNA Polymerase, Takara) following the manufacturer’s recommended protocol. The PCR products are purified and quantified, and a denaturing reaction is set up. The reannealed products are digested with T7 endo I (NEB) and incubated at 37  C for 15 min. 0.25 M EDTA is then added to

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stop the reaction, and the products are loaded immediately on a 1.5% agarose gel for visualization. 3.3 In Vitro Transcription of sgRNA

1. Adding the T7 promoter onto the sgRNA template via PCR Synthesize the specific T7 promoter-containing forward primers and universal reverse primer (see below for specific sequences). Set up a PCR reaction with a high-fidelity enzyme with an sgRNA containing pSpCas9(BB)-2A-GFP CRISPR/ Cas9 plasmid as the template. Next, run the PCR product on a 2% agarose gel. The correctly sized band (template size: ~100 bp) is then excised and purified with a Gel DNA Recovery kit (Zymo Research) and eluted with nuclease-free water. The concentration of each guide RNA is measured using a NanoDrop instrument (Thermo Fisher Scientific). Primer

Sequence

Target -Ai14-

TAATACGACTCACTATAGAAAGAATTGATTTGATACCGgttttagagct

L Target -Ai14-

TAATACGACTCACTATAGGTATGCTATACGAAGTTATTgttttagagct

R Universal

AAAAGCACCGACTCGGTGCCAC

reverse

Within the forward primers, the T7 promoter sequence is marked with capital letters; the sgRNA sequences (Ai14-L/R are sgRNA sequences targeting loxP sites in the genome of Rosa26-STOP-tdTomato mice) are marked in red; and the sequences that are annealed to the template are written in lowercase letters. 2. In vitro sgRNA transcription The gel purified PCR products are used as templates for in vitro transcription using a MEGAscript T7 Transcription kit (Thermo Fisher Scientific) following the manufacturer’s recommended protocol (see Note 1). 3. Purify RNA using phenol/chloroform extraction and isopropanol precipitation This is the most robust method for purifying transcripts (see Note 2). It will remove all enzymes as well as most of the free nucleotides. We strongly recommend that these purification steps are carried out under clean, RNA nuclease-free conditions, as directed by the stepwise instructions provided in the MEGAscript T7 Transcription kit manual. A brief description of the steps is listed below: (a) Add 115 μL of nuclease-free water and 15 μL of the ammonium acetate stop solution and mix thoroughly.

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(b) Add an equal volume of phenol:chloroform:isoamyl alcohol (25:24:1, pH < 5) and then add the same volume of pure chloroform. Recover the aqueous phase and transfer it to a new tube. (c) Precipitate the RNA by adding 1 volume of isopropanol and mixing well. (d) Chill the mixture for at least 15 min at 20  C. Centrifuge at 4  C for 15 min at 20,000  g to pellet the RNA. Carefully remove the supernatant solution and resuspend the RNA in nuclease-free water. (e) Measure the concentration with a NanoDrop instrument and prepare aliquots of the solution; store frozen at 80  C. 4. (Optional, but highly recommended) Analyze the sgRNA products by gel electrophoresis Polyacrylamide gels (4–5%) are better for sizing smaller transcripts than agarose gels. Since secondary structures formed in sgRNA transcripts may cause aberrant migration and/or multiple bands, the gel should be run under denaturing conditions. Denaturing urea polyacrylamide gels are prepared and run according to standard procedures (Molecular Cloning, A Laboratory Manual, 2012). A brief instruction for preparing and running denaturing acrylamide gels is supplied here. (a) 5% acrylamide/8 M urea gel l

Mix the following (15 mL is enough gel solution for one 13  15 cm  0.75 mm gel):

For 15 mL

Component

7.2 g

Urea (high quality)

1.5 mL

10 TBE (see Note 3)

1.9 mL

40% Acrylamide (19 acryl:1 bis-acryl)

To 15 mL

ddH2O

l

Stir at room temperature until the urea is completely dissolved and then add:

For 15 mL

Component

120 μL

10% Ammonium persulfate

16 μL

TEMED l

Mix briefly after adding the last two ingredients, which will catalyze polymerization and then pour gel immediately.

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(b) Sample preparation: To achieve good resolution of the guide, load ~1 μg of sgRNA per gel lane. For denaturing polyacrylamide gels, add an equal volume of Gel Loading Buffer II provided in the MEGAscript T7 Transcription kit and heat for 3–5 min at 80–90  C. (c) To stain the RNA with ethidium bromide during electrophoresis, add 10 μg/mL ethidium bromide to the RNA samples (d) Run the gel using 1 TBE as the gel running buffer (see Note 4) (e) View ethidium bromide stained gels on a UV transilluminator. Ideally, there will be a single band at the 100 bp molecular weight (Fig. 1) 3.4 Recombinant Cas9 Protein Expression and Purification

1. Cas9 protein expression using bacterial expression vector (a) The amplified full-length humanized SpCas9 gene is cloned into the pET21-(b) bacterial expression vector, which is driven by the T7 promoter and encodes a His-tag at the C-terminus (Fig. 2a). (b) To express the Cas9 protein, a pET21-(b) vector encoding Cas9 is transformed into E. coli BL21 cells, followed by plating onto LB agar medium containing 100 μg/mL ampicillin. (c) Pick single colony and incubate in 4 mL of LB broth containing 100 μg/mL ampicillin as a starter culture overnight at 37  C. (d) On the following day, the cells are diluted 1:100 into the LB broth and grown at 37  C until the OD600 value reaches around 0.6. (e) The culture is incubated at 18  C for 30 min, and IPTG is then added to a final concentration of 0.2 mM to induce cas9 expression at 18  C for 17 h. (f) After this incubation, cells are pelleted by centrifugation at 8000  g for 10 min at 4  C. 2. Cas9 protein purification (a) The cell pellet is washed with PBS and lysed by sonication (40% W, 3 s pulse, 6 s rest, for a total of 15 min, on ice) in a sonication buffer. (b) After lysis, the bacterial cell lysate is pelleted by centrifugation at 10,000  g for 40 min at 4  C, and the supernatant is separated and collected into a new container. (c) To initially prepare the nickel-NTA resin, 2 mL of the resin is washed twice with wash buffer and centrifuged at 100  g for 5 min at 4  C.

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Fig. 1 In vitro transcription of sgRNA (sgRNA-Ai14-L and Ai14-R). Add T7 promoter sequencing via PCR, followed by in vitro transcription. The purified sgRNA was analyzed with denaturing urea polyacrylamide gel electrophoresis. The arrow indicates the expected position of the sgRNAs

(d) The supernatant from the cell lysate samples is then incubated with the prepared nickel-NTA resin at 4  C for 4 h to capture His-tagged Cas9. (e) After co-incubation, the resin is transferred to a 25-mL column, which is washed once with 10 column volumes of wash buffer. (f) The proteins are eluted with 10 mL of elution buffer. Collect the eluent with 0.5 mL/tube. A sample aliquot from each tube is analyzed via SDS-PAGE with Coomassie

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Fig. 2 Recombinant Cas9 protein expression and purification. (a) Structure of the bacterial expression vector, pET21b, encoding the Cas9 gene. T7: T7 RNA polymerase promoter, hSpCas9: human codon-optimized Streptococcus pyogenes Cas9, NLS: nuclear localization signal, and 6 His-tag: hexahistidine tag. (b) SDS-PAGE of purified Cas9 protein. The arrow indicates the expected position of the Cas9 protein

blue staining (Fig. 2b), and the protein concentration is determined using a NanoDrop instrument. (g) The proteins are concentrated by dialysis. First, incubate the eluent in 2 L of dialysis buffer A for 2 h, followed by incubating in 2 L of dialysis buffer B for another 2 h. Finally, incubate the eluent in 1 L of dialysis buffer C overnight at 4  C. 3. Test Cas9 degradation rate in vitro cell line (optional) (a) Transfect cells with recombinant Cas9/sgRNA ribonucleoprotein (molar ratio is kept at 1:1) using Neon Transfection System according to product manual. (b) Harvest the cells at indicated time posttreatment (Fig. 3a) and lyse cells with 1 SDS loading buffer at 98  C for 10 min. (c) Perform western blot with prepared samples using anticas9 antibody (Active MOTIF) according to standard procedures (Fig. 3b). 3.5 Intradermal Injection and In Vivo Electroporation

1. Mice are anesthetized with isoflurane. 2. Premix the Cas9 protein with sgRNAs; the Cas9/sgRNA mixture is incubated at room temperature for 15 min. The molar ratio of gRNA to Cas9 protein is kept at approximately ~1:1, and the molar ratio among the two sgRNAs is kept at 1. For each newborn mouse, around 50 μg of purified recombinant Cas9 protein is mixed with 5 μg of sgRNA (2.5 μg/sgRNA) in a 20-μL volume with 1 DPBS. 3. Before injection, dynabeads (with a 1:30 bead dilution ratio, Invitrogen) are added into the Cas9/sgRNA mixture to mark the electroporation area for later analysis.

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Fig. 3 Time-course analyses of Cas9/sgRNA ribonucleoproteins degradation rate in vitro cell line. (a) Schematic diagram of electroporation with Cas9/sgRNA Ai14-L/R ribonucleoproteins within keratinocyte cell line generated from C57BL/6J mice. (b) Western blot analysis of keratinocyte cells transfected with Cas9/ sgRNA ribonucleoproteins via Neon Transfection System (two pulses of 30 ms each, 1150 V)

4. A 29-gauge needle (BD) is used to perform intradermal injection at the tail of a newborn mouse (see Note 5). 5. After intradermal injection, electroporation is performed using a square wave pulse generator (BTX, ECM 830), and 5-mm-diameter tweezertrodes (BTX, #45-0489) with the following conditions: ten pulses of 50 ms each, 18 V/mm, and 100 ms interval time. The distance between the two electrodes depends on the thickness of the tail diameter. For tail skin, two electroporation treatments are performed, with orientations perpendicular to each other. 6. Evaluate mutated cell from treated skin with T7 endo I assay. Or analyze the tail skin from treated mice with wholemount imaging (Fig. 4c–h). 7. (Optional) To determine whether epidermal stem cells are targeted successfully, the long-term fate of the labeled basal epidermis cells after one electroporation is performed and quantified with section staining (Fig. 4a, b).

4

Notes 1. In order to increase the yield of the reaction, increase the reaction time to 6–8 h and increase the amount of template to 250 ng per 20 μL reaction system. 2. The RNA can also be purified with other methods such as lithium chloride precipitation or spin column chromatography. 3. 10 TBE can be made on your own following standard protocols (Molecular Cloning, A Laboratory Manual, 2012). Alternatively, commercial 10 TBE solutions are available inform vendors (e.g., Life Technologies).

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Fig. 4 tdTomato keratinocyte colony analysis within the tail epidermis after one electroporation treatment with Cas9/sgRNA Ai14-L/R ribonucleoproteins. (a) Schematic diagram of electroporation with Cas9/sgRNA Ai14-L/ R ribonucleoproteins at the tails of newborn Rosa26-stop-tdTomato reporter mice. (b) Quantifications of basal cell number per colony after one electroporation treatment with Cas9/sgRNA ribonucleoproteins at 3 days, 20 days, 2 months, 4 months, and 13 months posttreatment. Representative wholemount images at 415 days posttreatment (c–e) and respective stimulated model images (f–h) using Imaris software are shown at the right. Scale bar: 20 μm

4. Pre-running of the gel without loading is performed for about 30 min, and the wells of urea-containing gels are rinsed immediately before loading of the samples. 5. Avoid bubbles within Cas9/sgRNA ribonucleoprotein mixtures. For intradermal injection in the tail of postnatal mice, a 20-μL injection volume is recommended. For adult mice, a

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50-μL tail injection volume is recommended. To keep variability to a minimum, we recommended that the same skilled operator should perform all of the injections. References 1. Horvath P, Barrangou R (2010) CRISPR/Cas, the immune system of bacteria and archaea. Science 327(5962):167 2. Bhaya D, Davison M, Barrangou R (2011) CRISPR-Cas systems in bacteria and archaea: versatile small RNAs for adaptive defense and regulation. Annu Rev Genet 45:273–297 3. Cong L et al (2013) Multiplex genome engineering using CRISPR/Cas systems. Science 339(6121):819 4. Jinek M et al (2013) RNA-programmed genome editing in human cells. Elife 2:e00471 5. Mali P et al (2013) RNA-guided human genome engineering via Cas9. Science 339 (6121):823 6. Doudna JA, Charpentier E (2014) Genome editing. The new frontier of genome engineering with CRISPR-Cas9. Science 346 (6213):1258096 7. Maddalo D et al (2014) In vivo engineering of oncogenic chromosomal rearrangements with the CRISPR/Cas9 system. Nature 516 (7531):423–427 8. Yin H et al (2014) Genome editing with Cas9 in adult mice corrects a disease mutation and phenotype. Nat Biotechnol 32(6):551–553 9. Swiech L et al (2015) In vivo interrogation of gene function in the mammalian brain using CRISPR-Cas9. Nat Biotechnol 33 (1):102–106 10. Chiou SH et al (2015) Pancreatic cancer modeling using retrograde viral vector delivery and in vivo CRISPR/Cas9-mediated somatic genome editing. Genes Dev 29(14):1576–1585 11. Bakondi B et al (2016) In vivo CRISPR/Cas9 gene editing corrects retinal dystrophy in the S334ter-3 rat model of autosomal dominant retinitis pigmentosa. Mol Ther 24(3):556–563 12. Long C et al (2016) Postnatal genome editing partially restores dystrophin expression in a mouse model of muscular dystrophy. Science 351(6271):400 13. Nelson CE et al (2016) In vivo genome editing improves muscle function in a mouse model of Duchenne muscular dystrophy. Science 351 (6271):403

14. Maresch R et al (2016) Multiplexed pancreatic genome engineering and cancer induction by transfection-based CRISPR/Cas9 delivery in mice. Nat Commun 7:10770 15. Yarmush ML, Golberg A, Sersa G, Kotnik T, Miklavcic D (2014) Electroporation-based technologies for medicine: principles, applications, and challenges. Annu Rev Biomed Eng 16:295–320 16. Titomirov AV, Sukharev S, Kistanova E (1991) In vivo electroporation and stable transformation of skin cells of newborn mice by plasmid DNA. Biochim Biophys Acta 1088 (1):131–134 17. Heller R et al (1996) In vivo gene electroinjection and expression in rat liver. FEBS Lett 389 (3):225–228 18. Aihara H, Miyazaki J (1998) Gene transfer into muscle by electroporation in vivo. Nat Biotechnol 16(9):867–870. 1087-0156 (Print) 19. Heller LC, Heller R (2006) In vivo electroporation for gene therapy. Hum Gene Ther 17 (9):890–897 20. Favard C, Dean D, Rols M-P (2007) Electrotransfer as a non viral method of gene delivery. Curr Gene Ther 7(1):67–77 21. Cemazar M, Sersa G (2007) Electrotransfer of therapeutic molecules into tissues. Curr Opin Mol Ther 9(6):554–562 22. Suzuki K et al (2016) In vivo genome editing via CRISPR/Cas9 mediated homologyindependent targeted integration. Nature 540 (7631):144–149 23. Wu W (2016) Efficient in vivo gene editing using ribonucleoproteins in skin stem cells of recessive dystrophic epidermolysis bullosa mouse model. Proc Natl Acad Sci U S A 114 (7):1660–1665 24. Ran FA et al (2013) Genome engineering using the CRISPR-Cas9 system. Nat Protoc 8 (11):2281–2308 25. Cong L, Zhang F (2015) Genome engineering using CRISPR-Cas9 system. Methods Mol Biol 1239:197–217

Methods in Molecular Biology (2019) 1879: 87–99 DOI 10.1007/7651_2018_171 © Springer Science+Business Media New York 2018 Published online: 20 July 2018

Isolation and Characterization of Cutaneous Epithelial Stem Cells Stephanie R. Gillespie and David M. Owens Abstract The outer layer of mammalian skin is a multilayered epithelium that perpetually renews multiple differentiated lineages. During homeostasis, the maintenance of skin epithelial turnover is ensured by regionalized populations of stem cells that largely remain dedicated to distinct epithelial lineages including squamous, follicular, sebaceous, Merkel, and sweat glands. Cutting edge developments in this field have focused on: (1) stem cell activation cues derived from a number of extrinsic sources including neurons, dermal fibroblasts and adipocyte, and immune cells; and (2) characterization of epithelial stem cell homeostasis via hierarchical versus stochastic paradigms. The techniques outlined in this chapter are designed to facilitate such studies and describe basic procedures for cutaneous stem cell isolation and purification, which are based on leveraging their unique expression of surface proteins for simultaneous targeting and purifying of multiple subpopulations in adult skin. In addition, protocols for assessment of in vitro and ex vivo progenitor capacity as well as techniques to visualize progenitor populations in whole skin are discussed. Keywords Epithelial lineage, Epithelial progenitor markers, Epithelial stem cell homeostasis, Skin differentiation, Skin reconstitution assay

1

Introduction The perpetual renewal of mammalian skin is known to be maintained by permanently residing stem cells that are able to sustain at least five principal differentiated lineages: the interfollicular epidermis (IFE), sebaceous gland (SG), hair follicle (HF), Merkel cells, and sweat glands [1–3]. While it has long been accepted that skin homeostasis is dependent on the ability of stem cells to replenish the turnover of these mature epithelial lineages, it is the technological advances in the areas of skin stem cell isolation and genetic drivers over the last decade that have significantly enhanced our ability to effectively characterize progenitor niches in the skin. These findings have dramatically changed our view of the cutaneous epithelial stem cell landscape rendering a highly compartmentalized epithelium maintained by multiple classes of phenotypically distinct regional niches [2].

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In some cases, progenitor niches have been accessed using mouse genetics approaches and characterized under normal conditions to be long-lived and able to sustain the cellular input to certain epithelial structures including the IFE [4, 5], sebaceous gland [6, 7], hair follicle [8–12], and touch dome [13, 14]. In other cases, antibodies against cell surface proteins have been utilized to mark and isolate epithelial progenitors located in the IFE [3, 5, 15] and HF [16–20]. These efforts have facilitated our understanding of the relative proliferative capacity of progenitor pools as well as their capacity to regenerate IFE, HF, SG, or Merkel lineages in surrogate assays. In recent years, studies have elucidated that regulation of skin homeostasis by epithelial stem cells depends on several types of extrinsic activation signals. Treg expression of Notch family member Jagged1 induces HFSC differentiation during the hair cycle [21]. Also, both FGF9 secretion by γδ T cells and CX3CR1 and TGFβ1 secretion by cutaneous macrophages support skin homeostasis by inducing stem cell-mediated hair follicle regeneration after injury [22, 23]. In alopecia areata, signaling by cytotoxic T lymphocytes disrupts hair follicle progenitor cells, stopping hair growth [24]. Dermal adipocytes have also been shown to play a role in hair follicle stem cell-activated regulation of the hair follicle growth cycle; they can both stimulate and inhibit hair growth through PDGF and BMP signaling [25–27]. Additionally, both bulge and touch dome stem cells require neuronal SHH signaling for their maintenance [28, 29]. Collectively, these studies have illustrated that epithelial progenitors maintain skin homeostasis and respond to insult through a complex crosstalk with a variety of external cues. As new biomarkers have been implemented to better define the profile of progenitor cell subsets in the IFE and HFs, the individual cell of interest becomes less frequent. This can be a major technical challenge to functional studies such as skin and hair reconstitution and clonogenic studies where a significant number of cells may be required. In this chapter, we will outline some basic methods for isolation and functional assessment of keratinocyte clonogenicity, multipotency, and self-renewal capabilities from freshly isolated single cell suspensions of murine epidermal keratinocytes that have been subjected to FACS sorting. In particular, we will focus on clonogenic and skin and hair reconstitution assays. Methodologies to establish cultures of epidermal keratinocytes at clonal densities have been established for more than three decades and were developed by Rheinwald and Green, whose colony forming efficacy (CFE) assay uses a feeder layer of mitotically arrested mouse 3T3 fibroblasts [30]. There have been many modifications added to this method over the years [31]. As such, we also describe new methodology that enhances the CFE assay by using serum-free media with additional extracellular matrix proteins. The development of

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the hair reconstitution assay [32, 33] revealed the shortcomings of in vitro assays, which typically do not account for stem cell potency. Importantly, we feel the ability to conduct skin and hair reconstitution assays from freshly isolated FACS-sorted keratinocyte subsets provides a robust platform to identify and distinguish unipotent, bipotent, and multipotent epithelial progenitors. We also include a protocol for whole mount skin immunolabeling, which enables the identification of less frequent cell types and the visualization of entire cell networks. This whole skin protocol can also be used with genetically labeled mice to visualize individual stem cells in their niches. This technique potentially allows for lineage-tracing studies in three-dimensional space, which can facilitate studies to determine whether skin homeostasis results from a hierarchical [34] or stochastic [4, 35] stem cell function [36].

2

Materials

2.1 Skin Cell Isolation Solutions

1. 2.5% Trypsin (Life Technologies) diluted to 0.25% in Hank’s balanced salt solution (HBSS) (Invitrogen) 2. 1
PBS, pH ¼ 7.6 (Invitrogen), sterilized 3. Fibroblast growth medium: DMEM (Invitrogen) supplemented with 10% donor bovine serum (Invitrogen) and 2% penicillin–streptomycin (Invitrogen) 4. Collagenase type I (Worthington Biochemical), 10 mg/ml stock solution in PBS 5. DNAse I (Worthington Biochemical), 20,000 units/ml stock solution in PBS 6. 70-μm cell strainer (Fisher Scientific) 7. Betadine 1% solution in water 8. 70% EtOH solution

2.2

Antibodies

1. α6 integrin (CD49f, BD Biosciences) (see Note 1) 2. Sca-1 (Ly6G, BD Biosciences) 3. CD34 (RAM 34, BD Biosciences) 4. CD200 (OX-2, BD Biosciences)

2.3 Clonogenic Assay

1. Nunclon 6-well dishes (Fisher Scientific) 2. 4% Paraformaldehyde in PBS, pH 6 3. Rhodamine B, 1% solution in H2O (Sigma) Classic CFE Assay 1. Complete FAD growth medium: three parts DMEM (Invitrogen), one part Ham’s F12 supplement (Invitrogen), 10%

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defined fetal bovine serum (HyClone), 10 ng/ml EGF (PeproTech), 0.5 mg/ml hydrocortisone (Sigma-Aldrich), 1010 M cholera enterotoxin (Sigma-Aldrich), 5 mg/ml insulin (SigmaAldrich), 1.8
104 M adenine (Sigma-Aldrich), 100 U/ml penicillin (Invitrogen), and 100 mg/ml streptomycin (Invitrogen) 2. 3T3 Fibroblasts (ATCC) mitotically arrested with either mitomycin c (Sigma) or γ-radiation 3. 0.25% Trypsin/1 mM EDTA stock solution (Invitrogen) Serum-Free CFE Assay 1. Cnt-PR serum-free medium (ZenBio)

2. VF coating medium (William’s E medium without L-glutamine (Gibco) with: 0.01 mg/ml fibronectin (Sigma), 0.03 mg/ml vitrogen collagen (Advanced Biomatrix), 25 μg/ml tenascin-C (R&D Systems) (see Note 2), 10% BSA, 1 M HEPES, and 116 mM CaCl2) 2.4 Skin Reconstitution Assay

1. Silicon culture chambers—Upper F2U #30-268; Lower F2L #30-269 (Renner GmbH) 2. Surgical instruments including forceps, curved scissors, stapler and staple remover (all from Temin); sterile drapes, alcohol swabs, and anesthetics 3. Immunodeficient mice (Nude mice) (NCR nude, male, 7–9 weeks old), supplied by Taconic or we prefer NSWNUM (homozygote females) (see Note 3) 4. Small heating pad

2.5 Whole Mount Staining Assay

1. 1
PBS, pH ¼ 7.6 (Invitrogen) 2. Depilatory cream (Surgi-Cream, American International Industries) 3. 4% Paraformaldehyde in PBS, pH 6 4. Wash solution (3% Triton X-100 in PBS) 5. Antibody solution (20% DMSO, 0.05% normal serum, 1% Triton X-100 in PBS) 6. Aqueous mounting media containing a nuclear counterstain such as DAPI

3

Methods

3.1 Epidermal Keratinocyte Isolation

1. Under a biological cabinet, submerge euthanized 8-week-old mice in 1% Betadine for 2 min and wash in sterile H2O. Submerge mice in 70% EtOH for 1 min and wash in sterile H2O.

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2. Under a biological cabinet, surgically excise the dorsal skin using sterile forceps and scissors and float skin dermis side up in sterile PBS in a sterile Petri dish. Scrape away the subcutaneous fat and muscle using a sterile scalpel and forceps. During scraping, move skin to a new Petri dish containing fresh PBS as necessary (see Note 4). 3. Float skin epidermis side up in 0.25% trypsin (in Ca++-free HBSS) in a 10-cm Petri dish for 1.5–2 h (see Note 5). 4. Aspirate trypsin and recover skins in 10-ml fibroblast growth medium. Gently detach epidermis from dermis using a scalpel and mince epidermal scrapings into small pieces. 5. Transfer scrapings into a sterile 100-ml bottle and add 30-ml fibroblast growth medium and a stir bar. Recover epidermal cells with gentle stirring for 30 min and filter cells through a 70-μm cell strainer into a 50-ml falcon tube. Spin cells for 10 min at 1000 rpm. Wash cells in 10 ml PBS, spin and resuspend cells in 10-ml fibroblast growth medium, and spin for 10 min at 1000 rpm. 6. For antibody labeling (see below), resuspend cell pellets at 5–10
106 cells/ml in fibroblast growth medium. Typically, 10–12
106 viable basal keratinocytes are harvested from a single dorsal skin. 3.2 Preparation of Highly Inductive Dermal Fibroblasts

1. Surgically excise the dorsal skin from euthanized postnatal day 1–2 mice using sterile forceps and scissors (see Note 6). 2. In a dry Petri dish, lay skins flat dermis side down with no folded edges. Slowly pour in ice-cold 0.25% trypsin/1 mM EDTA and avoid getting the tops of the skins wet. Incubate skins overnight at 4  C. 3. Remove skins, one at a time, from trypsin and place on dry p150 plate, dermis side up. Flatten it again and use fine forceps to separate epidermis from dermis, starting at one edge of skin and flipping the dermis up and off the epidermis, which should stay on the plate. 4. Transfer dermis one at a time to a plate containing 10 ml of media on ice. 5. For eight dermises, use 0.5-ml collagenase stock solution (10 mg/ml in H2O) plus 12 ml HBSS in a 50–100-ml sterile beaker. Transfer dermises into the beaker and mince into small pieces using sharp scissors. 6. Transfer to a 250-ml flask with a magnetic stir bar. Stir at 37  C for 30 min [optional: For the last 5 min, add 20 μl of DNAse I (stock at 20,000 U/ml in PBS)].

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7. Dilute three- to fourfold with media and filter through sterile gauze or 70-μm filter. Rinse the flask with 5-ml media and pass through the filter. 8. Spin the cells at 1500 rpm for 5 min at 4  C. 9. Resuspend and wash the pellet once in HBSS. 10. Count cells, use at 8
106 fresh dermal cells per graft, and cryopreserve or plate at 1
106 cells per 10-cm dish for later use. 3.3 Antibody Labeling and FACS Analysis

1. Select an appropriate panel of antibodies for the target cells of interest (please see Table 1 for examples of published progenitor marker profiles). When possible, select antibodies directly conjugated to fluorescent dyes. 2. Count cells and aliquot equal amounts into experimental and control (single-stained and unstained) tubes in complete growth medium. 3. Incubate antibodies at concentrations according to manufacturer guidelines for 30 min to 1 h in complete growth medium on ice (see Note 7). Spin and wash cells, and resuspend in growth medium supplemented with DAPI or an alternative nuclear stain. 4. Conduct sorting (see Note 8). Typically, 10–20
106 viable α6+ basal keratinocytes can be sorted from a single dorsal skin.

Table 1 Markers of epithelial progenitor cells in adult mouse skin Progenitor location

Markers

IFE

Lrig1+

IFE

α6

IFE and HF infundibulum

α6 CD34 Sca-1

IFE touch dome

α6+CD34Sca-1+CD200+

bright

References [5]

CD71 

+

dim

[15] +

[19]

+

[3] [5]

Junctional zone

Lrig1

HF isthmus and infundibulum

α6 MTS24(Plet-1) +

+

+

[18] [7]

Sebaceous duct

Blimp1

HF isthmus

α6lowCD34Sca-1 Lgr6+

[19] [11]

HF bulge

α6dim + brightCD34+ Krt15+

[16, 17] [37]

Lower HF bulge and hair germ

Lgr5+

[10]

Hair germ

P-cadherin

+

[38]

Methods for Skin Stem Cells

3.4 Clonogenic Analysis

3.4.1 Classic Colony Forming Efficacy Assay in FAD Medium

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Clonogenic assays typically require fewer cells compared to skin and hair reconstitution assays, and harvested cells from a single animal will usually suffice. Cells can be grown with a confluent layer of mitotically arrested 3T3 fibroblasts in complete FAD growth medium or defined serum-free medium in the absence of fibroblasts. We include both the classic protocol for the CFE assay that uses a feeder layer of mitotically arrested mouse 3T3 fibroblasts and our CFE assay that uses serum-free media with additional extracellular matrix proteins. The classic protocol was optimal to perform the CFE assay with human keratinocytes. While human keratinocytes form colonies in high-Ca++ conditions, mouse keratinocytes differentiate when exposed to high Ca++. The recent availability of low-Ca++ media enables our ability to perform the CFE assay with human and mouse keratinocytes in serum-free conditions and without co-culture with a feeder layer. 1. Preparing the 6-well plates 1 day in advance will allow the fibroblasts to fully attach, spread, and condition the growth medium. Plate 1
106 mitotically arrested 3T3 fibroblasts in 3-ml complete FAD growth medium per well. 2. The next day harvest human keratinocytes from a single mouse dorsal skin and FACS sort desired keratinocyte subpopulations as described above. The FACS instrument can be optimized towards purity and accuracy in counting since cell numbers will be in excess. Propidium iodide or DAPI should be used to exclude dead cells. Many of the dead cells are postmitotic suprabasal cells that are sensitive to the 70% ethanol washes during cell harvesting. Use either the FACSAria Automated Cell Deposition Unit (ACDU) function or manually seed 1
103 (a range of 0.5–2
103 cells can be used) sorted keratinocytes per well. 3. Incubate cells for 2 weeks at 32  C in a humidified incubator with 5% CO2 and change the medium every 48 h. 4. After 2 weeks, aspirate off the culture medium and replace with 3-ml Versene per well. After 1–2 min at room temperature, detach the feeder layer by repeated pipetting of Versene over the plate (keratinocytes will not detach). Gently wash plates two times with PBS (take care not to detach colonies). 5. Fix for 1 h at room temperature with 4% PFA. Gently wash plates two times with PBS. 6. Stain wells with Rhodamine B for 1 h (see Note 9) at room temperature (just enough Rhodamine solution to cover the cells is sufficient). Aspirate off Rhodamine solution and gently wash the wells three times with PBS. 7. Aspirate off the final wash and allow wells to completely dry by turning plates upside down. Afterwards, plates can be imaged

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and colonies may be manually counted. Typically, the total number of colonies and the number of colonies greater than 4 mm in diameter are counted and compared between keratinocyte subpopulations. 3.4.2 Serum-Free Colony Forming Efficacy Assay

1. Add 1 ml VF coating medium to each well of 6-well plate and incubate at 37  C for 1 h. Aspirate VF coating medium and add CnT medium to wells. 2. Harvest mouse or human keratinocytes from a single mouse dorsal skin and FACS sort desired keratinocyte subpopulations as described above. The FACS instrument can be optimized towards purity and accuracy in counting since cell numbers will be in excess. Propidium iodide or DAPI should be used to exclude dead cells. Many of the dead cells are postmitotic suprabasal cells that are sensitive to the 70% ethanol washes during cell harvesting. Use either the FACSAria Automated Cell Deposition Unit (ACDU) function or manually seed 1
103 (a range of 0.5–2
103 cells can be used) sorted keratinocytes per well. 3. Incubate cells for 2 weeks at 32  C in a humidified incubator with 5% CO2 and change the medium every 48 h. 4. After 2 weeks, aspirate off the culture medium, gently wash with PBS (take care not to detach colonies), and fix in 4% PFA for 1 h at room temperature. 5. Gently wash with PBS and stain wells with Rhodamine B for 1 h (see Note 9) at room temperature (just enough Rhodamine solution to cover the cells is sufficient). Aspirate off Rhodamine solution and gently wash the wells three times with PBS. 6. Aspirate off the final wash and allow wells to completely dry by turning plates upside down. Afterwards, plates can be imaged and colonies may be manually counted. Typically, the total number of colonies and the number of colonies greater than 4 mm in diameter are counted and compared between keratinocyte subpopulations.

3.5 Skin and Hair Reconstitution Assay

1. Clip hair with electric clippers, if necessary, and clean the dorsal skin with 1% Betadine and place anesthetized mice on heating pad. 2. Use scissors to make a small incision on the back of the mouse (approximately 1 cm in diameter). Better areas for chamber placement are interscapular or suprapelvic. Do not make incisions directly on the spinal protrusion. 3. Assemble the upper and lower grafting chambers together and insert through the incision so that the rims of the chamber are under skin (see Note 10).

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4. Secure the chamber to the skin with surgical stapler clips (two staples are usually enough). 5. Allow the chamber to adhere to the dorsal surface overnight prior to implanting cells. 6. Mix the desired number of epidermal cells and 2
106 dermal cells together as a slurry in HBSS. We have successfully implanted 1
105 to 6
106 epidermal cells per graft. Spin cells at 1000 rpm for 5 min, resuspend the pellet in 100 μl HBSS, and store on ice until use. 7. Gently mix cell suspensions before pipetting entire aliquot into chamber of hat, through the hole on top. 8. Replace each mouse in individual cages (on belly and away from the spout of the water bottle). 9. After 1 week (see Note 11), anesthetize mice and remove staples and gently tug on chamber to release it from mouse’s back. Use tweezers to loosen skin around edge of chamber. Grafted area may be moist and oozy, leave it be and replace mouse in cage, as before. 10. Chambers are retained, cleaned (soak overnight in soapy water), and autoclaved for reuse. 11. Grafts are usually biopsied at 5–10 weeks post-grafting. Hair usually appears after approximately 2–3 weeks. 3.6 Skin Whole Mount Immunolabeling

1. Shave the hair of the dorsal skin of euthanized mouse with electric clippers and apply depilatory cream to the shaved dorsal surface for 6 min. Rinse well with warm water and pat dry with paper towel. 2. Surgically excise the dorsal skin using forceps and scissors. Place the excised skin on a flat surface dermis side up. Quickly scrape away the subcutaneous fat and muscle using a scalpel. Do not allow the skin to dry. Transfer the scraped skin to a Petri with PBS and scrape off residual adipose tissue. 3. Secure dorsal skin epidermis side up onto a microscope slide using four small binder clips. Ensure that the skin is stretched and flat. Place slides into a container containing cold 4% PFA (up to three skins per 200 ml 4% PFA). Refrigerate at 4  C overnight. 4. Use a scalpel to remove the tissue from the slide, cutting a rectangle just inside the bounds of the clips. Cut this into two narrow strips and nick the top-right corner of each strip to indicate the epidermal side. 5. Place strips into 50-ml falcon tubes containing 45–50 ml wash solution for 4 h with shaking, changing solution every hour.

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6. Place strips into a 5-ml tube containing antibody solution. Add an appropriate panel of antibodies for the target cells of interest. Incubate with shaking for 48–72 h. 7. Wash in 50-ml falcon tubes containing 45–50-ml wash solution for 4 h with shaking, changing solution every hour. Place strips into a 5-ml tube containing antibody solution and secondary antibodies to the chosen panel. 8. Wash in 50-ml falcon tubes containing 45–50-ml wash solution for 4 h with shaking, changing solution every hour. 9. Arrange the strips on a slide so that the nick is on the upper right-hand corner. Mount strips onto a slide with approximately 100-μl aqueous mounting media containing a nuclear stain such as DAPI.

4

Notes 1. The use of directly conjugated antibodies is recommended whenever possible. 2. We have seen that adding tenascin-C to the VF coating medium augments clonogenic capacity. 3. In our hands, hairless immunodeficient mouse strains such as Nude are more amenable to skin grafting procedures. 4. We typically use three Petri dishes with clean PBS per skin. 5. After 1.5 h of trypsin digestion, check the skins for detachment by gently scraping the epidermis with a scalpel. If the epidermis is easily removed, then no further digestion is required. If the epidermis does not detach, check again every 15 min. 6. Euthanized pups are washed in sterile water once, followed by two washes in 70% EtOH. EtOH is removed completely and clean pups are placed in sterile Petri dish in the hood. When processing multiple pups, place detached skins in PBS until all skins are harvested. 7. Depending on affinity and purification, antibody labeling concentrations typically range from 0.25 to 1.0 μg per 1
106 cells. 8. A high-speed sorting device is required, i.e., BD FACSAria; however, many factors contribute to a successful cell sort that require consideration. The length of the trypsin digest required to separate the epidermal and dermal compartments renders the isolated keratinocytes fragile to further mechanical stress. Keratinocytes also have a tendency to aggregate and, although it is possible to prefilter the sample and gate out most aggregates electronically, they threaten the number of events that can

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be analyzed and sorted due to clogging of the sample line. To account for these issues, we prefer a larger size nozzle (100 μm) in order to obtain an uninterrupted sort. This on the other hand decreases the sheet pressure and limits the drop drive to around 24,000 drops/s on the BD FACSAria system. The resulting events that can be analyzed per second therefore are around 8000. If the population of interest is 3% of the total sample, 240 target events can be identified of which between 5 and 15% will be aborted electronically as they present a conflict to the purity of the sample. Thus, acquiring 1
106 target cells would require more than 1.5–3 h of efficient sorting time. If more cells are required, it may be necessary to sacrifice the animals and harvest the cells at multiple time points in order to maintain viable cells. 9. The cells can be stained from 1 h to 1 week in Rhodamine B. 10. Prior to implanting the chambers, make sure that there is a hole in top half of hat. A small hole punch can be used. 11. If necessary, the grafting caps can be left on the skin for 2 weeks.

Acknowledgment This work was supported by Columbia University Skin DiseaseResource-Based Center (EPICURE) funded by the NIH (5P30AR069632). References 1. Fuchs E, Tumbar T, Guasch G (2004) Socializing with the neighbors: stem cells and their niche. Cell 116:769–778 2. Yan X, Owens DM (2008) The skin: a home to multiple classes of epithelial progenitor cells. Stem Cell Rev 4:113–118 3. Woo S-H, Stumpfova M, Jensen UB, Lumpkin EA, Owens DM (2010) Identification of epidermal progenitors for the Merkel cell lineage. Development 137:3965–3971 4. Clayton E, Doupe DP, Klein AM, Winton DJ, Simons BD, Jones PH (2007) A single type of progenitor cell maintains normal epidermis. Nature 446:185–189 5. Jensen KB, Collins CA, Nascimento E, Tan DW, Frye M, Itami S, Watt FM (2009) Lrig1 expression defines a distinct multipotent stem cell population in mammalian epidermis. Cell Stem Cell 4:427–439 6. Ghazizadeh S, Taichman LB (2001) Multiple classes of stem cells in cutaneous epithelium: a

lineage analysis of adult mouse skin. EMBO J 20:1215–1222 7. Horsley V, O’Carroll D, Tooze R, Ohinata Y, Saitou M, Obukhanych T, Nussenzweig M, Tarakhovsky A, Fuchs E (2006) Blimp1 defines a progenitor population that governs cellular input to the sebaceous gland. Cell 126:597–609 8. Levy V, Lindon C, Harfe BD, Morgan BA (2005) Distinct stem cell populations regenerate the follicle and interfollicular epidermis. Dev Cell 9:855–861 9. Ito M, Liu Y, Yang Z, Nguyen J, Liang F, Morris RJ, Cotsarelis G (2005) Stem cells in the hair follicle bulge contribute to wound repair but not to homeostasis of the epidermis. Nat Med 11:1351–1354 10. Jaks V, Barker N, Kasper M, van Es JH, Snippert HJ, Clevers H, Toftgard R (2008) Lgr5 marks cycling, yet long-lived, hair follicle stem cells. Nat Genet 40:1291–1299

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11. Snippert HJ, Haegebarth A, Kasper M, Jaks V, van Es JH, Barker N, van de Wetering M, van den Born M, Begthel H, Vries RG, Stange DE, Toftga˚rd R, Clevers H (2010) Lgr6 marks stem cells in the hair follicle that generate all cell lineages of the skin. Science 327:1385–1389 12. Petersson M, Frances D, Niemann C (2013) Lineage tracing of hair follicle stem cells in epidermal whole mounts. Methods Mol Biol 989:45–60 13. Doucet YS, Woo SH, Ruiz ME, Owens DM (2013) The touch dome defines an epidermal niche specialized for mechanosensory signaling. Cell Rep 3(6):1759–1765 14. Maksimovic S, Nakatani M, Baba Y, Nelson AM, Marshall KL, Wellnitz SA, Firozi P, Woo S-H, Ranade S, Patapoutian A, Lumpkin EA (2014) Epidermal Merkel cells are mechanosensory cells that tune mammalian touch receptors. Nature 509:617–621 15. Tani H, Morris RJ, Kaur P (2000) Enrichment for murine keratinocyte stem cells based on cell surface phenotype. Proc Natl Acad Sci U S A 97:10960–10965 16. Trempus CS, Morris RJ, Bortner CD, Cotsarelis G, Faircloth RS, Reece JM, Tennant RW (2003) Enrichment for living murine keratinocytes from the hair follicle bulge with the cell surface marker CD34. J Invest Dermatol 120:501–511 17. Blanpain C, Lowry WE, Geoghegan A, Polak L, Fuchs E (2004) Self-renewal, multipotency, and the existence of two cell populations within an epithelial stem cell niche. Cell 118:635–648 18. Nijhof JG, Braun KM, Giangreco A, van Pelt C, Kawamoto H, Boyd RL, Willemze R, Mullenders LH, Watt FM, De Gruijl FR, van Ewijk W (2006) The cell-surface marker MTS24 identifies a novel population of follicular keratinocytes with characteristics of progenitor cells. Development 133:3027–3037 19. Jensen UB, Yan X, Triel C, Woo SH, Christensen R, Owens DM (2008) A distinct population of clonogenic and multipotent murine follicular keratinocytes residing in the upper isthmus. J Cell Sci 121:609–617 20. Soteriou D, Kostic L, Sedov E, Yosefzon Y, Steller H, Fuchs Y (2016) Isolating hair follicle stem cells and epidermal keratinocytes from dorsal mouse skin. J Vis Exp (110). https:// doi.org/10.3791/53931 21. Ali N, Zirak B, Rodriguez RS, Pauli ML, Truong HA, Lai K, Ahn R, Corbin K, Lowe MM, Scharschmidt TC, Taravati K, Tan MR, Ricardo-Gonzalez RR, Nosbaum A, Bertolini M, Liao W, Nestle FO, Paus R,

Cotsarelis G, Abbas AK, Rosenblum MD (2017) Regulatory T cells in skin facilitate epithelial stem cell differentiation. Cell 169 (6):1119–1129 22. Rahmani W, Liu Y, Rosin NL, Kline A, Raharjo E, Yoon J, Stratton JA, Sinha S, Biernaskie J (2018) Macrophages promote woundinduced hair follicle regeneration in a CX3CR1 and TGFβ1 dependent manner. J Invest Dermatol. https://doi.org/10.1016/j.jid.2018. 04.010. [Epub ahead of print] 23. Gay D, Kwon O, Zhang Z, Spata M, Plikus MV, Holler PD, Ito M, Yang Z, Treffeisen E, Kim CD, Nace A, Zhang X, Baratono S, Wang F, Ornitz DM, Millar SE, George C (2013) Fgf9 from dermal γδ T cells induces hair follicle neogenesis after wounding. Nat Med 19:916–923 24. Xing L, Dai Z, Jabbari A, Cerise JE, Higgins CA, Gong W, de Jong A, Harel S, DeStefano GM, Rothman L, Singh P, Petukhova L, Mackay-Wiggan J, Christiano AM, Clynes R (2014) Alopecia areata is driven by cytotoxic T lymphocytes and is reversed by JAK inhibition. Nat Med 20(9):1043–1049 25. Guerrero-Juarez CF, Plikus MV (2018) Emerging nonmetabolic functions of skin fat. Nat Rev Endocrinol 14:163–173 26. Zwick RK, Guerrero-Juarez CF, Horsley V, Plikus MV (2018) Anatomical, physiological, and functional diversity of adipose tissue. Cell Metab 27(1):68–83 27. Plikus MV, Mayer JA, de la Cruz D, Baker RE, Maini PK, Maxson R, Chuong CM (2008) Cyclic dermal BMP signalling regulates stem cell activation during hair regeneration. Nature 451(7176):340–344 28. Xiao Y, Thoresen DT, Williams JS, Wang C, Perna J, Petrova R, Brownell I (2015) Neural Hedgehog signaling maintains stem cell renewal in the sensory touch dome epithelium. Proc Natl Acad Sci U S A 112(23):7195–7200 29. Brownell I, Guevara E, Bai CB, Loomis CA, Joyner AL (2011) Nerve-derived Sonic hedgehog defines a niche for hair follicle stem cells capable of becoming epidermal stem cells. Cell Stem Cell 8(5):552–565 30. Rheinwald JG, Green H (1975) Serial cultivation of strains of human epidermal keratinocytes: the formation of keratinizing colonies from single cells. Cell 6:317–330 31. Morris RJ (1994) Procedure for harvesting epidermal cells from the dorsal epidermis of adult mice for primary cell culture in “high calcium” defined medium. In: Leigh IM, Watt FM (eds) Keratinocyte methods. Cambridge University Press, Cambridge, pp 25–31

Methods for Skin Stem Cells 32. Weinberg WC, Goodman LV, George C, Morgan DL, Ledbetter S, Yuspa SH, Lichti U (1993) Reconstitution of hair follicle development in vivo: determination of follicle formation, hair growth, and hair quality by dermal cells. J Invest Dermatol 100:229–236 33. Kamimura J, Lee D, Baden HP, Brissette J, Dotto GP (1997) Primary mouse keratinocyte cultures contain hair follicle progenitor cells with multiple differentiation potential. J Invest Dermatol 109:534–540 34. Mascre´ G, Dekoninck S, Drogat B, Youssef KK, Brohee´ S, Sotiropoulou PA, Simons BD, Blanpain C (2012) Distinct contribution of stem and progenitor cells to epidermal maintenance. Nature 489(7415):257–262 35. Lim X, Tan SH, Koh WL, Chau RM, Yan KS, Kuo CJ, van Amerongen R, Klein AM, Nusse R

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(2013) Interfollicular epidermal stem cells selfrenew via autocrine Wnt signaling. Science 342 (6163):1226–1230 36. Hsu YC, Li L, Fuchs E (2014) Emerging interactions between skin stem cells and their niches. Nat Med 20(8):847–856 37. Lyle S, Christofidou-Solomidou M, Liu Y, Elder DE, Albelda S, Cotsarelis G (1998) The C8/144B monoclonal antibody recognizes cytokeratin 15 and defines the location of human hair follicle stem cells. J Cell Sci 111:3179–3188 38. Greco V, Chen T, Rendl M, Schober M, Pasolli HA, Stokes N, Dela Cruz-Racelis J, Fuchs E (2009) A two-step mechanism for stem cell activation during hair regeneration. Cell Stem Cell 4:144–169

Methods in Molecular Biology (2019) 1879: 101–110 DOI 10.1007/7651_2018_146 © Springer Science+Business Media New York 2018 Published online: 07 July 2018

Interfollicular Epidermal Stem Cells: Boosting and Rescuing from Adult Skin Mariana T. Cerqueira, Ana M. Frias, Rui L. Reis, and Alexandra P. Marques Abstract Epidermal stem cells (EpSCs) isolation struggle remains, mainly due to the yet essential requirement of welldefined approaches and markers. The herein proposed methodology integrates an assemblage of strategies to accomplish the enrichment of the interfollicular EpSCs multipotent fraction and their subsequent separation from the remaining primary human keratinocytes (hKC) culture. Those include rapid adherence of freshly isolated hKC to collagen type IV through the β1-integrin ligand and Rho-associated protein kinase inhibitor (Rocki) Y-27632 administration to the cultures, followed by an immunomagnetic separation to obtain populations based in the combined CD49fbri/CD71dim expression. Flow cytometry is the supporting method to analyze the effect of the treatments over the expression rate of early epidermal markers keratins19/5/14 and in correlation to CD49fbri/CD71dim sub-populations. The step-by-step methodology herein described indulges the boosting and consecutive purification and separation of interfollicular epidermal stem cells, from human keratinocytes cultures. Keywords Collagen IV, Epidermal stem cells, Flow cytometry, Immunomagnetic separation, Rock inhibitor

1

Introduction Human Keratinocytes (hKC) have a limited lifespan in culture that constraints their proliferative capacity and consequently their clinical potential. The long-term function of the skin equivalents generated in a Regenerative Medicine context can be limited by the length of time needed to obtain epithelial sheets in vitro, during which patient is highly susceptible to infection, and also by extensive culture that may lead to terminal differentiation of the hKC to be grafted, thus compromising its success. Therefore, the use of epidermal stem cells (EpSCs) that play an important role in cellular regeneration, wound healing and neoplasm formation [1] for this purpose, enlarges the possibility of providing an alternative and clinically relevant active source of biological material. Despite a wide effort among stem cells biologists community [2–6], EpSCs isolation difficulty remains, mainly due to the insufficiency of molecular markers that distinguish these cells from other proliferative cells within skin basal layer, highlighting the need for

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defining approaches and a panel of markers to obtain specific and well-characterized cell populations. P63 is abundantly expressed by holoclones, and therefore recognized as being also present in EpSCs playing an important role in morphogenesis and in the expression pattern of the cultures [7]. It has been also proposed that EpSCs exhibit a characteristic keratin profile that includes the typical K5 and K14 expression of the basal layer but not of K1/ 10 of the suprabasal layer cells. K19 appears also as an EpSCs associated marker as it is expressed by cells present in the skin hair follicles bulge and in the deep epidermal rete ridges within thicker epidermis, being also expressed by a sub-population of hKC in the human basal layer during proliferative lateral skin expansion [8]. Interestingly, the molecules related with cell-substratum adhesion are naturally meaningful as potential EpSCs markers, supported by the hypothesis that EpSCs require strong adherence to the basement membrane to maintain their stem cell characteristics or their position in the stem cell niche [9]. Beta 1 integrin was firstly identified in highly proliferating KC (holoclones) and was used to distinguish EpSCs and other basal cells [5]. However, subsequent studies revealed that the majority of the cells of the basal layer in the human epidermis, EpSCs and transient amplifying cells, exhibit the expression of beta 1 integrin [10] and other putative markers such as the combination of CD49f (α6-integrin) and CD71 (transferrin receptor) [5]. Human epidermal cells have been thus divided into three different subsets, α6briCD71dim, α6briCD71bri, and α6dim expressing cells, the first being those with the highest proliferation rate and capability of long-term epidermal renewal [11], even at a limited dilution. Despite the high importance of EpSCs, they constitute between 1–10% of the basal layer cells and, independently of the standardization of a characteristic panel of markers, boosting this population in culture through enrichment methods is a major demand. The involvement of Rho-associated protein kinase (Rock) in tissue homeostasis, namely in the epidermis, is already recognized. Regardless of the unconsciousness of the exact timing of events, the key role that Rock plays in determining hKC fate was clearly demonstrated. By blocking Rock function, an inhibition of hKC terminal differentiation and an increase in cell proliferation was observed [12]. It has also been shown that Rock inhibitor (Rocki) leads to an increased number of hKC in primary cultures that can survive and grow forming healthy colonies, thus suggesting its effect in boosting the cells exhibiting stem cell behavior [13] yet retaining the ability to differentiate and to form a stratified epithelium in adequate organotypic models [14]. The herein proposed methodology describes an assemblage of strategies to accomplish enrichment and further purification of the EpSCs multipotent fraction present in hKC primary cultures [15]. The procedure combines the rapid adherence of primary hKC to β1integrin ligand in collagen type IV and the administration of Rocki

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Y-27632 to the culture, together with subsequent immunomagnetic separation of subpopulations combining CD49fbri/CD71dim expression.

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Materials

2.1 Labware (See Note 1)

Petri dishes (vWR, Cat. No. 391-2080) Forceps (vWR, Cat. No. 231-2177) Surgical scissors (vWR, Cat. No. 231-2177) Cell culture flasks (75 cm2, 150 cm2) (BD Falcon, Cat. No. 353136, 353028) 6-Well culture plates (BD Falcon, Cat. No. 353224) 15 mL Falcon tubes (BD Falcon, Cat. No. 352097) 50 mL Falcon tubes (BD Falcon, Cat. No. 352070) Flow cytometry tubes (BD Falcon, Cat. No. 352052) Pipettes (vWR, Cat. No. 734-1694) Cell strainers of 100 μm pore size (BD Falcon, Cat. No. 352360) Eppendorf tubes 1.5 mL (Laborspirit, Cat. No. 200400P) DynaMag™-2 magnet (Invitrogen, Cat. No. 123-21D) 0.22 μm pore membrane filters (Sarstedt, Cat. No. 83.1823.101)

2.2

Reagents

Phosphate buffer saline (PBS) (Sigma, Cat. No. P4417) Distilled water (diH2O) Antibiotic/antimycotic solution (Gibco, Cat. No. 15240062) Dispase (BD Biosciences, Cat. No. 354235) Trypsin–EDTA (Gibco, Cat. No. 25300-062) Keratinocyte-SFM medium kit with L-glutamine, EGF, and BPE (Gibco, Cat. No. 17005-075) Y-27632 dihydrochloride monohydrate (Rock inhibitor—Rocki) (Sigma, Cat. No. Y0503) Acetic acid (vWR, Cat. No. 20104.334) Human placenta collagen type IV (Sigma, Cat. No. C5533) Bovine serum albumin (BSA) (Sigma, Cat. No. A2153) Dynabeads M-450 Epoxy (Life Technologies, Cat. No. 14011) Sodium phosphate (Sigma, S0876) Permeabilization buffer (10) (eBioscience, Cat. No. 00-8333) CD49f-APC antibody (eBioscience, Cat. No. 17-0495-82) CD71-PE antibody (BD Biosciences, Cat. No. 555537) Cytokeratin 19-AF488 antibody (ExBio, Cat. No. A4-120-C100)

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Cytokeratin 14-FITC antibody (AbD Serotec, MCA890F) Keratin 5 antibody (Covance, Cat. No. PRB-160P) Alexa Fluor 488 goat anti-rabbit (Invitrogen, Cat. No. A-11008) Formaldehyde (vWR, Cat. No. ALFA33314K2) Sodium azide (Sigma, Cat. No. 13412) 2.3

Reagents Setup

1. Dispase stock solution (25 U/mL): dilute dispase, 1:2 in PBS (see Note 2). 2. Collagen IV stock solution (1 mg/mL): Add 5 mL of 0.25% acetic acid and let to dissolve overnight at 4  C (see Notes 2 and 3). 3. Rocki stock solution (1 mM): Reconstitute 1 mg of Y-27632 dihydrochloride monohydrate in 2.96 mL of diH2O. 4. Dynabeads buffer 1: prepare a buffer of 0.1 M sodium phosphate in diH2O and adjust pH to 7.4–8.0 (see Note 4). 5. Dynabeads buffer 2: make a 0.1% BSA solution in PBS and adjust the pH to 7.4. 6. Coating of immunomagnetic beads with CD71 and CD49f antibodies (see Note 5). 7. Permeabilization buffer: Dilute permeabilization buffer (10) in diH2O to obtain a 1 working solution, store at 4  C. 8. Labelling buffer: Prepare a 3% BSA solution in PBS. 9. Acquisition buffer: Make 1% formaldehyde and 0.1% sodium azide solution in PBS, filter (0.22 μm pore membrane), and store at RT. 10. Antibiotic/antimycotic solution: Make a 1% antibiotic solution in PBS. 11. Dispase working solution (2.5 U/mL): Dilute 1:10 of stock dispase solution in 1% solution of antibiotic/antimycotic in PBS. 12. Rocki working solution (10 μM): Dilute Rocki stock solution (1 mM) 1:100 in KSFM, in order to have KSFM supplemented with 10 μM Rocki.

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Methods

3.1 Isolation of Human Keratinocytes from Adult Skin 3.1.1 Processing Human Skin

1. Remove the exceeding fat tissue from the dermis with scissors and scalpel. 2. Wash the skin samples with antibiotic/antimycotic solution (20 s). 3. Cut skin into 0.5 cm2 pieces.

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Incubate skin pieces in dispase working solution (2.5 U/mL) overnight at 4  C, in a 250-mL flask. 1. After incubation, place the skin samples on a Petri dish and peel off epidermis from dermis using two pairs of forceps.

3.1.3 Isolation of Human Keratinocytes: Digestion of Epidermis with Trypsin

1. Place epidermis (dermal side up) in a new Petri dish. 2. Add 0.05% trypsin–EDTA. 3. Incubate the samples at 37  C for 5–7 min. 4. Add an equal amount of KSFM. 5. Scrape off cells carefully with a cell scraper. 6. Pipette rigorously up and down several times. 7. Pour cell suspension through a 100-μm pore size cell strainer into a 50-mL Falcon tube. 8. Wash with PBS, passing the liquid through the 100-μm pore size cell strainer. 9. Centrifuge for 5 min at 290  g. 10. Wash pellet with 5 mL of PBS. 11. Pour cells through a 100-μm pore size cell strainer into a 50-mL Falcon tube. 12. Centrifuge for 4 min at 290  g. 13. Resuspend cell pellet (hKC) in KSFM.

3.2 Epidermal Stem Cells Enrichment Strategies (See Note 6) 3.2.1 Rapid Adherence to Collagen IV

3.2.2 Rock Inhibitor

1. Coat tissue culture surface with collagen IV, by incubating 5 μg/cm2 at 37  C, for at least 1 h (see Note 3). 2. Wash with PBS. 3. Plate 2  104 hKC/cm2 in KSFM. 4. Change medium every 2–3 days and keep the culture until 80% confluency. 1. Plate 2  104 cells/cm2 in Rocki working solution (10 μM). 2. Change medium every 2–3 days, keep them in culture until 80% confluency.

3.2.3 Combined Approach

3.3 Intermediate Analysis

1. Perform steps described in Subheading 3.2.1 to 3.2.2 2. Proceed as described in Subheading 3.3. An intermediate analysis of a fraction of the obtained cells after each treatment should be performed in order to assess the effect in the increased fraction of interest—EpSCs fraction. Therefore, in this section a flow cytometry protocol using a combination of CD49fAPC and CD71-PE markers is described.

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1. Harvest the adherent cells cultured under the described conditions with trypsin–EDTA. 2. Transfer cells to a 15-mL Falcon tube and add labeling buffer up to 10 mL. 3. Centrifuge cell suspension at 200  g for 5 min. 4. Count cells using a hemocytometer. 5. Discard supernatant and resuspend cell pellet to a concentration of 0.5–106 cells/mL in fresh labelling buffer. 6. Add 100 μL of cell suspension to each flow cytometry tube (see Note 7). 7. Add 4 μL of CD71-PE and 2 μL CD49f-APC antibodies; reserve one tube per condition without antibody as control. 8. Incubate for 30 min at room temperature. 9. Wash by adding 2 mL of PBS per tube and centrifuge at 250  g for 3 min. 10. Resuspend cell pellets in 500 μL of acquisition buffer. 11. Acquire data in flow cytometer. 12. Analyze simultaneous expression of CD71 and CD49f (see Fig. 1).

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Fig. 1 Dot plots of CD49f/CD71 staining on human keratinocytes isolated from the same human sample and cultured without any treatment (a) and after epidermal stem cells (EpSCs) enrichment with Rho-associated protein kinase inhibitor (Rocki) (b), showing an increase of the population of interest by the differential expression of CD49fbri/CD71dim (3.44% in a and 10.37% in b)

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1. Wash CD71 coated beads by placing the tubes in a magnet for 1 min, discarding the supernatant, and adding 1 mL of Dynabeads buffer 2, twice. +

3.4.1 Depletion of CD71 Cells

2. Incubate the cells harvested in Subheading 3.3, step 1, and resuspend in Dynabeads buffer 2 with the washed CD71 beads for 30 min at 2–8  C with gentle tilting and rotation. 3. Place the tubes in a magnet for 2 min. 4. Transfer the supernatant containing the unbound cells to a fresh 15-mL Falcon tube (CD71 cell fraction). 5. Count cells using a hemocytometer. 6. Plate 2  104 cells/cm2 in new tissue culture vessels/flasks with the corresponding treatments (described in Subheading 3.2). 7. Culture cells until 80% confluency, by changing medium 2–3 days.

3.4.2 Positive Selection of CD49f+ Cells among CD71 Population

1. Harvest CD71 cells with trypsin–EDTA and resuspend them in Dynabeads buffer 2. 2. Wash CD49f coated beads by placing the tubes in a magnet for 1 min, discarding the supernatant, and adding 1 mL of Dynabeads buffer 2, twice. 3. Incubate CD71 cells for 20 min at 2–8  C with gentle tilting and rotation. 4. Place the tubes in a magnet for 2 min. 5. Discard the supernatant and gently wash the bead-bounded cells, four times, by adding 1 mL of Dynabeads buffer 2. 6. Place the tubes in the magnet for 1 min and discard the supernatant. 7. Resuspend the cells in fresh KSFM. 8. Plate 2  104 cells/cm2 in new tissue culture vessels/flasks with the corresponding treatments (described in Subheading 3.2) for further cell expansion and analysis.

Analysis

The analysis of the expression of the early epidermal markers on the obtained cell fraction by flow cytometry is advisable to validate the success of the employed strategies. Thus, this section comprises the protocol for identifying the expression of the intracellular markers using Cytokeratin 19-AF488 and Cytokeratin 14-FITC, and Keratin 5 (see Fig. 2), respectively, by direct and indirect staining.

3.5.1 Direct Staining

Follow the protocol from Subheading 3.3, steps 1–12, using 5 μL of Cytokeratin 19-AF488 and 4 μL of Cytokeratin 14-FITC in separate tubes.

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Counts

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Fig. 2 Expression fluorescence profile of Keratin 5 (early epidermal marker) in both CD71 /CD49f (16.05%) (a) and CD71-/CD49f+ cells (32,09%) (b), showing the higher expression of K5 in the fraction of interest, thus illustrating its early differentiation stage

3.5.2 Indirect Staining

1. Follow the protocol from Subheading 3.3, steps 1–6. 2. Incubate cells with 200 μL of Permeabilization Buffer for 10 min at RT. 3. Centrifuge for 5 min at 250  g and remove supernatant. 4. Resuspend cells in 200 μL of Keratin 5 antibody diluted 1:500 in labelling buffer. 5. Incubate for 1 h at room temperature. 6. Wash by adding 2 mL of PBS, centrifuge at 250  g for 5 min, and remove supernatant. 7. Resuspend cells in 200 μL of Alexa Fluor labelled secondary antibody diluted 1:500 in labelling buffer. 8. Incubate for 45 min at 4  C protected from light. 9. Wash by adding 2 mL of PBS, centrifuge at 250  g for 5 min, and remove supernatant. 10. Resuspend cell pellets in 500 μL of acquisition buffer. 11. Acquire data in flow cytometer.

4

Notes 1. All the labware has to be sterilized prior use. 2. It is recommended to make aliquots that should be kept at 20  C, avoiding repeated freeze-thawing.

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3. Coating with collagen IV can be previously performed by incubating solution overnight at 4  C, without drying. 4. Alternatively, a solution of 0.1 M sodium borate sulfate in diH2O, pH 7.4–8, can be used as buffer 1. 5. This protocol is adapted from the section “coupling of ligands to Dynabeads” of the Dynabeads m-450 Epoxy manufacturer’s instructions. (a) Transfer 10 μL of Dynabeads to an Eppendorf tube. (b) Place the tube in a magnet for a minute and discard the supernatant. Remove the tube from the magnet. (c) Resuspend the beads in 50 μL of Dynabeads buffer 1 and add 4 μL of CD71, or 2 μL of CD49f antibody. (d) Incubate for 16–24 h at room temperature with gentle tilting and rotation. (e) Repeat step 2 and resuspend beads in Dynabeads buffer 2. 6. Cells cultured, in these different treatments, present distinct adherence and proliferation rates. No specific treatment, in which cells are cultured in non-coated plates and in KSFM, should be performed as a control. hKC cultured in Rocki and no treatment take almost 1 week to first adhere. 7. The remaining cells should be used as described in Subheading 3.4 and kept in culture for comparison purposes in the end of the experiment. References 1. Blanpain C, Lowry WE, Geoghegan A, Polak L, Fuchs E (2004) Self-renewal, multipotency, and the existence of two cell populations within an epithelial stem cell niche. Cell 118(5):635–648 2. Lau K, Paus R, Tiede S, Day P, Bayat A (2009) Exploring the role of stem cells in cutaneous wound healing. Exp Dermatol 18(11):921–933 3. Watt FM, Lo Celso C, Silva-Vargas V (2006) Epidermal stem cells: an update. Curr Opin Genet Dev 16(5):518–524 4. Blanpain C, Fuchs E (2006) Epidermal stem cells of the skin. Annu Rev Cell Dev Biol 22:339–373 5. Li A, Simmons PJ, Kaur P (1998) Identification and isolation of candidate human keratinocyte stem cells based on cell surface phenotype. Proc Natl Acad Sci U S A 95 (7):3902–3907 6. Taylor G, Lehrer MS, Jensen PJ, Sun TT, Lavker RM (2000) Involvement of follicular stem cells in forming not only the follicle but also the epidermis. Cell 102(4):451–461

7. Pellegrini G, Dellambra E, Golisano O, Martinelli E, Fantozzi I, Bondanza S et al (2001) p63 identifies keratinocyte stem cells. Proc Natl Acad Sci U S A 98(6):3156–3161 8. Pontiggia L, Biedermann T, Meuli M, Widmer D, Bottcher-Haberzeth S, Schiestl C et al (2009) Markers to evaluate the quality and self-renewing potential of engineered human skin substitutes in vitro and after transplantation. J Invest Dermatol 129(2):480–490 9. Alonso L, Fuchs E (2003) Stem cells of the skin epithelium. Proc Natl Acad Sci U S A 100 (Suppl 1):11830–11835 10. Kaur P, Li A (2000) Adhesive properties of human basal epidermal cells: an analysis of keratinocyte stem cells, transit amplifying cells, and postmitotic differentiating cells. J Invest Dermatol 114(3):413–420 11. Schluter H, Paquet-Fifield S, Gangatirkar P, Li J, Kaur P (2011) Functional characterization of quiescent keratinocyte stem cells and their progeny reveals a hierarchical organization

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in human skin epidermis. Stem Cells 29(8):1256–1268 12. McMullan R, Lax S, Robertson VH, Radford DJ, Broad S, Watt FM et al (2003) Keratinocyte differentiation is regulated by the Rho and ROCK signaling pathway. Curr Biol 13(24):2185–2189 13. Terunuma A, Limgala RP, Park CJ, Choudhary I, Vogel JC (2010) Efficient procurement of epithelial stem cells from human tissue specimens using a rho-associated protein

kinase inhibitor Y-27632. Tissue Eng Part A 16(4):1363–1368 14. Chapman S, Liu X, Meyers C, Schlegel R, McBride AA (2010) Human keratinocytes are efficiently immortalized by a Rho kinase inhibitor. J Clin Invest 120(7):2619–2626 15. Cerqueira MT, Frias AM, Reis RL, Marques AP (2014) Boosting and rescuing epidermal superior population from fresh keratinocyte cultures. Stem Cells Dev 23(1):34–43

Methods in Molecular Biology (2019) 1879: 111–118 DOI 10.1007/7651_2018_165 © Springer Science+Business Media New York 2018 Published online: 13 June 2018

Whole-Mount Immunofluorescent Staining Coupled to Multicolor Lineage Tracing Model for Analyzing the Spatiotemporal Organization of Epidermal Stem Cells Edwige Roy and Kiarash Khosrotehrani Abstract An overall three-dimensional picture of the distribution of epidermal cells at a given time point is of importance to better characterize epidermal progenitors. We introduce a whole-mount immunofluorescent staining coupled to a multicolor lineage tracing model for analyzing the spatiotemporal organization of epidermal stem cells. Laser scanning confocal microscopy with multiple labelling allows for robust imaging of dorsal skin with excellent resolution. Keywords Confocal microscopy, Mouse dorsal skin, Multicolor lineage tracing model, Whole-mount immunofluorescent staining

1

Introduction Interfollicular epidermal (IFE) homeostasis is driven by the continuous proliferation, differentiation, and shedding of epidermal cells, the keratinocytes, throughout life. Despite progress in our understanding of stem cell populations in the skin, the cellular mechanisms associated with IFE maintenance have remained controversial [1–5]. How epidermal progenitor cells are able to provide the large numbers of keratinocytes needed to maintain the barrier function of the skin upon differentiation is a fundamental question. On the one hand, it is proposed that rare stem cells are the source of all epidermal cells in the IFE. However, a more recent alternative concept has emerged suggesting that all cells with proliferative capacity have a chance to act as a progenitor randomly, providing the cells that will then form the IFE. Lineage tracing is an essential tool to study cell fate and to address the question of the maintenance of the IFE homeostasis. Although traditional lineage tracing techniques have considerably advanced our understanding of stem cell behavior, they present important limitations, in particular in identifying and tracking over time the progeny of individual stem cells, and comparing their behavior. This is of importance given the well-established heterogeneity

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among stem cells both in terms of potentialities and proliferative capacities. The recent development of multicolor genetic reporters addressable to specific cell populations largely overcomes these issues [6]. These technologies provide increased resolution in clonal identification and offer the possibility to study the relative distribution, contacts, tiled arrangement, and competitive interactions among cells or groups of cells of the same type. Although lineage tracing over time is crucial to follow the fate of progenitor cells, an overall three-dimensional picture of the distribution of epidermal cells at a given time point is of importance to better characterize epidermal progenitors. Thus, by combining a multicolor lineage tracing model with a whole fluorescent staining we propose to conduct the analysis of spatiotemporal organization of epidermal cells at a single clone level to address the question of the maintenance of the IFE homeostasis. This chapter describes a protocol to stain whole mount of dorsal skin from a multicolor lineage tracing mouse model.

2

Materials Except the formaldehyde and sucrose, every solution is freshly prepared.

2.1 Collection and Preparation of Dorsal Skin

1. Skinbow adult mice [6] or other equivalent mice allowing multicolor fate tracing. 2. Scissors and forceps. 3. Shaver and depilatory cream. 4. Fixation solution: 4% formaldehyde. For 1 l of 4% formaldehyde, add 800 ml of 1 phosphate buffered saline (PBS) to a glass beaker on a stir plate in a ventilated hood. Heat while stirring to approximately 60  C. Take care that the solution does not boil. Add 40 g of paraformaldehyde powder to the heated PBS solution. The powder will not immediately dissolve into solution. Slowly raise the pH by adding 1 N NaOH dropwise from a pipette until the solution clears. Once the paraformaldehyde is dissolved, the solution should be cooled and filtered. Adjust the volume of the solution to 1 l with 1 PBS. Recheck the pH, and adjust it with small amounts of dilute HCl to approximately 6.9. The solution can be aliquoted and frozen or stored at 2–8  C for up to 1 month. 5. 20% Sucrose solution: For 200 ml of 20% sucrose, add 200 ml of 1 PBS to a glass beaker on a stir plate. Add 40 g of sucrose powder to the PBS solution. Stir until it is completely dissolved. Stored at 2–8  C for up to 1 month.

Whole-Mount Immunofluorescent Staining Coupled to Multicolor Lineage Tracing. . .

2.2 Whole-Mount Immunofluorescent Staining

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1. Blocking solution: 1 PBS, 0.5% TritonX, 2% bovine serum albumin (BSA), 20% normal goat serum (NGS), 1% dimethyl sulfoxide (DMSO), and 100 mM maleic acid (7.5 pH). For 10 ml of blocking solution, dissolve 0.2 g of BSA in 6.85 ml of PBS, then add 50 μl of TritonX, 2 ml of NGS, 100 μl of DMSO, and 1 ml of 1 M maleic acid (7.5 pH). 2. Diluent solution: 1 PBS, 0.5% TritonX, 2% BSA, 1% DMSO, and 100 mM maleic acid (7.5 pH). For 10 ml of diluent solution, dissolve 0.2 g of BSA in 8.85 ml of PBS, then add 50 μl of TritonX, 100 μl of DMSO, and 1 ml of 1 M maleic acid (7.5 pH). 3. Washing solution: 1 PBS, 0.5% TritonX, and 0.5% Tween 20. For 1 l of washing solution, in 990 ml of PBS add 5 μl of TritonX and 5 ml Tween 20. 4. 4,6-Diamidino-2-phenylindole (DAPI) solution: For 1 ml DAPI solution, add 2 μl of DAPI in 998 μl of 1 PBS (1/500 from a 14.3 M DAPI stock solution). 5. RapiClear®: commercial clearing solution (SunJin Lab, Hsinchu City, Taiwan).

2.3 Confocal Microscopy

3

1. Fluorescent mounting media. 2. Single concave microscope slide and cover slip.

Methods An overview of the main steps of the procedure is presented in Fig. 1, with the indication of the time required to perform each step. The whole procedure is described step-by-step below by giving a detailed description followed by background notes with comments.

3.1 Collecting and Preparing Specimen

1. Sacrifice mice that have been injected with BrdU following standard laboratory protocol and use the electric shaver to remove all hair from the back of an adult mouse. Shave as close to the skin as possible while being careful to avoid nicks. Then, apply depilatory cream on the shaved surface for 2 min (see Note 1). Rinse with water. Repeat if necessary to remove all the hair follicle. Dry gently the back skin with a soft tissue. 2. With scissors, make incision to collect a piece of back skin. Use the forceps to carefully peel off the skin in one piece from the mouse. 3. Place the skin epidermis side down on the dissecting pad and use the needles to pin down two adjacent sides of the skin.

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3.1

Collection and preparation specimen 3.1.1- 3.1.4

Collect dorsal skin

3.1.5

Tissue fixation 4% Formaldehyde 2hrs 4oC

3.1.6

20% Sucrose At least 3 days at 4oC

3.2

3.3

BrdU wholemount staining 3.2.1

Collect samples for staining

3.2.2

7% HCL 20min 37oC

3.2.3

Blocking solution 5hrs room temperature

3.2.4

anti-BrdU antibody 1/50 in diluent solution overnight 4oC

3.2.5

Washing step 3 times at least 1 hour

3.2.7

Secondary antibody Alexa 647 1/500 in diluent solution overnight 4oC

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Washing step 4 times at least 1 hour

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DAPI counterstain Overnight 4oc

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Clearing step RapiClear® 3-6hrs room temperature

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Flat mounting tissue on single concave slide

Confocal microscopy

Fig. 1 An overview of the main steps of the procedure

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4. Using the scalpel, gently scrape away the fat and blood vessels covering the dermis until the dermis is clearly and uniformly exposed. Note that the dermis has a dull appearance whereas fat is shiny and reflective (see Note 2). 5. Place explants in at least 5 ml of 4% formaldehyde in 1 PBS buffer for a 9-cm2 piece of back skin. Incubate at 4  C for 2 h. 6. Remove the skin from the formaldehyde solution and wash with 1 PBS for 5 min to get rid of formaldehyde. With a tissue, pat dry the excess of PBS and place the skin in at least 5 ml of sucrose 20%. Incubate at 4  C for at least 3 days (see Note 3). 3.2 BrdU WholeMount Staining

Immunofluorescent staining (six pieces of back skin, 4 mm2 each) 1. Place the skin in a petri dish with 1 PBS and, using the scalpel, prepare small pieces of back skin (4 mm2 maximum) for staining of specific antigens. 2. Place the samples in a 1.5-ml Eppendorf tube with 300 μl of HCl 7% solution for 20 min at 37  C. Then, remove the samples from the Eppendorf tube and rinse in PBS before to dry them on a soft tissue. 3. Place the samples in a 1.5-ml Eppendorf tube with 500 μl of blocking solution. Gently mix the samples and solution by rocking the tube on a tube roller mixer at room temperature for 5 h. Make sure that the samples are covered in solution before leaving to incubate. 4. Place the samples in a new Eppendorf tube containing 200 μl primary anti-BrdU antibody diluted at 1/50 in diluent solution. Gently mix the samples and solution by rocking the tube on a tube roller mixer at 4  C overnight. 5. Place the samples in new Eppendorf tubes with 300 μl of washing buffer. Place tubes on a tube roller mixer at room temperature for at least 1 hour. 6. Repeat step 5 twice. 7. Place the skin pieces in a new Eppendorf tube containing 200 μl secondary anti-rat antibody conjugated with Alexa 647 (to avoid emission overlap) diluted at 1/500 in diluent solution. Gently mix the samples and solution by rocking the tube on a tube roller mixer at 4  C overnight. 8. Place the samples in new Eppendorf tubes with 300 μl of washing buffer. Place tubes on a tube roller mixer at room temperature for at least 1 hour. 9. Repeat step 8 three times. 10. Place samples in a new Eppendorf tube with 300 μl of DAPI solution. Gently mix the samples and solution by rocking the tube on a tube roller mixer at 4  C overnight.

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11. Place samples in a new Eppendorf tube with four drops of RapiClear® for 3–6 h (see Note 4). Remove the samples from the RapiClear® and remove the excess using a tissue. 12. Samples are place in the center of a single concave microscope slide. Add 2–3 drops of fluorescent mounting media and apply a coverslip (see Note 5). 3.3 Confocal Microscopy

1. Set up appropriate lasers for fluorophores. Confocal images were acquired with a Zeiss LSM 710 microscope using a 405-nm diode laser for DAPI (detector 1, main beam filter MBS-405, 414–463-nm barrier filter), a 458-nm Argon line for Cerulean (detector 2, main beam filter MBS-458, 464–502-nm barrier filter), a 514-nm Argon line for YFP (detector 3, main beam filter MBS-458/514, 512–570-nm barrier filter), a 561-nm photodiode laser for dTomato (detector 3, main beam filter MBS-488/561, 562–611-nm barrier filter), and a 633-nm laser for Alexa 647 (detector 3, main beam filter MBS-488/561/633-nm barrier filter). 2. Image stacks for all four channels were acquired sequentially using a 10  0.45 objective, 20  0.8 objective, or 40  1.2 W objective. In all images, YFP was represented in yellow, dTomato in red, Cerulean in blue, BrdU in green, and DAPI in cyan (Fig. 2). 3. Use sequential scan tool to avoid or reduce cross talk in which all dyes in double- or triple-stained samples will be excited at the same time. In the sequential scan mode, images will be recorded in a sequential order.

4

Notes 1. Do not apply the Veet cream for more than 2 min to avoid damaging the skin. Reapply if necessary. 2. It is essential to remove all of the fat attached to the dermis in order for trypsin to easily penetrate the tissue and dissociate cells. If the fat has been insufficiently removed, staining of the cells will be affected. 3. After 3 days in sucrose, the sample can be stored at after removing the excess of sucrose.

80  C

4. Some samples need to be exposed longer to the clearing solution (anagen phase mouse skin). The level of clearing should be checked visually. 5. Apply light pressure on the coverslip to flatten the sample and remove bubbles if necessary.

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Fig. 2 Multiple optical section from a Z stack acquisition of whole-mount preparation of skin stained with antiBrdU (green). Small clones not attached to HF (two examples figured by dashed lines) can be tracked from the corneocyte layer down to the basement membrane where BrdU staining demonstrates their proliferation. YFP yellow, dTomato red, Cerulean blue, DAPI cyan. Scale: 20 mm

Acknowledgements This work was supported by the Australian Research Council Discovery Project DP130104777 and the National Health and Medical Research Council project grant 1145350.

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References 1. Potten CS, Kovacs L, Hamilton E (1974) Continuous labelling studies on mouse skin and intestine. Cell Tissue Kinet 7(3):271–283 2. Jones PH, Harper S, Watt FM (1995) Stem cell patterning and fate in human epidermis. Cell 80 (1):83–93 3. Mackenzie IC (1970) Relationship between mitosis and the ordered structure of the stratum corneum in mouse epidermis. Nature 226 (5246):653–655 4. Morris RJ, Fischer SM, Slaga TJ (1985) Evidence that the centrally and peripherally located

cells in the murine epidermal proliferative unit are two distinct cell populations. J Invest Dermatol 84(4):277–281 5. Clayton E et al (2007) A single type of progenitor cell maintains normal epidermis. Nature 446 (7132):185–189 6. Roy E et al (2016) Bimodal behaviour of interfollicular epidermal progenitors regulated by hair follicle position and cycling. EMBO J 35 (24):2658–2670

Methods in Molecular Biology (2019) 1879: 119–132 DOI 10.1007/7651_2018_131 © Springer Science+Business Media New York 2018 Published online: 27 March 2018

Isolation and Enrichment of Newborn and Adult Skin Stem Cells of the Interfollicular Epidermis Stefano Sol, Dario Antonini, and Caterina Missero Abstract The interfollicular epidermis regenerates from a heterogeneous population of basal cells undergoing either self-renewal or terminal differentiation, thereby balancing cell loss in tissue turnover or in wound repair. In this chapter, we describe a reliable and simple method for isolating interfollicular epithelial stem cells from the skin of newborn mice or from tail and ear skin of adult mice using fluorescence-activated cell sorting (FACS). We also provide a detailed protocol for culturing interfollicular epidermal stem cells and to assess their proliferative potential and self-renewing ability. These techniques are useful for directly evaluating epidermal stem cell function in normal mice under different conditions or in genetically modified mouse models. Keywords Adult stem cells, CD34, CD71, Clonogenic assay, Epidermis, Flow cytometry, Integrins, Isolation, Keratinocytes, Progenitor cells

1

Introduction The epidermis is a stratified squamous epithelium that provides a vital barrier function to the mammalian skin. It consists of several layers of keratinocytes that move from the innermost basal layer to the stratum corneum, becoming progressively differentiated. The basal layer is composed by undifferentiated proliferative progenitors that express high levels of keratins 5 and 14 (K5 and K14). In the basal layer, keratinocytes can both self-renew and sustain longterm tissue maintenance, and hence this layer is where bona fide adult stem cells reside [1]. Recent studies using sophisticated mouse reporter models have been instrumental to identify different cell populations in the epidermal basal layer. Several groups have reported that the epidermis is not maintained by a small population of slowly dividing stem cells, but rather basal keratinocytes largely divide stochastically along the basement membrane to generate daughter cells that can acquire different fates [1–6]. Stem cells can generate either identical basal progenitors with extended self-renewing capacity, two progenitors committed to differentiation, or one basal daughter that remains attached to the basement membrane, and one committed

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daughter that eventually delaminates undergoing terminal differentiation [2, 4–7]. In the basal layer, committed progenitors may still have some proliferative potential, and for this reason they are also called transit amplifying cells. However, at the molecular level, committed progenitors can be distinguished by the expression of dual differentiation markers of the basal (K5, K14) and suprabasal (K1, K10, involucrin) compartments [4, 8]. During embryonic development, epidermal appendages including hair follicles, sebaceous glands, sweat glands, and nails are generated from the ectoderm. Each population is endowed with specialized stem cells, residing in different niches along the basement membrane, and generally contributes only to its own differentiation program except when participating in wound healing (reviewed in [1]). In this chapter, we will focus specifically on the isolation and enrichment of newborn and adult skin stem cells of the interfollicular epidermis. Basal epidermal cells of the interfollicular epidermis and of the epidermal appendages adhere to the basement membrane mainly through integrins α3β1 and α6β4, the major epidermal receptors for laminin-5. Lower expression levels of integrins have been shown to correlate with a decreased proliferative potential and increased differentiation [4, 8–11]. Mouse keratinocytes characterized by high levels of α6 integrin and low to undetectable expression of the transferrin receptor (CD71) cells have a stem cell-like behavior being highly clonogenic and self-renewing for a long period of time in culture [11, 12]. Bulge stem cells share this expression pattern and are enriched in the α6bright, CD71dim population. To distinguish between interfollicular and bulge stem cells, expression of the hematopoietic progenitor cell antigen CD34 is evaluated, since bulge stem cells specifically express CD34 [13]. In addition we will describe in details a simple procedure to determine the number of progenitors with high proliferative potential and clonogenic ability that form large clones and that can be propagated by serial passages [12], a typical feature of cells with high self-renewing capacity.

2

Materials All the solution should be 0.2 μm filter sterilized.

2.1 Epidermal Cell Isolation

1. C57BL/6J or any other mouse strain. All animal studies have been approved by the Italian Ministry of Health (311/2016-PR). 2. Ethanol 70% solution in water. 3. Surgical instruments: Iris scissors (FST), serrated curved forceps, and Dupont fine forceps.

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4. Dispase solution in HBSS: Dispase 0.80 U/mL (Thermo Fisher), HEPES 10 mM, Na-bicarbonate 0.075%, HBSS, antibiotic/antimycotic 1
(Thermo Fisher). 5. Hank’s Balanced Salt Solution (HBSS) (Thermo Fisher). 6. Trypsin (2.5%), no phenol red (Thermo Fisher). 7. Phosphate buffer saline (PBS) Ca2+, Mg2+-free. 8. Falcon® 70 μm cell strainer. 2.2 Preparation of Chelated Fetal Bovine Serum (FBS)

1. Prepare 150 g Chelex 100 resin (Biorad; 100–200 mesh, sodium form) in 2 L glass-distilled H2O. Cover and stir slowly overnight at room temperature (rt). 2. Adjust pH to 7.4 using 10 N HCl. Stir for 15 min, readjust pH with 10 N HCl, and repeat a few times as needed until pH remains stable for more than 20 min. 3. Carefully decant H2O, and eliminate the remaining water by filtering (do not let the Chelex resin dry too much). 4. Add 500 mL characterized FBS (Hyclone). 5. Stir at rt for 1 h with a low speed set to minimize bubbles. 6. Sit undisturbed at 4  C for 60 min to allow the Chelex resin to form a compact pellet. 7. Recover supernatant and filter serum through a 0.20 μm filter. 8. Aliquot and freeze at 20  C.

2.3

FACS Analysis

1. Falcon® 40 μm cell strainer or Corning/Falcon round-bottom polystyrene test tube with cell strainer snap cap. 2. Round-bottom polystyrene test tubes for FACS. 3. FACS staining solution: 1
Ca2+, Mg2+-free PBS, 1 mM EDTA, 25 mM HEPES pH 7.0, 1% chelated FBS. 4. PE (phycoerythrin) rat anti-CD49f (α6 integrin) clone G0H3 (BD Biosciences). 5. APC rat anti-mouse CD71 (transferrin receptor) clone R17217 (RI7 217.1.4) (Thermo Fisher). 6. FITC rat anti-mouse CD34 clone RAM34 (BD Biosciences). 7. SYTOX blue nucleic acid stain (Thermo Fisher), or DAPI (40 ,6diamidino-2-phenylindole; Sigma), or 7-AAD (7-amino-actinomycin D) (Thermo Fisher).

2.4 Clonogenic Assay

1. Matrigel matrix (Corning). Thaw the Matrigel on ice in the cold room overnight. Once Matrigel matrix is thawed, swirl vial to ensure the material is dispersed. Aliquot it using prechilled tubes, and store it in aliquots at 70  C. Repeated freezing and thawing is not recommended. Thaw aliquots overnight on ice before coating.

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2. CnT-07 progenitor (CELLnTEC).

cell

medium

with

supplements

3. Crystal violet solution: 1% crystal violet (Sigma) in 20% methanol. Store solution in the dark at rt. Use within 2 months. 4. PYREX® 6
8 mm Cloning Cylinders (Corning). 5. Dow Corning® silicone high vacuum grease (Sigma). Sterilize by placing a small amount in the bottom of a glass Petri dish and autoclaving under standard conditions. 6. ImageJ: image processing software (https://imagej.nih.gov/ij/).

3

Methods

3.1 Isolation of Mouse Epidermal Keratinocytes

Here we will focus on the isolation of newborn epidermis, and on adult epidermis from ears and tail, since isolation of adult dorsal skin has been described in details previously [14, 15]. Ears and tails are of interest because the epidermis is thicker and hair follicles are sparser as compared to dorsal skin, and they are the preferred sites for basal cell carcinoma in various mouse models.

3.1.1 Isolation of Newborn Epidermis

1. Place newborn mice between 0 and 3 days old in a Petri dish, in an ice bucket under ice for 45–60 min (see Note 1). 2. Rinse newborn mice once with distilled water and then twice with 70% ethanol vigorously shacking for 30 s. Remove ethanol completely, and place mice under a tissue culture hood. 3. Place one mouse at the time in a sterile Petri dish, and remove a large portion of the limbs cutting in the middle of the proximal limbal segment with sterile surgical scissors. The remaining proximal segment will be useful to hold the mouse body while removing the skin. Remove the tail and the region around the anus cutting the ventral skin above the anus to avoid major bacterial contamination. 4. Make a longitudinal cut of the skin along the dorsal midline with a scalpel leaving the internal tissues intact. Remove the skin gently tearing it apart from the dorsal midline using serrated curved forceps, then turn the mouse on the ventral side, and remove the skin from the truncated hind limb and on the abdomen by holding onto a truncated hind limb (see Note 2). The last portion that is removed is the head skin. Place isolated skins in PBS while separating the others (Fig. 1a). 5. Flatten the skins with dermis down in an empty Petri dish (10–12 skins for a 150 mm dish) with the back of the forceps. It is crucial to stretch entirely the skin all around the edges; otherwise peeling will be difficult. Work quickly to avoid drying the skin.

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Fig. 1 Mouse epidermal isolation. (a) Newborn skin is isolated from the entire body and carefully stretched on a dish with the epidermal side up. (b, c) After incubation in dispase overnight, the epidermis (ep) is removed from the dermis (d) as indicated. (d–f) After incubation in EDTA, adult tail skin is stretched on a dish with the epidermal side up and is separated from dermis as shown. (g–i) To isolate the two sheaths of adult ear skin, the lower sheath of the skin and the ear cartilage are kept with thick forceps, whereas the upper sheath of the skin is carefully peeled with thin forceps. (i) The two sides of the skin are isolated from each other, although one side remains adherent to the cartilage. Subsequently, both sides are incubated in EDTA to separate epidermis from dermis (not shown)

6. Add approximately 20 mL of dispase solution to each 150 mm Petri dish, and incubate o/n at 4  C or 2 h at 37  C. The skin should be floating on the dispase solution and should not sink. 7. After the incubation, place one skin at the time on an empty Petri dish, flatten it again epidermis up, and use forceps to separate the epidermis from the dermis, starting from one corner of the skin, making sure to collect the entire epidermis (Fig. 1b, c). The epidermis should easily peel away in a single piece. 3.1.2 Isolation of Adult Tail Epidermis

1. Isolate mouse tail and remove the tip. Quickly rinse in 70% ethanol. 2. Cut the tail longitudinally with a scalpel from the base to the tip.

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3. Use forceps to grab the expose tail bone at the base, and grasp the skin with another forceps. Gently peel the skin tissue from the tail bone (see Note 3). 4. Cut the tail skin in three pieces, rinse in sterile PBS, and stretch them on a Petri dish. 5. Incubate o/n in dispase solution as above (see Note 4). Use the minimum amount of solution just enough to cover the entire dish and to have the skin floating on top. 6. After the incubation, place one skin at the time on an empty Petri dish, flatten it again epidermis up, and use forceps to separate the epidermis from the dermis, starting from one corner of the skin. The epidermis should separate easily from the underlying dermis (Fig. 1d–f). 3.1.3 Isolation of Adult Ear Epidermis

1. Cut the ear off at the base with scissors. Quickly rinse in 70% ethanol. 2. Hold the open tissue close to the cartilage dish with forceps while gently peeling the dorsal skin along the cartilage dish with fine forceps (Fig. 1g, h). 3. Rinse the isolated skin in sterile PBS (Fig. 1i). 4. Incubate for 2 h in 3 mM EDTA in PBS at 37  C. 5. Separate epidermis from the dermis as above.

3.2 Isolation of Single Cells from the Epidermis

1. Place each epidermis in a 60 mm Petri dish and add 2.0 mL of cold trypsin solution (0.125% trypsin/0.1 mM EDTA solution in HBSS). Tilt the Petri dish at a ~30 angle, and quickly mince epidermis with scissors as much as possible, until epidermis is reduced in small fragments. This step can be also performed by pooling two epidermis. 2. Place the minced tissue in a sterile falcon tube and incubate for 50 –80 at 37  C gently shacking to dissociate cells (see Note 5). 3. Add 10 mL of Ca2+, Mg2+-free HBSS +10% chelated FBS to each tube to inactivate trypsin (see Note 6). Filter cell suspension through 70-μm cell strainer by pouring slowly the cell suspension and distributing homogeneously over the entire area of the filter. This step gets rid of pieces of undigested epidermis and large clamps of cells. For subsequent FACS analysis, cell suspension should be filtered again through a 40-μm cell strainer to eliminate small cell aggregates. 4. Spin down the cells at 1000
rpm for 5 min at 4  C and resuspend in Ca2+, Mg2+-free PBS. Place on ice.

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3.3 Isolation of Basal Progenitor Cells by FluorescenceActivated Cell Sorting (FACS)

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This is an optional step to isolate and/or enrich for clonogenic progenitors: 1. Wash cells in 5–10 volumes of PBS before incubating with appropriate antibodies. Use a hemocytometer to determine cell numbers. Typically, 3–4
106 viable keratinocytes can be obtained from a single P2-P3 skin. Epidermal samples can be pooled together as needed. 2. Resuspend the cell pellet in a concentration of 1
107 cells per mL in FACS staining solution. The sample must be prepared and processed as soon as possible to minimize cell stress as sorting is a relatively harsh process. 3. Select an appropriate panel of antibodies directly conjugated to fluorescent dyes for the target cells of interest. To enrich for epidermal progenitors, staining is performed with anti-CD49f (α6 integrin) that enriches for basal cells and anti-CD71 that is more elevated in transit amplifying and differentiating cells. Alternatively, to enrich for interfollicular epidermal cells, an antibody anti-CD34, a marker of hair follicle stem cells, is used to exclude hair follicle cells. A triple staining scheme can also be used (Table 1). The choice of fluorophore will depend on the FACS/cell sorting available. 4. Add fluorescently conjugated antibodies at concentrations according to manufacturer’s guidelines. Antibodies should be titrated in preliminary experiments to find the optimal amount for an ideal fluorescence intensity compared to a negative control. 5. Include crucial controls necessary to set proper compensation: (a) unstained cells (for each cell type examined); (b) non-specific staining control for each fluorophore or dye and each cell type (“stain” cells with the fluorophore isotype

Table 1 FACS/cell sorting strategy Cell population

Antibodies

Basal cells of the interfollicular epidermis (CD49fbright/CD34)

PE CD49f FITC CD34

Highly clonogenic basal cells of the epidermis (CD49fbright/CD71dim)

PE CD49f APC CD71

Highly clonogenic basal cells of the interfollicular epidermis (CD49fbright/CD71dim/CD34)

PE CD49f APC CD71 FITC CD34

Basal cells of the hair follicle and bulge stem cells (CD49fbright/CD34+)

PE CD49f FITC CD34

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control); and (c) cells stained with only one antibodyfluorophore (or other dye) at a time (for each cell type and each fluorophore or dye). This latter control is essential to accurately collect and analyze data from samples stained with more than one fluorophore or dye. 6. Incubate for 30 min on ice protected from light. Mix cell suspension by gentle tapping every 10 min to prevent cells from settling at the bottom of the tube. 7. Wash the cells twice with ice-cold 1–2 mL of FACS staining solution. 8. Addition of a dye that enables exclusion of dead cells in the sort (i.e., DAPI or SYTOX or 7-AAD) should be added a few minutes before cell sorting (see Note 7). 9. Transfer to FACS tubes by filtering the cell suspension through cell strainer caps to ensure single cell suspension. Remember to keep the samples on ice and sort as soon as possible. 10. Sort using the setup described in Fig. 2 starting from the controls (see Note 8). Gate cells excluding cell debris, dead cells, and doublets. Select α6bright/CD34 and/or α6bright/ CD71dim (Table 1). 11. Collect cells in pre-coated collecting tubes containing FBS or PBS containing at least 10% FBS (see Note 9). In order to prevent cells sticking to the sides of the tubes, pre-coat the tubes filling them with the serum for at least 30 min before sorting. 12. After cells are sorted into collection tubes, centrifuge cells to remove diluted buffer, and replenish with fresh culture media. Plate as soon as possible. 13. A purity check can be performed after sorting by rerunning a small fraction of the sorted population, followed the samples lines are rinsed with bleach and ethanol to prevent carryover from previous sorting sample. 3.4 Clonogenic Assay

1. Coat culture dishes with Matrigel matrix. For a 35 mm dish, add 1 mL of Matrigel diluted in cold CnT-07 medium (0.5 mg/mL). Incubate at room temperature for 1–2 h. 2. Remove the coating solution without letting it dry. Seed cells shortly after sorting is completed in CnT-07 medium supplemented with antibiotics/antimycotic at a concentration of 103 cells/cm2 (see Note 10). 3. Incubate at 37  C with 5% CO2. When comparing the clonogenic capacity of cells under different experimental conditions, make sure that the same number of cells has attached to the dish the day after plating.

Isolation and Culture of Mouse Epidermal Stem Cells

c

FSC-H (x 1.000) 50 100 150 200 250

SYTOX (x 1.000) 50 100 150 200 250

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105 104 103 102 101 0

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0101 102

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Fig. 2 FACS strategy for the isolation of epidermal cells with high clonogenic potential. (a) A density plot with forward scatter area (FSC-A; a measurement of cell size) versus side scatter area (SSC-A; a measurement of cell complexity/granularity) is used to exclude debris, and to select for relatively small cells with relatively low complexity, corresponding to less differentiated cells. (b) A forward scatter area (FSC-A) versus forward scatter height (FSC-H) density plot is used to isolate single cells from doublets. (c) Staining for dead cells using SYTOX blue or DAPI staining. (d) Basal cells with high clonogenic potential are isolated as CD49fbright/CD71dim to exclude transient amplifying (CD CD49fbright/intermediate/CD71bright) and differentiated keratinocytes (CD49fdim/CD71bright). (e) Basal cells of the interfollicular epidermis are selected as CD49fbright/CD34, excluding the hair bulge cell population CD49fbright/CD34+

4. Change medium every 2 days, and culture for about 3–4 weeks depending on the size of the colonies. During this period of time, clones derived from stem cells/progenitors will have a dense healthy appearance with no signs of differentiation (Fig. 3a). 5. At the end of the experiment, stain clones with crystal violet. Remove the medium and wash twice with PBS. Remove PBS completely and stain with 2 mL of crystal violet solution for 10 min. Filter crystal violet solution through a 0.22-μm filter to remove any precipitates before using. 6. Wash dish in tap water by immersion in a large beaker until the water is clear. Be careful not to lift off cells. Change tap water between washes. 7. Drain upside down on paper towels. Allow the dish to dry completely before visualizing with a stereomicroscope (Fig. 3b).

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Fig. 3 Clonogenic assay. (a) Mouse keratinocyte clone shape and size depend on the proliferative potential of the cell from which they originate. Left panel: rapidly growing clone of highly proliferative keratinocytes 10 days after plating. Right panel: clone with irregular shape, containing large, flattened, differentiated cells 10 days after plating. (b) Clonogenic assay in a 35-mm dish 3 weeks after plating stained with crystal violet. (c) Graphic representation of clonogenic assay performed with freshly isolated mouse keratinocytes cultured for 3 weeks. % of colonies with different size (2 mm) are represented as mean  SD

8. Colonies can be counted manually or using ImageJ (an open source image processing program), with the freely available ImageJ-plugin “ColonyArea.” 9. Count number of colonies dividing them in three categories: smaller than 1.5 mm, between 1.5 and 2 mm, and larger than 2 mm (Fig. 3c). Larger and densely packed colonies derive from stem cells, whereas smaller colonies and/or sparse colonies derive from transit amplifying cells that have lost their proliferating capacity. 3.5 Secondary and Tertiary Clonogenic Assay

1. To test the self-renewing ability of progenitor cells, select dense and well-isolated colonies of average size. Draw a circle around them on the bottom of the dish with a pen marker. 2. Remove the growth medium. Rinse cells twice with PBS to remove any floating cells.

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3. Using sterile forceps pick up a cloning cylinder. Gently press the flat bottom of the cylinder into the smooth silicone grease, and remove with a sudden vertical motion. If done properly, this will give even distribution of grease on the bottom of cylinder. Set the cylinders around each colony, and press firmly against the dish using sterile forceps (Note 11). 4. Verify positioning of the cylinder over the colony under a microscope. Make sure there are no other colonies within the sealed area in the cylinder. 5. Add about 0.2 mL of the 0.25% trypsin to the cloning cylinder. 6. Incubate the dish at 37  C for 4–5 min. Then examine cells under the microscope every 2–3 min until the cells have begun to round up and come off the dish bottom. Add a few drops of growth medium to the cylinder, and pipette up and down to generate single cell suspension. 7. Transfer the single cell suspension from each cloning cylinder separately into an individual micro-centrifuge tube. Wash the area within the cylinder two to three times with approximately 250 μL of supplemented CnT-07 medium to collect all remaining cells, and transfer the wash from each cylinder into the respective micro-centrifuge tube. 8. Centrifuge the cells at 500 rpm on tabletop centrifuge for 5 min. 9. Remove the supernatant and resuspend the cell pellet in 3.0 mL fresh supplemented CnT-07 medium. Transfer 1.5 mL of cell suspension (containing all the cells from individual colonies) from each tube into two 35 mm Matrigel pre-coated dishes. Add the appropriate volume of medium. Incubate cells as above. 10. After growing secondary clones for 3 weeks, clones can be stained with crystal violet, or the procedure can be repeated to obtain tertiary colonies.

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Notes 1. Hypothermia is an acceptable method of euthanasia for fetuses and neonates up to 7 days of age as long as direct contact with ice/cold surfaces is avoided. Hypothermia leads to anesthesia first and then death. Newborn mice are resistant to hypoxia; therefore, CO2 as a euthanasia agent is not recommended but requires a second physical method such as decapitation. The epidermis of mice older than 3 days is difficult to peel off because the growing hair shafts anchor the epidermis to the dermis.

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2. Newborn are relatively fragile and by tearing the skin apart on the ventral side, the peritoneum may break. If the intestine is intact, carefully proceed to separate the skin; otherwise discard the skin and change Petri dish and forceps to avoid contaminating the epidermal cell preparation. 3. In contrast to adult dorsal skin, there is no need to remove fat from the dermis before incubating in trypsin, and doing so may actually damage the skin and compromise the following steps. 4. As an alternative protocol, newborn and adult tail skin can be incubated overnight in 0.25% trypsin in HBSS with antibiotics/antimycotic at 4  C to separate epidermis from dermis. However, trypsin is a harsh treatment (see Note 5). 5. When using trypsin, special care must be taken to minimize the exposure time, and ensure rapid inactivation, since trypsin can compromise some antibody epitopes and cell viability. In addition if cells are left in trypsin for too long, genomic DNA may entrap live cells, significantly reducing the purification efficiency. 6. Keratinocytes have the tendency of clumping together; therefore, after trypsinization it is essential to use Ca2+, Mg2+-free HBSS or PBS and to chelate the serum to avoid stabilizing adherens junction. 7. The choice for the dead cell indicator depends on the FACS analyzer and fluorophores used. The options listed in this section are compatible for the proposed fluorophores: SYTOX Blue stain Excitation/Emission (nm) (444/480) (it can be excited at 405 nm), DAPI (500–1000 ng/mL) Excitation/Emission (nm) (360/460) (it can be excited at 405 nm), and 7-AAD (2.5 μm) Excitation/Emission (nm) (546/647) (it can be excited at 488 nm). For more information on the fluorescence spectrum and the use of multiple fluorophores on specific FACS analyzers, see http://www. bdbiosciences.com/us/s/spectrumviewer. 8. Cells must be at the proper concentration in order for the sorter to function optimally. Cells that are too concentrated will have a lower recovery due to cell clumping and coincidence aborts (two cells that are too close together will be rejected by the machine in order to ensure purity), and cells that are too dilute will have a longer processing time. For large adherent cells such as keratinocytes 5–9 x 106 may be a good concentration to start with. 9. It is preferable to use serum or PBS with serum instead than medium, because phosphate-buffered medium (i.e., FACS sheath fluid) can cause precipitation of salts when mixed with carbonate-buffered medium (RPMI, MEM, etc.) leading to reduced viability for sensitive cells.

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10. CnT-07 medium is a fully defined medium ideal for growing mouse epidermal progenitors in colonies. Alternatively mouse epidermal progenitors can be cultured in colonies in a low-calcium DMEM-/FAD-based medium in the presence of feeder cells. However, feeder cells tend to detach in low calcium medium and have to be replaced every few days. 11. Be careful not to slide the cylinder across the colony. This will smear the silicone grease over the cells and prevent the trypsin from contacting them.

Acknowledgments This work was supported by the Telethon grant GGP16235 and the Italian Association for Cancer Research (AIRC IG2011N.11369). References 1. Gonzales KAU, Fuchs E (2017) Skin and its regenerative powers: an alliance between stem cells and their niche. Dev Cell 43(4):387–401. https://doi.org/10.1016/j.devcel.2017.10. 001 2. Clayton E, Doupe DP, Klein AM, Winton DJ, Simons BD, Jones PH (2007) A single type of progenitor cell maintains normal epidermis. Nature 446(7132):185–189. https://doi. org/10.1038/nature05574 3. Doupe DP, Klein AM, Simons BD, Jones PH (2010) The ordered architecture of murine ear epidermis is maintained by progenitor cells with random fate. Dev Cell 18(2):317–323. https://doi.org/10.1016/j.devcel.2009.12. 016 4. Mascre G, Dekoninck S, Drogat B, Youssef KK, Brohee S, Sotiropoulou PA, Simons BD, Blanpain C (2012) Distinct contribution of stem and progenitor cells to epidermal maintenance. Nature 489(7415):257–262. https:// doi.org/10.1038/nature11393 5. Rompolas P, Mesa KR, Kawaguchi K, Park S, Gonzalez D, Brown S, Boucher J, Klein AM, Greco V (2016) Spatiotemporal coordination of stem cell commitment during epidermal homeostasis. Science 352(6292):1471–1474. https://doi.org/10.1126/science.aaf7012 6. Sada A, Jacob F, Leung E, Wang S, White BS, Shalloway D, Tumbar T (2016) Defining the cellular lineage hierarchy in the interfollicular epidermis of adult skin. Nat Cell Biol 18

(6):619–631. https://doi.org/10.1038/ ncb3359 7. Lim X, Tan SH, Koh WL, Chau RM, Yan KS, Kuo CJ, van Amerongen R, Klein AM, Nusse R (2013) Interfollicular epidermal stem cells selfrenew via autocrine Wnt signaling. Science 342 (6163):1226–1230. https://doi.org/10. 1126/science.1239730 8. Asare A, Levorse J, Fuchs E (2017) Coupling organelle inheritance with mitosis to balance growth and differentiation. Science 355 (6324). pii: eaah4701). https://doi.org/10. 1126/science.aah4701 9. Jensen UB, Lowell S, Watt FM (1999) The spatial relationship between stem cells and their progeny in the basal layer of human epidermis: a new view based on whole-mount labelling and lineage analysis. Development 126(11):2409–2418 10. Jones PH, Watt FM (1993) Separation of human epidermal stem cells from transit amplifying cells on the basis of differences in integrin function and expression. Cell 73(4):713–724 11. Tani H, Morris RJ, Kaur P (2000) Enrichment for murine keratinocyte stem cells based on cell surface phenotype. Proc Natl Acad Sci U S A 97(20):10960–10965 12. Ferone G, Thomason HA, Antonini D, De Rosa L, Hu B, Gemei M, Zhou H, Ambrosio R, Rice DP, Acampora D, van Bokhoven H, Del Vecchio L, Koster MI, Tadini G, Spencer-Dene B, Dixon M,

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Dixon J, Missero C (2012) Mutant p63 causes defective expansion of ectodermal progenitor cells and impaired FGF signalling in AEC syndrome. EMBO Mol Med 4(3):192–205. https://doi.org/10.1002/emmm. 201100199 13. Blanpain C, Lowry WE, Geoghegan A, Polak L, Fuchs E (2004) Self-renewal, multipotency, and the existence of two cell populations within an epithelial stem cell niche. Cell 118(5):635–648. https://doi.org/10.1016/j. cell.2004.08.012

14. Jensen KB, Driskell RR, Watt FM (2010) Assaying proliferation and differentiation capacity of stem cells using disaggregated adult mouse epidermis. Nat Protoc 5 (5):898–911. https://doi.org/10.1038/ nprot.2010.39 15. Nowak JA, Fuchs E (2009) Isolation and culture of epithelial stem cells. Methods Mol Biol 482:215–232. https://doi.org/10.1007/ 978-1-59745-060-7_14

Methods in Molecular Biology (2019) 1879: 133–138 DOI 10.1007/7651_2018_132 © Springer Science+Business Media New York 2018 Published online: 27 March 2018

Isolation and Cultivation of Epidermal (Stem) Cells Xusheng Wang, Shiyang Dong, and Yaojiong Wu Abstract Recent studies have shown that epidermal stem cells derived from the epidermis of are able to form hair follicles in the presence of hair follicle-inducing cells. Here we describe the method that we have used to isolate and cultivate epidermal stem cells. Keywords Cryopreservation, Cultivation, Epidermal stem cells, Isolation

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Introduction Epidermal stem cells (Epi-SCs) inhabit in the basal layer of the interfollicular epidermis. They express high levels of cytokeratin (CK)5, CK14, CD29 (integrin β1), and CD49f (integrin α6) [1]. Epi-SCs divide and move upward to form new keratinocytes in the epidermis to replenish the dead cells, thus maintaining the homeostasis of the structure. Epi-SCs in the epidermis are preferable cells for skin bioengineering for their advantages in availability and proliferation potential in culture. Autologous human epidermal cells after culture expansion have been used to generate skin substitutes in combination with autologous dermal fibroblasts. Following transplantation into cutaneous wounds in patients, the skin substitute is able to form lasting epidermis, suggesting the existence of self-renewable epidermal stem cells in the graft. However, this skin substitute does not regenerate the hair follicle and other appendages [2]. Recently, Epi-SCs in the epidermis have been suggested to participate in the regeneration of the hair follicle following skin wounding in mice [3]. In our recent study, we show that Epi-SCs derived from the adult human foreskin and scalp retain the capacity to form de novo hair follicles and hairs, even after culture expansion, when engrafted into excisional skin wounds in combination with hair follicle-inducing cells, the skin-derived precursors (SKPs) [4].

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Materials

2.1 Reagent Preparation

1. Phosphate-buffered saline (PBS), sterile. 2. 70% ethanol. 3. RPMI 1640 medium (GIBCO). 4. Keratinocyte (K)-SFM (GIBCO) (basal), supplemented with 5 μg/ml gentamycin before use. 5. Growth K-SFM (500 ml): basal K-SFM supplemented with the following before use: (a) 20 μl penicillin–streptomycin (GIBCO), which contains penicillin and streptomycin at 100 U/ml. (b) Fungizone (GIBCO) at 7.5 μg/ml. (c) Defined keratinocyte-SFM growth supplement (GIBCO). (d) Dulbecco’s phosphate buffer saline (DPBS, GIBCO). 6. Dispase II (Sigma): Power stored at 4  C and dissolved into right concentration using PBS. 7. 0.05% trypsin/0.53 Mm EDTA (GIBCO). 8. 0.025% trypsin/0.53 mM EDTA (GIBCO). 9. Gentamycin (GIBCO). 10. 2% (W/V) iodine solution. 11. EDTA (GIBCO). 12. Soybean trypsin inhibitor (Life Technologies). 13. Serum-free cell freezing medium (Cellbanker 2, AMS Biotechnology). 14. Epidermal growth factor (EGF) (R&D). 15. Cholera toxin (GIBCO). 16. CNT-07 medium (500 ml, basal) (CELLnTEC) contains low concentration of calcium (0.07 mM); growth medium is prepared before use: basal CNT-07 medium supplemented with CnT-07.S (supplements, CELLnTEC), 20 μl penicillin– streptomycin, which contains penicillin and streptomycin at 100 U/ml, and Fungizone (GIBCO) at 7.5 μg/ml. 17. Collagen–fibronectin solution: bovine plasma fibronectin (Sigma) at 10 μg/ml and type I collagen (Sigma) at 30 μg/ml in DMEM containing 25 mM HEPES.

2.2 Equipment Preparation

1. 100 mm tissue culture dishes (treated) (Corning): for subculture of Epi-SCs, dishes are coated with collagen–fibronectin before being used. 3 ml of collagen–fibronectin solution is added to a 100 mm cell culture plate in a tissue culture hood and incubated for 1 h at room temperature. Then the solution

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is aspirated and plates are air dried in the hood. Coated plates are rinsed three times with PBS to neutralize the acidity before seeding cells. 2. Tissue culture incubator. 3. 80 mesh filter (Utah biodiesel supply). 4. Hemacytometer. 5. Surgical scissors. 6. Forceps. 7. Pipettes. 8. Centrifuge. 9. Programmable cooler (Thermal Fisher). 10. Cell freezing container (Thermal Fisher).

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3.1 Isolation and Culture of Human Keratinocytes

The protocol is adopted from previous publications [5] with modifications. 1. Collect human scalp or foreskin tissues in serum-free RPMI 1640 medium containing 5 μg/ml gentamycin and stored at 4  C. 2. In a 6-well plate, add to 5 wells, respectively, 5 ml iodine solution, 5 ml of 70% ethanol, and 3 wells of 5 ml DPBS without Ca or Mg containing 10 μg/ml gentamicin. 3. Briefly submerge the skin sample in iodine solution, 70% ethanol, and then the DPBS without Ca or Mg for a few seconds each, and then submerge in each of the three gentamicin solutions, each for 15 min (see Note 1). 4. Cut skin tissues into halves or quarters, depending on the size of the tissue; transfer the pieces, dermis side down, to a petri dish containing 25 units/ml (3 mg/ml) dispase II and 10 μg/ml gentamicin in K-SFM (basal); and incubate 18 to 24 h at 4  C. 5. Using suitable forceps, gently peel away the epidermis from the dermis and pool in a 60 mm culture dish containing 5–7 ml of 0.05% trypsin/0.53 mM EDTA in DPBS. Incubate the epidermis at 37  C for 15–20 min with gentle pipetting to aid in tissue dissociation. 6. Stop trypsin activity by addition of soybean trypsin inhibitor (10 mg/ml in DPBS) at 1:1 ratio (trypsin: inhibitor, weight), and add 5 ml K-SFM with supplements (growth medium). 7. Filter cells through an 80 mesh filter. Centrifuge cell suspension at 400  g, 5 min, 25  C.

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8. Remove supernatant and resuspend the pellet in 5 ml of fresh K-SFM growth medium. 9. Count cells using a hemacytometer. 10. Seed cells into tissue culture dishes at 1.5  105 per ml growth K-SFM (10 ml medium per 100 mm dish), and culture cells in an incubator with 5% CO2 at 37  C. Change medium every 2–3 days (see Note 2). 11. Subculture: when cells grow to 75% confluence, remove the medium. Wash culture with 10 ml DPBS. Add 2 ml 0.025% trypsin/0.53 mM EDTA (for a 100 mm dish). Incubate cells at 37  C until they become round (~5 min). Remove trypsin, and incubate cells at 37  C until they detach from the culture surface with gentle tapping (~5 min). Add 2 ml 10 mg/ml soybean trypsin inhibitor solution to the culture (100 mm dish). Collect cells by centrifuging at 400  g, for 5 min, and at 25  C. Seed the cell into a 100 mm tissue culture dish at 1  106 cell density per 10 ml K-SFM medium (see Note 3). 3.2 Isolation and Culture of Murine Keratinocytes

The protocol is adopted from previous publications [6] with modifications. 1. Prepare 0–2-day-old newborn C57 mice. 2. Clean the dorsal skin with iodine solution and 70% ethanol. 3. Make a longitudinal incision from tail to snout, and then harvest the dorsal skin using sterile forceps and surgical scissors. 4. Wash the skin sample for 10 min in RPMI 1640 medium containing ten times the normal concentrations of penicillin and streptomycin. 5. After removal of fat and membranous materials from the dermal side, cut the skin into 10-mm-wide strips. Transfer the pieces, dermis side down, to a petri dish containing 25 units/ml (3 mg/ml) dispase II and 10 μg/ml gentamicin in K-SFM (without growth supplement) and incubate for 40 min at 37  C. 6. Using suitable forceps, gently peel away the epidermis from the dermis, pool in 60 mm culture dishes containing 5–7 ml of 0.05% trypsin/0.53 mM EDTA in DPBS, and incubate at 37  C for 15–20 min with gentle pipetting to aid in tissue dissociation. 7. Repeat steps 6–12 in protocol 3.1, except that the keratinocyte-SFM growth medium for murine keratinocytes is further supplemented with 10 ng/ml EGF and 10 10 M cholera toxin.

3.3 Cell Cryopreservation

Resuspend epidermal cells in freezing medium to a concentration of 106–107 cells per ml. Aliquot cells into cryogenic storage vials. Place vials on wet ice or in a 4  C refrigerator, and start the freezing

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procedure within 5 min. Cells are frozen slowly at 1  C/min. This can be achieved using a programmable cooler or by placing vials in an insulated freezing container placed in a 70 to 90  C freezer, and then transferring to liquid nitrogen storage. 3.4 Isolation and Culture of Epidermal Stem Cells

The protocol is adopted from previous publications [5, 6] with modifications. 1. Isolate the epidermis from the mouse skin following steps 1–5 in Sect. 3.2. 2. Using suitable forceps, gently peel away the epidermis from the dermis and transfer to an Erlenmeyer flask containing CNT-07 medium and a magnetic stir bar. Stir the medium on the magnetic stirrer for 1 h in medium speed, 37  C. 3. Filter cells through a sterile Teflon mesh (70–100 μm). Centrifuge cell suspension at 400  g, for 5 min, and at 25  C. 4. Remove supernatant and resuspend the pellet in CNT-07 medium. 5. Count cells using a hemacytometer. 6. Seed 3  106 cells in 4 ml CNT-07 medium into a 100 mm tissue culture dish (uncoated). Place the plate in an incubator in 5% CO2 at 37  C. 7. After 10 min, aspirate the culture medium to remove the detached epidermal stem cells. Wash the plate with 5 ml CNT-07 medium gently. The cells attached to the plate are largely Epi-SCs, which express CD49fhigh and CD29high. Add 10 ml fresh CNT-07 medium and incubate the cells in a humidified 5% CO2 atmosphere at 37  C. 8. Check cells daily under microscope for cell density, and change medium every 48 h. 9. When culture becomes 70–75% confluence, cells are split into new plates. The medium is removed, and the culture is raised with PBS three times. Add 2 ml of Accutase and incubate for 30 min at 37  C. Check cells under microscope to make sure all cells become detached. Add 5 ml of CNT-07 medium to the plate. Collect the cell suspension into a 15 ml centrifuge tube, and pellet cells by centrifuging at 400  g for 5 min, room temperature. 10. Resuspend cells in CNT-07 medium, and seed cells into new 100 mm plates which are pre-coated with collagen–fibronectin at a density of 1  106 cells per plate in 10 ml CNT-07 medium. Incubate the cells in a humidified 5% CO2 atmosphere at 37  C. 11. Repeat 8–9.

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Notes 1. If skin sample is too large, before submerging, cut into smaller pieces (1 cm wide) with a surgical scalpel, and also remove subcutaneous fat by scraping with the scalpel. For small or thin samples, the iodine and ethanol steps may be too stringent; an alternative is to soak the sample for 10 min in RPMI 1640 medium containing ten times the normal concentrations of penicillin and streptomycin. 2. Use of collagen- or fibronectin-coated plates may enhance keratinocyte attachment and growth. 3. Trypsinization times are critical for the performance of any keratinocyte medium. Avoid excess trypsinization. Alternatively, 0.025% trypsin/0.53 mM EDTA can be replaced by 2 ml Accutase, which is milder and may create less cell damages.

References 1. Blanpain C, Fuchs E (2019) Epidermal homeostasis: a balancing act of stem cells in the skin. Nat Rev Mol Cell Biol 10(3):207–217 2. Boyce ST et al (2002) Cultured skin substitutes reduce donor skin harvesting for closure of excised, full-thickness burns. Ann Surg 235 (2):269–279 3. Ito M et al (2007) Wnt-dependent de novo hair follicle regeneration in adult mouse skin after wounding. Nature 447(7142):316–320

4. Wang X et al (2016) Hair follicle and sebaceous gland de novo regeneration with cultured epidermal stem cells and skin-derived precursors. Stem Cells Transl Med 5(12):1695–1706 5. Wang X et al (2017) Macrophages induce AKT/beta-catenin-dependent Lgr5(+) stem cell activation and hair follicle regeneration through TNF. Nat Commun 8:14091 6. Bickenbach JR (2005) Isolation, characterization, and culture of epithelial stem cells. Methods Mol Biol 289:97–102

Methods in Molecular Biology (2019) 1879: 139–148 DOI 10.1007/7651_2018_117 © Springer Science+Business Media New York 2018 Published online: 25 February 2018

One-Step Simple Isolation Method to Obtain Both Epidermal and Dermal Stem Cells from Human Skin Specimen Hua Qian, Xue Leng, Jie Wen, Qian Zhou, Xin Xu, and Xunwei Wu Abstract Stem cells play a crucial role in maintaining and repairing tissues during homeostasis and following injury. The efficient procurement of high quantity and quality of skin stem cells is important for both laboratory studies and clinical applications. Here, we describe a one-step isolation procedure to efficiently obtain both epidermal and dermal cell population from human skin specimen, based on the different influence of the Rho kinase inhibitor Y27632 on the growth of epidermal and dermal cells during the initial culture. Compared with the conventional methods, our protocol shows that it is simpler and less time consuming and can efficiently obtain the high quality of skin stem cells and can maintain the stem cell features after culture expansion. Keywords Adult, Culture, Dermal fibroblasts, Epidermal stem cells, Human, Y-27632

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Introduction Primary epidermal and dermal cells have been widely used for both laboratory researches to study skin diseases and clinical applications to repair damaged skin [1–3]; therefore, it has been always important to efficiently isolate primary cells from the skin specimen and to maintain their stem cell potential after culture expansion [1–4]. The conventional method to isolate primary skin cells from the whole skin involves a two-step enzymatic digestion [5]. Briefly, the tissue is firstly digested with dispase for separation of the epidermis from the dermis; then, the separated epidermis is digested with trypsin to obtain dissociated epidermal cells and the dermis is digested by type I collagenase to obtain dermal cells. This method works nicely for neonatal foreskin tissues but has given low cell recovery rate, and reduced cell viability when isolating cells from adult skin tissues. We recently developed a simple isolation procedure to obtain both epidermal and dermal cell population at the same time by applying Y-27632, an inhibitor of Rho-associated protein kinase (ROCK), into the inoculation medium [6–8]

Hua Qian and Xue Leng contributed equally to this work.

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Summary of the isolation procedure Homogenizing-15min

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Fig. 1 One-step isolation procedure to obtain both epidermal and dermal cells. The protocol does not require the separation procedure of epidermis from dermis, and the skin specimen is directly homogenized and digested with mixture of enzymes to obtain the dissociated cells. After inoculation, the dissociated cells with 2-day treatment of Y2762 grow as epidermal cells; the other part without Y27632 treatment grows dermal cells

(summarized in Fig. 1). Our new method is not only to efficiently procure substantial cells from the isolation but also maintain the stem cell features of culture-expanded skin cells, indicated by highlevel integrin expression and the ability to form colony for primary epidermal cells, and multi-differentiation potential for dermal cells. Notably, our method does not require a separation of epidermis from dermis and will be suitable to acquire cells from different types of skin tissues [7]. Here, we document the protocol including isolation and culture procedure in detail. We believe that our new protocol will be very useful to obtain primary skin stem cells from the adult human tissues.

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Materials

2.1 Materials for Skin Stem Cell Isolation and Culture

1. Adult human foreskin [The procedure for obtaining human foreskin tissue from discarded hospital specimens without any personal identity information was approved by the Medical Ethical Committee of the School of Stomatology Shandong University (No. 2015120401, Date: 12-05-2015)] 2. Disinfection solution: 70% ethanol, 2 antibiotics PBS: 1% penicillin (100 U/ml) and streptomycin (100 mg/l) in PBS 3. Enzyme mixture: 2.5 mg/ml Dispase and 2.5 mg/ml type I collagenase in DMEM 4. Digestion solution: 0.05% and 0.25% Trypsin, Dnase I (10 mg/ml) in H2O 5. Neutralization solution: 10% FBS in DMEM with 1% penicillin/streptomycin 6. Inoculation medium (Growth factors containing medium, the so-called G-medium): DMEM/F12 (3:1) containing 1% penicillin/streptomycin, 40 μg/ml Fungizone, 40 ng/ml FGF2,

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20 ng/ml EGF, and 2% B27 supplement. This medium is also used for dermal cell culture 7. Y-27632, 10 mM in distilled water 8. Freezing medium: ice cold FBS with 10% DMSO 9. Epidermal cell culture medium: CNT07 (CELLnTEC) plus 0.1% penicillin/streptomycin 10. Materials for tissue processing: Forceps, autoclaved scalpel holder, sterile blades, 50 ml falcon tube, 100 μm cell strainers, and 100 mm tissue culture dishes 2.2 Materials for Cell Characterization

1. Osteogenic differentiation medium (10% FBS in DMEM supplemented with 10 nM dexamethasone, 10 mM β-glycerophosphate, and 50 mg/ml ascorbate phosphate) 2. Adipogenic differentiation medium [10% FBS in DMEM supplemented with 1 mM dexamethasone, 0.2 mM indomethacin, 0.01 mg/ml insulin, and 0.5 mM isobutyl-methylxanthine (IBMX)] 3. Myogenic differentiation medium (10% horse serum–DMEM supplemented with 10 μM 5-Aza-CR) 4. Antibodies: Mouse monoclonal anti-Integrin alpha 6 [MP 4F10] (ab20142, Abcam); Rabbit polyclonal anticalponin (ab46794, Abcam); Rabbit polyclonal anti-osteocalcin(ab93876, Abcam); and Rabbit polyclonal anti-FABP4 (D25B3) XP (#3544S,CST) 5. Flow cytometry

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Methods

3.1 Tissue Processing

1. Collect a fresh adult foreskin tissue, which could be kept in fridge for no more than 72 h (see Note 1). 2. Wash the skin tissue with PBS once and weigh the tissue. 3. Rinse skin tissue with 70% ethanol for 30 s for disinfection. 4. Incubate tissue with 2 antibiotics cold PBS for 2 times to wash the ethanol out and for further disinfection, 5 min for each time. 5. Transfer the disinfected tissue to another sterile dish and homogenize the skin tissue thoroughly by using scalpel blades (see Note 2).

3.2

Tissue Digestion

1. Transfer the homogenized skin tissue into a 50-ml falcon tube and add the enzyme mixture into the tube, 10 ml enzyme mixture for 1 g tissue.

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2. Incubate the tissue in a 37  C water bath with shaking at the speed of 80 g/min for 1.5 h (see Note 3). 3. Add 1/5 volume of 0.25% trypsin into the digestion solution and incubate for another 30 min with shaking (see Note 4). 4. Add Dnase I solution (10 mg/ml) to the mixture at 1:100 ratio and incubate for another 5 min at 37  C with shaking (see Note 5). 5. Add equal volume of neutralization medium to the digestion solution and mix well to stop the digestion. 6. Pipette the solution up and down for about 20 times by using 10 ml serological pipette to obtain dissociated cells (see Note 6). 3.3 Dissociated Cells Collection and Primary Cell Culture

1. Filter the dissociated-cell solution through a 100-μm cell strainer into a new 50-ml falcon tube (see Note 7). 2. Centrifuge at 1000 r.p.m. for 5 min at room temperature to collect the cell pellet. 3. Remove the supernatant from the tube, wash the cell pellet with the neutralization medium once, then resuspend the cells with 20 ml of inoculation medium and count the cells. 4. Divide the cells equally and inoculate into two 100 mm culture dishes. 5. Add 10 μM Y-27632 into one dish, the other dish without adding Y-27632 (see Note 8). 6. Culture cells at 37  C incubator with 5% CO2. 7. Two days later, the dish with Y-27632 is replaced with CNT07 medium, which is for the epidermal stem cell culture. Change the medium every 2 days till the cells reach 70–80% confluency. The image of primary epidermal stem cells at day 2 and day 7 after inoculation is shown Fig. 2a. 8. The other dish without adding Y-27632 is used for the dermal stem cell culture, and the medium will be changed at day 5 with the same inoculation medium, and change the medium every 5 days. The image of primary dermal stem cells at day 2 and day 7 after inoculation is shown Fig. 2b.

3.4 Passaging and Freezing Cells

1. Passage when primary epidermal stem cells reach 80% confluence (usually around day 7 after inoculation) (see Note 9).

3.4.1 Epidermal Cell Passaging

2. Remove the medium, wash by cold PBS, add 2 ml 0.05% trypsin (for one 100 mm culture dish), and incubate the cells in the incubator (37  C and 5% CO2) for 2 min (see Note 10). 3. Aspirate 2 ml trypsin and wash the cells with 5 ml PBS once. 4. Add another 2 ml 0.05% trypsin for another 6–10 min at 37  C incubator (see Note 11).

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Fig. 2 The representative images of epidermal and dermal cells of primary culture (passage 0). (a) The image of epidermal cells at day 2 and day 7 after inoculation; (b) The image of dermal cells at day 2 and day 7 after inoculation

5. Stop the digestion by adding 8 ml neutralization medium. 6. Collect cells and centrifuge at 1000 rpm for 5 min. 7. Remove the supernatants slowly; resuspend the cell pellet with CNT07 medium. 8. Count cells and seed 1  106 cells into one 100 mm dish. 3.4.2 Dermal Cell Passaging

Follow the standard passage protocol, nothing is special for passaging the dermal cells, except using the trypsin concentration 0.05%.

3.4.3 Cell Freezing

Follow the standard cell freezing protocol. Both epidermal and dermal stem cells can be frozen with the freezing medium. The frozen cells can be kept in liquid nitrogen for long-term use.

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3.5 Characterization of Cultured Skin Stem Cells

The colony formation ability is a gold standard to evaluate the characterization of epidermal stem cells, and holocloneforming cells possess epidermal stem cell features such as self-renewing and regeneration potential [9–11]. The epidermal cells prepared from this protocol trend to grow as colony morphology, and 75% of passage 3 epidermal stem cells will be able to form holoclones (arrows, Fig. 3a). In the skin epidermis, only the basal cells express integrins, and the epidermal stem cells usually express high level of integrins α6 and β1, which are well-recognized markers for epidermal stem

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Fig. 3 The culture-expanded epidermal cells maintain the stem cell features. (a) The colony formation analysis of passage 1 (P1) and passage 3 (P3) epidermal cells. Both P1 and P3 cells were able to form holoclones (arrows). (b) FACS analysis of a6 integrin (FITC-A) expression. The left panel is negative control without antibody

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cells [1, 9, 11]. FACS analysis revealed that more than 90% of passage 3 epidermal cells express α6 integrin (see Fig. 3b). 2. Multiple-differentiation ability of dermal cells The multiple-differentiation ability is another important feature of stem cells [12, 13]. To test whether our cultured dermal cells have stem cell potential, we induced cultured dermal cells into different lineages with different condition media. The passage 3 dermal cells (left panel, Fig. 4) were seeded at 5.0–8.0  104 cells/well in a 24-well plate. When the cells reached 80% confluency with growth medium, we changed the

Fig. 4 The culture-expanded dermal cells possess multi-differentiation ability. (a) The representative image of passage 3 dermal cells. (b–d) The immunofluorescence analysis of differentiation markers of different lineage cells induced from passage 3 dermal cells after cultured with corresponding condition media. Osteocalcin for osteoblast (red, b), FABP4 for adipocytes (green, c) ,and calponin for myocytes (red, d). DAPI (blue) for nuclear staining

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culture medium to the following conditioned media: the osteogenic-inducing medium for osteocytes, adipogenic medium for adipocytes, and myogenic differentiation media for myocytes. Immunofluorescence staining of differentiation markers were performed, and our culture-expanded dermal cells at passage 3 can be induced to differentiate into three different lineages (see Fig. 4), suggesting that they maintain the stem cell potential after expansion. 3. The cultured skin stem cells are able to regenerate a normal human skin in vivo. For this characterization, please see our previous publication [7].

4

Notes 1. The skin tissue, collected from clinics, should be avoided from strong shaking and contamination during the transportation, and it should be always kept in cold DMEM or PBS before used for isolating cells. We have tested that the skin tissue can be stored in fridge up to 72 h without significantly affecting the cell viability. In this protocol, we used the foreskin tissue as an example to demonstrate the isolation and culture procedure, and all data in figures are from analysis of the skin cells derived from the foreskin tissues. We found that this protocol also worked well for isolating cells from other type skin tissues such as scalp tissue and abdominal tissues [7]. 2. The sufficient homogenization is crucial for obtaining high recovery of dissociated skin cells. Usually, the homogenizing procedure of 1 g tissue takes 10–15 min. In order to prevent the tissue from drying during homogenization, add 200 μl PBS every 5 min. 3. The incubation time for enzymatic digestion of tissue depends on how the tissue was well homogenized. Usually, this step takes one and half hours. Manually mix cells well every half hour. The incubation is completed when there are no major pieces of the tissues to be seen by eyes and the solution looks translucent. 4. After adding the trypsin into the digestion solution, the incubation time should stick to the half hour; otherwise, the viability of cells will clearly decrease. 5. Deoxyribonuclease I (DNase I) is used for degradation of nucleic acids that leak into the digestion solution, which could increase viscosity of digested mixture. DNase I will not damage intact cells.

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6. Pipetting is another crucial step to affect the recovery of dissociated cells. 10 ml serological pipette is recommended, and roughly pipetting is avoided. 7. To get the maximum recovery of dissociated cells, we recommend that one cell strainer be used for filtration of no more than 20 ml cell solution. 8. Y-27632 has been shown to enhance epidermal cell growth and inhibit dermal cell growth, and 2–3 days of Y-27632 treatment are sufficient to selectively obtain the epidermal stem cell growth [7]. 9. To maintain the undifferentiation status of epidermal stem cells, it is important to passage the epidermal cells when it reaches 70-80% confluency. 10. This short-time trypsin step is to remove the contaminated dermal cells. The trypsin time should be no more than 3 min in order to avoid detaching any epidermal cells. After this step, we could get pure epidermal cells with little dermal cell contamination at passage 1 [7]. 11. Always use 0.05% trypsin to passage the epidermal cells and never use 0.25% trypsin. The digestion time of trypsin should never be more than 10 min.

Acknowledgement This work was supported by National Key Research and Development Program of China (2017YFA0104604), General Program of National Natural Science Foundation of China (81772093), and Shandong Taishan Scholar Award (tshw201502065). References 1. Prodinger CM, Reichelt J, Bauer JW, Laimer M (2017) Current and future perspectives of stem cell therapy in dermatology. Ann Dermatol 29 (6):667–687 2. Gonzales KAU, Fuchs E (2017) Skin and its regenerative powers: an alliance between stem cells and their niche. Dev Cell 43(4):387–401. https://doi.org/10.1016/j.devcel.10.001 3. Hirsch T, Rothoeft T, Teig N, Bauer JW, Pellegrini G, De Rosa L, Scaglione D, Reichelt J, Klausegger A, Kneisz D, Romano O, Secone Seconetti A, Contin R, Enzo E, Jurman I, Carulli S, Jacobsen F, Luecke T, Lehnhardt M, Fischer M, Kueckelhaus M, Quaglino D, Morgante M, Bicciato S, Bondanza S, De Luca M (2017)

Regeneration of the entire human epidermis using transgenic stem cells. Nature 551 (7680):327–332 4. Terunuma A, Limgala RP, Park CJ, Choudhary I, Vogel JC (2010) Efficient procurement of epithelial stem cells from human tissue specimens using a rho-associated protein kinase inhibitor Y-27632. Tissue Eng A 16 (4):1363–1368. https://doi.org/10.1089/ ten.TEA.2009.0339 5. Aasen T, Izpisua Belmonte JC (2010) Isolation and cultivation of human keratinocytes from skin or plucked hair for the generation of induced pluripotent stem cells. Nat Protoc 5 (2):371–382. https://doi.org/10.1038/ nprot.2009.241

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6. Zou D, Pan J, Zhang P, Wu X (2016) A new method to isolate human epidermal keratinocytes. J Clin Dermatol 6:424–429. (in Chinese) 7. Wen J, Zu T, Zhou Q, Leng X, Wu X (2017) Y-27632 simplifies the isolation procedure of human primary epidermal cells by selectively blocking focal adhesion of dermal cells. J Tissue Eng Regen Med. https://doi.org/10. 1002/term.2526 8. Zhou Q, Zhang Q, Leng X, Zu T, Qin J, Wen J, Wu X (2017) Culture-expanded adult dermal cells derived from scalp tissue are multidifferentiation potential. Smart healthcare, 3 (3):1-8 DOI:https://doi.org/10.19335/j. cnki.2096., (in chinese) 9. Watt FM (1998) Epidermal stem cells: markers, patterning and the control of stem cell fate. Philos Trans R Soc Lond B Biol Sci 353 (1370):831–837. https://doi.org/10.1098/ rstb.1998.0247

10. Barrandon Y, Green H (1987) Three clonal types of keratinocyte with different capacities for multiplication. Proc Natl Acad Sci U S A 84 (8):2302–2306 11. Chen S, Lewallen M, Xie T (2013) Adhesion in the stem cell niche: biological roles and regulation. Development 140(2):255–265. https:// doi.org/10.1242/dev.083139 12. Bartsch G, Yoo JJ, De Coppi P, Siddiqui MM, Schuch G, Pohl HG, Fuhr J, Perin L, Soker S, Atala A (2005) Propagation, expansion, and multilineage differentiation of human somatic stem cells from dermal progenitors. Stem Cells Dev 14(3):337–348. https://doi.org/10. 1089/scd.2005.14.337 13. Chen FG, Zhang WJ, Bi D, Liu W, Wei X, Chen FF, Zhu L, Cui L, Cao Y (2007) Clonal analysis of nestin( ) vimentin(+) multipotent fibroblasts isolated from human dermis. J Cell Sci 120(Pt 16):2875–2883. https://doi.org/ 10.1242/jcs.03478

Methods in Molecular Biology (2019) 1879: 149–152 DOI 10.1007/7651_2018_141 © Springer Science+Business Media New York 2018 Published online: 08 May 2018

Isolation and Cultivation of Skin-Derived Precursors Xiaoxiao Wang, Shiyang Dong, and Yaojiong Wu Abstract Skin-derived precursors (SKPs) have been shown recently to be capable of inducing hair genesis and hair follicle regeneration. Here, we describe a protocol for SKP isolation and culture based on our experience and previous publications. Keywords Cultivation, Isolation, Skin-derived precursor, SKPs

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Introduction The formation of hair follicles relies on signals derived from the dermis during skin morphogenesis and regeneration number [1, 2]. Previous studies indicate that dermal papilla (DP) cells in the hair follicle are able to induce hair genesis [3]. However, the application of DP cells is limited by their availability. Recently, multipotent skin-derived precursors (SKPs) have been isolated from the dermis of adult skin of different anatomic sites and shown to be capable of inducing hair genesis and hair follicle regeneration [4–7], implying a potential application as induction cells in hair genesis and skin bioengineering.

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Materials

2.1 Reagent Preparation

1. Phosphate buffer saline (PBS). 2. DPBS (Corning). 3. Saline. 4. 70% ethanol. 5. 2% (W/V) iodine solution. 6. 100 Penicillin/streptomycin solution (Gibco). 7. Gentamycin. 8. SKP growth medium: DMEM: F12, 3:1 mixture (Corning), supplemented with 20 μl/ml B27, 40 ng/ml basic fibroblast

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growth factor (bFGF), 20 ng/ml epidermal growth factor (EGF), and 1% penicillin/streptomycin solution. 9. RPMI 1640 medium (Gibco). 10. TripLE™ Express (Gibco). 11. EGF (PeproTech): stock solution is prepared as 100 ng/μl EGF in sterilized diluent (0.1% BSA in PBS), which is aliquoted into PCR tubes and stored at 20 to 80  C. 12. bFGF (PeproTech): stock solution is prepared as 100 ng/μl EGF in sterilized diluent (0.1% BSA in PBS), which is aliquoted into PCR tubes and stored at 20 to 80  C. 13. B27 (Gibco). 14. Collagenase I (Sigma): powder stored at 20  C and dissolved into right concentration using PBS, then filter sterilizing using a syringe filter (0.22 μm). 15. Dispase II (Sigma): power stored at 4  C and dissolved into right concentration using PBS, then filter sterilizing using a syringe filter (0.22 μm). 2.2 Equipment Preparation

1. 0.22 μm filter (Millipore). 2. Hemocytometer. 3. 15 and 50 ml polypropylene centrifuge tube (Corning). 4. 100-mm petri dishes (non-treated) (JET BIOFIL). 5. 6-Well plate. 6. Surgical scissors. 7. Forceps. 8. Pipettes. 9. Centrifuge. 10. Tissue culture incubator. 11. 80 mesh filter (Utah biodiesel supply). 12. Syringes.

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Methods

3.1 Isolation and Culture of Human Skin Divided Precursors

1. Collect human scalp or foreskin tissues (in full thickness) and store in serum-free RPMI 1640 medium containing 5 μg/ml gentamycin at 4  C (see Note 1). 2. In a 6-well plate, add to five wells, respectively, 5 ml 2% iodine solution, 5 ml of 70% ethanol, and three wells of 5 ml DPBS without Ca or Mg containing 10 μg/ml gentamicin. 3. Briefly submerge the skin sample in iodine solution, 70% ethanol, then the DPBS without Ca or Mg for a few seconds each,

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and then submerge in each of the three gentamicin solutions, each for 15 min (see Note 2). 4. Cut skin tissues into halves or quarters, depending on the size of the tissue, and transfer the pieces, dermis side down, to a petri dish containing 25 U/ml (3 mg/ml) dispase II and 10 μg/ml gentamicin in DMEM: F12, 3:1 mixture (without supplements) and incubate 8–14 h at 4  C. 5. Using suitable forceps, gently peel away the epidermis from the dermis (see Note 3). 6. Cut the dermis into pieces as small as possible using a pair of scissors. 7. Add 5–10 ml 1% collagenase I to the tissue and incubate for 1–2 h at 37  C (see Note 4). 8. Filter the cell suspension through an 80 mesh filter. Collect cells by centrifuging the suspension at 400  g for 5 min at room temperature (see Note 5). 9. Remove the supernatant, and resuspend the pellet with SKP growth medium. 10. Count cells using a hemocytometer. Seed 6–10  105 cells in 10 ml SKP growth medium into a 10-cm Petri dish. Incubate cells in a tissue culture incubator with 5% CO2 at 37  C. 11. Add cytokines (bFGF at 40 ng/ml and EGF at 20 ng/ml) to the culture twice a week. 12. Cells grow as spheres in suspension. When spheres reach ~100 μm, it is time to split them. 13. Collect cells and the culture medium into a 15-ml centrifuge tube. Pellet cells by centrifuging at 400  g for 5 min at room temperature. Keep the supernatant as conditional medium for subsequent culture use. 14. Resuspend the cell pellet in 0.5 ml TripLE™ Express and incubate for ~2 min in a 37  C water bath. Separate cell aggregates by gentle pipetting. 15. Add DMEM: F12, 3:1 mixture (without supplements) to the cells immediately when cells are separated and centrifuge the cell suspension at 400  g for 5 min at room temperature to pellet cells. 16. Discard the supernatant and resuspend cells in 75% fresh SKP growth medium and 25% conditional medium from earlier culture. Seed cells into F127-coated Petri dishes and incubate cells in an incubator with 5% CO2 at 37  C. 17. Repeat steps 11 and 12.

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Notes 1. For isolation of SKPs from newborn mouse skin, make a small incision in the head. Insert scissors to the created incision to dissociate the dorsal skin gently. 2. If skin sample is too large, before submerging, cut into smaller pieces (0.5–1 cm wide) with a surgical scalpel and also remove subcutaneous fat by scraping with the scalpel. For small or thin samples, the iodine and ethanol steps may be too stringent; an alternative is to soak the sample for 10 min in RPMI 1640 medium containing ten times the normal concentrations of penicillin and streptomycin. 3. Skin samples from neonatal mice are fragile and delicate. Thus samples should be treated gently with forceps. 4. Use 0.3% collagenase I for dermal tissues from neonatal mice and incubate for 30–40 min at 37  C. 5. When tissues are sufficiently digested, there are no large pieces left over on the filter after filtration.

References 1. Ito M et al (2007) Wnt-dependent de novo hair follicle regeneration in adult mouse skin after wounding. Nature 447(7142):316–320 2. Morris RJ et al (2004) Capturing and profiling adult hair follicle stem cells. Nat Biotechnol 22:411–417 3. Jahoda CA et al (1984) Induction of hair growth by implantation of cultured dermal papilla cells. Nature 311:560–562 4. Biernaskie J et al (2009) SKPs derive from hair follicle precursors and exhibit properties of adult dermal stem cells. Cell Stem Cell 5(6):610–623

5. Wang X et al (2016) Hair follicle and sebaceous gland de novo regeneration with cultured epidermal stem cells and skin-derived precursors. Stem Cells Transl Med 5(12):1695–1706 6. Toma JG et al (2001) Isolation of multipotent adult stem cells from the dermis of mammalian skin. Nat Cell Biol 3:778–784 7. Toma JG et al (2005) Isolation and characterization of multipotent skin-derived precursors from human skin. Stem Cells 23:727–737

Methods in Molecular Biology (2019) 1879: 153–163 DOI 10.1007/7651_2018_151 © Springer Science+Business Media New York 2018 Published online: 11 October 2018

Magnetic-Based Cell Isolation Technique for the Selection of Stem Cells Petek Korkusuz, Sevil Ko¨se, Nilgu¨n Yersal, and Selin O¨nen Abstract Magnetic-activated cell sorting (MACS) is the technology that is recently used as a magnetic-based cell isolation/purification technique. This technique enables the isolation and selection of germ, hematopoietic, and somatic stem cells including skin stem cells (SkSCs). Here, we have tried to describe the isolation of stem cells by MACS using CD34 antigen for SkSCs, again CD34 for hematopoietic stem cells (HSCs) and Thy-1 for spermatogonial stem cells (SpSCs). MACS allowed the isolation of CD34+, CD34+, and Thy-1+ human SkSCs, HSCs, and SpSCs with minimum 98% purity. Keywords Hematopoietic stem cell, MACS, Skin stem cell, Spermatogonial stem cell

1

Introduction Magnetic-activated cell sorting (MACS) uses magnetic particles to isolate the cell subsets which are defined [1]. Several types of paramagnetic beads have been developed and made commercially available for usage in cell sorting purposes. Between those, the most widely used ones are the MACS® beads (nm size; Miltenyi Biotec, Germany) [2]. MACS system uses particles consisting of polysaccharide and iron oxide conjugated to antibodies and it is based on high-gradient magnetic cell sorting. MACS has been extensively and preferably used for the isolation of rare cells and/or the large cell numbers for both research and clinical applications [3]. The flow rate depends on the size of the hole at the base of the column and also it can be governed by an attached needle which is depending on the type of column. Because of the paramagnetic iron particles bound to the antibodies, the technique requires a highgradient magnetic cell separation column filled with ferromagnetic steel wool matrix to retain the labeled cells in a magnetic field generated by an external magnet. When the magnetic field is turned off (separation column is discarded from magnetic field), the matrix can no longer retain the labeled cells, which can then be eluted and isolated from heterogeneous suspension [4] (Fig. 1).

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Fig. 1 The schematic view of basic magnetic-activated cell sorting (MACS) technology positive (A) and negative cell selection steps (B) consisting of MACS conjugated antibodies, a high-gradient magnetic separation column filled with ferromagnetic stainless steel wool and a strong, external permanent magnet

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A removal reagent for the beads is available for using in the removal of MACS beads enzymatically. It allows the cells to be relabeled with another marker and sorted again. Even so, the positive selection is an efficient way to a fast isolation of the cells with high purity and yield, and negative cell selection or positive selection with appropriate combination of antibodies is highly preferred by researchers. In immune-magnetic sorting, the cell doublets which contain only the desired phenotype can be retained and those that contain unwanted cells can be removed at the same time by using a double sorting method. In first step, the unwanted cell type should be labeled and removed, and in second step the unlabeled target cells should be labeled and positively sorted.

2

Materials Prepare all solutions with ultrapure water (should be prepared by purifying the deionized water) and cell culture or analytical grade reagents. Prepare all reagents at room temperature and store in accordance with their own storage conditions. Diligently follow all the waste disposal regulations when disposing waste materials. We do not add sodium azide to reagents but we sterilize them by filtration using 0.2 mm membrane filter in general.

2.1 Materials for Skin Stem Cells

Dulbecco’s modified Eagle medium (DMEM) (Gibco Life Technologies, Gaithersburg, MD, USA) Antimycotic agents: 15 U/ml penicillin, 15 mg/ml streptomycin, 30 μg/ml neomycin (Sigma, St Louis, MO, USA), and 100 μg/ml Normosin™ (InvivoGen, San Diego, CA, USA) Accumax™: 2% (Chemicon International Inc., CA, USA) Dispase: 0.4 mg/ml (Sigma) FBS (Gibco 10500): Heat inactivated fetal bovine serum, ready to use. Aliquots are stored at 20  C Antibiotics (Hyclone SV30010): 10,000 lin–10,000 μg/ml of streptomycin

U/ml

penicil-

HBSS (Sigma H6648): Hank’s balanced salt solution, ready to use Collagenase type IA: 2 mg/ml; Sigma Medium 199/Ham’s F12 nutrient mixture (M199/F12): 1:1, Sigma Human EGF: 10 ng/ml, Sigma Human FGF: 20 ng/ml, Sigma Human insulin: 5 mg/ml, Sigma Hydrocortisone: 0.5 mg/ml, Sigma Human transferrin: 2 mg/ml, Sigma Bovine serum albumin: 125 μg/ml, Sigma

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Defined keratinocytes serum-free medium: Gibco Life Technologies Trypsin–EDTA (Thermo Fisher Scientific, Gibco™, 25200056): Trypsin–EDTA (0.25%), phenol red, ready to use 2.2 Materials for Hematopoietic Stem Cells

HBSS (Sigma H6648): Hank’s balanced salt solution, ready to use

2.3 Materials for Spermatogonial Stem Cells

A. Reagent for Isolation of Spermatogonial Stem Cells (SpSCs)

Ficoll-Paque media: Biocoll (#L6113/5, Merck Millipore, Germany)

HBSS (Sigma H6648): Hank’s balanced salt solution, ready to use PBS (Calbiochem 524620): Dissolve one tablet in 1 L of deionized water. Store at 4  C DNase (Sigma DN25): DNase I solution is prepared at 7 mg/ml in HBSS and sterilized by filtration using 0.22 μm membrane filter Trypsin–EDTA (Thermo Fisher Scientific, Gibco™, 25200056): Trypsin–EDTA (0.25%), phenol red, ready to use Antibiotics (Hyclone SV30010): 10,000 lin–10,000 μg/ml of streptomycin

U/ml

penicil-

FBS (Gibco 10500): Heat inactivated fetal bovine serum, ready to use. Aliquots are stored at 20  C Sodium pyruvate (Sigma P2256): Stock solution is prepared at 100 mM sodium pyruvate in DW, sterilized by filtration using 0.22 μm membrane filter, and stored at 2–8  C L-Glutamine

(Hyclone SH30034.01): 200 mM (100 solution). Aliquots are stored at 20  C

D-(+)-Glucose

(Sigma 6152): Store at room temperature

Percoll solution (Nidacon, PureSperm 40/80): 30% Percoll solution is prepared in PBS containing 1% FBS, 50 U/ml penicillin, and 50 mg/ml streptomycin. Percoll solution is sterilized by filtration using a 0.22-μm membrane filter and stored at 4  C Thy1.2 Microbeads, mouse (Miltenyi Biotec, 130-049-101) B. Stock Solution of Reagents for Spermatogonial Stem Cells (SpSCs) Culture BSA (Sigma A3803): Albumin from bovine serum HEPES (Sigma H0887): 1 M, pH 7.0–7.6, ready to use and store at 4 C Holo-Transferrin (Sigma T1283): Stock solution is prepared at 10 mg/ml in PBS and stored at 20  C Na2SeO3 (Sigma 21, 448-5): Stock solution is prepared at 3  104 M in DW and stored at 20  C

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2-ME (Sigma M7522): 100 mM solution is prepared freshly each time medium is made Insulin (Sigma I5500): Stock solution is prepared at 10 mg/ml in 10 mM HCl and stored at 20  C Putrescine (Sigma P5780): Stock solution is prepared at 100 mM in PBS and stored at 20  C Recombinant human GDNF (BioLegend, 711002): Stock solution is prepared at 5 μg/ml in PBS containing 0.1% BSA and stored at 70  C Recombinant rat GFRa1/Fc chimera (R&D Systems, 560-GR-100): Stock solution is prepared at 50 μg/ml in PBS containing 0.1% BSA and stored at 70  C Recombinant human FGF-basic (BioLegend, 710304): Stock solution is prepared at 10 μg/ml in PBS containing 0.1% BSA and stored at 70  C C. Reagents for STO Feeder Layers Medium for STO cells: Dulbecco’s modified Eagle’s medium, high glucose (DMEM; Gibco/Invitrogen 5796) supplemented with 10% FBS, 2 mM glutamine, 50 U/ml penicillin, and 50 mg/ml streptomycin. SSM/STO is sterilized by filtration using a 0.22μm membrane filter and stored at 4  C. Mitomycin C (Sigma M4287): Stock Mitomycin C solution is prepared at 200 mg/ml in PBS and stored at 70  C. Gelatin (Sigma G2500): 0.1% Gelatin in DW is autoclaved for sterilization and stored at room temperature. 2.4 Materials for Magnetic-Activated Cell Sorting Procedure

PBS-M: PBS is supplemented with 1% FBS, 10 mM HEPES, 1 mg/ ml glucose, 1 mM pyruvate, 50 U/ml penicillin, and 50 μg/ml of streptomycin. Thy1.2 Microbeads, mouse (Miltenyi Biotec, 130-049-101): Ready to use. Store at 4  C. CD 34 Microbeads, human (Miltenyi Biotec, 130-046-702): Ready to use. Store at 4  C.

3

Methods

3.1 Isolation of Skin Stem Cells from Human Scalp

1. Transport the tissue sample to the laboratory in DMEM containing antimycotic agents. 2. Rinse the tissue in HBBS for the removal of blood and debris and place it in a 100-mm plastic tissue culture dish (Nunc, Rochester, NY, USA). Clear connective tissue and fat tissue (see Note 1).

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3. Mince the rest of the tissue with sterile scalpels into 1-2 mm fragments and digest with Accumax™ and Dispase in DMEM containing 10% FBS plus antibiotic and antimycotic agents for 30 min at 37  C on a shaking incubator. 4. Wash the tissue with sterile HBBS, digest with collagenase type IA in DMEM with 10% FBS plus antibiotic and antimycotic agents for 30 min at 37  C on a shaking incubator. 5. Pipette vigorously the tissue digest after gentle centrifugation (2500  g, 5 min), remove the supernatant, resuspend the pellet in medium 199/Ham’s F12 (M199/F12; 1:1) supplemented with human EGF, FGF, insulin, hydrocortisone, human transferrin, bovine serum albumin, 10% FBS, and antibiotic and antimycotic agents, and distribute on a 25-cm2 flask (see Note 2). 6. After 3 days, change the culture medium of the hair follicle cells attached on the flask to serum-free medium (without FBS) and transfer the suspended cells on the flask to 75 cm2 flask with complete culture medium. 7. Change the medium to serum-free medium (without FBS) after 3 days and place it for 14 days at 37  C in a humidified atmosphere containing 5% CO2. 8. Carefully remove each culture medium every 3 days and replace with fresh culture medium. 9. After 7 days in culture, change all culture media to defined keratinocytes serum-free medium (Gibco Life Technologies). 10. After 2 weeks in primary culture, collect the cells by incubation with trypsin–EDTA solution, for 5 min at 37  C. 11. During the incubation period, the release of clusters of cells from the hair follicles has occurred. Harvest the cells by centrifugation at 300  g at 4  C for 10 min. 3.2 Isolation of Mononuclear Cells from Bone Marrow

1. Add 2 ml of defibrinated or anticoagulant treated bone marrow and the same volume of HBSS into a 15-ml-volumed centrifuge tube (final volume 4 ml) (see Note 3). 2. By the inversion of the tube several times or drawing the mixture in and out of a pipette, mix the blood and buffer (see Note 3). 3. Add 3 ml of Ficoll-Paque media to the centrifuge tube. 4. Layer the 4 ml of diluted blood sample onto the Ficoll-Paque media solution carefully (see Note 4). 5. Centrifuge the tube at 400  g for 40 min at 20  C and brake should be turned off (see Note 5).

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6. Discard the upper layer which contains plasma and platelets by using a sterile pipette, without disturbing the mononuclear cell layer at the interface (see Note 6). 7. Transfer the mononuclear cell layer into a new sterile centrifuge tube by using a sterile pipette. 8. Estimate the volume of the mononuclear cells that transferred into the new tube and add at least 3 volumes (~6 ml) of HBSS to the mononuclear cells in the centrifuge tube. 9. Suspend the cells by gently drawing them in and out of a pipette. 10. Centrifuge the tube at 400  g for 10 min at 20  C. Then, remove the supernatant (see Note 7). 11. Resuspend the mononuclear cells in 8 ml HBSS. 12. Centrifuge at 400  g for 10 min at 20  C. After the centrifuge, remove the supernatant. 13. Resuspend the cell pellet for the MACS. 3.3 Isolation of Spermatogonial Stem Cells

1. Remove testes from 5 C57BL/6 pups using sterilized forceps and scissors and collect the testes in a 50-mm sterile petri dish containing 5 ml of ice-cold HBSS. SpSCs appear since after birth, between 0 and 6 days. Meiosis of primary spermatocytes begins on days 8–10, thus 6-day-old mice testis is suitable to isolate SpSCs (Fig. 2). 2. Transfer the testes to another petri dish containing ice-cold HBSS and remove tunica albuginea under a dissecting microscope using forceps. 3. Transfer the seminiferous tubules to a 15-ml polypropylene conical centrifuge tube (Kırgen KG2611) containing room temperature 0.5 ml of 7 mg/ml DNase I solution and room temperature 4.5 ml of trypsin–EDTA (0.25%). 4. Pipette up and down using p1000 pipette to break the aggregated seminiferous tubules and incubate at 37  C for 5 min. 5. After pipetting with p1000 pipette several times, incubate the tube at 37  C for an additional 3 min in order to obtain single cell suspension. 6. Stop digestion by adding 0.7 ml of fetal bovine serum (FBS) and add 0.5 ml of 7 mg/ml DNase to digest genomic DNA from dead cells. 7. Pipet digested cells with p1000 pipette several times. If cells remain clumped, they can be digested by adding 0.5 ml of 7 mg/ml DNase. 8. Filter the cell suspension through a 40-μm pore nylon cell strainer (Falcon) to eliminate clumps and debris and rinse the cell strainer with HBSS.

Fig. 2 Isolation, separation, and fluorescent labeling of spermatogonial stem cells (SpSCs) from 6-day-old C57BL/6 mouse testis. The isolation protocol consists of four steps: removal of testes from mouse (A), digestion of pup testes (B), Percoll gradient (C), and MACS separation (D). SpSCs are 100% immunolabeled with Thy1.2 (E, F) 400 (E) and 630 (F) SpSC colonies from 6-day-old mouse on day 5 (G) and on day 10 (H) of culture

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9. Centrifuge the cell suspension at 600  g for 7 min at 4  C. 10. Remove supernatant carefully and add 10 ml of PBS to resuspend cells (PBS is supplemented with 1% FBS, 10 mM HEPES, 1 mg/ml glucose, 1 mM pyruvate, 50 U/ml penicillin, and 50 μg/ml of streptomycin). Count cell number. 11. To remove debris and erythrocytes, gently overlay 5 ml of cell suspension onto 2 ml of 30% Percoll solution in a 15-ml polypropylene conical centrifuge tube. 12. Centrifuge at 600  g for 7 min at 4  C with the brake off. 13. Carefully collect the cells and debris at the interface between the cell suspension and the 30% Percoll solution. Then, remove all aqueous phases containing PBS and 30% Percoll solution. Leave the pellet at the bottom of the tube. 14. Resuspend the cell pellet gently in 2 ml of PBS. Count cell number. 15. Centrifuge cell suspension at 600  g for 7 min at 4  C. 16. Resuspend the pellet in 90 μl of PBS and pipette well (see Note 8). 3.4 MagneticActivated Cell Sorting Procedure (See Note 9)

1. Add 10 μl of magnetic microbeads conjugated with anti-CD34 antibody for the skin stem cells (SkSCs) and hematopoietic stem cells (HSCs) and anti-Thy-1.2 antibody (Miltenyi Biotec 130-049-10) for the SpSCs into the 90 μl of the cell suspension and mix well by gentle pipetting (see Note 10). 2. Incubate the cell suspension for 20 min in the dark at 4  C. Mix gently by tapping every 5 min (see Note 11). 3. Add 2 ml of PBS-M to the centrifuge tube to dilute magnetically labeled with or CD34+ cells or Thy1.2+ cells and centrifuge at 300  g for 7 min at 4  C. Aspirate the supernatant completely and resuspend in 1 ml of PBS-M. 4. Place an MS MACS column (MS Column; Miltenyi Biotec 130-042-201) into the magnetic field of the MACS cell separator. At the beginning, prepare the column by rinsing with 0.5 ml of ice-cold PBS-M (Fig. 2). 5. Load cell suspension into the MACS column. After the cell suspension has passed through the column and the column reservoir has become empty, wash the column three times with 0.5 ml of ice-cold PBS-M (see Note 12). 6. Remove the MS column from the MACS cell separator and place the column into a new 15-ml-volumed polypropylene centrifuge tube. 7. Add 1 ml of ice-cold serum-free culture medium and elute the magnetically CD34 or Thy1.2 labeled cells using the plunger supplied with the column.

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8. Centrifuge the tube which contains the cells at 600  g for 7 min at 4  C and resuspend the cell pellet with 1 ml of mouse serum-free medium. Repeat this step once. 9. After the last rinsing step, suspend cells in 0.5 ml of serum-free medium and determine the cell number. 10. The cells obtained by MACS protocol need to be confirmed in one instance by a method such as immunofluorescence labeling (Fig. 2G, H) or FACS to determine the purity rates (Fig. 3).

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Notes 1. The entire process must be carried out aseptically by using a sterile laminar flow hood. 2. Flask should not be moved at all for 3 days. Otherwise, the adhesion of the cells may be delayed and on day 3 the cells may be lost when the medium is changed. 3. The coagulation of the bone marrow and dilution with HBSS affects the success of the gradient centrifugation process. 4. Do not mix the Ficoll-Paque media with diluted bone marrow suspension when layering the sample. If you mix in case, gradient centrifugation process will not be successful or cell loss will occur. 5. In the case of centrifugal braking, the centrifuge is repeated without braking and without any interference to the solution. 6. The upper layer containing the plasma might be saved for later use.

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7. Although the centrifugation of the tube at high speed increases the recovery of the mononuclear cells, to get rid of platelets, a lower speed centrifuge is recommended (60–100  g). 8. It is very important that the cells disperse homogeneously in the solution without aggregating. Otherwise, the labeling and penetration activity of the antibody will decrease. 9. All MACS procedure should be performed on ice and the used solutions for these steps should be ice cold. If the cells are to be cultured later, the process should be done in sterile environment. 10. Magnetic microbeads conjugated with antibody should be removed at +4  C immediately before use and pipetted gently before use. 11. Tapping is an important step in preventing cells from becoming obstructed. If possible, a horizontal automatic mixer can be used at medium level at +4  C. 12. Care should be taken not to keep the colony dry.

Acknowledgments The experiments presented in this protocol chapter were supported by grants from Technical and Research Council of Turkey (#113S819) and Hacettepe University Research Funds (#013D04101005, #THD-2017-13430). References 1. Grutzkau A, Radbruch A (2010) Small but mighty: how the MACS-technology based on nanosized superparamagnetic particles has helped to analyze the immune system within the last 20 years. Cytometry A 77(7):643–647. https://doi.org/10.1002/cyto.a.20918 2. Abts H, Emmerich M, Miltenyi S, Radbruch A, Tesch H (1989) CD20 positive human B lymphocytes separated with the magnetic cell sorter (MACS) can be induced to proliferation and antibody secretion in vitro. J Immunol Methods 125(1–2):19–28 3. Will B, Steidl U (2010) Multi-parameter fluorescence-activated cell sorting and analysis of

stem and progenitor cells in myeloid malignancies. Best Pract Res Clin Haematol 23 (3):391–401. https://doi.org/10.1016/j. beha.2010.06.006 4. Kose S, Aerts-Kaya F, Kopru CZ, Nemutlu E, Kuskonmaz B, Karaosmanoglu B, Taskiran EZ, Altun B, Uckan Cetinkaya D, Korkusuz P (2018) Human bone marrow mesenchymal stem cells secrete endocannabinoids that stimulate in vitro hematopoietic stem cell migration effectively comparable to beta-adrenergic stimulation. Exp Hematol 57:30–41.e31. https:// doi.org/10.1016/j.exphem.2017.09.009

Methods in Molecular Biology (2019) 1879: 165–174 DOI 10.1007/7651_2018_152 © Springer Science+Business Media New York 2018 Published online: 05 May 2018

Isolation of Human Skin Epidermal Stem Cells Based on the Expression of Endothelial Protein C Receptor Meilang Xue, Suat Dervish, and Christopher J. Jackson Abstract Skin epidermis is a continuous self-renewal tissue maintained by interfollicular epidermal stem cells (IESCs) that reside in the basal layer of epidermis. IESCs also contribute to the repair and regeneration of the epidermis during wound healing. The great plasticity and easy accessibility afforded by IESCs make them a promising source of stem cells for scientific research and clinical applications. Thus, simple methods to isolate and define pure and viable IESCs are a valuable resource. Here, we provide a method for isolating IESCs from human skin epidermis. This method relies exclusively on selecting cells with a higher expression of the endothelial protein C receptor, using fluorescence-activated cell sorting. Keywords Endothelial protein C receptor, Epidermal stem cell, Fluorescence-activated cell sorting, Isolation, Skin

1

Introduction Skin epidermis undergoes continuous self-renewal throughout one’s lifetime and has an extensive ability to repair wounds. A constant replenishment of epidermal cells is generated by proliferation and differentiation of interfollicular epidermal stem cells (IESCs) that reside in the innermost (basal) layer of skin epidermis [1, 2]. Recently, human transgenic IESCs were used to regenerate a fully functional epidermis to treat epidermolysis bullosa [3]. Due to their great plasticity and easy accessibility, IESCs can be used for scientific research and novel regenerative medicine applications not only for skin epidermis but also for other epithelia such cornea and urethra [4]. The identification and isolation of human IESCs therefore is of fundamental importance. Several different isolation methods for IESCs, based on β1 integrin expression [5, 6], clonal analysis [7], high expression of α6 integrin and low expression of CD71 [8, 9], cell size and collagen type IV [10], or Hoechst 33342 exclusion combined with cell size [11], have been reported. Although these methods have greatly improved the isolation and enrichment of IESCs, no single method has been universally adopted for identifying and isolating pure viable IESCs.

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Additionally, these approaches are generally time-consuming and result in a poor yield and recovery of IESCs [10], which has made it difficult to establish IESC identity and function. The search for new IESC markers and isolation approaches that allow to more efficiently isolate IESCs is continuing. We have recently identified endothelial protein C receptor (EPCR) as a potential human IESC marker and demonstrated that epidermal cells that express highest levels of EPCR also expressed highest levels of other well-established IESC markers such as p63 and integrin β1 [12]. Interestingly, mouse bone marrow cells isolated on the basis of EPCR expression alone are highly enriched hematopoietic stem cells, showing the levels of engraftment in vivo comparable to that of stem cells purified using the most effective conventional methods [13]. Moreover, hematopoietic stem cell activity is always associated with EPCR-expressing cells [13]. Here, we describe a human IESC isolation method based on EPCR expression. When these fluorescence-activated cell sorting (FACS)-isolated IESC populations are placed in culture, they give rise to colonies that display stem cell-like properties and are able to regenerate epidermis in vitro [12]. The isolation technique is particularly advantageous because it relies exclusively on the expression of a single cell surface protein, EPCR, and can be easily performed by flow cytometry. Additionally, it allows for differentiation of epidermal cells into distinct subpopulations based on EPCR expression, and then functional assessment of their stem cell properties using a variety of assays [12].

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Materials (See Note 1)

2.1 Cell Culture Media and Buffers

1. Keratinocyte-serum-free medium (K-SFM): with matched growth supplements bovine pituitary extract and epidermal growth factor (EGF) 2. EpiLife™ medium: with 60 μM calcium and matched growth supplements HKGS 100 3. TrypLE Express (1) liquid™: without phenol red 4. Trypsin power 5. Fetal bovine serum (FBS), horse serum, penicillin–streptomycin (100, liquid), phosphate buffered saline (PBS, 0.137 M NaCl, 0.05 M NaH2PO4, pH 7.4, without calcium and magnesium) 6. 10 μg/mL Propidium iodide (PI) in PBS stored at 4  C in the dark

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1. Polyclonal goat anti-human EPCR antibody; polyclonal goat IgG control 2. Donkey anti-goat IgG antibody, Alexa 488 conjugated

2.3 Instruments and Supplies

1. Dissecting forceps, iris scissors, and disposable scalpels No. 10 2. 40- and 70-μm cell strainers, Millipore Millex 0.22 μM PVDF syringe filters 3. Tissue culture flasks (75 cm2), 24-well flat bottom plates, 100 mm  20 mm culture dishes 4. 50 mL urine pots, 15 and 50 mL centrifuge tubes (RNase/ DNase free, non-pyrogenic), 5 mL round-bottom cap tubes with and without cell strainer attached 5. Betadine antiseptic liquid, trypan blue solution (0.4%) 6. Hemocytometer 7. Flow cytometer (FACSCalibur, FACSVantage BD Biosciences)

2.4

Tissues

2.5 Preparation of Culture Media and Buffers

Neonatal foreskin or adult skin 1. Complete media: (a) neonatal foreskin epidermal cell culture: K-SFM containing 5 ng/mL recombinant human EGF, 50 μg/mL bovine pituitary extract, 100 IU/mL penicillin, and 100 IU/mL streptomycin; (b) adult skin epidermal cell culture: EpiLife™ containing 1 HKGS, 100 IU/mL penicillin, and 100 IU/mL streptomycin 2. Conditioned medium: culture medium collected from ~50% confluent neonatal epidermal cells, filtered with a prewet (to remove fibers) 0.22-μM filter (see Note 2) 3. 1 mg/mL Trypsin stock: dissolve 100 mg trypsin power in 100 mL PBS and filter through a 0.22-μM filter, store 5 mL aliquots at 20  C 4. 0.1 mg/mL Trypsin: 2 mg/mL trypsin stock in 18 mL PBS 5. Chelated FBS: swell 100 g Chelex resin in 500 ml distilled water, adjust to pH 7.4 with HCl while stirring, and then filter through Whatman no. 1 paper. Scrape the slurry resin into 500 ml FBS and incubate overnight at 4  C while stirring. On the following day, filter the mixture of Chelex resin and FBS through Whatman no. 1 paper, and discard the resin. Filter the Chelex-treated FBS through a 0.22-μm filter to sterilize FBS and store it at 20  C (see Note 3) 6. FACS buffer: 5% chelated FBS in PBS (v/v), filtered through a 0.22-μM filter

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Methods (See Note 4) Tissue Collection

Neonatal foreskins are obtained following newborn circumcision and adult skin tissues from routine surgical procedures. Each sample is placed into a 50-mL urine pot that contains 30 mL sterile PBS and stored at 4  C until use. Usage of human skin tissues should be in accordance with the local ethics committee.

3.2 Sterilization and Digestion of Skin Samples

1. Transfer skin tissue to a new urine pot that contains 6 mL betadine and 24 mL PBS, and immerse the tissue into the liquid for 5 min.

3.1

2. Wash tissue three times with 30 mL PBS each. 3. Transfer the sterile tissue to a culture dish cover with epidermis facing down (the rim of the cover is lower which provides better access to the tissue). 4. Using a fine tweezer and a scalpel, gently scrape away the fat, loose connective tissues, and blood vessels covering the dermis until the dermis is clearly and uniformly exposed (see Note 5). 5. Wash the trimmed tissue with PBS and transfer it to a new cover of culture dish. 6. Cut skin into 0.5–1 cm2 pieces (keep skin in moisture with a few drops of PBS). 7. Transfer skin pieces into a new urine pot that contains 20 mL of 0.1 mg/mL trypsin (for 2 cm  2 cm skin tissue), 400 IU/mL penicillin, and 400 IU/mL streptomycin (see Note 6). 8. Incubate the tissue pieces at 4  C overnight (16–22 h) (see Note 7). 3.3 Isolation of Epidermal Cells

1. The following day, pour the mixture of tissue and liquid within the urine pot to a culture dish with dermal side facing down. 2. Separate the epidermis from the dermis by gently pulling the tissues apart using forceps. 3. Remove the dermis from the dish and briefly mince the epidermal pieces using a scalpel and forceps. 4. Release epidermal cells by pipetting the suspension up and down using a 10-mL pipet until the mixture can be easily pipetted without tapping (see Note 8). 5. Place a 70-μM cell strainer on top of a 50-mL tube, and pass the mixture through the strainer. 6. Wash the strainer with 5 mL PBS to collect the leftover cells. 7. Quench trypsin activity by adding 2 mL FBS into the tube, and pellet cells at 180  g for 10 min at room temperature (RT) (see Note 9).

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8. Wash cell pellet with 20 mL cold FACS buffer and pass the resulting cell suspension over a 40-μm cell strainer and collect into a new 50-mL tube. 9. Pellet cells at 180  g for 10 min at 4  C, resuspend cells in 5 mL cold FACS buffer, and place cells on ice. 10. For culturing of the isolated cells, replace cold FACS buffer in steps 8 and 9 with complete medium. 11. Determine cell density and cell viability by mixing 20 μL cell suspension with 20 μL of trypan blue solution and count non-stained cells and total cells using a hemocytometer. Cell yield expected: ~2  106 cells/cm2 tissue for both neonatal and adult skin. 12. Cells are ready for IESC isolation or can be expanded by culture. 3.4 Culture of Epidermal Cells

1. Seed cells into culture flasks (75 cm2) at a density of approximately 2  106 for neonatal and 3  106 for adult cells per flask in 15 mL complete K-SFM or EpiLife™ medium. 2. Change medium twice a week. 3. Primary cultures normally reach ~70% confluence ~2–3 weeks after isolation. 4. Rinse cells with 20 mL PBS, discard it, and add 3 mL TrypLE Express solution. 5. After 1 min, remove the TrypLE and replace with 3 mL fresh TrypLE solution. Incubate cells for 5 min at RT. 6. Gently tap the flask bottom surface to release cells, then collect cells into a 50 mL tube containing 2 mL FBS. 7. To obtain the remaining cells in flask, add 4 mL fresh PBS and 1 mL fresh TrypLE into the flask and continuously incubate for 2–5 min at RT. 8. Repeat step 6, collecting cells into the same 50 mL tube. Then, add 5 mL PBS to rinse the flask and observe the cells under a microscope. If there are still cells attached the flask surface, continue the incubation for an additional 2–5 min and repeat step 6 (see Note 10). 9. Pellet cells at 180  g for 10 min at RT. 10. Resuspend cells in cold FACS buffer, place cells on ice, and determine cell density and cell viability. 11. Cells are ready for IESC isolation.

3.5 Staining Cells with Anti-endothelial Protein C Receptor Antibody (See Note 11)

1. Pellet cells isolated from skin or collected from cultures and transfer 5  106 cells to a 1.5-mL Eppendorf tube, and wash cells twice with 1 mL cold FACS buffer.

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2. Suspend cells in 0.5 mL cold FACS buffer containing 10% horse serum (V/V) and block cells for 15 min at RT. 3. Add 1 mL cold FACS buffer and pellet cells at 250  g at 4  C for 5 min and repeat another two washes with 1 mL FACS buffer. 4. Resuspend cells in 0.5 mL FACS buffer, divide cells into four tubes with 300 μL cells for anti-EPCR antibody staining, 100 μL cells for IgG isotype staining, 50 μL cells for PI single staining, and 50 μL cells as a non-stained control (see Note 12). 5. Incubate cells with antibody or IgG isotype control (100 μL diluted antibody for 1  106 cells) for 2 h at 4 C with gentle mixing by flicking with the index finger every 10–15 min to prevent cells from settling at the bottom of the tube. (a) Incubate cells with anti-EPCR antibody: 3 μL anti-EPCR antibody in 9 μL FACS buffer, mix well and take 3 μL diluted antibody mixed with 300 μL cells to reach final 1:400 dilution. (b) Incubate cells with IgG isotype control: 3 μL IgG in 27 μL FACS buffer, mix well and take 2.5 μL diluted IgG mixed with 100 μL cells to reach final 1:400 dilution. 6. Wash cells with FACS buffer as in step 3 and resuspend cells in FACS buffer in original volumes of 300 and 100 μL, respectively. 7. Incubate cells with Alexa 488 conjugated donkey anti-goat IgG antibody at 1:1000 dilution (5 μL antibody in 95 μL FACS buffer, mixed well and then put 3 μL in the tube with cells stained with anti-EPCR antibody, 1 μL in the tube with cells stained with IgG control) for 1 h at 4  C. Gently mix cells, and cover with aluminium foil to protect from light for all subsequent steps. 8. Wash cells as in step 3. 9. Resuspend all cells in 0.5 mL cold FACS buffer, transfer cell suspension into 5 mL cell strainer round-bottom cap tubes by passing through the cap (see Note 13). 10. Add 5 μL PI solution to tubes with cells except the non-stained control, and mix well. 11. Cells are ready for IESC isolation by FACS. 3.6 Flow Cytometry Isolation of Interfollicular Epidermal Stem Cells

1. Set up the FACS and ensure that the configuration is applicable to the fluorochromes used. 2. Run the unstained control cells and adjust the forward scatter (FSC) and side scatter (SSC) voltage to ensure that the bulk of the large epidermal cells are on scale. The FSC and SSC detectors should be set to a linear scale, while all fluorescence

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Fig. 1 EPCR high cells and their growth potential. (a) EPCRhigh cells sorted by fluorescence-activated cell sorting (FACS) observed under a fluorescent microscope. (b) The growth potential of unsorted, EPCRhigh and EPCRlow cells seeded into a 24-well plate at 1000 cells/well in 50% conditioned and 50% complete medium for 48 h. Images are representative of three independent experiments

channels should be set to logarithmic scales. Ensure that FSC/SSC height and width parameters are recorded to discriminate doublets. 3. Run a small amount from all samples to ensure that the voltages for each of the fluorescent detectors are optimal and on scale. 4. Collect compensation controls if required from single stained cells for each fluorochrome to be used, calculate and apply compensation. 5. Acquire data from all unstained and control tubes including the isotype. 6. Gate all cells based on FSC/SSC to exclude debris and noise, then gate single cells based on FSC/SSC height and width parameters. 7. Exclude dead cells by gating live cells based on the viability dye. 8. Using the isotype and the stained sample as a guide, identify EPCR high cells as a scant population that have a higher fluorescence than the majority of the epithelial cells and have a tendency to be lower, rather than higher in FSC and SSC when compared to the complete range of values (Fig. 1a). Normally, EPCRhigh cells are less than 0.05% of whole cell population (see Notes 14 and 15). 9. Cells collected for culture can be sorted directly into 24- or 96-well plates or a tube that contains 50% complete and 50% conditioned medium (see Note 16). 3.7 Potential Applications of Isolated Interfollicular Epidermal Stem Cells

1. Colonegenesis assay as described by Xue et al. [12] or cell proliferative potential as shown in Fig. 1b 2. RNA isolation: IESCs can be collected for subsequent RNA isolation by sorting cells directly into lysis buffer (for example, in TRI Reagent® or similar)

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3. Genetic manipulation: IESCs can be used to generate induced pluripotent stem cells [14] or transduced with correct gene to produce epidermal grafts [4, 15] for treatment of recessive dystrophic epidermolysis bullosa, junctional epidermolysis bullosa (JEB) and other skin disorders 4. In vitro and in vivo epidermal regeneration: IESCs isolated by FACS can be directly seeded on dermal equivalents for Organotypic cultures [12], or regeneration of the human epidermis to treat JEB, chronic wounds, or severe burns

4

Notes 1. All buffers, media, solutions, and consumables are sterile to eliminate any potential contamination. 2. Conditioned medium is collected from neonatal epidermal cells (passage 0) when culture reaches ~50% confluence. This medium has growth-stimulatory properties [16] and can improve the recovery of sorted IESCs. 3. Free Ca2+ is removed from FBS after Chelex treatment. Skin epidermal cells are exquisitely sensitive to calcium, and it is essential to carefully control the calcium levels to which the cells are exposed at all times. 4. All procedures, except tissue collection, are performed under sterile conditions in a laminar flow hood. This will help minimize the chance of contamination if cells are to be used for culture experiments. 5. It is essential to remove all of the fat and loose tissue attached to the dermis. This will ensure the easy separation of the epidermis from the dermis on the following day and minimize the contamination of non-epidermal cells. 6. Urine pots are used here as they are easy to keep in a freestanding position and have a large surface area to ensure that the skin is freely floating with an unsubmerged epidermis. 7. Less than 16 h incubation may result in difficulties in separating the epidermis from the dermis. In such cases, incubate tissues for an additional 1–2 h at 4  C prior to subsequent separation. 8. To ensure a clean and high yield of healthy epidermal cells, it is not recommended to incubate tissue with trypsin at 37  C. 9. After pelleting cells, it is normal for the supernatant to be cloudy as it contains dead cells and other debris which does not pellet efficiently. Carefully aspirate supernatant using 10 mL pipette to avoid the dislodgement of cell pellet.

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10. Prolonged trypsin treatment can result in the cleavage of membrane associated proteins [17], damage cellular membranes, and delay the first cell division [18]. 11. Handling cells gently and keeping cells on ice at all times helps to maintain cell viability. 12. Small numbers of cells are used to create an unstained and isotype control for calibration of cell sorting. 13. This step is to eliminate any cell clumps. The preparation of a high quality, single cell suspension is the most essential component for successful flow cytometry. 14. Different fractions of epidermal cells can be collected based on their expression of EPCR, which may differentiate IESCs from transit amplifying (TA) cells and differentiated cells, similar to that as shown by β1 integrin [5]. 15. Although the use of anti-EPCR antibody alone is sufficient to isolate distinct and substantially enriched populations of IESCs by FACS, the addition of other IESC markers (β1 integrin, α6integrin, and CD71) may also be used in conjunction with EPCR to guarantee that EPCRhigh are derived from the epidermis and do not represent contaminating cells from other non-epithelial tissues and increase the purity of IESCs; however, these markers do not completely overlap with EPCR [12]. 16. Keep sorted cells on ice at all times, and place them in culture as soon as possible. Culture sorted cells in 50% complete medium plus 50% conditioned medium for the first week to maximize cell recovery. References 1. Fuchs E, Raghavan S (2002) Getting under the skin of epidermal morphogenesis. Nat Rev Genet 3(3):199–209. https://doi.org/10. 1038/nrg758 2. Blanpain C, Fuchs E (2014) Stem cell plasticity. Plasticity of epithelial stem cells in tissue regeneration. Science 344(6189):1242281. https:// doi.org/10.1126/science.1242281 3. Hirsch T, Rothoeft T, Teig N, Bauer JW, Pellegrini G, De Rosa L, Scaglione D, Reichelt J, Klausegger A, Kneisz D, Romano O, Secone Seconetti A, Contin R, Enzo E, Jurman I, Carulli S, Jacobsen F, Luecke T, Lehnhardt M, Fischer M, Kueckelhaus M, Quaglino D, Morgante M, Bicciato S, Bondanza S, De Luca M (2017) Regeneration of the entire human epidermis using transgenic stem cells. Nature 551 (7680):327–332. https://doi.org/10.1038/ nature24487

4. Jackson CJ, Tonseth KA, Utheim TP (2017) Cultured epidermal stem cells in regenerative medicine. Stem Cell Res Ther 8(1):155. https://doi.org/10.1186/s13287-017-05871 5. Jones PH, Watt FM (1993) Separation of human epidermal stem cells from transit amplifying cells on the basis of differences in integrin function and expression. Cell 73(4):713–724 6. Zhou JX, Chen SY, Liu WM, Cao YJ, Duan EK (2004) Enrichment and identification of human ‘fetal’ epidermal stem cells. Hum Reprod 19(4):968–974. https://doi.org/10. 1093/humrep/deh166 7. Papini S, Cecchetti D, Campani D, Fitzgerald W, Grivel JC, Chen S, Margolis L, Revoltella RP (2003) Isolation and clonal analysis of human epidermal keratinocyte stem cells in long-term culture. Stem Cells 21

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(4):481–494. https://doi.org/10.1634/ste mcells.21-4-481 8. Li A, Simmons PJ, Kaur P (1998) Identification and isolation of candidate human keratinocyte stem cells based on cell surface phenotype. Proc Natl Acad Sci U S A 95 (7):3902–3907 9. Kaur P, Li A (2000) Adhesive properties of human basal epidermal cells: an analysis of keratinocyte stem cells, transit amplifying cells, and postmitotic differentiating cells. J Invest Dermatol 114(3):413–420. https://doi.org/ 10.1046/j.1523-1747.2000.00884.x 10. Li J, Miao C, Guo W, Jia L, Zhou J, Ma B, Peng S, Liu S, Cao Y, Duan E (2008) Enrichment of putative human epidermal stem cells based on cell size and collagen type IV adhesiveness. Cell Res 18(3):360–371. https://doi. org/10.1038/cr.2007.103 11. Dunnwald M, Tomanek-Chalkley A, Alexandrunas D, Fishbaugh J, Bickenbach JR (2001) Isolating a pure population of epidermal stem cells for use in tissue engineering. Exp Dermatol 10(1):45–54 12. Xue M, Dervish S, Chan B, Jackson CJ (2017) The endothelial protein C receptor is a potential stem cell marker for epidermal keratinocytes. Stem Cells 35(7):1786–1798. https:// doi.org/10.1002/stem.2630 13. Balazs AB, Fabian AJ, Esmon CT, Mulligan RC (2006) Endothelial protein C receptor (CD201) explicitly identifies hematopoietic stem cells in murine bone marrow. Blood 107 (6):2317–2321

14. Matsumura W, Fujita Y, Nakayama C, Shinkuma S, Suzuki S, Nomura T, Abe R, Shimizu H (2018) Establishment of integrationfree induced pluripotent stem cells from human recessive dystrophic epidermolysis bullosa keratinocytes. J Dermatol Sci 89 (3):263–271. https://doi.org/10.1016/j. jdermsci.2017.11.017 15. Mavilio F, Pellegrini G, Ferrari S, Di Nunzio F, Di Iorio E, Recchia A, Maruggi G, Ferrari G, Provasi E, Bonini C, Capurro S, Conti A, Magnoni C, Giannetti A, De Luca M (2006) Correction of junctional epidermolysis bullosa by transplantation of genetically modified epidermal stem cells. Nat Med 12 (12):1397–1402. https://doi.org/10.1038/ nm1504 16. Kratz G, Haegerstrand A, Dalsgaard CJ (1991) Conditioned medium from cultured human keratinocytes has growth stimulatory properties on different human cell types. J Invest Dermatol 97(6):1039–1043 17. Wu Y, Wu J, Lee DY, Yee A, Cao L, Zhang Y, Kiani C, Yang BB (2005) Versican protects cells from oxidative stress-induced apoptosis. Matrix Biol 24(1):3–13. https://doi.org/10. 1016/j.matbio.2004.11.007 18. Hirai H, Umegaki R, Kino-Oka M, Taya M (2002) Characterization of cellular motions through direct observation of individual cells at early stage in anchorage-dependent culture. J Biosci Bioeng 94(4):351–356

Methods in Molecular Biology (2019) 1879: 175–185 DOI 10.1007/7651_2018_147 © Springer Science+Business Media New York 2018 Published online: 28 June 2018

Decellularized bSIS-ECM as a Regenerative Biomaterial for Skin Wound Repair Mahmut Parmaksiz, Ays¸e Eser Elc¸in, and Yas¸ar Murat Elc¸in Abstract Tissue engineering-based regenerative applications can involve the use of stem cells for the treatment of non-healing wounds. Multipotent mesenchymal stem cells have become a focus of skin injury treatments along with many other injury types owing to their unprecedented advantages. However, there are certain limitations concerning the solo use of stem cells in skin wound repair. Natural bioactive extracellular matrixbased scaffolds have great potential for overcoming these limitations by supporting the regenerative activity and localization of stem cells. This chapter describes the use of bone marrow mesenchymal stem cells together with decellularized bovine small intestinal submucosa (SIS), for the treatment of a critical-sized full-thickness skin defect in a small animal model. Keywords Critical-sized skin defect, Extracellular matrix (ECM), Full-thickness wound repair, Mesenchymal stem cells (MSCs), Small intestinal submucosa (SIS), Xenogenic biomaterials

1

Introduction The human skin is a complex organ, which is responsible for protecting the body from environmental influences. The skin consists of three main layers, i.e., the epidermis, dermis, and the hypodermis, each having complex structural and biological properties [1, 2]. The skin can regenerate itself after minor trauma. However, non-healing wounds, such as critical-size and chronic injuries, are usually beyond the innate self-healing capacity of the skin. If these wounds are not treated properly, skin cannot fulfill its important functions and further complications may arise [1, 3]. For this reason, development of effective and functional treatment strategies for non-healing wounds is imperative [4, 5]. Among the strategies, stem cells receive attention due to their ability to differentiate into different cell types, regenerate themselves, and release bioactive factors that can play a role during repair [1, 6]. Stem cells, especially the circulating mesenchymal and hematopoietic stem cells, are already involved in the early stages of natural healing after injury. In addition, the skin stem cells are also involved in the healing process [7]. Besides, stem cells and progenitors specific to the skin appendages, such as interphilic

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epidermis, hair follicles, sebaceous gland, and sweat gland, are also involved in the wound repair and homeostasis. Studies have shown that the sweat gland stem cells can renew themselves and repair the epidermis during wound healing. The cells located in the hair follicles and in the intrafollicular epidermis are known to migrate to the wound area. It is also reported that the hair follicle stem cells are present only at the wounds [8]. It is not clear how the epidermal stem cell populations balance the proliferation, differentiation, and migration dynamics during wound healing. Stem cells possessing multiple differentiation potencies compared to tissue-specific stem cell types are preferred in cellular therapies, based on their regenerative and differentiation capacities at the wound site. While pluripotent stem cells have great potential for the future, their clinical use is currently very limited, due to safety and/or ethical concerns. Thus, stem cell-based regenerative approaches are mainly built on the basis of mesenchymal stem cells (MSCs), in regard to their well-known biological properties and relatively easy accessibility. MSCs are multipotent cells which can be isolated from various tissue types, especially from the bone marrow and adipose tissue [9, 10]. Along with their successful application in a number of indications, they are also known to accelerate skin wound repair based on their differentiation and immunomodulatory properties [11, 12]. MSCs are known to accelerate angiogenesis and the granulation tissue formation. They may also contribute to the formation of extracellular matrix (ECM) and skin appendages during the wound healing process [10, 13, 14]. MSCs have the potential to repair the wounds both physically and functionally. However, direct injection of MSCs can cause rapid cell death, thus reducing the potential for use in therapy. Furthermore, it is difficult to localize the stem cells at the wound site by direct injection. Such that the cells have different natural microenvironments that help maintain their biological functions, such as signaling and proliferation [15], it is critical to take into consideration the microenvironment of the cells to be transplanted. Within this context, scaffolds as functional bioactive stem cell carriers may be instrumental to provide effective regeneration and repair [16]. Tissue engineering approach involves the combined use of scaffolds, cells, and bioactive molecules, such as the growth factors to repair the wounded tissues [17]. Tissue engineering scaffolds can guide the adhesion, proliferation, and signalization of stem cells during the healing process. ECM components, such as the collagen, hyaluronic acid, or biocompatible synthetic polymers, are frequently used to develop the scaffold structures. However, these biomaterials alone usually do not suffice considering the complex composition of the complete natural ECM. Thus, natural tissues contain a large number of ECM molecules and bioactive components.

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Decellularization technology is based on the principle removal of cells and DNA from allogenic or xenogenic tissues while preserving the ECM proteins and their natural structure. Decellularized matrices have the advantage to retain high bioactive content and have the potential for use as a wound regeneration biomaterial alone, or in combination with stem cells. In particular, decellularized small intestinal submucosa (SIS) is known to act as a biological modulator, inducing angiogenesis and epithelialization, and contributing to the repair of the ECM [18]. In this chapter, we present a method for the use of decellularized bovine SIS-based ECM (bSIS-ECM) scaffold and MSCs for the repair of a critical-sized full-thickness skin defect in a small rodent model.

2 2.1

Materials Equipments

1. Magnetic stirrer. 2. Micropipettes. 3. Surgical scissors, clamps, forceps, sutures, and blades. 4. Centrifuge. 5. Microscope. 6. Automated cell counter. 7. Ultralow temperature freezer (
86  C). 8. Carbon dioxide incubator. 9. Laminar flow cabinet. 10. Pipette pump. 11. Nontoxic and nonpyrogenic plate inserts. 12. Histology sectioning microtome.

2.2 Reagents and Solutions

1. Bone Marrow Mesenchymal Stem Cell Growth Medium: DMEM F12 + 10% FBS + 2 mM L-glutamine, 1% Pen-Strep (see Note 1). 2. Dulbecco’s Phosphate Buffered Saline; DPBS with calcium and magnesium. 3. Xylazine-ketamine solution. 4. Iodopovidone solution. 5. Trypsin-EDTA solution (see Note 2). 6. Hematoxylin and Eosin Y solution. 7. Periodic acid solution. 8. Schiff’s reagent. 9. Tissue freezing medium.

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Methods

3.1 Isolation and Culture of Rat Bone Marrow MSCs

Rat MSCs can be isolated and cultured using standard protocols [9]. Briefly, the steps followed during isolation and culture are presented below. 1. In order to isolate bone marrow MSCs, anesthetize donor rats with intraperitoneal injection of xylazine-ketamine. Disinfect the operation area, and collect the femurs and tibias in sterile buffer solution (see Note 3). 2. Form an incision at the lower and upper parts of the bones with sterile scissors and flush the marrow in a 15 mL centrifuge tube by using a syringe containing 5 mL of growth medium (see Note 4). 3. Centrifuge the collected bone marrow tissues at 250  g for 10 min. Wash the pellet with the growth medium and centrifuge at 250  g for 5 min. 4. Resuspend the cell pellet in DMEM F12 medium containing 2 mM L-glutamine, 1% penicillin-streptomycin, and 10% fetal bovine serum and transfer to flat culture dishes (see Note 5). 5. Incubate the cell cultures at 37  C under ambient conditions of 5% CO2, 95% air, and 90% humidity. Change the medium of the cultures every 2 days during the incubation.

3.2 Seeding of MSCs on Decellularized bSIS-ECM

1. When the cells reach 70–75% confluence, collect the rat bone marrow MSCs from the culture flasks by using trypsin-EDTA solution. 2. Briefly, wash the cultures twice with 10 mL of sterile PBS after draining the waste medium (see Note 6). 3. Treat the cells with 5 mL of 0.05% Trypsin-0.53 mM EDTA solution for 2–3 min inside the incubator at 37  C or 8–10 min at room temperature. 4. Add 10 mL of serum-containing medium and collect the detached cells from the surface by pipetting into 50 mL conical tube. 5. Centrifuge the collected cell suspension at 250  g for 7 min and discard the supernatant. 6. Add 1 mL of the growth medium on the MSC pellet and then resuspend the cells by gentle pipetting (see Note 7). 7. Before cell seeding, add 10 μL trypan blue on 10 μL of cell suspension for cell counting. 8. Add the 10 μL of cell suspension treated with trypan blue onto the cell counting slide and measure % viability and the total cell number.

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9. Seed the rat bone marrow MSCs (at a density of ~1.0  106 cells/cm2) on decellularized bovine SIS-ECM membranes which have been placed in UV-sterilized 6 well inserts (Cell-crown™), and then culture the cells for 14 days (see Note 8). 10. In addition, seed 2.0  105 rat bone marrow MSCs onto each well of 6 well plates, as the two-dimensional standard control group. 3.3 Creation of the Critical-Sized FullThickness Wound Defect

The in vivo studies should be conducted in accordance with the international guidelines. 1. Use healthy adult Wistar rats (age 8–10 weeks; weight 200–250 g) to create the skin wound defects. 2. Secure the rats and anesthetize by intraperitoneal injection of xylazine (10 mg/kg) and ketamine hydrochloride (60 mg/kg) (see Note 9). 3. Upon anesthesia, shave the operation area and prepare in accordance with the asepsis and antisepsis rules. Shave the dorsal side of the subjects and swab with iodopovidone solution (see Note 10). 4. Resect the full-thickness skin (approximately 7 cm2) including the epidermis and dermis at the dorsal site with a sterile scissors (see Note 11) (Fig. 1). 5. Fix the resected skin in 2.5% glutaraldehyde solution prepared in phosphate buffered 0.9% sodium chloride solution. Then, store the fixed skin at +4  C for further histopathological characterization study as control. 6. After resection, clean the surgical wound area with sterile saline and prepare for the graft implantation procedure.

3.4 Implantation of the Stand-Alone or Stem Cell-Laden bSISECM Grafts

1. Place either the stand-alone or the MSC-laden bSIS-ECM membrane on the surgically created wound of the rat (Fig. 1). 2. Then, stabilize the bSIS-ECM grafts using a 5-0 nonabsorbable (e.g., silk) suture with several knots from the sides (see Note 12). 3. Cover the graft with an additional bSIS-ECM membrane and stabilize it with four separate knots to protect the graft from external damage (see Note 13). 4. Keep the wound area wet with sterile PBS during the surgical procedures. 5. Follow the changes at the wound site, such as the wound size, inflammation, fluid leakage, swelling, erythema, and epithelization, and record daily, in the course of the healing process (Fig. 2).

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Fig. 1 Macroscopic images retrieved at different stages of the in vivo study. (A) Critical-sized full-thickness skin defect, approximately 7 cm2 on the dorsal area, (a) lyophilized decellularized bSIS-ECM membrane, (b) the excised skin is used as the control for histopathological analyses; (B–D) wound closure with decellularized bSIS-ECM; (E) 0 h post-operation

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Fig. 2 (a) Graph showing wound contraction percent versus time of the defects treated with stand-alone bSISECM, cell-laden bSIS-ECM, and the control. The use of decellularized bSIS-ECM (with and without MSCs) accelerated wound healing about twofold. (b) Macroscopic images showing the wound healing process in the groups. Wounds in both the stand-alone and MSC-laden bSIS-ECM groups were completely closed after 49 days, however, with less scarring in the MSC-laden group (dashed circles ϕ ¼ 7 mm) [2] 3.5 Explantation of the Tissue Samples

1. To evaluate the healing process, anesthetize the rats at predetermined time points by intraperitoneal injection of xylazine (10 mg/kg) and ketamine hydrochloride (60 mg/kg). 2. Explant the graft implantation site from the dorsal skin, then fix the tissue in 2.5% glutaraldehyde solution prepared in PBS, and then keep the explants at +4  C until further processing. 3. Euthanize the unconscious rats by overdose anesthesia.

3.6 Histopathological Evaluation

1. Embed the fixed tissue samples in tissue freezing medium and transfer to an ultrafreezer at
80  C. Later, section the samples at a thickness of ~8–10 μm using a microtome, and place the sections on microscopic glass slides.

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Fig. 3 Histological comparison of the wound healing process in the stand-alone and MSC-laden bSIS-ECM groups after 28 days: (a) H&E and (b) PAS staining results. Findings showed that epithelialization was almost complete in both bSIS-ECM treated groups. However, MSC-laden bSIS-ECM group demonstrated highly orientated ECM and collagen, with new-formed skin appendages (scale bars ¼ 200 μm) [2]

2. Stain the samples with hematoxylin and eosin (H&E) and periodic acid–Schiff (PAS) solutions using standard method and examine under the light microscope (Fig. 3). The staining protocols are briefly given below. 3.7 Hematoxylin and Eosin (H&E) Staining

1. Soak the histology section slides in distilled water for 2–3 min and then treat with hematoxylin dye solution for 8 min. 2. Wash under running water for 10 min to remove the hematoxylin remnants. 3. Wash with 95% ethanol. 4. Incubate the sections with Eosin Y for 30 s–1 min, and then wash with 95% ethanol. 5. Finally, examine the samples using the light microscope.

3.8 Periodic Acid–Schiff (PAS) Staining

1. Wash the histology section slides with distilled water for 5 min. 2. Incubate the sections with periodic acid solution for 5 min, and then rinse with tap water.

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3. Treat the samples with Schiff’s reagent for 15 min and then wash with tap water for 10 min. 4. Incubate the samples with Gill’s hematoxylin solution for 1 min, and wash with tap water. 5. Dehydrate the samples using 95% and 100% ethanol. 6. Finally, examine the samples under a light microscope.

4

Notes 1. The serum must be stored at
20  C. Serum should be inactivated at 56  2  C for 30 min after dissolving at room temperature, before use in cell culture. 2. The main stock of Trypsin-EDTA solution should be stored in portions at
20  C. 3. Disinfection of the operation zone should be carried out from the center to the periphery. Femur and tibia heads should not be separated during collection. 4. This step of isolation should be performed inside a laminar flow cabinet. To prevent cell loss, avoid severe washing. Instead, perform gentle repeated washes. 5. All media, buffers, and the supplements used during the culture steps should be brought to room temperature before use to protect the cells from cold shock. 6. At this stage, culture should be washed twice to remove the serum completely. It should be noted that remaining serum will inhibit trypsin activity. 7. Pipette slowly and repeatedly to avoid damage to the cells. Failure to obtain cells as single-cell suspensions will reflect negatively on cell viability analysis, which is the next step. 8. Avoid tearing the bSIS-ECM membranes while placing in the insert. By using the plate insert, it is possible to keep the seeded cell population without significant loss. 9. Permanently check the depth of anesthesia during the procedure. In case of a possible awakening, suspend the operation and re-anesthetize the subject. It should be kept in mind that rats can be anesthetized for a wide time range of 20–50 min (i.p. injection of 80–100 mg/kg of ketamine, 5–10 mg/kg of xylazine). 10. The operation should be performed under nominal aseptic conditions. The rats should be placed on an isothermal support to keep their body temperature. All surgical instruments should be sterile.

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11. After shaving of the subjects, the planned circular incision area should be marked with a surgical pen. This will ensure the created wounds have the same size. Avoid damaging the muscle tissue during incision by lifting the skin with the forceps. 12. Try to limit suturing and knot tying during graft stabilization, since this will create secondary defects at the wound periphery. 13. It is possible to use other closure materials, such as collagen or gelatin sheets; however, decellularized bSIS-ECM membrane provides adequate mechanical durability.

Competing Interests The authors have intellectual properties related to decellularized tissues. Y.M.E. is the founder and director of Biovalda, Inc. (Ankara, Turkey). References 1. Tartarini D, Mele E (2015) Adult stem cell therapies for wound healing: biomaterials and computational models. Front Bioeng Biotechnol 3:206 2. Parmaksiz M, Elcin AE, Elcin YM (2017) Decellularization of bovine small intestinal submucosa and its use for the healing of a critical-sized full-thickness skin defect, alone and in combination with stem cells, in a small rodent model. J Tissue Eng Regen Med 11 (6):1754–1765 3. Bielefeld KA, Amini-Nik S, Alman BA (2013) Cutaneous wound healing: recruiting developmental pathways for regeneration. Cell Mol Life Sci 70(12):2059–2081 4. Metcalfe AD, Ferguson MW (2007) Tissue engineering of replacement skin: the crossroads of biomaterials, wound healing, embryonic development, stem cells and regeneration. J R Soc Interface 4(14):413–437 5. Ho J, Walsh C, Yue D, Dardik A, Cheema U (2017) Current advancements and strategies in tissue engineering for wound healing: a comprehensive review. Adv Wound Care (New Rochelle) 6(6):191–209 6. Ojeh N, Pastar I, Tomic-Canic M, Stojadinovic O (2015) Stem cells in skin regeneration, wound healing, and their clinical applications. Int J Mol Sci 16(10):25476–25501 7. Chen M, Przyborowski M, Berthiaume F (2009) Stem cells for skin tissue engineering and wound healing. Crit Rev Biomed Eng 37 (4–5):399–421

8. Ito M, Liu Y, Yang Z, Nguyen J, Liang F, Morris RJ, Cotsarelis G (2005) Stem cells in the hair follicle bulge contribute to wound repair but not to homeostasis of the epidermis. Nat Med 11(12):1351–1354 9. Odabas S, Elc¸in AE, Elc¸in YM (2014) Isolation and characterization of mesenchymal stem cells. Methods Mol Biol 1109:47–63 10. Isakson M, de Blacam C, Whelan D, McArdle A, Clover AJP (2015) Mesenchymal stem cells and cutaneous wound healing: current evidence and future potential. Stem Cells Int 2015:831095 11. An Y, Wei W, Jing H, Ming L, Liu S, Jin Y (2015) Bone marrow mesenchymal stem cell aggregate: an optimal cell therapy for fulllayer cutaneous wound vascularization and regeneration. Sci Rep 5:17036 12. Su N, Gao PL, Wang K, Wang JY, Zhong Y, Luo Y (2017) Fibrous scaffolds potentiate the paracrine function of mesenchymal stem cells: a new dimension in cell-material interaction. Biomaterials 141:74–85 13. Wu Y, Chen L, Scott PG, Tredget EE (2007) Mesenchymal stem cells enhance wound healing through differentiation and angiogenesis. Stem Cells 25:2648–2659 14. Hocking AM (2012) Mesenchymal stem cell therapy for cutaneous wounds. Adv Wound Care (New Rochelle) 1(4):166–171 15. Dogan A, Parmaksız M, Elc¸in AE, Elc¸in YM (2016) Extracellular matrix and regenerative

Decellularized bSIS-CM for Wound Repair therapies from the cardiac perspective. Stem Cell Rev 12(2):202–213 16. Duscher D, Barrera J, Wong VW, Maan ZN, Whittam AJ, Januszyk M, Gurtner GC (2016) Stem cells in wound healing: the future of regenerative medicine? A mini-review. Gerontology 62:216–225

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17. Elc¸in YM (2004) Stem cells and tissue engineering. Adv Exp Med Biol 553:301–316 18. Parmaksiz M, Dogan A, Odabas S, Elc¸in AE, Elc¸in YM (2016) Clinical applications of decellularized extracellular matrices for tissue engineering and regenerative medicine. Biomed Mater 11(2):022003

Methods in Molecular Biology (2019) 1879: 187–200 DOI 10.1007/7651_2018_142 © Springer Science+Business Media New York 2018 Published online: 28 April 2018

Protocols for Full Thickness Skin Wound Repair Using Prevascularized Human Mesenchymal Stem Cell Sheet Lei Chen, Daniel Radke, Shaohai Qi, and Feng Zhao Abstract Split thickness skin grafts (STSGs) are one of the standard treatments available for full thickness wound repair when full thickness grafts (FTGs) are not viable, such as in the case of wounds with large surface areas. The donor sites of STSGs may be harvested repeatedly, but STSG transplants are still limited by insufficient blood supply at the early stages of wound healing. Prevascularized human mesenchymal stem cell (hMSC) sheets may accelerate wound healing and improve regeneration by providing preformed vessel structures and angiogenic factors to overcome this limitation. This book chapter provides the protocol of co-culturing hMSCs and endothelial cells to attain a prevascularized hMSC cell sheet (PHCS). The protocols for implantation of the prevascularized stem cell sheet for full thickness skin wound repair in a rat autologous skin graft model as well as the evaluation of the wound healing effects are also provided. Keywords Full thickness skin wound, Human mesenchymal stem cells, Prevascularization, Split thickness skin grafts, Wound repair

1

Introduction Burn injuries represent a leading cause of full thickness skin wounds, accounting for approximately 15,000–20,000 hospitalizations per year in the USA [1]. Transplantation of an autologous full thickness graft (FTG) or a skin flap remains the current gold standard of treatment due to immunological acceptance and close match to native skin color and texture [2]. However, the limited supply of donor skin and unavoidable donor site injury restricts the ability to treat extensive wounds with FTGs. An autologous split thickness skin graft (STSG) on the other hand, in which only the epidermis and a portion of the dermis are harvested rather than the full skin thickness, can be harvested repeatedly to cover larger wounds and can be used under conditions that would cause an FTG to fail, such as low vasculature or moderate infection at the wound site [3]. The disadvantage is that STSG can contract significantly during healing, leading to poor cosmetic outcome, physical disability, and reduced pliability [4]. Including engineered dermal substitutes with the STSG can reduce contraction, but the limited blood supply at early stages of transplantation cause

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these grafts to experience relatively long hypoxic and ischemic periods after surgery and suffer from degeneration and necrosis [5]. Sufficient blood supply is the primary factor that determines the quality of a transplanted STSG. The graft becomes engorged by plasmatic fluid during the first 48 h of transplantation, and a poorly vascularized bed hinders plasmatic diffusion [6]. Effective delivery of oxygen, nutrients, and growth factors can be achieved by ensuring that the transplanted graft has sufficient vascular support at these early stages [7]. The inclusion of a prevascularized mesenchymal stem cell (MSC) cell sheet may provide this essential early support while also promoting graft integration to improve wound healing and enhance therapeutic outcome [8, 9]. Our previous study has shown that a uniform human MSC (hMSC) cell sheet (HCS) can be obtained by growing hMSCs under a physiologically low oxygen concentration (2% O2), which helps to maintain the stemness of the cells [10]. We have also achieved a prevascularized hMSC cell sheet (PHCS) by co-culturing endothelial cells (ECs) on top of the HCS under a normoxic condition to promote angiogenesis and neovascularization [11]. After implantation of the PHCS in a rat autologous full thickness skin wound model, we have found that the PHCS could significantly reduce skin contraction and improve cosmetic appearance relative to the STSG control group [12]. This book chapter provides the protocol of co-culturing hMSCs and endothelial cells (ECs) to attain an HCS and a PHCS. The protocols for implantation of the prevascularized stem cell sheet for full thickness skin wound repair in a rat autologous skin graft model as well as the evaluation of the wound healing effects are also provided in detail.

2

Materials

2.1 Cell Sheet Culture

1. hMSCs are expanded in complete α-minimum essential medium (α-MEM) with 20% fetal bovine serum, 1% L-glutamine, and 1% penicillin/streptomycin (Life Technologies, Rockville, MD). Passage 3–5 hMSCs are used for cell sheet culture (see Note 1). 2. Endothelial cells (ECs) are expanded in endothelial cell growth medium (EGM-2, BulletKit, Lonza). Passage 3–5 ECs are used for cell sheet culture (see Note 2).

Animals

1. Sprague Dawley (SD) rats (weighing 180–200 g) are used in the animal study under the approval of University Institutional Animal Care and Use Committee (IACUC).

2.3 Solutions for Tissue Fixation and Staining

1. Cell sheet fixation solution: 4% paraformaldehyde (PFA), commercially available.

2.2

2. Phosphate-buffered saline (PBS), commercially available.

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3. Cell sheet permeabilization solution: 0.2% Triton X-100 in PBS. 4. Blocking buffer: 1.0% bovine serum albumin (BFA) in 0.2% Triton X-100 in PBS. 5. Cell nuclei staining solution: 40 ,6-diamidino-2-phenylindole (DAPI) solution, commercially available. 6. Hematoxylin solution, commercially available. 7. 1% Eosin Y solution, commercially available. 8. Biebrich scarlet–acid fuchsin, commercially available. 9. Working phosphotungstic/phosphomolybdic acid solution: mix 1 volume of phosphotungstic acid solution and 1 volume of phosphomolybdic acid solution with 2 volumes of deionized water (DI H2O). The solution is discarded after one use. 10. Aniline blue solution, commercially available. 11. Acetic acid, commercially available. 12. Sodium citrate buffer solution (pH 6.0), commercially available. 13. VECTASTAIN® Elite ABC Reagent, commercially available. 14. 3,30 -Diaminobenzidine (DAB) substrate kit, commercially available. 15. Antigen unmasking solution: sodium citrate buffer solution (pH 6.0), commercially available.

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Methods

3.1 Preparation of Human Mesenchymal Stem Cell Sheet

1. Seed hMSCs on collagen-1-coated cover glasses at a density of 10,000 cells/cm2. 2. Culture the cells under 2% O2 for 4 weeks in complete α-MEM medium. The medium is changed every 2 days. 3. Harvest hMSC sheets by gently peeling the cell layers off the cover glass. The size of the cell sheet is decided by the size of the glass cover slip (see Note 3). 4. For evaluation of thickness, fix the detached hMSC sheet in 4% PFA and measure with confocal microscope.

3.2 Preparation of Prevascularized Human Mesenchymal Stem Cell Sheet

1. Seed ECs on top of hMSC sheets at a density of 20,000 cells/ cm2. 2. Culture under 20% O2 for 1 week in EGM-2. Change medium every other day. 3. Harvest the PHCS by gently peeling the cell layers off the cover glass.

3.3 Microvessel Observation

1. Wash PHCS with PBS for three times. 2. Fix the cell sheet with 4% PFA.

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3. Permeabilize the sample with 0.2% Triton X-100 in PBS. 4. Block the sample with 1.0% BFA in 0.2% Triton X-100 in PBS. 5. Incubate the sample with primary anti-CD 31 antibody (see Note 4). 6. Incubate the sample with DyLight 488 secondary antibody (see Note 5). 7. Stain cell nuclei with DAPI solution. 8. Image with fluorescence microscope (see Note 6). 3.4 Full Thickness Excision Wound Creation and Graft Transplantation

1. Put the rats in anesthesia box and anesthetize rats with inhaled gas anesthesia (O2, 2 L/min; isoflurane, 2%). 2. Scrub the dorsal skin of the rat with Betadine Veterinary Surgical Scrub twice and use a 70% alcohol gauze to remove Betadine, wait for 1 min to let the operation area dry up. 3. Use iris scissors to create a round full thickness excisional wound with a diameter of 20 mm on the dorsum. Make sure that all procedures are done under sterile conditions. 4. Use iris scissors to remove the deep partial dermis and panniculus carnosus away from the round excised skin to attain the STSG. Protect the dermal side of the STSG with saline gauze (Fig. 1a). 5. Use a sharp tipped scalpel to pierce the skin and create evenly spaced fenestrations on the grafts for secretion drainage (the length of every fenestrations should be less than 2 mm). 6. Transplant three cell sheets onto the wound site.

Fig. 1 Animal surgery. (a) A round full thickness excisional wound with a diameter of 20 mm on the dorsum. (b) Split thickness skin grafts (STSG) with spaced fenestrations (yellow rectangular box) pierced by a sharp tipped scalpel. The STSG was approximated to the adjacent wound margin and sutured with interrupted stitches (yellow arrow)

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(a) Discard the culture medium by use of Pap dropper. Make sure that the cultured cell sheets are untouched. (b) Rinse the cell sheets with PBS twice. (c) Use sharp tipped tweezers to pry the glass cover from the bottoms of culture plate. Then, set it on the wound bed of the rat, cell sheet side up. (d) Use sharp tipped tweezers to strip off the cell sheet. (e) Remove the glass cover. 7. Lay the autologous STSG on top of the cell sheets (see Note 7). 8. Approximate the grafts to the adjacent wound margin and suture with interrupted stitches. The stitches have to be applied 2–3 mm apart from each other. After knotting, leave the thread uncut (Fig. 1b). 9. Take photos for the grafts in the same exposure setting, from the same distance and angle. 10. Clean the grafts and the skin around by use of saline gauze. 11. Apply a layer of sterilized, transparent, plastic, standard measuring film on the top of each graft. 12. Delineate the shape of the graft by use of fine tipped surgical marker. 13. Clean the grafts and the skin around by use of saline gauze. 14. Apply folded cotton gauze on the grafts and affix the gauze with wrapped elastic bandage. 15. Apply a padded bolster (folded cotton gauze) on top of the skin grafts and use the threads to affix it (knot the threads that are in a diagonal) to provide appropriate pressure (the bolster is affixed by the knotted threads and cannot be easily lifted from the skin) and prevent scratching. 16. House animals in individual cages post-surgery. 17. Change dressings 3 days after surgery and remove on day 7. 3.4.1 Changing Wound Dressings (Days 3 and 7)

1. On post-operational day 3, put the rats in anesthesia box and anesthetize rats with inhaled gas anesthesia (O2, 2 L/min; isoflurane, 2%). Use sharp tipped scissors to cut off the threads. Gently remove all the dressing from the graft. 2. Take photos for the grafts in the same exposure setting, from the same distance and angle. Apply a layer of sterilized, transparent, plastic, standard measuring film on the top of each graft. 3. Delineate the shape of the graft by use of fine tipped surgical marker. 4. Clean the grafts and the skin around by use of saline gauze.

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5. Apply folded cotton gauze on the grafts and affix the gauze with wrapped elastic bandage. 6. On post-operational day 7, put the rats in anesthesia box and anesthetize rats with inhaled gas anesthesia (O2, 2 L/min; isoflurane, 2%). 7. Gently remove all the dressing from the graft. 8. Clean the grafts and the skin around by use of saline gauze. 9. Put the rats back in their own cages. 3.4.2 Wound Site Observation (See Note 8)

1. At each observation time points, put the rats in anesthesia box and anesthetize rats with inhaled gas anesthesia (O2, 2 L/min; isoflurane, 2%). 2. Take photos for the grafts in the same exposure setting, from the same distance and angle. 3. Clean the grafts and the skin around by use of saline gauze. 4. Apply a layer of sterilized, transparent, plastic, standard measuring film on the top of each graft. 5. Delineate the shape of the graft by use of fine tipped surgical marker. 6. Clean the grafts and the skin around by use of saline gauze. 7. For post-operational day 0 and 3 rats, apply folded cotton gauze on the grafts and affix the gauze with wrapped elastic bandage. 8. For post-operational day 7, 14, 21, and 28 rats, wait for the rats to regain consciousness and put the rats back to their own cages, or sacrifice the rats. 9. Cut the measuring film along the mark. 10. Weigh the measuring film. 11. Measure skin graft contraction via gravitational planimetry (see Note 9).

3.4.3 Tissue Collection

1. Sacrifice six rats in each group for each time point and harvest grafts and surrounding normal tissue (including the skin and subcutaneous tissue). 2. At each time point, put the rats in anesthesia box and anesthetize rats with inhaled gas anesthesia (O2, 2 L/min; isoflurane, 2%). 3. Apply cervical dislocation method to sacrifice the rats. 4. Use sharp tipped scissors to cut off the graft and its surrounding normal skin (about 0.5 cm wide). 5. Cut samples into two parts, make sure that they are of approximate size. 6. Immerse samples in 10% formalin and fix the tissue at room temperature for 24 h (see Note 10).

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1. Put tissues in a pencil labeled cassette. 2. Immerse cassette in 70% ethanol for 1 h. 3. Immerse cassette in 95% ethanol (95% ethanol/5% methanol) for 1 h. 4. Immerse cassette in absolute ethanol for 1 h. 5. Immerse cassette in absolute ethanol for 1½ h. Perform this procedure twice with fresh solution each time. 6. Immerse cassette in absolute ethanol for 2 h. 7. Immerse cassette in clearing agent (xylene or substitute) for 1 h. Perform this procedure twice with fresh solution each time. 8. Immerse cassette in wax (Paraplast X-tra) at 58  C for 1 h. Perform this procedure twice with fresh solution each time. 9. Open cassette to view the skin sample and choose a mold that best corresponds to the size of the sample. Discard cassette lid. 10. Put small amount of molten paraffin in mold, dispensing from paraffin reservoir. Use warm forceps to transfer skin sample into mold, placing cut side down, as it was placed in the cassette. 11. Transfer mold to cold plate. Add the labeled tissue cassette on top of the mold as a backing when the tissue is in the desired orientation. 12. Add hot paraffin to the mold. Make sure that there is enough paraffin to cover the face of the plastic cassette. 13. Fill cassette with paraffin while cooling, keeping the mold full until solid (about 30 min). 14. Pry mold when the wax is completely cooled and hardened (Fig. 2a).

3.5 Pathology Analysis: Staining, Imaging, and Analysis

1. Turn on the water bath and adjust the temperature to 35–37  C.

3.5.1 Section the Paraffin-Embedded Tissue Blocks

3. Place the blocks face down on an ice block for 10 min.

2. Use a microtome to section paraffin-embedded tissues. 4. Place a fresh blade on the microtome; insert the block into the microtome chuck (placing cut side of the sample parallel with the blade (Fig. 2b)). 5. Set the dial to cut several 10 μM sections in order to plane the block; once it is cutting smoothly, set to 5 μM and cut. 6. Pick tissue sections with 5 μm thickness up with fine paint brush and float them on the surface of the 37  C water bath. 7. Float the sections onto the surface of clean glass slides.

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Fig. 2 Tissue section. (a) A paraffin-embedded tissue (yellow arrow) block. (b) A block inserted into the microtome chuck. (c) A typical skin tissue section

8. Place the slides with paraffin sections on the warming block in a 65  C oven for 20 min to melt the wax (to bond the tissue to the glass) (Fig. 2c). 9. Use for staining. 3.5.2 Dewax and Rehydrate Samples (See Note 12)

1. Dry heat the slides under 60  C for 2 h. 2. Immerse the slides in xylene for 5 min. Perform this procedure thrice with fresh solution each time. 3. Immerse the slides in 100% ethanol for 5 min. Perform this procedure twice with fresh solution each time. 4. Immerse the slides in 80% ethanol for 5 min. 5. Immerse the slides in DI H2O for 5 min.

3.5.3 Hematoxylin and Eosin (HE) Staining and Masson’s Trichrome Staining (See Note 13) HE Staining (See Note 14)

1. Put the slides into DI H2O and leave the slides there for 5 min. 2. Put the slides into a staining jar containing 210 mL hematoxylin solution and leave the slides there for 5 min. 3. Put the slides into tap water, then leave the staining jar under running tap water for 10 min, make sure that the running tap water does not directly hit the slides.

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4. Put the slides into DI H2O and leave the slides there for 2 min. 5. Put the slides into a staining jar containing 210 mL 1% Eosin Y solution for 30 s. 6. Put the slides into tap water, then leave the staining jar under running tap water for 10 min, make sure that the running tap water does not directly hit the slides. 7. Dehydrate the slides with 95% ethanol for 1.5 min twice, with fresh solution each time. 8. Further dehydrate the slides with absolute ethanol for 1.5 min twice, with fresh solution each time. 9. Immerse the slides in xylene for 1.5 min three times, with fresh xylene each time. 10. Drop one drop of resinous mounting medium and seal the section using glass cover. 11. Observe and image the tissue morphology (see Note 15). Masson’s Trichrome Staining (See Note 16)

1. Immerse the slides in hematoxylin solution for 5 min. 2. Put the slides into a staining jar containing tap water, then leave the staining jar under running tap water for 5 min, make sure that the running tap water does not directly hit the slides. 3. Put the slides into DI H2O and leave the slides there for 5 min. 4. Put the slides into a staining jar containing 210 mL Biebrich scarlet–acid fuchsin for 5 min. 5. Put the slides into DI H2O and leave the slides there for 5 min. Put the slides into a staining jar containing 210 mL phosphotungstic/phosphomolybdic acid solution for 5 min. 6. Place slides into a staining jar containing 210 mL aniline blue solution for 5 min. 7. Place slides into a staining jar containing 210 mL acetic acid for 2 min. Discard solution. 8. Put the slides into DI H2O and leave the slides there for 1 min twice, with fresh DI H2O each time. 9. Dehydrate the slides with 95% ethanol for 1.5 min twice, with fresh solution each time. 10. Further dehydrate the slides with absolute ethanol for 1.5 min twice, with fresh solution each time. 11. Immerse the slides in xylene for 1.5 min three times, with fresh xylene each time. 12. Drop one drop of resinous mounting medium and seal the section using glass cover. 13. Observe and image the tissue morphology (see Note 15).

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3.5.4 Immunohistochemistry (IHC), Ki67, and NM95 Staining (See Note 17)

1. Antigen unmasking: add 200 mL sodium citrate buffer solution (pH 6.0) in plastic staining jar. Place it in Black and Decker vegetable steamer and steam for 30 min. Put all sections into the jar when the sodium citrate buffer solution is 95–98  C, 10 min. 2. Take out the staining jar and leave it at room temperature for 60 min to slowly cool down the slides. This will prevent tissue destruction caused by drastic temperature change. 3. Incubate the sections for 30 min in 0.3% H2O2. 4. Wash in 10 mM sodium phosphate (PBS), pH 7.5, for 5 min. 5. Apply background buster on slides, incubate for 30 min at room temperature. Drain excess buster. No wash. 6. Apply rabbit anti-CD68 or diluted rabbit anti-Ki67 and incubate for 45 min at room temperature. Immerse slides two times in PBS, 1 min each, drain excess PBS. 7. Apply biotinylated secondary antibody (goat anti-rabbit) and incubate for 30 min. Repeat wash as in step 6. 8. Apply Vectastain Elite ABC and incubate for 30 min, repeat wash as in step 7. 9. Apply DAB Chromogen and incubate for 5 min, wash in three changes of DI H2O. Rinse in tap water for 2 min. 10. Counterstain in hematoxylin for 30 s. Rinse slides in tap water until clear. 11. Blue nuclei by dipping four times in ammonium water, rinse in tap water for 2 min. 12. Immerse the slides in xylene for 1.5 min three times, with fresh xylene each time. 13. Drop one drop of resinous mounting medium and seal the section using glass cover. 14. Observe and image the tissue morphology (see Note 15).

3.6 Image Analysis (See Note 18) 3.6.1 Count (Microvessels, Hair Follicles, and Cells) (See Note 19) 3.6.2 Measure Epidermal Thickness

Open ImageJ ! hit “File” ! hit “Open” ! open one photo ! hit “Type” ! hit “8-bit” ! hit “Image” ! hit “Adjust” ! hit “Threshold” ! hit “Auto” ! hit “Apply” ! hit “Analyze Particles” ! choose “summarize”.

Open ImageJ ! hit “File” ! hit “Open” ! open one photo ! hit “Straight Line Selection” ! measure the scale bar in the photo ! hit “Analyze” ! hit “Tool” ! hit “Scale Bar” ! chose any color you like ! hit “OK” ! use the mouse to draw a line that starts from the epidermal basement membrane and stops at the surface of the epidermis (make sure that the line is perpendicular to the epidermal basement membrane).

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3.6.3 Measure Collagen Index

Open ImageJ ! hit “File” ! hit “Open” ! open one photo ! hit “Plugins” ! hit “Analyze” ! hit “Measure RGB” ! write the “means” value from the “Results” window into excel file ! use formula: collagen index ¼ sum (blue mean value, green mean value)/sum(2 * red mean value, blue mean value, green mean value).

3.6.4 Vessel Area Quantification (See Note 20)

Open ImageJ ! hit “File” ! hit “Open” ! open one photo ! hit “Straight Line Selection” ! measure the scale bar in the photo ! hit “Analyze” ! hit “Set Scale” ! enter the length of the scale bar in “known distance,” enter the unit of the scale bar in unit of length ! hit “Global” ! hit “OK” ! hit “rectangular” ! use the mouse to draw a rectangular frame ! hit “Analyze” ! hit “Analyze Measurements” ! hit “Area” ! the result will be the area of the entire photo ! enter it in excel ! hit “Freehand Selections” ! use the mouse to track the shape of every vessels ! hit “Analyze” ! hit “Analyze Measurements” ! hit “Area” ! the result will be the area of every vessel ! enter it in excel ! sum up all measured data of vessel area and divide it by the total area of the photo.

4

Notes 1. The hMSCs that we use are from Texas A&M University Health Sciences Center. 2. The ECs that we use are human umbilical vein endothelial cells (HUVECs). The cells are from Lonza (Walkersville, MD). 3. The strong interaction between the cells as well as the cells and the ECM in the cell sheet can overcome the cell adhesion force between the cells and the glass cover slip. Thus, the cell sheet can be easily peeled off from glass cover slip. 4. The primary anti-CD 31 antibody that we use is obtained from Abcam (Cambridge, MA). The other antibodies anti-CD68 and anti-Ki67 mentioned in this protocol are also from Abcam. 5. The DyLight 488 secondary antibody we use is obtained from VECTOR Laboratories (Burlingame, CA). The biotinylated secondary antibody (goat anti-rabbit) mentioned in the text is also from VECTOR Laboratories. 6. A vessel network was defined as a contiguous length of interconnected capillaries. Six low-magnification (20) images on three individual samples in each group were used for calculation. 7. STSG transplantation without a cell sheet is used as a control. 8. In our study, we observe wound site at days 0, 3, 7, 14, 21, and 28.

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9. Since the material density and the thickness of the measuring film are the same, the skin graft contraction can be measured by gravitational planimetry (graft size * thickness * material density ¼ weight of the measuring film) and expressed as a percentage of the remaining skin graft size to its original wound size (relative skin graft size % ¼ (remaining skin graft size/Original skin graft size) * 100%). 10. Tissue fixation is a critical step in the preparation of histological sections by which biological tissues are preserved from degradation and putrefaction. Fixation terminates any ongoing biochemical reactions and may also increase the mechanical strength or stability of the treated tissues. For skin tissue fixation, neutral buffered formalin solution is normally used. 11. Dehydrate fixed samples through a graded series of ethanol wash and embed in paraffin. The tissue samples are dehydrated to displace water contained in the samples. Paraffin is used to infiltrate tissues. The paraffin-embedded tissues can be stored for many years. 12. To stain the sections, the wax needs to be removed and the samples need to be rehydrated. During the dewax and rehydration processes, the sections need to go through multiple changes of xylene, series ethanol solutions (100–0%), and finally DI H2O. 13. The HE and Masson’s trichrome staining are conducted according to the manufacturer’s standardized protocols. 14. The HE staining is the most commonly used staining method for tissue sections. Hematoxylin stains nuclei of cells and a few other objects, such as keratohyalin granules. The color is blue. The Eosin Y stains eosinophilic and other structures in various shades of red, pink, and orange. This staining technique is essential for recognizing various tissue types and their morphologic changes. 15. At least five stained sections from each group at each time point should be observed. At least three random views should be imaged. 16. Masson’s trichrome is suited for distinguishing cells from surrounding connective tissue and for evaluating collagen deposition (collagen index) as well as morphologic change. 17. Ki67 is a nuclear antigen expressed in proliferating cells. It is used to evaluate the proliferative activity of cells in the tissues. NM95 recognizes an antigen associated with the nucleoli in human cells. It is used to determine whether implanted human cells (hMSCs and ECs) actually participated in the process of skin wound healing after grafting. 18. The HE staining images can be used to measure epidermal thickness and count microvessels (Fig. 3a), hair

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Fig. 3 Identified structure in stained tissues. (a) Blood vessels (black dashed circle). (b) Hair follicle (yellow star) and sebaceous glands (yellow dashed rectangle). (c) Epidermis (yellow triangle) and epidermal basement membrane (yellow dashed line). (d) Collagen fibers (yellow arrow)

follicles (Fig. 3b) as well as cells and skin tissues (Fig. 3c) in high-power field (HPF) using ImageJ software. The collagen index can be measured using the images from Masson’s trichrome staining (Fig. 3d). 19. Microvessels are identified as small circles of endothelial cells with red blood cells in them. 20. Quantification is defined as the sum of all vessel segments/HPF, using ImageJ.

Acknowledgements This study was supported by the National Institutes of Health (1R15CA202656 and 1R15HL115521-01A1) and the National Science Foundation (1703570) to F.Z. It is also supported by the National Natural Science Foundation of China (81601703) to L.C.

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References 1. Mendez-Eastman S (2005) Burn injuries. Plast Surg Nurs 25:133–139 2. Dreifke MB, Jayasuriya AA, Jayasuriya AC (2015) Current wound healing procedures and potential care. Mater Sci Eng C 48:651–662 3. Shimizu R, Kishi K (2012) Skin graft. Plast Surg Int 2012:563493 4. Wood FM (2014) Skin regeneration: the complexities of translation into clinical practise. Int J Biochem Cell Biol 56:133–140 5. Zografou A, Papadopoulos O, Tsigris C, Kavantzas N, Michalopoulos E, Chatzistamatiou T, Papassavas A, StavropoulouGioka C, Dontas I, Perrea D (2013) Autologous transplantation of adipose-derived stem cells enhances skin graft survival and wound healing in diabetic rats. Ann Plast Surg 71:225–232 6. Seyhan T (2011) Split-thickness skin grafts. Skin Grafts Marcia Spear, IntechOpen, Croatia 7. Chua AWC, Khoo YC, Tan BK, Tan KC, Foo CL, Chong SJ (2016) Skin tissue engineering advances in severe burns: review and therapeutic applications. Burns Trauma 4:3 8. Kang Y, Ren L, Yang Y (2014) Engineering vascularized bone grafts by integrating a biomimetic

periosteum and β-TCP scaffold. ACS Appl Mater Interfaces 6:9622–9633 9. Lesman A, Habib M, Caspi O, Gepstein A, Arbel G, Levenberg S, Gepstein L (2009) Transplantation of a tissue-engineered human vascularized cardiac muscle. Tissue Eng A 16:115–125 10. Zhao F, Veldhuis JJ, Duan Y, Yang Y, Christoforou N, Ma T, Leong KW (2010) Low oxygen tension and synthetic nanogratings improve the uniformity and stemness of human mesenchymal stem cell layer. Mol Ther 18:1010–1018 11. Zhang L, Xing Q, Qian Z, Tahtinen M, Zhang Z, Shearier E, Qi S, Zhao F (2016) Hypoxia created human mesenchymal stem cell sheet for prevascularized 3D tissue construction. Adv Healthc Mater 5:342–352 12. Chen L, Xing Q, Zhai Q, Tahtinen M, Zhou F, Chen L, Xu Y, Qi S, Zhao F (2017) Pre-vascularization enhances therapeutic effects of human mesenchymal stem cell sheets in full thickness skin wound repair. Theranostics 7:117

Methods in Molecular Biology (2019) 1879: 201–210 DOI 10.1007/7651_2018_173 © Springer Science+Business Media New York 2018 Published online: 12 August 2018

Cultivation of Adipose-Derived Stromal Cells on Intact Amniotic Membrane-Based Scaffold for Skin Tissue Engineering Ehsan Taghiabadi, Bahareh Beiki, Nasser Aghdami, and Amir Bajouri Abstract Application of cell-based skin substitutes has recently evolved as a novel treatment for hard-to-heal wounds. Here, we focus on the development of a novel skin substitute by seeding human adipose-derived stromal cells (ASCs) on acellular human amniotic membrane (HAM). This construction is probably associated with higher rates of host cell infiltration and implanted cell engraftment. ASCs are achieved by separation of stromal cells from lipoaspirates using collagenase digestion and acellular HAM was obtained by separation of outer membrane of the chorion and removing its epithelial cells. Keywords Adipose-derived stromal cells, Amniotic membrane, Skin substitutes, Stem cell, Tissue engineering, Wound healing

1

Introduction Human amniotic membrane (HAM) as the innermost layer of fetal membrane has been widely used for wounds due to its biological and mechanical characteristics, since 1910 [1]. It contains an epithelial cell layer, a basilar membrane layer, and a compact layer. The extracellular matrix of basilar membrane contains collagen IV, heparan sulfate, proteoglycan, collagen I, collagen III, collagen IV, and fibronectin that provide mechanical support and proper protection for wounds [2, 3]. Moreover, HAM secretes high levels of different growth factors [2] that play important roles in tissue regeneration [4] through their anti-inflammatory [5], anti-fibrotic [6], and anti-microbial effects; also, HAM has low immunogenicity [7]. In recent decades, concurrent utilization of bio-scaffolds and stem cells has been considered to increase the rates of host cell infiltration and implanted cell engraftment [8]. Research in wound cell-based therapies is mainly focused on transplantation of mesenchymal stem cells, which could be derived from different sources such as bone marrow, adipose tissue, menstrual blood, cord blood, and dental pulp [9–14]. It seems that these cells can differentiate into endothelial and epithelial cells, and exert paracrine effects

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through secretion of soluble growth factors and cytokines. Adipose-derived stromal cells (ASCs) have been recently come under the spotlight as they have high proliferation rates and induce hematopoiesis and wound regeneration with minimal harvesting risk [15]. Here, we describe a straightforward strategy for production of a two-dimensional (2D) skin substitute from human ASCs and intact HAM. Acellular HAM is obtained by separation of the outer membrane of the chorion and removing its epithelial cells. ASCs have shown to have the ability to differentiate into adipocytes and osteocytes, adhere to plastic, and express specific surface antigen markers (e.g., CD73, CD105, CD90, and CD 44) in the absence of some other markers (e.g., CD45 and CD14) [15]. The intact HAM are seeded by cultured ASCs and characterized before transplantation.

2 2.1

Materials Tissue

1. The human amniotic membrane is obtained from human placentas following the approval of the Ethics Committee. 2. Lipoaspirate is commonly obtained from healthy donors. Before cell isolation, the lipoaspirate can be kept at 4  C, overnight (see Note 1).

2.2 Reagents and Medium

1. Phosphate buffered saline (PBS) without Ca2+ or Mg2+ (at pH 7.4) (Gibco, USA). 2. Dimethyl sulfoxide (DMSO) (Sigma, USA). 3. Dulbecco’s Modified Eagle’s Medium/Nutrient Mixture F-12 HAM solution (DMEM/F12) (Gibco, USA). 4. L-Glutamine (Gibco, USA). 5. 10,000 U/ml penicillin and 10,000 mg/ml streptomycin (Gibco, USA). 6. Fetal Bovine Serum (FBS; Gibco-BRL). 7. Collagenase A type I (Cat. No. C-0130, Sigma-Aldrich). 8. Falcon 70 μm cell strainers (Becton Dickinson). 9. Plastic conical tubes (50 and 15 ml) (TPP, Germany). 10. Disposable plastic pipettes (50, 25, and 10 ml) (TPP, Germany). 11. Anti-CD45 FITC-conjugated mouse anti-human monoclonal antibody (BD, USA). 12. Anti-CD90 FITC-conjugated mouse anti-human monoclonal antibody (BD, USA). 13. Anti-CD105 FITC-conjugated mouse anti-human monoclonal antibody (BD, USA).

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14. Anti-CD34 FITC-conjugated mouse anti-human monoclonal antibody (BD, USA). 15. Anti-CD73 FITC-conjugated mouse anti-human monoclonal antibody (BD, USA). 16. Paraformaldehyde solution 4% (Sigma, USA). 17. Anti-vimentin mouse anti-human monoclonal antibody (Abcam, USA).

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Methods

3.1 Separation of the Stromal Vascular Fraction from Lipoaspirate 3.1.1 Lipoaspirate Washing

The lipoaspirate is washed with PBS (three times) under aseptic conditions to remove erythrocytes and leukocytes, as much as possible. 1. Place a maximum volume of 30 ml of lipoaspirate into 50 ml plastic conical tubes. 2. Allow the adipose tissue to float above the blood fraction. 3. Remove the blood fraction using a sterile 25 ml pipette. 4. Add an equivalent volume of PBS supplemented with penicillin/streptomycin and amphotericin B and shake vigorously for 5–10 s. 5. Set the conical tubes on the bench and allow the adipose tissue to float above the PBS (1–5 min). 6. Remove the PBS using a 50 ml pipette. 7. Repeat the above washing procedure (steps 4–6) three times.

3.1.2 Collagenase Digestion

Dispersion of adipose tissue with high cell viability is achieved by collagenase type I digestion. 1. Prepare collagenase solution at final concentration of 0.2% in PBS just prior to digestion; then, filter the solution into the remaining working volume. 2. Add collagenase solution into 50 ml plastic conical tubes containing washed adipose tissue. 3. Resuspend the adipose tissue while vigorously shaking the conical tube for 5–10 s. 4. Incubation at 37  C for 2 h followed by vigorous manual shaking of the conical tube for 5–10 s every 15 min. 5. After completion of the digestion period, the digested adipose tissue should have a “soup-like” consistency. 6. Add DMEM/F12 containing 10% FBS, to stop collagenase activity (see Note 2).

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3.1.3 Separation of the Stromal Vascular Fraction

1. After digestion, 70 μm cell strainers (Falcon®) are used to separate any undigested adipose tissue; then, centrifuge at room temperature at 300  g for 10 min. 2. After centrifugation, use a 50 ml pipette to aspirate the floating adipocytes, lipids, and the digestion medium. Leave the stromal vascular fraction (SVF) pellet in the tube.

3.1.4 Culture of AdiposeDerived Stromal Cells

1. Resuspend 105 freshly isolated stromal stem cells in 2 ml of DMEM: F12 (Gibco, USA) medium containing 10% FBS and 1% penicillin/streptomycin (Gibco, USA). 2. Add the cell suspension to a 75 cm2 cell culture flask. 3. Incubate in a humidified incubator at 37  C with 5% CO2 for 2 min. 4. Cells should be daily observed using an inverted phase contrast microscope. 5. It usually takes several days before cells forming a fibroblastic morphology and starting to divide. 6. Estimate the percentage of cells which adhere to the plastic surface and form a fibroblastic-like morphology after 8 days of culture (see Note 3).

3.1.5 Subculture of Adipose-Derived Stromal Cells

1. Wash confluent primary culture of adipose-derived stromal cells with PBS (two times). 2. Add 2 ml of trypsin/EDTA 0.05% to 75 cm2 culture flasks (TPP, Germany). 3. Incubate in a humidified incubator at 37  C with 5% CO2. 4. After completion of the isolation period, add 2 ml of DMEM: F12 (Gibco, USA) medium containing 10% FBS and 1% penicillin/streptomycin (Gibco, USA) to cell suspension for enzyme inactivation. 5. Centrifuge each cell suspension at 300  g for 5 min then, resuspend the cell pellet in normal saline, and count the cells using a hemocytometer (Bright-Line™, USA). 6. Resuspend 5  105 freshly isolated adipose-derived stromal cells in 15 ml of DMEM: F12 (Gibco, USA) medium containing 10% FBS and 1% penicillin/streptomycin (Gibco, USA) and seed them in a 150 cm2 culture flask. 7. Incubate ASCs at 37  C in a humidified atmosphere with 5% CO2 for up to three passages (Fig. 1).

3.2 Flow Cytometry Analysis for Further Characterization of Cultured Cells

1. Resuspend 105 human adipose-derived stromal cells at passage 3 in a 3% bovine serum albumin (BSA; Sigma-Aldrich, USA) solution and incubate for 20 min at room temperature.

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Fig. 1 Human adipose-derived stromal cells at passage 3. (Adopted from cellbased skin substitutes accelerate regeneration of extensive burn wounds in rats by Motamed et al. [14])

2. Wash ASCs with PBS twice (each time for 5 min); then, centrifuge at 300  g for 5 min. 3. Add antibodies for specific CD markers including CD90, CD105, CD73, CD45/34 and IgG1 isotype control (Sigma, USA) at 1:100 dilution to each tube and incubate for 25 min at 4  C, separately (see Notes 4 and 5). 4. Centrifuge tubes containing ASCs at 300  g for 5 min. 5. Wash cells with PBS (Gibco, USA), centrifuge and analyze using a FACSCalibur flow cytometer (BD Biosciences, USA) using the CELL Quest software version 3.3 (Fig. 2). 3.3 Differentiation Potential of AdiposeDerived Stromal Cells 3.3.1 Osteogenic Differentiation

1. Harvest the cells in an osteogenic differentiation medium containing L-DMEM, 10% FBS, 0.1 μM dexamethasone (SigmaAldrich), 200 μM L-ascorbic acid-2-phosphate (SigmaAldrich), and 10 mM β-glycerol phosphate (Sigma-Aldrich) for 21 days and confirm the osteogenic induction by Alizarin red S staining. 2. Alizarin red-positive cells (differentiated ASCs with calcium deposits) seen bright orange-red under the microscope, while negative cells (undifferentiated ASCs without calcium deposits) appear as faintly reddish cells (Fig. 3a).

3.3.2 Chondrogenic Differentiation

1. Harvest cells in a chondrogenic differentiation medium containing Dulbecco’s Modified Eagle’s Medium, high (4.5 g/L) glucose (DMEM-HG, Invitrogen) supplemented with 10% ITS + Premix Tissue culture supplement (Becton Dickinson),

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G1-FITC/G1-PE

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Fig. 2 Dot plots and histograms corresponding to the analysis of human adipose-derived stromal cells suspension. Characterization of the adipose-derived stromal cells in terms of CD markers: CD90, CD105, CD73, and CD45/34. The bars on the peak levels of the red histogram (Isotope control) and green histogram are a measure of positive expression of CD markers: CD90, CD105, CD73, and CD45/34. (Adopted from cellbased skin substitutes accelerate regeneration of extensive burn wounds in rats by Motamed et al. [14])

7–10 M dexamethasone (Sigma), 1 μM ascorbate-2-phosphate (Wako, Richmond, VA), 1% sodium pyruvate (Invitrogen), and 10 ng/ml transforming growth factor-beta 1 (TGF-β1, PeproTech, Rocky Hill, NJ), for 21 days and confirm the chondrogenic induction using dexamethasone. Observation of dexamethasone-positive cells under microscope proves the chondrogenic differentiation of ASCs (Fig. 3b). 3.3.3 Adipogenic Differentiation

Seed cells into a medium consisting of DMEM/10% FBS, 50 μmol/L indomethacin, 10 μM insulin, 1 μmol/L dexamethasone, and 0.5 mM 3-isobutyl-1-methyl-xanthine (all from SigmaAldrich) for 2 weeks and confirm the adipogenic induction using Oil red O staining according to the standard method. Observation of Oil red droplets in microscopic evaluation proves the adipogenic differentiation of ASCs (Fig. 3c).

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Fig. 3 (a) Adipogenic differentiation, (b) Chondrogenic differentiation, and (c) Osteogenic differentiation. (Adopted from cell-based skin substitutes accelerate regeneration of extensive burn wounds in rats by Motamed et al. [14]) 3.4 Preparation of Human Amniotic Membrane 3.4.1 Collection of HAMs

3.5

HAM Preparation

In this experimental study, after obtaining informed consents from the subjects, human placentas are taken from HAMs bank, Royan Institute public cord blood bank, following the approval of the Ethics Committee. All placenta donors are serologically negative for human immunodeficiency virus, hepatitis virus type B, hepatitis virus type C, and syphilis. The placentas are washed 3 times with PBS (pH 7.4; Gibco, USA) in class 2 laminar flow. After separation of amniotic membrane from the underlying chorion and cutting into approximately 10  10 cm2 pieces, the pieces are stored in PBS containing 1.5% dimethyl sulfoxide (DMSO) at 70  C for up to 5 months. HAMs are prepared from HAMs bank, Royan Institute public cord blood bank. 1. Incubate 10  10 cm2 pieces of HAMs in trypsin–EDTA (ethylene-diamine-tetraacetic acid) solution at 37  C for 5 min. 2. Slowly remove HAM epithelial cells by a cell scraper, then wash for three times with PBS (Gibco, USA).

3.6 Cultivation of Human ASCs on Skin Substitute

1. Seed human ASCs on two 10  10 cm2 pieces of the acellular amniotic membrane at a density of 5  105/cm2 and incubate in a humidified incubator at 37  C with 5% CO2, for 7 days. 2. Add complete medium containing Dulbecco’s Modified Eagle’s Medium/Nutrient Mixture F-12 HAM solution (DMEM/F12) (Gibco, USA), 10% fetal bovine serum (Gibco, USA), 1% L-glutamine (Gibco, USA), and 1% penicillin/streptomycin (Gibco, USA). 3. Change the medium every 3 days (Fig. 4).

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Fig. 4 Human adipose-derived stromal cells (black arrows) 8 days after seeding on the human amniotic membrane. (Adopted from cell-based skin substitutes accelerate regeneration of extensive burn wounds in rats by Motamed et al. [14]) 3.7 Immunohistochemical (IHC) Analysis of the Skin Substitute

1. Fix the skin substitute for IHC staining using 4% paraformaldehyde (Sigma, USA) for 5 min (see Note 6). 2. Cultivate the adipose-derived stromal cells on the amniotic membrane and wash two times (each time for 5 min) with Dulbecco’s phosphate buffered saline (PBS/Tween (PBST) 0.05%; Sigma, USA) (see Note 6). 3. Determine the cell membrane permeability by adding Triton X-100 (MERK, Germany, 0.2%) and incubating at room temperature for 10 min. 4. Wash cells twice with PBS/Tween 0.05% (each time for 5 min) (see Note 6). 5. Use blocking buffer containing 10% goat serum (Sigma, USA) to block non-specific antibody binding and then incubate at 37  C for 20 min. 6. Wash ASCs twice (each time for 5 min) with PBS/Tween 0.05% (see Note 6). 7. Add primary antibodies including vimentin (Sigma, USA) at 1:100 dilution and incubate at 4  C for 15 h (see Note 4). 8. Wash the cells twice (each time for 10 min) with PBS/Tween 0.05% (see Note 6). 9. Add fluorescein isothiocyanate (FITC)-conjugated goat antimouse (Dako, USA) secondary antibody at 1:200 dilution and incubate at 37  C for 1 h (see Notes 4 and 5). 10. Finally, wash the cells with PBST and examine under a fluorescent microscope. Use DAPI dye for nuclear staining (Fig. 5).

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Fig. 5 Immunostaining of human adipose-derived stromal cells cultivated on HAM for 8 days (a) more than 90% of cells are positive for vimentin staining. (b) DAPI dye was used for nuclear staining. (Adopted from cellbased skin substitutes accelerate regeneration of extensive burn wounds in rats by Motamed et al. [14])

4

Notes 1. For maximum viability, liposuction sample should be processed as soon as possible after surgery. However, if necessary, tissue can be kept at 4  C for 24 h without considerable loss of viability. The yield of ASC per 1 ml of liposuction sample depends on the health status and age of the donor. Younger donors generate more stem cells than older ones. 2. If the liposuction sample is not completely digested, keep the tube at room temperature and remove the undigested fat tissues using sterile cell strainers mesh. 3. The adipose-derived stromal cells may not be sufficiently adherent cells at 24 h. Wait for at least 3 days and allow cells to adhere to the bottom of cell culture flask. 4. The serial concentration of primary antibody during immunostaining and flow cytometry procedures should be checked by a researcher to reach the optimal concentration. 5. The correct isotype-specific secondary antibody should be used for each primary antibody. 6. Do not let cells dry out during immunostaining procedure.

Acknowledgments The protocols described here were developed with the support of Royan Institute grant.

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References 1. Litwiniuk M, Grzela T (2014) Amniotic membrane: new concepts for an old dressing. Wound Repair Regen 22:451–456 2. Niknejad H, Peirovi H, Jorjani M, Ahmadiani A, Ghanavi J, Seifalian AM (2008) Properties of the amniotic membrane for potential use in tissue engineering. Eur Cell Mater 15:88–99 3. Taghiabadi E, Nasri S, Shafieyan S, Jalili Firoozinezhad S, Aghdami N (2015) Fabrication and characterization of spongy denuded amniotic membrane based scaffold for tissue engineering. Cell J 16:476–487 4. Lo V, Pope E (2009) Amniotic membrane use in dermatology. Int J Dermatol 48:935–940 5. Hao Y, Ma DH, Hwang DG, Kim WS, Zhang F (2000) Identification of antiangiogenic and antiinflammatory proteins in human amniotic membrane. Cornea 19:348–352 6. Tseng SC, Li DQ, Ma X (1999) Suppression of transforming growth factor-beta isoforms, TGF-beta receptor type II, and myofibroblast differentiation in cultured human corneal and limbal fibroblasts by amniotic membrane matrix. J Cell Physiol 179:325–335 7. Stock SJ, Kelly RW, Riley SC, Calder AA (2007) Natural antimicrobial production by the amnion. Am J Obstet Gynecol 196:255. e1–255.e6 8. Kim SS, Song CK, Shon SK, Lee KY, Kim CH, Lee MJ et al (2009) Effects of human amniotic membrane grafts combined with marrow mesenchymal stem cells on healing of full-thickness skin defects in rabbits. Cell Tissue Res 336:59–66

9. Marquez-Curtis LA, Janowska-Wieczorek A, McGann LE, Elliott JA (2015) Mesenchymal stromal cells derived from various tissues: biological, clinical and cryopreservation aspects. Cryobiology 71:181–197 10. Jin HJ, Bae YK, Kim M, Kwon SJ, Jeon HB, Choi SJ et al (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 14:17986–18001 11. Faramarzi H, Mehrabani D, Fard M, Akhavan M, Zare S, Bakhshalizadeh S et al (2016) The potential of menstrual bloodderived stem cells in differentiation to epidermal lineage: a preliminary report. World J Plast Surg 5:26–31 12. Ledesma-Martinez E, Mendoza-Nunez VM, Santiago-Osorio E (2016) Mesenchymal stem cells derived from dental pulp: a review. Stem Cells Int 2016:4709572 13. Arno A, Smith AH, Blit PH, Shehab MA, Gauglitz GG, Jeschke MG (2011) Stem cell therapy: a new treatment for burns? Pharmaceuticals 4:1355–1380 14. Motamed S, Taghiabadi E, Molaei H, Sodeifi N, Hassanpour SE, Shafieyan S et al (2017) Cell-based skin substitutes accelerate regeneration of extensive burn wounds in rats. Am J Surg 214:762–769 15. Chen JY, Mou XZ, Du XC, Xiang C (2015) Comparative analysis of biological characteristics of adult mesenchymal stem cells with different tissue origins. Asian Pac J Trop Med 8:739–746

Methods in Molecular Biology (2019) 1879: 211–219 DOI 10.1007/7651_2018_135 © Springer Science+Business Media New York 2018 Published online: 08 May 2018

Amniotic Membrane Seeded Fetal Fibroblasts as Skin Substitute for Wound Regeneration Ehsan Taghiabadi, Bahareh Beiki, Nasser Aghdami, and Amir Bajouri Abstract Various natural and synthetic biomaterials have been applied as skin substitutes for regenerating damaged skin. Here, we describe a straightforward method for fabrication of a tissue-engineered skin substitute by seeding human fetal fibroblasts on acellular human amniotic membrane (HAM). Fetal fibroblasts are achieved from the skin of normal and non-macerated fetus of 11–14 weeks old after spontaneous pregnancy termination. Acellular HAM is obtained by separation of the outer membrane of the chorion and removing its epithelial cells. Keywords Amniotic membrane, Fetal fibroblast, Skin substitutes, Stem cell, Tissue engineering, Wound healing

1

Introduction Considering the importance of fetal fibroblasts in accelerating fetal wound healing, these cells are an attractive choice for cell-based engineered skin substitutes [1, 2]. Fetal fibroblasts possess paracrine activities which are mediated through secretion of several growth factors and cytokines such as basic fibroblast growth factor (bFGF), vascular endothelial growth factor (VEGF), keratinocyte growth factor (KGF), hepatocyte growth factor (HGF), and transforming growth factor beta (TGF-β), which have been demonstrated to accelerate wound regeneration [3]. Moreover, fetal fibroblasts have high expansion capacity under simple culture condition and do not evoke immunological responses in the recipient site [4]. During the past decades, because of its biological and mechanical characteristics, human amniotic membrane (HAM) has been widely applied for wound regeneration. Amniotic membrane contains an epithelial cell layer, a basilar membrane layer, and a compact layer. The extracellular matrix of basilar membrane that contains collagens I, III, and IV, heparan sulfate, proteoglycan, and fibronectin provides mechanical support for HAM [5, 6]. Moreover, by secreting high levels of various growth factors [5], HAM exerts

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anti-inflammatory [7], anti-fibrotic [8], and antimicrobial [9] properties that are beneficial to wound regeneration. Here, we explain a method for fabrication of a two-dimensional (2D) skin substitute from fetal fibroblasts and intact human amniotic membrane. Fetal fibroblasts are achieved from skin of normal and non-macerated fetus of 11–14 weeks old after spontaneous pregnancy termination. Acellular HAM is obtained by separation of the outer membrane of the chorion and removing its epithelial cells. Finally, human fetal fibroblasts are seeded on pieces of the acellular amniotic membrane and incubated.

2 2.1

Materials Tissue

1. Human amniotic membrane was obtained from human placentas following ethics committee approval. 2. Fetal fibroblasts were obtained from the Royan cell bank, Tehran, Iran. Fetal skin of normal and non-macerated fetus of 11–14 weeks old was collected after spontaneous pregnancy termination. None of the donors had human immunodeficiency virus, hepatitis B and C, or syphilis. They all signed an informed consent before donating the fetus.

2.2 Reagents and Medium

1. Phosphate buffered saline (PBS), without Ca pH ¼ 7.4 (Gibco, USA).

2+

or Mg

2+

at

2. Dimethyl sulfoxide (DMSO; Sigma, USA). 3. Dulbecco’s Modified Eagle’s Medium/Nutrient Mixture F-12 HAM solution (DMEM/F12) (Gibco, USA). 4. 10,000 U/ml penicillin and 10,000 mg/ml streptomycin (Gibco, USA). 5. Fetal bovine serum (FBS; Cat. No. 10099-141, Gibco-BRL) was aliquoted, stored frozen, and thawed at 4  C. 6. Collagenase A type I (Cat. No. C-0130, Sigma-Aldrich). 7. Falcon 70 μm cell strainers (Becton Dickinson). 8. Plastic conical tubes (50 and 15 ml; TPP, Germany). 9. Disposable plastic pipettes (50, 25, and 10 ml; TPP, Germany). 10. Mouse anti-vimentin monoclonal antibody (Abcam, USA). 11. Goat anti-mouse fluorescein conjugated (Dako, USA).

isothiocyanate

(FITC)-

12. Mouse anti-human CD90 FITC-conjugated monoclonal antibody (BD, USA). 13. Mouse anti-human CD73 PE-conjugated monoclonal antibody (BD, USA). 14. Mouse anti-human CD29 PE-conjugated monoclonal antibody (BD, USA).

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Methods

3.1 Primary Human Fetal Fibroblast Culture

1. Human fetal fibroblasts were obtained from Department of Regenerative Medicine of Royan Institute cell bank, Tehran, Iran. 2. Resuspend 5  105 thawed fetal fibroblasts (passage 1) in 3 ml of DMEM/F12 (Gibco, USA) medium containing 10% FBS and 1% penicillin/streptomycin (Gibco, USA). 3. Add the cell suspension to a 25-cm2 cell culture flask. 4. Incubate the cells in a humidified incubator at 37  C with 5% CO2. 5. Observe the cells daily using an inverted phase contrast microscope. 6. Usually, after 2–3 days, cells with a fibroblastic morphology start dividing. 7. After 24 h, estimate the percentage of cells which adhere to the plastic surface (see Note 1). 8. The 25-cm2 culture flasks become confluent within 6 days.

3.2 Subculture of Fetal Fibroblasts

1. Wash confluent primary culture of human fetal fibroblasts with PBS (two times). 2. Add 2 ml of trypsin/EDTA 0.05% into 75-cm2 culture flasks. 3. Incubate the cells in a humidified incubator at 37  C with 5% CO2 for 2 min. 4. After completion of the isolation period, add 2 ml of DMEM/ F12 (Gibco, USA) medium containing 10% FBS and 1% penicillin/streptomycin (Gibco, USA) to cell suspension for enzyme inactivation. 5. Each cell suspension should be centrifuged at 1500 rpm for 5 min; then, resuspend cell pellet in normal saline for cell counting using a hemocytometer (Bright-Line™, USA). 6. Resuspend 5  105 freshly isolated adipose-derived stem cells in 10 ml of DMEM/F12 (Gibco, USA) medium containing 10% FBS and 1% penicillin/streptomycin (Gibco, USA), and seed them into 75-cm2 culture flasks. 7. Incubate fetal fibroblasts at 37  C in a humidified atmosphere with 5% CO2, for up to three passages (Fig. 1).

3.3 Morphological Analysis

In vitro expanded fetal fibroblasts at passage 3 were evaluated under inverted microscope Olympus CKX-41.

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Fig. 1 Appearance of fibroblasts cultured on tissue culture plastic at passage 3 (Adopted from Cell-based skin substitutes accelerate regeneration of extensive burn wounds in rats by Motamed et al. 2017)

3.4 Flow Cytometry Analysis for Further Characterization of Cultured Cells

1. Resuspend 105 human fetal fibroblasts at passage 3, in a 3% bovine serum albumin (BSA; Sigma-Aldrich, USA) solution, and incubate for 20 min at room temperature. 2. Wash human fetal fibroblasts twice (each time for 5 min) with PBS; then, centrifuge at 1500 rpm for 5 min. 3. Add specific fetal fibroblast markers including vimentin, CD29, CD73, CD90, and IgG1 isotype control (Sigma, USA) at 1/100 dilution to each tube, and incubate for 25 min at 4  C (see Notes 3 and 4). 4. Centrifuge the tubes that contain fetal fibroblasts at 1500 rpm for 5 min. 5. Wash the cells with PBS, centrifuge, and analyze by a FACSCalibur Flow Cytometer (BD Biosciences, USA) using the CELL Quest software version 3.3 (Fig. 2).

3.5 Immunocytochemical (ICC) Analysis for Characterization of Cultivated Fetal Fibroblasts

1. Fix human fetal fibroblasts for ICC staining using paraformaldehyde 4% (Sigma, USA) for 5 min (see Note 2). 2. Wash fetal fibroblasts twice with Dulbecco’s PBS (PBS/Tween (PBST); 0.05%, Sigma, USA) for 5 min. 3. Determine the permeability of the cells by adding Triton X-100 (MERK, Germany, 0.2%) followed by incubation for 10 min at room temperature. 4. Wash fetal fibroblasts twice (each time for 5 min) with PBS/Tween 0.05%.

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Fig. 2 Flow cytometry diagrams. Dot plot and histogram analysis of human fetal fibroblasts cytoplasmic markers: CD29, CD73, CD90, and vimentin. The peak levels of black and red histograms are isotope control and markers expression, respectively (Adopted from Cell-based skin substitutes accelerate regeneration of extensive burn wounds in rats by Motamed et al. 2017)

5. Block the cells by blocking buffer containing 10% goat serum (Sigma, USA), and then incubate for 20 min at 37  C, to block non-specific antibody binding. Then, wash twice each time for 5 min. 6. Add primary antibody for vimentin (Sigma, USA), CD29, CD73, and CD90 at 1/100 dilution, and incubate for 2 h at room temperature (see Note 4). 7. Wash the cells twice (each time for 10 min) with PBS/Tween 0.05%.

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Fig. 3 Fetal fibroblasts cultured on 4-well culture plates and immunostained with vimentin markers. This representative figure illustrates the cultured cells positive for fibroblast marker (vimentin). Fluorescent compounds, (a) vimentin (green) and (b) DAPI (Blue), were used to stain the cells and nucleus

8. Add goat anti-mouse FITC-conjugated (Dako, USA) secondary antibody at 1/200 dilution, and incubate for 1 h at room temperature (see Notes 3 and 4). 9. Add 10 μl DAPI dye for nuclear staining. 10. Wash the fetal cells with PBS and examine under a fluorescent microscope (Fig. 3). 3.6 Gene Expression Analysis by Reverse Transcriptase Polymerase Chain Reaction (RT-PCR)

The expression of type I collagen in fibroblasts was evaluated by RT-PCR technique that is presented in Fig. 4. Expression level of the targeted gene was normalized against GAPDH. The following primers, specific for GAPDH sequences, were used for RT-PCR: forward, 50 ATGCCTGGTGAACGTGGT30 , and reverse, 0 5 AGGAGAGCCATCAGCACCT30 . Targeted primer was designed with primer 3. 1. Extract the total RNA from fibroblasts using trayzol (Sigma, USA). 2. Incubate the extracted RNA with 1 U/ml of RNase-free DNase I (EN0521, Fermentas, Germany) per 1 mg of RNA, in order to eliminate residual DNA in the presence of 40 U/ml of ribonuclease inhibitor (E00311, Fermentas, Germany) and 1 reaction buffer with MgCl2(Sigma, USA), for 30 min at 37  C. 3. Add 1 ml of 25 mM EDTA (Sigma, USA) and incubate at 65  C for 10 min to inactivate DNase I. 4. Perform standard reverse transcriptase (RT) reactions with 2 μg total RNA using oligo (dt) (Fermentas, Germany) as a primer and a Revert Aid TM First Strand cDNA synthesis kit (K1622, Fermentas, Germany) based on the manufacturer’s instructions.

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Fig. 4 For different passages of human fetal fibroblasts, namely, passages 1, 2, and 3, no difference was observed in terms of the expression of collagen I RNAs as compared with normal human skin tissue. NTC, no template control; Coll 1, collagen I expression at passage 1; Coll 2, collagen I expression at passage 2, Coll 3, collagen I expression at passage 3, RT, negative control; and GAPDH, glyceraldehyde 3-phosphate dehydrogenase

5. For every reaction set, prepare one RNA sample without Revert Aid TMM MuLV RTreaction to provide a negative control in the subsequent PCR. 6. To minimize variation in the RT reaction, all RNA samples from a single experimental setup should simultaneously undergo reverse transcription. Reaction mixtures for PCR include 2 ml cDNA, 1 PCR buffer (AMSTM, CinnaGen Co., Tehran, Iran), 200 mM dNTPs, 0.5 mM of each antisense and sense primers (AMSTM, CinnaGen Co., Tehran, Iran), and 1 U Taq DNA polymerase (AMSTM, CinnaGen Co., Tehran, Iran). 7. The accession number of primer is NM-000088.3 and length of ladder is 50 base pairs (bp). 8. Use the following primers, specific for human collagen type I chain sequences, for RT-PCR: Forward: 50 TTGCCGACAGGATGGAGAAGGA30 Reverse: 50 AGGTGGACAGCGAGGCCAGGAT30 3.7

HAM Preparation

HAMs were prepared in HAM bank which is part of the public cord blood bank, Royan Institute. 1. Incubate 10  10 cm2 pieces of HAMs in trypsin/EDTA (ethylenediaminetetraacetic acid) solution at 37  C for 5 min. 2. Slowly remove HAM epithelial cells by a cell scraper, and then wash three times with PBS (Gibco, USA).

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Fig. 5 (a) Human fetal fibroblasts 8 days after seeding on human amniotic membrane. (b) Immunostaining of human fetal fibroblasts cultured on HAM for 8 days; more than 90% of cells are positive for vimentin staining. (c) DAPI dye was used for nuclear staining (Adopted from Cell-based skin substitutes accelerate regeneration of extensive burn wounds in rats by Motamed et al. 2017)

3.8 Cultivation of Human Fetal Fibroblasts on Human Amniotic Membrane

1. Seed human fetal fibroblasts on two pieces (10  10 cm2) of the acellular amniotic membrane at a density of 5  105/cm2, and incubate in a humidified incubator at 37  C with 5% CO2 for 7 days. 2. Add complete medium containing Dulbecco’s Modified Eagle’s Medium/Nutrient Mixture F-12 HAM solution (DMEM/F12) (Gibco, USA), 10% fetal bovine serum (Gibco, USA), 1% L-glutamine (Gibco, USA), and 1% penicillin/streptomycin (Gibco, USA). 3. Change the medium every 3 days (Fig. 5a).

3.9 Immunohistochemical (IHC) Analysis for Characterization of Skin Substitute

1. Fix skin substitute for IHC staining using paraformaldehyde 4% (Sigma, USA) for 5 min (see Note 2). 2. Wash fetal fibroblasts cultivated on amniotic membrane, twice (each time for 5 min) using Dulbecco’s PBS [PBS/Tween (PBST) 0.05%; Sigma, USA]. 3. Determine the permeability of the cells by adding Triton X-100 (MERK, Germany, 0.2%) followed by incubation for 10 min at room temperature. 4. Wash the cells twice (each time for 5 min) with PBS/Tween 0.05%. 5. Block the cells by blocking buffer containing 10% goat serum (Sigma, USA); then, incubate for 20 min at 37  C, to block non-specific antibody binding. 6. Wash fetal fibroblasts twice (each time for 5 min) with PBS/Tween 0.05%. 7. Add primary antibodies including vimentin (Sigma, USA) at 1/100 dilution, and incubate for 15 h at 4  C (see Note 4).

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8. Wash the cells twice (each time for 10 min) with PBS/ Tween 0.05%. 9. Add FITC-conjugated goat anti-mouse (Dako, USA) secondary antibody at 1/200 dilution, and incubate for 1 h at 37  C (see Notes 3 and 4). 10. Finally, wash the cells with PBST and examine under a fluorescent microscope. Use DAPI dye for nuclear staining (Fig. 5b, c).

4

Notes 1. Not many fetal fibroblast might be achieved within 24-h adherent cells at 24 h. Allow the cells, for at least 3 days, to adhere to the bottom of cell culture flask. 2. Do not let cells dry out during immunostaining procedure. 3. Serial concentration of primary antibody during immunostaining and flow cytometry procedures should be checked to determine the optimal concentration. 4. The correct isotype-specific secondary antibody should be used for each primary antibody.

Acknowledgments The protocols described here were developed under the support of a Royan institute grant. References 1. Pouyani T, Papp S, Schaffer L (2012) Tissueengineered fetal dermal matrices. In Vitro Cell Dev Biol Anim 48:493–506 2. Motamed S, Taghiabadi E, Molaei H, Sodeifi N, Hassanpour SE, Shafieyan S et al (2017) Cellbased skin substitutes accelerate regeneration of extensive burn wounds in rats. Am J Surg 214:762–769 3. Litwiniuk M, Grzela T (2014) Amniotic membrane: new concepts for an old dressing. Wound Repair Regen 22:451–456 4. Zuliani T, Saiagh S, Knol AC, Esbelin J, Dreno B (2013) Fetal fibroblasts and keratinocytes with immunosuppressive properties for allogeneic cell-based wound therapy. PLoS One 8:e70408 5. Niknejad H, Peirovi H, Jorjani M, Ahmadiani A, Ghanavi J, Seifalian AM (2008) Properties of the amniotic membrane for potential use in tissue engineering. Eur Cell Mater 15:88–99

6. Taghiabadi E, Nasri S, Shafieyan S, Jalili Firoozinezhad S, Aghdami N (2015) Fabrication and characterization of spongy denuded amniotic membrane based scaffold for tissue engineering. Cell J 16:476–487 7. Hao Y, Ma DH, Hwang DG, Kim WS, Zhang F (2000) Identification of antiangiogenic and antiinflammatory proteins in human amniotic membrane. Cornea 19:348–352 8. Tseng SC, Li DQ, Ma X (1999) Suppression of transforming growth factor-beta isoforms, TGF-beta receptor type II, and myofibroblast differentiation in cultured human corneal and limbal fibroblasts by amniotic membrane matrix. J Cell Physiol 179:325–335 9. Stock SJ, Kelly RW, Riley SC, Calder AA (2007) Natural antimicrobial production by the amnion. Am J Obstet Gynecol 196(255):e1–e6

Methods in Molecular Biology (2019) 1879: 221–241 DOI 10.1007/7651_2018_161 © Springer Science+Business Media New York 2018 Published online: 24 May 2018

Skin Wound Healing: Refractory Wounds and Novel Solutions Gabriel M. Virador, Lola de Marcos, and Victoria M. Virador Abstract This overview of the current state of skin wound healing includes in vitro and in vivo approaches along with some recent clinical trials. From an introduction to wound healing, to tissue engineering as applied to the skin, we cover the basis for the current wound care techniques as well as novel and promising approaches. Special emphasis is given to refractory wounds which include wounds in diabetic patients. Natural compounds have been ever present in wound healing, and so we devote a section to highlighting current attempts to understand their mechanisms and to use them in novel ways. Keywords Biomaterials, Cellular therapies, Natural compounds, Refractory wounds, Stem cells, Tissue engineering, Wound healing

1

Introduction

1.1 Skin Wound Healing: Definition and History

In mammals, the skin is the primary protection barrier against physical damage. As wounding disrupts epithelial tissue and its local environment, wound-healing mechanisms become activated at the cellular and tissue levels. A PubMed search of the term wound, which returned more than a million results, is an indication of the extent to which scientists and physicians have tried to understand, mimic, and accelerate wound healing for the benefit of patients. While healing the skin using traditional medicines has been used for generations, the practice has evolved to include the current tissue engineering scientific disciplines as well as a greater understanding of the mechanisms of action of traditional remedies. Here we review several aspects of this ancient art while highlighting some exciting developments in tissue engineering and in the application of natural compounds to wound healing.

1.2 Phases of Wound Healing

Four phases have been described in wound healing: hemostasis, inflammation, proliferation, and maturation (described in Medscape, http://emedicine.medscape.com/article/884594-overview. Accessed 11 March, 2018). As soon as tissue is damaged, platelets start the clot formation of hemostasis. Damaged cells are removed

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during the inflammatory phase, while the proliferative phase is starting in a sequential, yet overlapping, manner, to achieve epithelialization, fibroplasia, and angiogenesis. Formation of granulation tissue during the proliferative phase (3–5 days following the wound) is the most critical element of wound healing as it sets the stage for proper contraction and maturation. This phase, which encompasses synthesis, deposition, and cross-linking of collagen, results in the contraction of the wound. The wound closes toward the center at a rate which depends on its shape and tissue density. Maturation is the final phase in skin wound healing, and it mainly involves the development of an epithelial barrier layer [1]. Fibroblasts are key in the generation of new extracellular matrix, deposition of collagen, and wound contraction, all important aspects of the granulation tissue, and therefore have potential in cellular therapies [2]. They comprise various specialized populations which secrete fibrous materials such as collagen, elastin, fibronectin, glycosaminoglycans, and proteases. Fibroblasts thus contribute to the remodeling of the wound microenvironment by providing tensile strength to the scar. Other cells present in the granulation tissue include inflammatory cells, of crucial importance in wound healing as they can favor or impair regeneration [3], as well as cells involved in the formation of new blood vessels [4]. Melanocytes do not seem to migrate into granulation tissue, and thus this tissue is typically hypopigmented with variable repigmentation of cutaneous wounds depending on the wound type [5]. Myofibroblasts, a differentiated subpopulation of fibroblasts, drive wound closure by their interactions with components of the extracellular matrix and undergo apoptosis when no longer needed [2]. 1.3 Classes of Wounds

There is no single system for wound classification. For example, war wounds have been classified following Red Cross guidelines [6] by shape and depth of the wounds. Other guidelines revolve around their infectibility. The American College of Surgeons, cited by the Centers for Disease Control and Prevention (CDC), classifies surgical wounds into four categories based on whether they are clean, contaminated, or infected [7]. But the validity of the surgical wound classification has been questioned based on its inconsistent application in different institutions [8], and the CDC issued recent guidelines about preventing surgical site infections [9]. Depending on the time to heal, wounds are referred to as acute or chronic, but these are descriptive terms, rather than a classification, as all chronic wounds start as acute wounds. Complicated wounds, defined as those that include tissue disruption and infection, seem to be in a class by themselves (https://emedicine.medscape.com/article/ 1129806-overview. Accessed 11 March, 2018).

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1.4 Refractory Wounds

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Poor or altered wound healing occurs in several medical conditions such as in the following examples: Diabetics are at higher risk of developing diminished peripheral blood flow, resulting in delayed or impaired healing. Diabetic leg ulcers constitute a major complication of this metabolic disease, affecting 15% of those with diabetes [10], and type 2 diabetes is the leading cause of amputation in the developed world. With over 30 million diabetics in the United States alone (https://www.cdc. gov/diabetes/new/index.html, accessed March 11, 2018), they are an important demographic to consider when researching new ways to treat wounds. Diabetic wound models aim to recreate an impaired blood flow and a hypoxic wound environment, where there is decreased collagen production and immune function. Significant differences are found between diabetic and nondiabetic wounds at the cellular level as detailed in Fig. 1. Keloids and hypertrophic scarring of the skin are associated with excessive fibrosis [11]. It has long been held that inflammationpromoted fibrosis causes scarring and limits wound regeneration, as is seen in the bariatric population which suffers of chronic wounds, with a prolonged inflammatory phase, abnormally high levels of proteases, and low levels of growth factors [12]. Interestingly, fetal wounds and oral mucosa heal without scarring, and therefore it is of great interest to understand the mechanisms involved [13, 14]. Wounds in autoimmune diseases such as rheumatoid arthritis, lupus, and scleroderma are prone to complicated infections [15]. Epidermolysis bullosa is a rare genetic blistering disease which presents multiple challenges in wound care [16]. Often, delayed wound healing is associated with infection by the opportunistic pathogen Staphylococcus aureus; according to a recent study, patients with EB, unlike the general population, are susceptible to infection by more than one S. aureus type [17]. Clearly the challenges in treatment of refractory wounds require novel approaches which are partly being met by a number of issued patents [18].

Skin Wound Healing and Cellular Therapies

2.1 Skin Grafts, Acellular Tissue, and Scaffolds

Grafts of a patient’s own tissue have been used for many years, with early descriptions of autologous skin flaps dating as far back as 600 BC [19]. In the case of autologous grafts, new treatments in tissue engineering usually begin by obtaining skin and tissue samples from an unwounded portion of the patient’s body using a variety of protocols. Cells from these samples are put in tissue culture, formed into tissue constructs, and then placed or infused onto the wounded areas. These methods can decrease healing time and aid in the treatment of difficult wounds such as ulcers by reducing risk of infection, increasing rate of wound closure, and aiding in pain

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Fig. 1 Mechanisms of wound healing in healthy versus diabetic wounds. In healthy individuals (left), the acute wound-healing process is guided and maintained through integration of multiple signals (in the form of cytokines and chemokines) released by keratinocytes, fibroblasts, endothelial cells, macrophages, and platelets. During wound-induced hypoxia, VEGF released by macrophages, fibroblasts, and epithelial cells induces the phosphorylation and activation of eNOS in the bone marrow, resulting in an increase in NO levels, which triggers the mobilization of bone marrow EPCs to the circulation. The chemokine SDF-1a promotes the homing of these EPCs to the site of injury, where they participate in neovasculogenesis. In a murine model of diabetes (right), eNOS phosphorylation in the bone marrow is impaired, which directly limits EPC mobilization from the bone marrow into the circulation. Moreover, SDF-1a expression is decreased in epithelial cells and myofibroblasts in the diabetic wound, which prevents EPC homing to wounds and therefore limits wound healing. Reproduced, with permission, from Brem et al., J Clin Invest. 2007;117(5):1219–1222. doi:https://doi. org/10.1172/JCI32169

management. The use of autologous cells is ideal as it lessens the risk of immune rejection or graft versus host disease compared to the use of donor tissue [20]. De-cellularized tissue has been used in skin wounds in an effort to provide physiological coverage. It has also been used in bioink as reviewed in [21]. Recently, Jakus et al. published an interesting study in which they described the fabrication of de-cellularized biomaterials from various porcine tissues. The authors made paper matrices using dried extracellular matrix deposited by specific cells. This bioactive tissue paper can be made from various organs and is pliable enough to fold into origami

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structures. More importantly, the authors demonstrate the papers’ ability to support growth of the corresponding primary cells thus conceivably providing the best homing material for viable 3D tissue reconstructs [22]. 2.2 Platelet-Rich Plasma and Other Blood-Related Products

Platelet-rich plasma (PRP) has seen a wide variety of uses over the past two decades including injection into joints to treat osteoarthritis symptoms [23] and to enhance healing in different kinds of wounds. PRP is autologous serum with a high concentration of platelets which can be obtained via various commercially available methods [24]. PRP contains elevated levels of growth factors [25] which contribute to wound healing [26]. Among the advantages of its use, compared to therapies with other autologous cells, is the relative ease with which it can be concentrated from peripheral blood and the fact that it does not need time for cell expansion. The effect of PRP on wound healing has been studied in both animal and human models. Yang et al. used plasma combined with heparin-conjugated fibrin in full-thickness skin wounds in mice [27], while Law et al. tested the effect of PRP enriched with human donor keratinocytes and fibroblasts on circular wounds created on nude mice [28]. In this study wound progress was monitored for 14 days, and it was demonstrated that of the three groups, only the PRP plus donor cells group had achieved full and efficient reepithelialization and wound closure by day 14 compared to controls with only saline or PRP-containing media. In a recent case study, Suthar et al. [29] selected 24 patients aged 18–85 with various types of chronic or non-healing ulcers (full thickness and of at least 4-week duration). They administered a one-time autologous PRP treatment by drawing peripheral blood then isolating PRP by centrifugation. After debridement, 3–4 ml of PRP, extracted from the platelet layer, were injected subcutaneously and peripherally, while 2–3 ml were sprayed onto the wound simultaneously with activator solution to form an autologous platelet gel. Healing was assessed by the appearance of granulation tissue and reduction in wound size by week 4. Overall, wound healing was observed in all patients, with reduction in pain and serous discharge within 1 week of application of the treatment, and no adverse effects were observed throughout the 24-week follow-up. PRP may have potential in the development of new woundhealing methods but with a few caveats. After many randomized controlled trials, it is still not clear that application of PRP to wounds promotes better healing than standard treatments [30]. And despite a large number of available techniques and devices for the preparation of autologous PRP, a standard preparation method has yet to be established. Recent animal studies have demonstrated that the addition of macrophages or monocytes to murine wounds can significantly accelerate wound healing. Cells were isolated from peripheral

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blood and mixed in a pullulan-collagen composite dermal hydrogel [31]. The treatments gave positive results both in normal and diabetic mice. In another study rat wounds were treated with topical application of complement C3 resulting in significant increase in wound-healing rate [32]. 2.3 Stem Cells, Wound Healing, and Animal or Clinical Trials

While grafts have traditionally been prepared with differentiated cells like keratinocytes and fibroblasts, less differentiated cells such as hair follicle stem cells and mesenchymal stem cells are also being used to develop cell-based treatments for wounds. Some adult stem cells such as epidermal stem cells (and intestinal crypt stem cells) proliferate almost continuously throughout the organism’s life, while others such as muscle stem cells proliferate only in response to injury [33]. Mesenchymal stem cells can be used in wound healing because they give rise to tendon, cartilage, bone, and fat progenitors [34]. For instance, bone marrow-derived mesenchymal stem cells have been shown to increase wound-healing rates in both diabetic rats [35] and in humans [36]. Mesenchymal stem cells found in the dermis among undifferentiated neonate fibroblasts accelerated healing rate in mice [37], Fig. 2. The extraction and expansion of mesenchymal stem cells from abundant tissues such as adipose hold promise for the development of cellular therapies for refractory wounds and have been reported in animal models. This has been demonstrated in a diabetic rat

Fig. 2 Engraftment and differentiation of mesenchymal cells in vivo. A mesenchymal population (E) originating in murine neonate dermis was added on a circular wound and compared to control epidermal cells (Epi, DHF) or dermal, non-mesenchymal (D). Shown is week 3 post-grafting with silicon chambers. Arrows point at graft area in all four panels. A very dark healed wound in DHF contained numerous hair follicles and cysts as observed in the Fontana-Masson panel below. Animals grafted with the mesenchymal population (E) healed faster and displayed a dark ring around the edges of the wound site; pigmented nests were deep within the host dermis, as seen in Fontana-Masson panel below. Reproduced, with permission, from Crigler et al., FASEB J. 2007 July;21(9):2050–2063

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model by Kato et al. [38] and in a mouse model by Kim et al. [39]. Adipose-derived stem cells have also been used clinically, as they yield 500 times the number of cells in the same amount of tissue compared to bone [40]. One recent study [41] described treatment of ten diabetic patients with leg ulcers with tissue grafts derived from autologous adipocytes. The group used the Coleman technique [42] to harvest abdominal adipose. After centrifugation and fat isolation, micrografts of the fat were injected around and under the wound site. The primary goal of the study was to observe wound closure rate, and the authors reported significant differences in the rate of wound closure for the nine patients who remained in the study. The group concluded that autologous adipose tissue seems to accelerate wound healing, most likely due to the transfer of adipose-derived stem cells while reducing pain and promoting the growth of good quality skin. Similar efforts have been made and are underway to experiment with the use of ADSCs in diverse situations such as the treatment of bones, scars, or autoimmune diseases; for review, see [40]. On the other side of the debate, it has been pointed that despite all the advances in the stem cell field and all the resources devoted to it, there is limited accumulated evidence to date that cellular therapies can provide effective treatment. The Food and Drug Administration (FDA) has expressed concern that most stem cell discoveries have not been shown to be clinically useful, while a growing number of “stem cell clinics” use testimonials to advertise services instead of solid clinical data [43]. 2.4 Developing Tissue Constructs for Skin Wound Care

Grafts are protective skin substitutes whose function is to provide temporary cover for a wound site, prevent fluid loss, and lower the risk of contamination. They can be made of various materials, such as silicone (Mepilex™) [44]. But, as mentioned before, skin grafts have been one of the most common uses of autologous (or allogeneic) tissues in the field of wound healing to cover wound sites. Commercially available acellular skin substitutes, such as Alloderm™ and Integra™, are used for the same purpose. They use acellular collagen to cover wounded areas and provide a scaffold for cells to move into [45, 46]. Tissue-engineered skin substitutes are a recent alternative that involves the culturing of patient or donor cells for use in therapies (for a review, see [47]). Tissue engineering itself has seen comparatively little widespread clinical use for a variety of economic and feasibility factors including the fact that tissue constructs require angiogenesis for viability in vivo. This has led to the development of new prevascularization techniques, which aim to generate vascular networks in tissue constructs before implantation in vivo (for a review, see [48]). The authors of a recent animal study [49]

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developed prevascularized mucosal cell sheets for use in oral wounds. They cultured rat keratinocytes and fibroblasts from oral mucosa and endothelial progenitor cells from peripheral blood in vitro. In order to compare healing rates with and without prevascularization, oral mucosal cell sheets were prepared. Fibrin glue was prepared from each rat’s plasma to form a scaffold on which to layer the keratinocytes. Control sheets (K sheets) were created solely with keratinocytes, while the prevascularized sheets (PV sheets) contained endothelial cell progenitors and submucosal fibroblasts mixed into the fibrin. Deep oral wounding was performed on the rats, and wound closure was tracked over the next 21 days in a group without cell sheets, a K group and a PV group. The PV sheet group consistently had earlier wound closure and smaller wound sizes than both other groups. Their results support further research into the use of autologous endothelial cells for prevascularization when developing tissue constructs for wound care. 2.5 Tissue Bioprinting

This is a technique used to fabricate living tissues using bioink, a combination of cellular components and vehicle materials that is printed on selected 3D patterns. Bioprinters can be laser-based, inkjet-based, or extrusion-based; the most commonly used bioprinters use either thermal inkjet printing or direct ink writing (extrusion). Early inkjet tissue bioprinters [50] have undergone technical variations depending on the tissue that is mimicked and on the particular focus of the authors [21, 51]. Medical and scientific 3D printing files can be shared among researchers and clinicians with the NIH 3D print exchange (https://3dprint.nih.gov, last accessed March 12, 2018). Based on an earlier published study demonstrating the feasibility of using liposome-like droplets to create tissue [52], Graham et al. recently reported constructs consisting of layers bioprinted in a sequential manner and in very small increments. The authors are moving toward commercializing the technique (OxSyBio), with cell-containing bioink droplets which are added into a lipid-containing platform and coalesce in a manner that resembles membrane formation by lipid bilayers [53]. Naturally, tissue bioprinting is proposed as a way to obtain large amounts of tissue for grafting or transplantation [54] and will have potential for clinical use if and when the system is scaled up to cm-size constructs rather than the current micrometer sizes achieved. Moving toward this goal, researchers of Jorcano’s group described the formation of up to 100 cm2 of printed skin in record time [55] by bioprinting skin substitutes made of plasma-based bilayers of human fibroblasts and keratinocytes. Bioprinting provided a standardized and faster alternative to manual approaches which had been previously established by the group.

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Natural Products for Skin Wound Healing In the absence of pharmaceutical products and laboratory research, our ancestors used natural compounds to assist the treatment of disease. The wound-healing capacities of certain natural substances are undeniable. With the current prevalence of diabetes and cardiovascular diseases, chronic wound complications are of great concern, which makes research on natural wound healers of particular interest. Natural compounds comprise a wide variety of chemical substances, each with its specific properties and particular place in the wound-healing process. Here we offer a limited sample of natural healers that participate in either microbial protection (such as honey), anti-inflammatory regulation (such as olive oil), or angiogenesis (such as vitamin D).

3.1

Honey

We have benefitted from honey’s healing properties since antiquity. Apart from its role in treating infection, pain, or gastrointestinal tract illnesses, honey has been one of the most important natural compounds in wound healing. Honey displays antimicrobial properties and the ability to heal wounds without scarring [56], providing nutrients and chemicals that speed up wound healing and reducing pus, odors, and pain [57]. These qualities justify a growing interest on its therapeutic applications in the recovery of surgical wounds or after grafting, burns, abscesses, and chronic leg ulcers. An evident example of this tendency is the commercial use of honey in postoperative bandages, (Medihoney™) [58]. Most publications on honey are in vitro studies with a focus on honey’s antimicrobial properties. One of the reasons for renewed research on natural antibacterials such as honey is the emergence of Methicillin-resistant Staphylococcus aureus (MRSA). In fact, honey is classified according to the strength of its antimicrobial function with the Unique Manuka Factor, a unit of measurement of honey’s activity against S. aureus, compared to other artificial antibacterial compounds. Since honey is “manufactured” from locally available flowers, it is understandable that honeys throughout the world exhibit significantly different characteristics [59, 60]. As suggested by several in vitro studies, all honeys worldwide possess antiseptic properties to different extents, with some types even reaching the effectiveness of sulfonamide antibiotics (Iranian honey) [61]. Manuka honey has synergistic effects with various antibiotics against MRSA [62]; acts against other common microorganisms, such as P. aeruginosa and E. coli; and inhibits the growth of many other relevant bacteria, like S. pyogenes (a potential cause of impetigo and cellulitis in humans). Manuka honey is also active against dermatophytes and has been tested as a therapeutic agent to heal

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skin rashes and treat dermatophytosis and infected wounds (for a review see [63]). For the above reasons, it has been argued that the MFU is not a reliable method to test antimicrobial function because it only considers the compound’s ability to fight S. aureus infections, not other microorganisms. As with other natural compounds, some of the substances responsible for biological actions of honey have been isolated. These include methylglyoxal, the main antibacterial in Manuka honey [64]. Honey is not only a powerful antiseptic but also an agent for immunomodulation and reepithelialization in wounds. This immunomodulatory activity is complex as honey’s many compounds may have several potential immunoregulatory targets. Some in vitro research has shown that when exposed to honey or its components, human macrophages and monocytes activate or inhibit the release of cytokines (such as tumor necrosis factor-α, interleukin-1β, interleukin-6), depending on the wound stage and condition. A similar phenomenon occurs in neutrophils, fibroblasts, and endothelium, which are all activated by honey or derivatives (reviewed in [65]). Because these cell types are involved in chemotaxis, cell migration, and regeneration of the collagen matrix, it can be observed that honey also plays a significant role in the reepithelialization of scars. An in vitro study concluded that all different types of honeys tested increased reepithelialization rates and chemoattractant activity when applied to HaCaT keratinocytes to a similar extent. On the other hand, the ability to induce epithelial to mesenchymal transition (EMT) varied among the honeys studied [66]. A limited number of clinical trials can be found in the literature, mostly studying the effects of natural compounds in healing of diabetic chronic ulcers. For example, research conducted in 2012 investigated the effects of dressings impregnated with honey in the recovery of neuropathic foot ulcers in type 2 diabetic patients, compared to a group treated with conventional dressings. Although the number of wounds did not significantly differ between the two groups, the healing rate increased among the patients treated with honey (healing time in treated group, 31  4 days; healing time in control group, 43  3 days (P < 0·05), as well as in the rate of ulcer disinfection [67]. 3.2

Propolis

Propolis is another bee product with wound-healing capacity. It is also known as the “bee glue” because it seals open spaces in the hive, and it is a mixture of plant resins and bees’ waxy secretions. Apart from its mechanical purpose, the bee-produced part of propolis grants it a chemical function against microbial threats to the hive, making it a powerful antiseptic. This factor contributes to the generation of a favorable microenvironment for reepithelialization. Propolis displayed antioxidant activity in fibroblast-like mouse cells (L929) injured with hydrogen peroxide by activation of antioxidant genes (such as HO-1, GCLM, and GCLC), and it

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decreased degradation of collagen mRNA [68]. Similarly to honey, propolis can activate keratinocyte reepithelialization in injured rats [69]. Moreover, the authors of this study were able to demonstrate good delivery of the propolis ointment to the wound bed, thus suggesting propolis and its components as good candidates for further testing in human wounds. Most of the research involving propolis has been in vitro, but its potential for wound healing has been demonstrated in a pilot study of diabetic foot ulcers [70]. 3.3

Olive Oil

Natural compounds of vegetal origin have a long history in folk wound healing. Olive oil is an example of such compounds. Apart from its nutritional benefits and its important role in the Mediterranean diet, olive oil has been historically used for the treatment of skin diseases, such as psoriasis, small burns, wounds, and damage after sun exposure, or to ameliorate stretch marks after pregnancy. This is because olive oil has significant anti-inflammatory properties, supposedly due to its fatty acid content and high levels of polyphenols, chemical components common to honey, olive oil, and other remedies used in traditional wound healing. Olive oil is often classified according to its acid content (particularly free oleic acid, its main fatty acid component). As olive oil acts as a natural anti-inflammatory and seems to decrease cholesterol levels, current research not only focuses on its use for treating leg ulcers but also on its effects on atherosclerosis. Beneficial effects of olive oil on wound-healing and antibacterial activity were demonstrated in a recent in vitro study on human keratinocytes based on improved closure in scratch assays as well as decreased infectivity of a large panel of bacterial strains involved in skin infections [71]. Yet another study assessed the ability of olive oil and of some of its phenolic compounds to reduce lipid accumulation in human endothelial cells after exposure to oxidative stress. After treatment with phenolic compounds, damaged endothelial human cells activated anti-inflammatory pathways as compared to control untreated cells, as seen by the expression of ICAM-1, release of NO, and the activation of NF-kB. The same treatment improved lipid metabolism in rat hepatoma cells demonstrated by increased lipid desaturase suggesting improved hepatic lipid metabolism [72]. In contrast with research on honey, the majority of reports on olive oil are not in vitro but animal trials. In this context, it has been proven that olive oil has the ability to accelerate the chemical pathways that improve the inflammatory state of animal wounds. For example, reactive oxygen species (ROS) are known to act as secondary messengers for immune cells involved in the wound-healing process [73]. One such study showed that olive oil applied to pressure ulcers in mice improved their wound healing mainly through stimulation of nitric oxide and reactive oxygen species synthesis [74] (Fig. 3).

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Fig. 3 Olive oil helps heal pressure ulcers. Pressure ulcers in mice were ameliorated by the application of olive oil through induction of reactive oxygen species and nitric oxide synthesis. Improved wound closure and dermal regeneration were found in the study. Reproduced, with permission, from Donato Trancoso, et al., J Dermatol Sci. 2016 July;83(1):60–9. doi:https://doi.org/10.1016/j.jdermsci.2016.03.012. Epub 2016 Apr 1

Some authors have focused on identification of the specific properties that make olive oil, as compared to other fatty acid oils, particularly beneficial for wound healing. For example, experiments conducted both on damaged mice fibroblasts and in vivo suggest that olive oil improves the rate of healing compared to the effects of fish oil with improvement in wound contraction, reepithelialization, vascular endothelial growth factor expression, and numbers of macrophages and neutrophils, whereas skin wounds exposed to fish oils did not [75]. It seems that phenolic compounds have many beneficial activities in vitro, but there is still limited clinical evidence of their effectiveness [76]. Due to their complex chemistry, it is difficult to compare effectiveness of a mixture of phenolic compounds to that of individual components, and more studies are needed. 3.4

Calcitriol

Humans are able to produce their own natural wound healers, like calcitriol (or 1,25-dihydroxycholecalciferol), a metabolite of vitamin D. Although the human body generates the inactive form of vitamin D, sunlight is required for its activation in the skin and its further metabolism in the liver and kidney. Calcitriol has powerful therapeutic effects in the treatment of diseases related to calcium imbalances, such as osteoporosis, osteomalacia, or hypocalcemia

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[77]. Besides, calcitriol has been used as an antiseptic for the recovery of diabetic leg ulcers and for other wounds. However, recent research is now being conducted to understand the effects of calcitriol in the wound-healing process, particularly in angiogenesis. The active form of vitamin D is known to induce antimicrobial peptides in keratinocytes from diabetic foot ulcers, but it could also have angiogenic properties. Trujillo et al. tested in vitro keratinocytes involved in type 2 diabetes-related wounds, which showed abnormal expression of angiogenic factors like VEGF, HIF-1α, and angiogenin. But, when exposed to a medium with calcitriol, the levels of these molecules returned to normal, allowing proper angiogenesis and recovery of the injury. This was compared to a control group with a non-calcitriol medium, and the difference suggested statistical significance. Furthermore, it was also shown that the treated cells healed in a time comparable to cells treated with exogenous VEGF or EGF [78]. Thus, in vitro calcitriol was found to regulate the expression of growth factors, promoting angiogenesis.

4

Wound Dressings: Materials, Scaffolds, and Devices for Skin Wound Healing External interventions, involving surgical procedures, wound dressings, and other mixed approaches, are required for the management of acute or chronic wounds with the goal of preparing a clean tissue bed for reepithelialization. It has long been recognized that both air exposure and wound closure have important functions in epithelialization [79], and much of our current standard of care is based on research on these mechanisms. Wound management usually starts with debridement, the removal of dead tissue in the wound in order to increase the regeneration of the living tissue [80]. Outside of surgical debridement, various other methods have been classically used such as mechanical, chemical, or biological (e.g., using maggots). Wound dressings provide nonselective debridement when they are changed after the wound has transitioned from moist to dry, thus cleaning the wound by removal of necrotic tissue. It is recommended that wound assessment be made at each dressing change [81]. Occlusive and semiocclusive dressings, applied in the absence of infection, can produce autolysis, a more selective debridement using the body’s own enzymes and moisture to liquefy necrotic tissue and achieve epithelialization [79]. This autolysis curves excess inflammation that would produce hyperplasia (for a comprehensive review on occlusive dressings, see [82], and for reviews on dressings for diabetic foot ulcers, see [83, 84]). Enzymatic debridement is a proactive method that uses enzymes to more quickly and effectively remove necrotic tissue. This technique is mostly used in

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the management of burns. The enzymes used can be derived from microorganisms such as Clostridium histolyticum or from plants. Most common enzymes include collagenase, papain, and bromelain [85]. 4.1 Biomaterials Used in Clinical Care

There are various biomaterials used in clinical wound treatment. For instance, oxidized regenerated cellulose/collagen was initially shown to promote wound closure in a diabetic mouse model [86] and in acute wounds in rats [87]. This material has been adopted in wound care with overall good results as assessed by a recent panel sponsored by Acelity [88] and by other independent studies [89]. Bacterial celluloses have also been proposed as useful materials for wound dressings as they can absorb fluid better than other cellulose sources and display low toxicity [90]. Despite the fact that bacterial celluloses are expensive and difficult to obtain in large scale, they may be well suited for this more limited application.

4.2 Novel Biomaterials in Preclinical Research for Wound Healing

Various biomaterials are under development in tissue regeneration to serve as scaffolds onto which relevant cells are seeded or as vehicles for targeted drug release. These scaffolds represent the topology of the tissue of origin and provide topological cues that will help tissue regeneration. Several natural or synthetic polymers, generically termed hydrogels, are being developed and evaluated in preclinical studies. Films made of hydrocolloids or hydrogels are biodegradable and, as such, capable of integrating into the patient’s wound as it regenerates. Silk, a material long accepted for sutures, is finding its place in the formulation of dressings and scaffolds. Modifications of the native silk polymer have been described with added properties such as a change in electric charge or pH [91]. The raw material has also been modified so as to produce a nanoparticle for drug delivery (e.g., see [92–94]). Scheibel’s group has produced modified recombinant silk proteins with amino acid substitutions or changes in posttranslational modifications to improve their functionality in tissue bioengineering [95, 96]. Silk proteins’ monomers are also used in the formulation of hydrogels, for example, Shi et al. formulated a hydrogel for bone regeneration which incorporates a biomineralization step [97]. In the case of wound regeneration, it has been demonstrated that there is an increased adhesion and proliferation of dermal fibroblasts and of keratinocytes to modified silk fibroin [98]. The combination of silk fibroin with polyvinyl alcohol polymers has shown promising results when tested in a diabetic rabbit model and could be useful in dressings for diabetic patients; in the study the dressings healed wounds faster with a more organized extracellular matrix deposition [99]. Elastin, added exogenously in the form of recombinant human tropoelastin, encourages the deposition of elastin by fibroblasts contributing elastic properties to the healing wound as reported

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recently by the Weiss lab [100]. If further preclinical research confirms this in vitro data, it would be feasible to combine exogenous stimulation of elastin in the patient with approved dermal substitutes [101]. Gelatin-, alginate-, and acrylate-based materials are used in combination for bioprinting. Gelatin proteins are cross-linked by oxidation of the alginate, which results in a final material of the right consistency for bioprinting [91, 102]. Alginate-based materials are also considered in tissue regeneration for arthritis because of alginate’s known antioxidant and anti-inflammatory properties [103]. Other authors propose the use of cross-linked acrylatebased materials which provide improved biomechanical properties [104]. Similarly, a cell-laden biodegradable cross-linked polymer was proposed by the Yang group at Stanford [105]. Chitosan, a biopolymer derivative of chitin, is used in various tissue regeneration models because of its biomimetic properties and known effects in accelerating wound healing [106, 107]. Chitosan combined with a collagen precursor has also been recently tested in a rat model of diabetic wounds with promising results [108]. 4.3 Materials Combined with Drugs or Drug Delivery Combined (Tunable Systems)

Bioengineers seek optimal tunable release of therapeutics in various tissues in the body. Both tunable and tailorable systems must combine optimal materials with adequate pharmacokinetics for drug release (for differences and examples, see recent review from [109]). There are recent studies of materials with the ability to provide discontinuous drug release. Zhang et al. propose a hydrogel loaded with superoxide dismutase providing antioxidant properties to the wound bed [110]. Also, a Johns Hopkins group recently reformulated Valsartan, an angiotensin type I receptor blocker used to treat high blood pressure, to treat the dysregulation of the reninangiotensin system which affects chronic wounds in some diabetics. The results of the study both in a mouse and pig model suggest potential for application to human wound healing [111]. Other authors report fibroblasts and insulin-loaded modified chitosan polymers for wound dressings. The proposed hydrogels provide drug or cell release that is induced by pH or glucose changes in the wound [112].

4.4 Materials Combined with Antimicrobials

Bacterial biofilms represent a major wound complication because the biofilm structure provides antibiotic resistance and causes destruction of the tissues in the wound. The Sen lab at Ohio State has been working on wireless electroceutical dressings (use of electrical currents generated in situ because of a “conductive wound exudate”). Ions present in the bandages can produce weak electric fields to disrupt bacterial biofilm formation. This helps prevent infections, combat antibiotic resistance, and enable healing in

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infected burn wounds [113]. According to the Ohio State web page, this FDA-approved technology could soon go on to clinical trials. As in therapies for other chronic diseases, the management of infected refractory wounds can be helped by combining successful individual approaches such as traditional herbal remedies with antimicrobial approaches when antibiotic therapy alone is not indicated. Laboratories at the CSIR-Institute of Himalayan Bioresource Technology have been screening cellulose from various native plants for this purpose and recently published a combination of cellulose nanocrystals with silver nanoparticles. The composite materials exhibited good wound-healing properties and infection control in diabetic mice [114]. 4.5

5

Surgical Sensors

Finally, materials such as surgical implants need to be biodegradable and able to rely information about the implant [115] or local drug delivery via dermal patches [116]. Sensors are being developed for the monitoring of surgical implants. These include Bluetoothoperated biodegradable electronics. It is important to find materials able to both provide homing signals for regenerative cells and allow those cells to migrate to natural wound sites. There are some studies that add to these features, those of releasing helpful compounds in the site of the wound, for example [117].

Conclusion In this review, we attempt to give an overview of skin wound healing while highlighting a handful of related fields where basic in vitro research and animal studies demonstrate progress. We found limited examples of clinical trials in the aspects covered. Obviously, wound healing is not among the top critical challenges in medicine, but it has a major impact on the quality of life of patients and on the aging population, and we hope progress will continue with tangible results for patients. We thank all our colleagues involved in this worthy cause, and we apologize if we omitted their work due to space limitations.

Acknowledgments Our undergraduate research (course number SCIR297) is conducted with the support of Montgomery College. We thank our students Nhi Luu and Rachel Tannenbaum for contributing ideas to the project. We also thank Jung Wha Lee and Lauren Kimlin (Virador and Associates) for their support. We appreciate our lively discussions with Jacqueline Muller and her critical reading of the manuscript.

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96. Petzold J, Aigner TB, Touska F et al (2017) Surface features of recombinant spider silk protein eADF4(κ16)-made materials are well-suited for cardiac tissue engineering. Adv Funct Mater 27:1701427 97. Shi L, Wang F, Zhu W et al (2017) Selfhealing silk fibroin-based hydrogel for bone regeneration: dynamic metal-ligand selfassembly approach. Adv Funct Mater 27:1700591 98. Karahalilog˘lu Z, Ercan B, Denkbas¸ EB et al (2015) Nanofeatured silk fibroin membranes for dermal wound healing applications. J Biomed Mater Res A 103:135–144 99. Chouhan D, Janani G, Chakraborty B et al (2018) Functionalized PVA-silk blended nanofibrous mats promote diabetic wound healing via regulation of extracellular matrix and tissue remodelling. J Tissue Eng Regen Med 12(3):e1559–e1570 100. Wang Y, Mithieux SM, Kong Y et al (2015) Tropoelastin incorporation into a dermal regeneration template promotes wound angiogenesis. Adv Healthc Mater 4:577–584 101. Mithieux SM, Weiss AS (2017) Design of an elastin-layered dermal regeneration template. Acta Biomater 52:33–40 102. Rao KM, Rao KSVK, Ramanjaneyulu G et al (2014) Biodegradable sodium alginate-based semi-interpenetrating polymer network hydrogels for antibacterial application. J Biomed Mater Res A 102:3196–3206 103. Kerschenmeyer A, Arlov Ø, Malheiro V et al (2017) Anti-oxidant and immunemodulatory properties of sulfated alginate derivatives on human chondrocytes and macrophages. Biomater Sci 5:1756–1765 104. Moghadam MN, Pioletti DP (2016) Biodegradable HEMA-based hydrogels with enhanced mechanical properties. J Biomed Mater Res B Appl Biomater 104:1161–1169 105. Elomaa L, Pan C-C, Shanjani Y et al (2015) Three-dimensional fabrication of cell-laden biodegradable poly(ethylene glycol-co-depsipeptide) hydrogels by visible light stereolithography. J Mater Chem B Mater Biol Med 3:8348–8358 106. Gnavi S, Barwig C, Freier T et al (2013) The use of chitosan-based scaffolds to enhance regeneration in the nervous system. Int Rev Neurobiol 109:1–62 107. Mekhail M, Tabrizian M (2014) Injectable chitosan-based scaffolds in regenerative medicine and their clinical translatability. Adv Healthc Mater 3:1529–1545

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Methods in Molecular Biology (2019) 1879: 243–256 DOI 10.1007/7651_2018_116 © Springer Science+Business Media New York 2018 Published online: 27 March 2018

Isolation, Culture, and Motility Measurements of Epidermal Melanocytes from GFP-Expressing Reporter Mice Lina Dagnino and Melissa Crawford Abstract In this article, we provide a method to isolate primary epidermal melanocytes from reporter mice, which also allow targeted gene inactivation. The mice from which these cells are isolated are bred into a Rosa26mT/mG reporter background, which results in GFP expression in the targeted melanocytic cell population. These cells are isolated and cultured to >95% purity. The cells can be used for gene expression studies, clonogenic experiments, and biological assays, such as capacity for migration. Melanocytes are slow moving cells, and we also provide a method to measure motility using individual cell tracking and data analysis. Keywords Cell migration, Cre recombinase, GFP, Melanocyte

1

Introduction Melanocytes are highly specialized pigment-producing cells mainly found in the epidermis and epidermal appendages, such as the hair follicles [1]. Additional populations of melanocytes exist in the inner ear, the nervous system, and the heart [1], emphasizing the importance of these cells in the function and homeostasis of multiple tissues. In mammals and other vertebrates, cutaneous melanocytes arise as a result of a complex process that involves migration and cell fate decisions of early embryonic precursors [2–4]. For example, in mice, between 9.5 and 10.5 days of gestation, a subset of embryonic neural crest cells become melanoblasts, which proliferate and migrate dorsolaterally, and colonize the epidermis around 15.5 days of gestation. Some melanoblasts also colonize the hair follicles, where they establish a population of amelanotic stem cells, which can give rise to pigmented, mature melanocytes that endow hair with its pigment during postnatal life [3]. Follicular melanocyte stem cells can migrate to the interfollicular epidermis in response to wounding and ultraviolet radiation, where they

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differentiate into mature, pigment-producing melanocytes [5]. Importantly, resident amelanotic cells, which serve as progenitors of fully differentiated melanocytes, have also been described in the postnatal interfollicular epidermis [6]. Melanocytes are dendritic cells and synthesize melanin in lysosome-like organelles termed melanosomes [1]. Melanosomes likely originate from the endoplasmic reticulum and mature through four stages. Maturation of melanosomes is associated with trafficking along microtubules to the tip of the dendrites. In response to a variety of stimuli, melanocytes exocytose melanosomes, which are then internalized by adjacent keratinocytes in the epidermis [7]. Oncogenic transformation of melanocytes gives rise to melanoma, a very aggressive and metastatic human tumor type. Collectively, these properties make melanocytes excellent models to study organelle transport, pigmentation disorders, cell–cell interactions, and carcinogenesis mechanisms. However, epidermal melanocytes only constitute about 5% of all epidermal cells, and studies on this cell type require efficient isolation and enrichment procedures. In this protocol, we describe the isolation of epidermal melanocytes from neonatal reporter mice in which a gene of interest can be targeted for inactivation through Cre recombinase. Targeted melanocytes isolated from these mice can be readily identified through Cre-mediated expression of green fluorescent protein (GFP). The use of growth medium supplemented with endothelin-3 in this protocol allows for efficient expansion of melanocyte cultures to 95% purity, which remain exponentially proliferative for about 2 months [8]. We also describe protocols for transient transfection and for single-cell analysis of melanocyte motility.

2 2.1

Materials Mice

The experiments we describe are conducted with mice generated by breeding the reporter strain Gt(ROSA)26Sortm4(ACTB-tdTomato,EGFP)Luo /J (hereafter termed ROSAmT/mG) [9] with B6.Cg-Tg (Tyr-cre/ERT2)13Bos/J (hereafter termed Tyr::CreERT2) [10], both available from The Jackson Laboratory. Our protocol uses melanocytes isolated from mice that are homozygous for the ROSAmT/mG allele and hemizygous for Tyr::CreERT2. These reporter mice can be further bred into other backgrounds in which a gene of interest is flanked by loxP sites and can therefore be used to generate GFP-tagged melanocytic cells in which a gene of interest has also been inactivated upon administration of tamoxifen [8].

Mouse Primary Melanocyte Isolation and Culture

2.2 Isolation, Culture, and Tamoxifen Treatment of Primary Mouse Melanocytes

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1. MGM-4 Basal Medium (Lonza) (see Note 1) 2. MGM-4 SingleQuot Kit with Supplements and Growth Factors (Lonza) 3. Lyophilized Endothelin 3 (ET3) (Lonza) 4. Melanocyte growth medium: MGM-4 Basal Medium (500 ml) supplemented with one SingleQuot Kit and 130 μg ET3 freshly dissolved in MGM-4 Basal Medium 5. Sterile Ca2+- and Mg2+-free phosphate-buffered saline (PBS) 6. Dispase II (Roche Applied Science). 7. PBS 8. 0.025% Trypsin/0.01% EDTA (Life Technologies, Thermo Fisher Scientific) 9. Trypsin Neutralizing Solution (Life Technologies, Thermo Fisher Scientific) 10. 0.4% Trypan blue stain (ICN BioMed), dissolved in 0.85% NaCl 11. 95% Ethanol 12. 70% Ethanol 13. 4-Hydroxy-tamoxifen (4OHT, Abcam) 14. Sterilizing syringe filters (0.2-μm pore size, Filtropur, Sarstedt) 15. Bacterial grade 100-mm Petri dishes (Sarstedt) 16. Scissors, two forceps, and scalpel (Fine Science Tools) 17. 50-ml Conical tubes (Sarstedt) 18. 100-μm Pore size nylon strainer (Corning) 19. Neubauer hemocytometer (Hausser Scientific) 20. 60-ml Sterile syringes with BD Luer-Lok Tip (BD Biosciences) 21. BioLite T175 flasks, cell culture treated (Thermo Fisher Scientific) 22. Tissue culture plates

2.3 Dispase II Working Solution

Use only freshly prepared solution. Dissolve 70 U lyophilized Dispase II in 14 ml PBS, to produce a solution containing 5 U/ml. Sterilize by filtration through 0.2-μm pore size syringe filter (see Note 2).

2.4 4Hydroxytamoxifen Stock Solution

Dissolve 10 mg 4OHT in 0.8 ml of 95% ethanol, to obtain a 32 mM stock solution. Store in 20-μl single-use aliquots at 20  C for up to 1 year.

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2.5 Transfection Reagents

1. Lipofectamine 3000 and P3000 Reagent (Thermo Fisher) 2. mCardinal-C1 (Addgene Plasmid # 54799), at a concentration of 0.5–5 μg/μl (see Note 3) 3. Melanocyte growth medium 4. Sterile Ca2+- and Mg2+-free PBS 5. Sterile 1.5-ml microfuge tubes 6. 24-Well tissue culture plates

2.6 Laminin-332 Matrix

1. 804G Rat bladder epithelial cells (obtain from Dr. J.C.R. Jones, Washington University; see Note 4) 2. T75 culture flasks 3. Dulbecco’s Modified Eagle’s Medium (DMEM) supplemented with 8% fetal bovine serum (FBS) 4. Serum-free DMEM 5. Sterilizing filters (0.4-μm pore size, PALL Life Sciences # 4654)

2.7 Single-Cell Motility Assays

1. 35-mm μ-Dishes (ibidi) 2. Inverted microscope equipped with 10 and 20 phase contrast objective, camera for time-lapse video recording, and culture chamber to maintain cells at 37  C in a 5% CO2 humidified atmosphere 3. Volocity software for cellular imaging and analysis (PerkinElmer), or equivalent 4. ImageJ image processing software (version 1.49v, Fiji, National Institutes of Health) 5. Manual Tracking plug-in for ImageJ 6. “Chemotaxis and Migration Tool” plugin for ImageJ software (version 2.0; ibidi)

3

Methods

3.1 Isolation of Melanocytes from Newborn Mouse Skin

1. Euthanize 3-day-old mice by CO2 inhalation, and following the appropriate institutionally approved Animal Use procedures (see Note 5). 2. Wash the mice by immersion in 70% ethanol for 10 min. 3. In a sterile laminar flow hood, wash the animals by immersion in sterile PBS. 4. Place a mouse on a sterile 100-mm Petri dish. 5. Using scissors, amputate the head, limbs, and tail.

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6. Make a superficial longitudinal dorsal incision with the scalpel throughout the entire skin, avoiding deep incisions that would result in exposure of the organs. 7. Using two sets of forceps, gently separate the skin from the carcass, starting at the incision point. Remove any subcutaneous fat. 8. Rinse the skin with sterile PBS. 9. Place the skin dermis side down in a clean Petri dish, ensuring that the tissue is fully spread with no folds. 10. For each skin, add 2 ml Dispase II working solution, allowing the fully spread tissues to float. 11. Incubate at 37  C for 1.5 h. 12. Transfer skin to a clean Petri dish, dermis side down, and gently pull off the epidermis with the tip of a sterile forceps, while holding the dermis down with another set of sterile forceps (see Note 6). 13. Place the epidermis in a clean Petri dish. Pool the epidermal tissues from all skins harvested this way. 14. With sterile scissors or two scalpels, mince the epidermal tissues isolated to ~2 mm  2 mm fragments and transfer to a 50-ml conical tube containing 0.025% trypsin/0.01% EDTA (0.5 ml/full epidermis). Alternatively, the epidermal tissues can be placed directly in a 50-ml conical tube containing 4–5 ml 0.025% trypsin/0.01% EDTA and minced with sterile, long scissors, after which the volume of trypsin is adjusted to 0.5 ml/epidermis. 15. Incubate the epidermal tissues and the trypsin at 37  C for 10 min with gentle rocking (see Note 7). 16. Add two volumes of Trypsin Neutralizing Solution. To completely release the melanocytes from epidermal fragments as single cells, use a sterile pipet to pass the cell mixture several times (see Note 8). 17. Filter the cell suspension through a sterile 100-μm pore size cell strainer to remove cornified envelope fragments and tissue debris. 18. Centrifuge at 200  g for 10 min. 19. Aspirate the supernatant carefully to avoid disturbing the cell pellet. 20. Resuspend the cells in melanocyte growth medium, using 2.5 ml for each epidermis obtained. 21. Mix a 10-μl aliquot of the melanocyte suspension with 10 μl of Trypan blue stain and determine the number of viable, Trypan

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blue-excluding hemocytometer.

cells/ml

of

suspension,

using

a

22. Plate the cells on culture dishes at a density of 1  105 cells/cm2. This will yield a culture 40–60% confluent the following day. 23. Place the culture in a humidified 37  C, 5% CO2 incubator. 24. One day after plating, remove the culture medium and nonadherent cells, and replace with fresh growth medium. Replace the culture medium every 72 h thereafter. 25. Ensure maintenance of cell confluence between 30 and 90% at all times. 3.2 Subculture and Propagation of Melanocytes

1. Subculture melanocytes 7 days after initial plating, and every 7 days thereafter (see Note 9). 2. Remove the growth medium by aspiration, rinse the cultures with Ca2+-free PBS, and add 3 ml of 0.025% trypsin/ 0.01% EDTA. 3. Incubate at 37  C for about 5 min and examine the culture under the microscope. 4. Once melanocytes have detached, but prior to detachment of keratinocytes (which will appear round, but still firmly attached to the culture dish), add two volumes of Trypsin Neutralizing Solution (see Note 10). 5. Transfer the cell suspension into a sterile 50-ml conical tube. 6. Centrifuge at 22  C, 200  g for 5 min. 7. Carefully remove the supernatant by aspiration without disturbing the cell pellet. 8. Resuspend the cells in 10 ml of melanocyte growth medium. 9. Mix a 10-μl aliquot of the melanocyte suspension with 10 μl of Trypan blue stain and determine the number of viable, Trypan blue-excluding cells/ml of suspension, using a hemocytometer. 10. Plate at a density of 2.5  104 cells/cm2 (see Note 11).

3.3 Induction of Cre-mediated Gene Inactivation

1. Thaw on ice an aliquot of 4OHT stock solution (32 mM). 2. Just prior to use for cell culture, dilute 12.5 μl of 32 mM 4OHT stock solution with 987.5 μl 95% ethanol, to obtain a working solution of 400 μM 4OHT. 3. Prepare the needed volume of a master mix used to treat the melanocytes, containing 2.5 μl of 400 μM 4OHT mixed with 997.5 μl of melanocyte growth medium. This solution contains a final concentration of 1 μM 4OHT and 0.25% ethanol (see Note 12).

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4. Remove the growth medium in the melanocyte cultures by aspiration. 5. Add the freshly prepared 4OHT-containing growth medium to the cells. 6. Culture at 37  C in a humidified 5% atmosphere incubator for 48 h (see Note 13). 7. Remove the 4OHT-containing growth medium by aspiration. 8. Rinse the cultures once with Ca2+-free PBS. 9. Add fresh melanocyte growth medium and continue culturing as appropriate for the experiment (see Note 14). 3.4 Transient Transfection

1. One day prior to transfection, seed P2 or P3 melanocytes in 24-well culture plates, at a density of 7.5  104 cells/2-cm2 well (see Note 15). 2. Just prior to transfection, prepare DNA-Lipofectamine 3000 mix: In a sterile microfuge tube, mix well 50 μl of melanocyte growth medium with 3 μl of Lipofectamine 3000. 3. In a separate sterile microfuge tube, dilute 1-μg plasmid DNA with 50 μl of melanocyte growth medium and then add 2 μl of P3000 Reagent. Mix well (see Note 16). 4. Add the diluted DNA from step 3 to the Lipofectamine 3000 mix prepared in step 2. Mix well and let stand for 15 min at 22  C, to allow formation of DNA–lipid complex. 5. Add the DNA-Lipofectamine mix dropwise to the cells, swirling the culture plate to ensure thorough mixing. 6. Culture the cells in the presence of DNA-Lipofectamine mix for 18 h (see Note 17). 7. Remove the DNA-containing medium. 8. Rinse the cells once with Ca2-free PBS. 9. Add fresh melanocyte growth medium and culture the cells for 24–48 h, and process as appropriate.

3.5 Preparation of Laminin-332 Matrix and Coating of Culture Surfaces

1. Seed 1  106 G804 rat bladder epithelial cells in a T75 culture flask. 2. Culture the cells in normal growth medium (DMEM with 8% FBS) for 48 h. At this time, a completely confluent monolayer should be present. 3. Remove the growth medium, rinse with Ca2-free PBS thrice, and add 10–15 ml of serum-free DMEM. Culture the cells for 48 h. 4. Transfer the G804 conditioned medium (which now contains laminin-332 matrix secreted by the G804 cells) to a conical tube. Centrifuge at 1000  g for 5 min to remove cells and debris.

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5. Filter sterilize the conditioned medium through a 0.2-μm pore size filter. Use immediately to coat melanocyte growth surfaces or store at 4 C (see Note 18). 6. To coat plastic or glass surfaces with laminin-332 matrix, incubate with sufficient G804 conditioned medium to completely cover the surface for 1–2 h at 37 C. Completely remove the conditioned medium just prior to plating melanocytes on the treated surface. No rinsing prior to seeding the cells is necessary. 3.6

Motility Assays

1. Coat a 35-mm μ-Dish with laminin-332 matrix. 2. Seed 2.5  104 melanocytes (P2 or P3) onto the precoated μ-dish. Allow the cells to fully attach and spread by culture at 37  C in a humidified 5% CO2 incubator for 24 h. 3. Monitor cell motility using an automated microscopy system, with a 10 or 20 objective, and define the number of examined cells per well, along with their XY positions. Take time-lapse images for 16 h at 10-min intervals (see Fig. 1a) (see Note 19). 4. Analyze the data using the procedure described in Subheadings 3.7 and 3.8.

3.7

Cell Tracking

1. Open ImageJ and select File > Import > Image Sequence. 2. Select Plugins > Manual Tracking Plugin and the main Tracking window will open. 3. Calibrate the program by inputting the time interval and the x/y calibration (see Note 20). 4. To begin tracking a cell, select Add Track and click on the center of the nucleus. A separate window will open with the tracking results. 5. At the end of the time-lapse image sequence, select End Track. 6. To begin tracking another cell, select Add Track and the results will appear in the same window, together with the tracking results from the first cell. 7. To identify those cells that have been tracked, open the main Tracking window and select Dots. A new window will appear with colored dots showing the location of the cells that have been tracked. 8. To save the results of the analysis, click on the tracking results window and select File > Save as.

3.8 Analysis of Motility

1. Open the Chemotaxis and Migration Tool for ImageJ by selecting Plugins > Chemotaxis Tool (see Note 21).

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Fig. 1 Analysis of melanocyte motility. (a) Still-frame micrograph of melanocytes cultured in the absence or presence of laminin-332 matrix. Time-lapse videomicroscopy images were acquired for 16 h. The cells tracked are indicated with the colored circles. Bar, 100 μm. (b) Cell trajectory plots of random motility of melanocytes tracked for 16 h, shown in panel (a). The origin in the plot represents the starting position of each cell at time zero

2. In the bottom right-hand corner, select Import Data and import the tracking results file saved in step 8, under Subheading 3.7. Once the data are imported, they will appear in the “Imported datasets” section (see Note 22). 3. Select the drop-down bar next to “Number of Slices” and select use slice range from..to.. and enter the corresponding values (see Note 23). 4. Select Add Dataset. 5. Select Settings to calibrate software as in step 3 (Subheading 3.7). Use drop-down bars to adjust units. 6. In the top-left corner, check the box “Selected Dataset 1” and use the drop-down bar to select the dataset that will be analyzed (see Note 24). 7. Select Apply Settings (see Note 25).

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8. Selecting Show Info immediately next to Apply Settings will open a separate window that shows the maximum, minimum, and mean values for all parameters. 9. To obtain the parameters for each individual cell tracked, select Statistic Feature. In the bottom half of the screen, select the parameter of interest to open a window with the values for each cell. Click on window and select File > Save as to save as an excel spreadsheet. 10. To generate trajectory plots, select Plot Feature. When analyzing random motility, select No Marking from the dropdown bar. All tracks will appear as black lines (see Fig. 1b) (see Note 26). 11. Select Set Axis Scaling to adjust scale of graph (see Note 27). 12. Once all settings are configured, select Plot Graph. A separate window will open. 13. Select File > Save As > to save the plot.

4

Notes 1. There are several manufacturers of growth medium developed for culturing human melanocytes. In our experience, the use of MGM-4 medium (Lonza) supplemented with MGM-4 SingleQuot Kit Supplements and Growth Factors (Lonza) and ET3 support well the growth of neonatal mouse melanocytes. However, the manufacturer does not seem to consistently maintain a steady supply of MGM-4 medium, and shortages can last several months. We have also used PromoCell Basal Melanocyte Medium supplemented with SingleQuot Kit Supplements and Growth Factors (Lonza) and ET3, finding that it also supports the growth of primary mouse melanocytes. 2. For each skin, use 2 ml Dispase II at a concentration of 5 U/ ml. Depending on their size, up to 7 skins can be digested in a single Petri dish containing 14 ml Dispase II solution. Dispase II solutions of a lower concentration require considerably longer times to achieve optimal tissue digestion. 3. Melanocytes with a ROSAmT/mG background may exhibit fluorescence both in the 488 nm and the red fluorescence range. To visualize exogenously expressed proteins in these live cells, tagging with a protein that exhibits fluorescence in the far-red region is necessary. mCardinal exhibits fairly intense and stable fluorescence in far-red regions. 4. G804 rat bladder epithelial cells secrete laminin-332 matrix into their culture medium [11]. Conditioned medium from these cells can be used to efficiently coat cell culture surfaces

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that promote melanocyte migration. Commercially available laminin-332 produced by 184A1 mammary epithelial cells is available from Kerafast (Boston, MA). 5. This protocol has been optimized for isolation of melanocytes from 3-day-old animals. At this stage, there are still substantial numbers of melanocytes in the interfollicular epidermis. The epidermis can be easily harvested and separated from the dermis and is sufficiently large to yield adequate numbers of melanocytes for expansion and subsequent experiments. The interfollicular melanocyte population rapidly declines in mice older than 4 days of age [12]. 6. The epidermis should readily separate as an intact sheet after 1.5 h of digestion in the Dispase II working solution. If this does not occur, continue digestion at 37  C in the Dispase working solution for 15–30 additional minutes. Forceful separation of epidermis from dermis in partially digested skin will reduce melanocyte yield and increase contamination of cultures with dermal fibroblasts. 7. Trypsin digestion of epidermal tissues for intervals longer than 10 min will result in substantial loss of melanocyte viability. 8. Pipetting the cell mixture up and down ten times leads to optimal cell yields, without compromising melanocyte viability. 9. Initial epidermal isolates contain a mixture of melanocytes and keratinocytes. Because melanocytes constitute only 3–5% of epidermal cells, the majority of the cells at this stage will be keratinocytes. The growth medium used favors proliferation of melanocytes, but not of keratinocytes. Seven days after initial isolation, these cultures typically contain 30% melanocytes and 70% keratinocytes. 10. During the first subculture, enrichment of melanocytes is facilitated by differential trypsinization. This method is based on the stronger attachment of keratinocytes to the cell culture dish, relative to melanocytes. As a result, it is possible to achieve full detachment of melanocytes before that of keratinocytes. The addition of Trypsin Neutralizing Solution at this time prevents keratinocyte detachment from the culture dish, readily allowing the formation of a highly enriched melanocyte cell suspension. 11. We term cultures obtained after the first subculture as Passage (P) 1 cells. P1 cultures typically consist of approximately 70% melanocytes, whereas P2 cultures are >95% melanocytes [8]. 12. A concentration of 1 μM 4OHT in the culture medium is sufficient for effective nuclear target DNA excision by the Cre-ERT2 protein. Excised DNA can be amplified with appropriate primers by polymerase chain reaction, and amplicons

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corresponding to the targeted DNA can typically be detected as early as 16 h following 4OHT addition to the cells. 13. In our experience, treatment of melanocytes with 4OHT concentrations higher than 1 μM or for longer than 48 h does not increase targeting efficiency but results in substantial loss of cell viability. Depending on the background strain, targeting efficiency of cultured melanocytes in response to 4OHT may vary, as we have observed a range of 40–60% GFP-positive melanocytes following this treatment. 14. GFP fluorescence is readily detected in ~50% of melanocytes isolated from mice with a ROSAmT/mG; Tyr::CreERT2 background 2–5 days after 4OHT treatment. However, cells can exhibit dual GFP and mTomato fluorescence as late as 10 days following 4OHT treatment, likely associated with the turnover characteristics of mTomato mRNA and protein. mTomato fluorescence has been reported to persist in hepatocytes as late as 9 days following tamoxifen treatment in vivo [9]. 15. Seeding 7.5  104 cells/2 cm2 will yield wells with 80–90% confluence 24 h later. At this density, transfection efficiency is approximately 20% and decreases substantially if the transfected cultures are less than 80% confluence. 16. Using 3-μl Lipofectamine 3000/μg DNA results in better transfection efficiency, relative to lower Lipofectamine:DNA ratios. Increasing the amount of DNA used to 2 μg/2 cm2 or higher results in loss of cell viability. 17. Incubation of melanocytes in the presence of the DNA-Lipofectamine mix for periods longer than 16 h does not enhance transfection efficiency but can reduce cell viability. We routinely observe about 20% transfection efficiency using this protocol. 18. Coating of culture surfaces with freshly prepared conditioned medium containing laminin-332 matrix or medium stored at 4 C for up to a week is equally effective [13]. 19. The length of time intervals depends on the number of cells examined, and the culture conditions. In the absence of stimuli that stimulate migration, melanocytes are slow migrating cells, and imaging for several hours is necessary [8]. 20. The x/y coordinates are represented in pixels, therefore the size of one pixel in microns needs to be defined. Use the Line Tool in ImageJ to draw a line the same size as the scale bar. Select Analyze > Measure to obtain the length of scale bar in pixels. Divide the length of the scale bar in pixels by the number of microns to obtain the length of one pixel in microns. Input this value into the x/y calibration box.

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21. The Chemotaxis and Migration Tool is a plugin for ImageJ that is used to generate cell trajectory plots as well as to determine parameters such as distance travelled and the speed of each cell. This plugin can be downloaded from https://ibidi. com/manual-image-analysis/171-chemotaxis-and-migrationtool.html. 22. Each tracking file contains information for all cells tracked, thus cell trajectory plots will show tracks of every cell on one plot. This sometimes makes it difficult to determine the path of each individual cell. If it is necessary to analyze a single-cell trajectory, select Import Data > Show Original Data, delete all other cell tracking data, and save the raw data for the selected cell under a different file name (e.g., “Results for Cell 1”). This is the only way the Chemotaxis and Migration Tool plugin will recognize the data. If the tracking file is adjusted in excel, this plugin will not open the file. 23. Acquiring time-lapse images for 16 h at 10-min intervals yields 96 frames or slices. To analyze all 96 frames, select Use slice range from ..to.. and add the values 1 and 96 in the corresponding boxes. 24. Multiple datasets can be imported and subsequently analyzed. Select which specific dataset is to be analyzed by using the drop-down bar. 25. It is essential to select Apply Settings each time a dataset is selected for analysis. If this is not done, the calculated values will be inaccurate. 26. Cell tracks can be colored based on their movement when studying chemotaxis. To color cell tracks based on direction (e.g., left vs. right or up vs. down), use the drop-down bar (see Fig. 1a). 27. The program will automatically determine the scale of the axis, but when comparing melanocyte motility (for example, on different ECM substrates) it is best to adjust the scale to ensure consistency. This will enable accurate representation of differences in motility.

Acknowledgments This work was supported by grants to LD from the Canadian Institutes of Health Research, the National Science and Engineering Research Council, and the Cancer Research Society. MC is the recipient of an Ontario Graduate Scholarship from the Ontario Ministry of Advanced Education and Skills Development.

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References 1. Cichorek M, Wachulska M, Stasiewicz A, Tyminska A (2013) Skin melanocytes: biology and development. Postepy Dermatol Alergol 30(1):30–41 2. Li A, Machesky LM (2012) Melanoblasts on the move: Rac1 sets the pace. Small GTPases 3 (2):115–119 3. Mort RL, Jackson IJ, Patton EE (2015) The melanocyte lineage in development and disease. Development 142(4):620–632 4. Luciani F, Champeval D, Herbette A, Denat L, Aylaj B, Martinozzi S, Ballotti R, Kemler R, Goding CR, De Vuyst F, Larue L, Delmas V (2011) Biological and mathematical modeling of melanocyte development. Development 138 (18):3943–3954 5. Chou WC, Takeo M, Rabbani P, Hu H, Lee W, Chung YR, Carucci J, Overbeek P, Ito M (2013) Direct migration of follicular melanocyte stem cells to the epidermis after wounding or UVB irradiation is dependent on Mc1r signaling. Nat Med 19(7):924–929 6. Glover JD, Knolle S, Wells KL, Liu D, Jackson IJ, Mort RL, Headon DJ (2015) Maintenance of distinct melanocyte populations in the interfollicular epidermis. Pigment Cell Melanoma Res 28(4):476–480 7. Haass NK, Smalley KS, Li L, Herlyn M (2005) Adhesion, migration and communication in

melanocytes and melanoma. Pigment Cell Res 18(3):150–159 8. Crawford M, Leclerc V, Dagnino L (2017) A reporter mouse model for in vivo tracing and in vitro molecular studies of melanocytic lineage cells and their diseases. Biol Open 6 (8):1219–1228 9. Muzumdar MD, Tasic B, Miyamichi K, Li L, Luo L (2007) A global double-fluorescent Cre reporter mouse. Genesis 45(9):593–605 10. Bosenberg M, Muthusamy V, Curley DP, Wang Z, Hobbs C, Nelson B, Nogueira C, Horner JW 2nd, Depinho R, Chin L (2006) Characterization of melanocyte-specific inducible Cre recombinase transgenic mice. Genesis 44(5):262–267 11. Baker SE, DiPasquale AP, Stock EL, Quaranta V, Fitchmun M, Jones JCR (1996) Morphogenetic effects of soluble laminin-5 on cultured epithelial cells and tissue explants. Exp Cell Res 228:262–270 12. Hirobe T (1984) Histochemical survey of the distribution of the epidermal melanoblasts and melanocytes in the mouse during fetal and postnatal periods. Anat Rec 208(4):589–594 13. Dagnino L, Ho E, Chang WY (2010) Expression and analysis of exogenous proteins in epidermal cells. Methods Mol Biol 585:93–105

Methods in Molecular Biology (2019) 1879: 257–266 DOI 10.1007/7651_2018_144 © Springer Science+Business Media New York 2018 Published online: 05 June 2018

Melanoblasts as Multipotent Cells in Murine Skin Tsutomu Motohashi and Takahiro Kunisada Abstract Melanoblasts (MBs) are melanocyte precursors that are derived from neural crest cells (NCCs). We recently demonstrated the multipotency of MBs; they differentiate not only into pigmented melanocytes but also other NCC derivatives. We herein describe methods for the isolation of MBs from mouse skin by flow cytometry. Methods to culture isolated MBs that retain their multipotency and isolation methods for MBs using gene-modified mouse are also described. Keywords CD45, Flow cytometer, KIT, Melanoblasts, Multipotency, SOX10

1

Introduction Melanoblasts (MBs) are committed and undifferentiated transitamplifying cells that only differentiate into pigmented melanocytes (Ms). In mice, MBs migrate ventrally through the developing mouse dermis from embryonic day 10.5 (E10.5), begin to invade the overlying epidermis on E11.5, and then migrate into the developing hair follicles from E15.5, in which they continue to proliferate and differentiate before starting to synthesize pigment [1, 2]. MBs differentiate from multipotent neural crest cells (NCCs) that emerge from the dorsal region of the fusing neural tube. NCCs lose their multipotency after their generation and become committed to their fate, changing into tripotent, bipotent, or monopotent precursors during their migration. For example, NCCs that dorsolaterally migrate in the embryo and finally colonize skin generate MBs: the monopotent precursor that only differentiate into mature melanocytes and are not multipotent [3, 4]. Multipotent precursor cells were recently identified in the skin of fetal and adult animals, and these cells have a similar differentiation capability to that of NCCs [5–7]. Previous studies have reported that cells in adult hair follicles are also capable of differentiating into derivatives of NCCs, and these cells are considered to be derived from NCCs based on the findings of cell lineage analysis [8–10]. Other studies demonstrated that the cells that differentiated from NCCs also have a multipotential cell fate [11–15]. These findings suggest that NCC-derived

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cells show unstable NCC phenotypes and may dedifferentiate and then redifferentiate into other NCC derivatives [15]. In this context, we reinvestigated the differentiation of MBs and showed that they can differentiate into not only pigmented melanocytes (Ms) but also into neurons (Ns) and glial cells (Gs), which are cells not previously considered to be generated from MBs [16]. This finding suggests that the MBs, which are considered to be restricted in their fate to Ms., actually have a multipotential cell fate even when they have already migrated toward the target sites in the skin [16]. In this chapter, we describe methods for the flow cytometric isolation of MBs from embryonic and neonatal skin and hair follicles as well as methods for culturing MBs that retain their multipotency. MBs isolated as SOX10-positive, KIT-positive, and CD45-negative cells from the skin and cultured on monolayers of ST2 stromal cells differentiate not only into pigmented Ms. but also into Ns and Gs.

2

Materials

2.1 Preparation of Cell Suspensions from Mouse Skin

1. Dispase II solution: Dilute 10
Dispase II (Sanko Junyaku Co., Ltd., Japan) with Ca- and Mg-free phosphate buffer saline (PBS) and sterilize by passage through a ϕ 0.20-μm membrane filter. 2. Staining medium (SM): PBS containing 3% fetal calf serum (FCS). 3. 0.25% Collagenase type I: Dilute collagenase type I (Sanko Junyaku Co., Ltd., Japan) with Ca- and Mg-free PBS and sterilize by passage through a ϕ 0.20-μm membrane filter. 4. Hanks’ balanced salt solution containing 0.005% DNaseI (Roche) and 20% FCS. 5. Cell-dissociation buffer (Gibco). 6. 0.05% Trypsin solution: 0.05% trypsin/0.5 mM EDTA (Gibco). 7. Binocular microscope (such as Carl Zeiss, Stereomicroscope DV4).

2.2 Immunostaining of Skin Cell Suspension for Flow Cytometric Analysis

1. Rat anti-mouse Fc gamma receptor (2.4-G2; BD Bioscience). 2. Phycoerythrin (PE)-conjugated (30-F11; BD Bioscience).

rat

anti-mouse

CD45

3. Allophycocyanin (APC)-conjugated rat anti-mouse Kit (2B8; BD Bioscience).

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4. PI solution: SM containing 3 μg/mL propidium iodide (Calbiochem). 5. Flow cytometer (such as Becton-Dickinson FACS Vantage, FACSAria). 2.3 Culture of Isolated MBs

1. 0.05% Trypsin solution: 0.05% trypsin/0.5 mM EDTA (Gibco). 2. Medium for maintenance of ST2 stromal cells: RPMI-1640 (Gibco) supplemented with 5% FCS, 50 μM 2-mercaptoethanol, 50 μg/mL streptomycin, and 50 U/mL penicillin. Store at 4  C and use within 2 months. 3. Dexamethasone (Dex; Sigma): Store original stock solution at 102 M in ethanol at 70  C. Prepare the working stock solution at 103 M in ethanol at 4  C and dilute from this stock solution for each use. Stable for at least 1 year (stock at 70  C) or 3 months (stock at 4  C). 4. Human recombinant fibroblast growth factor-2 (bFGF; R&D Systems): Store the stock solution at 200 nM in 0.1% bovine serum albumin (BSA)/PBS at 70  C. Store at 4  C after thawing. Stable for at least 6 months (stock at 70  C) or 1 month (stock at 4  C). 5. Cholera toxin (CT; Sigma): Store the stock solution at 50 nM in distilled water at 70  C. Store at 4  C after thawing. Stable for at least 1 year (stock at 70  C) or 1 month (stock at 4  C). 6. Human recombinant endothelin-3 (EDN3; Peptide Institute, Inc., Japan): Store the stock solution at 100 μg/mL in 0.1% acetic acid solution at 70  C. Store at 4  C after thawing. Stable for at least 1 year (stock at 70  C) or 1 month (stock at 4  C). 7. Medium for differentiation of melanoblasts: αMEM (Gibco) supplemented with 10% FCS, 107 M Dex, 20 pM bFGF, 10 pM CT, 100 ng/mL EDN3, 50 μg/mL streptomycin, and 50 U/mL penicillin. Store at 4  C and use within 2 months.

2.4 Immunohistochemical Analysis of Cultured MBs

1. 4% PFA (paraformaldehyde) in PBS: pH ¼ 7.0–7.5 (see Note 1). 2. 0.1% Triton-X100 (Gibco) in 0.5% BSA PBS. 3. Blocking solution: 3% goat serum or 5% BSA in PBS. 4. Primary antibodies: mouse anti-neuronal class III β-tubulin (TuJ-1; BABCO), rabbit anti-mouse glial fibrillary acidic protein (GFAP; Z0334, DakoCytomation). 5. Secondary antibodies: Texas Red-conjugated anti-mouse IgG (Molecular Probes), Alexa-Fluor 488-conjugated anti-rabbit IgG (Molecular Probes). 6. Fluorescence microscope (such as Olympus, IX-71).

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Methods

3.1 Maintenance of ST2 Stromal Cells and Preparation for Use

ST2 stromal cells are used as feeder cells for isolated MBs and are derived from mouse bone marrow stroma cells. ST2 stromal cells are maintained in medium for maintenance of ST2 stromal cells (see Note 2). 1. Obtain confluent ST2 cells in a dish or flask and trypsinize them with 0.05% trypsin solution at 37  C for 4 min. 2. Add medium, dissociate cells by pipetting, centrifuge them at 200
g for 4 min, and split cells 1:4 into 100-mm dishes. 3. Maintain cells by regularly passing them every 3 or 4 days. 4. In order to prepare feeder layers, seed 25% of the confluent cells from a 100-mm dish equally onto one plate type, i.e., 6-well or 96-well plates. Two days later, cells reach confluence and are ready for use for the differentiation of MBs. Neither irradiation nor a treatment with mitomycin C is needed.

3.2 Preparation of Cell Suspensions from Mouse Embryonic Skin, Neonatal Skin, and Hair Follicles

1. Place male mice and female mice in the same cage in the evening and allow them to mate overnight (see Note 3). 2. Check for the vaginal plug in female mice the next morning. Noon of the day that the plug is detected is designated as day 0.5 of gestation (E0.5, see Note 4).

3.2.1 Preparation of Pregnant Mice 3.2.2 Skin of E12.5E16.5 Embryos

1. The skin is removed from the dorsal lateral trunk region with fine forceps. 2. Skin samples are incubated at 37  C for 6 min in Dispase II solution (see Note 5). 3. Gently dissociate cells by passing them through an 18- to 21-G needle. 4. Add 2 volumes of SM to quench digestion, and then centrifuge the cells at 200
g for 4 min (see Note 6).

3.2.3 Skin of E17.5E19.5 Embryos and P0.5P6 Neonates

1. The back skin is removed with scissors and incubated at 37  C for 40 min in 0.25% collagenase type I in PBS (see Note 7). 2. Epidermal sheets are peeled mechanically with fine forceps from digested dermal tissues during observations through a binocular microscope. 3. The peeled epidermal sheet and dermal tissues are each washed with 5 mL of Hanks’ balanced salt solution. 4. Samples are cut finely and then incubated in cell-dissociation buffer at 37  C for 10–15 min (see Note 8).

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5. Dissociate cells by passage through an 18- to 21-G needle (see Note 9). 6. Add 2 volumes of SM to quench digestion, and then centrifuge the dissociated cells at 200
g for 4 min (see Note 10). 3.2.4 Vibrissa Follicles

1. The upper lip containing the vibrissa pad of neonates is cut to expose its inner surface. 2. Vibrissa follicles are gently dissected from the vibrissa pad under a microscope. 3. Isolated follicles are then washed and incubated at 37  C for 30 min in 0.05% trypsin solution (see Note 11). 4. Samples are gently dissociated by passage through 21-G needles. 5. Add 2 volumes of SM to quench the digestion, and then centrifuge dissociated cells at 200
g for 4 min (see Note 12).

3.3 Isolation of MBs by Flow Cytometry and the Culture of the Cells

3.3.1 Immunostaining of Skin Cell Suspensions for Flow Cytometry

Dissociated cells are immunostained with anti-KIT antibodies and anti-CD45 antibodies. The KIT molecule is an MB marker, and CD45 is a hematopoietic cell-specific marker. Although KIT molecules are a good MB marker, they are also expressed on hematopoietic cells in the skin [17]. Therefore, in order to eliminate hematopoietic cells, use anti-CD45 antibodies and isolate KIT-positive and CD45-negative cells as MBs from fetal skin by flow cytometry (Fig. 1a). Our recent analysis revealed that the populations of KIT-positive and CD45-negative cells were not sufficiently purified MBs. In order to further isolate purified MBs, we generated Sox10-IRES-Venus mice [18]. In these mice, the VENUS GFP variant gene [19] under the control of the internal ribosomal entry site (IRES) has been placed downstream of the Sox10 stop codon (Fig. 1a in [20]). SOX10 is an NCC marker and MB marker. Using these mice, MBs are isolated in the SOX10positive, KIT-positive, and CD45-negative cells (Fig. 1b). We herein describe the isolation of MBs from Sox10-IRES-Venus mice with a flow cytometer. 1. Dissociated cells are washed twice with SM. 2. Add the rat anti-mouse Fc gamma receptor and incubate cells on ice for 30–40 min (see Note 13). 3. Wash cells twice with SM. 4. Add the PE-conjugated rat anti-mouse CD45 and APC-conjugated rat anti-mouse KIT (see Notes 14 and 15). 5. Incubate on ice for 30–40 min. 6. Wash cells twice with SM. 7. Resuspend cells in PI solution (see Note 16).

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Fig. 1 Analysis and isolation of skin MBs by flow cytometry. (a) Flow cytometric analysis of Kit+/CD45 cells in skin from E12.5 embryos. Skin cells were stained with anti-Kit-APC antibodies, anti-CD45-FITC antibodies, and PI. The analysis is performed using PI-negative cells (line gate). The numbers show the average percentage of Kit+/CD45 cells (red square gate). (b) Flow cytometric analysis of SOX10+/KIT+/CD45 cells in skin from E12.5 embryos. Skin cells derived from Sox10-IRES-Venus mice were stained with anti-KitAPC antibodies, anti-CD45-PE antibodies, and PI. The analysis is performed using CD45-negative/PI-negative cells (black square gate). The numbers show the average percentage of SOX10+/KIT+ cells (red square gate)

3.3.2 Isolation of MBs by Flow Cytometry and the Culture of the Cells

The immunostained cell suspensions prepared in Subheading 3.3.1 are analyzed and sorted with a flow cytometer (see Note 17). 1. Isolate SOX10-positive, KIT-positive, and CD45-negative cells (SOX10+/KIT+/CD45 cells) from the skin cell suspension by performing flow cytometry (Fig. 1b) (see Note 18). 2. In order to culture sorted cells, directly inoculate the sorted cells (100–200 or single cells) into the wells of culture plates by using the flow cytometry system (see Note 19). The wells need to have previously been seeded with ST2 stromal cells and contain medium for differentiation of melanoblasts (see Note 20). 3. Incubate sorted cells in a 5% CO2 incubator at 37  C. Medium is changed every 2 days. The day on which sorted cells are seeded onto the ST2 monolayers is defined as day 0.

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Fig. 2 SOX10+/KIT+/CD45 cells differentiate into melanocytes, neurons, and glial cells. SOX10+/KIT+/ CD45 cells were sorted by flow cytometry (Fig. 1b, red square gate) from E12.5 skin, and inoculated at 100 cells/well onto ST2 monolayers in a 6-well dish. After 21 days in culture, colonies were immunostained for the neuronal marker β-tubulin (TuJ-1) and glial marker (GFAP). Melanocytes were detected as pigmented cells. Merge indicates the merged image. The same visual field is shown in each photo. Scale bar ¼ 200 μm

3.4 Culture and Immunohistochemical Analysis of Isolated MBs

3.4.1 Immunohistochemical Analysis of Isolated MBs

After being cultured for 10–14 days, SOX10+/KIT+/CD45 cells form colonies that contain pigmented Ms. (Fig. 2). After 21 days, some colonies contain TuJ-1-positive Ns and GFAP-positive Gs together with Ms. (Fig. 2). SOX10+/KIT+/CD45 cells also form colonies composed of 2 cell types (M/N, M/G, and N/G) and of 1 cell type (M, N, and G). Therefore, it is conceivable that formerly designated MBs have the potential to differentiate into multilineage cells.

1. Aspirate the culture medium from cultures and wash them three times with PBS. 2. Add 4% PFA in PBS to fix cells (see Note 21). 3. Incubate at room temperature for 15 min. 4. Wash three times with PBS.

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5. Add 0.1% Triton-X100 in 0.5% BSA PBS to render cells permeable (see Note 22). 6. Incubate at room temperature for 30 min. 7. Wash three times with PBS. 8. Add a mixture of 3% goat serum or 5% BSA in PBS to block the nonspecific binding of antibodies. 9. Incubate at room temperature for at least 30 min. 10. Wash three times with PBS. 11. Dilute the primary antibody (TuJ-1) in 0.5% BSA PBS (1:500, see Note 15), and then add it to cells at room temperature (see Note 23). 12. Wash three times with PBS. 13. Dilute the secondary antibody (Texas Red-conjugated antimouse IgG) in 0.5% BSA PBS (1:1000, see Note 15), and then add it to react with the primary antibody. 14. Wash cells three times with PBS, and react them with the antiGFAP (1:500, see Note 15), and then with the Alexa-Fluor 488-conjugated anti-rabbit IgG (1:1000, see Note 15) in the same manner. 15. Wash stained cells three times with PBS, and examine the colonies under the fluorescence microscope (see Note 24).

4

Notes 1. Do not make a stock solution. Prepare fresh solution when needed. 2. ST2 cultures sometimes exhibit a change in appearance (i.e., change to a more dendritic shape or senescent appearance) when cultured for long periods. Discard these cultures and use freshly thawed ST2 cells from the frozen stock. 3. We mainly use C57BL/6 mice in our experiments. 4. The developmental stages of embryos are judged by their morphological appearance, as described in “The Mouse” [21]. All animal experiments need to be performed in accordance with the appropriate regulations for animal experiments. 5. We use 1 mL of Dispase II solution for five embryonic skins and incubate them for up to 15 min for dissociation. 6. Approximately 5
106 cells are collected from eight embryonic skins. 7. We use 10 mL of collagenase solution for three neonatal skins and incubate them for up to 60 min for dissociation.

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8. We use 5 mL of cell-dissociation buffer for three neonatal skins and incubate them for up to 15 min for dissociation. 9. Passing the sheet several times through a blue-tip Pipetman with its tip cut back approx. 2–5 mm before using the needle helps the dissociation process using the needle. 10. Approximately 2
107 cells are collected from three neonatal skins. 11. We use 500 μL of trypsin solution for each neonatal follicle. 12. Approximately 3
106 cells are collected from the vibrissa follicles of one neonate. 13. Antibodies against the Fc gamma receptor block the nonspecific cell-surface binding of antibodies. Add an appropriate volume of the diluted antibody solution according to the supplier’s recommendations. 14. Keep aside a small amount of sample without the addition of antibodies. This sample is used as a negative control in the flow cytometric analysis. 15. Add an appropriate volume of the diluted antibody solution according to the supplier’s recommendations. 16. Only dead cells are stained by propidium iodide solution (Fig. 1b). 17. We use FACS Vantage or FACSAria from Becton Dickinson. 18. Exclude PI-positive dead cells (Fig. 1b). 19. One hundred-two hundred and single sorted cells are introduced into 6-well and 96-well plates, respectively. 20. Medium volumes are 2 mL/well and 200 μL/well in 6-well and 96-well plates, respectively. 21. Fixative volumes are 1 mL/well and 200 μL/well of 4% PFA for 6-well and 96-well plates, respectively. 22. Triton-X volumes are 500 μL/well and 200 μL/well for 6-well and 96-well plates, respectively. 23. Incubate at room temperature for more than 30 min or at 4  C overnight. 24. We use an Olympus IX-71 fluorescence microscope.

Acknowledgements This study was supported by a grant from the program Grants-inAid for Scientific Research (C) from the Japan Society for Promotion of Science.

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References 1. Luciani F, Champeval D, Herbette A et al (2011) Biological and mathematical modeling of melanocyte development. Development 138:3943–3954 2. Larue L, de Vuyst F, Delmas V (2013) Modeling melanoblast development. Cell Mol Life Sci 70:1067–1079 3. Le Douarin NM, Kalcheim C (1999) The neural crest, 2nd edn. Cambridge University Press, Cambridge 4. Wilson YM, Richards KL, Ford-Perriss ML et al (2004) Neural crest cell lineage segregation in the mouse neural tube. Development 131:6153–6162 5. Fernandes KJ, McKenzie IA, Mill P et al (2004) A dermal niche for multipotent adult skin-derived precursor cells. Nat Cell Biol 6:1082–1093 6. Toma JG, McKenzie IA, Bagli D et al (2005) Isolation and characterization of multipotent skin-derived precursors from human skin. Stem Cells 23:727–737 7. Wong CE, Paratore C, Dours-Zimmermann MT et al (2006) Neural crest-derived cells with stem cell features can be traced back to multiple lineages in the adult skin. J Cell Biol 175:1005–1015 8. Sieber-Blum M, Grim M, Hu YF et al (2004) Pluripotent neural crest stem cells in the adult hair follicle. Dev Dyn 231:258–269 9. Amoh Y, Li L, Katsuoka K et al (2005) Multipotent nestin-positive, keratin-negative hairfollicle bulge stem cells can form neurons. Proc Natl Acad Sci U S A 102:5530–5534 10. Yu H, Fang D, Kumar SM et al (2006) Isolation of a novel population of multipotent adult stem cells from human hair follicles. Am J Pathol 168:1879–1888 11. Dupin E, Glavieux C, Vaigot P et al (2000) Endothelin 3 induces the reversion of melanocytes to glia through a neural crest-derived glial-melanocytic progenitor. Proc Natl Acad Sci U S A 97:7882–7887

12. Trentin A, Glavieux-Pardanaud C, Le Douarin NM et al (2004) Self-renewal capacity is a widespread property of various types of neural crest precursor cells. Proc Natl Acad Sci U S A 101:4495–4500 13. Real C, Glavieux-Pardanaud C, Le Douarin NM et al (2006) Clonally cultured differentiated pigment cells can dedifferentiate and generate multipotent progenitors with selfrenewing potential. Dev Biol 300:656–669 14. Dupin E, Real C, Glavieux-Pardanaud C et al (2003) Reversal of developmental restrictions in neural crest lineages: transition from Schwann cells to glial-melanocytic precursors in vitro. Proc Natl Acad Sci U S A 100:5229–5233 15. Real C, Glavieux-Pardanaud C, Vaigot P et al (2005) The instability of the neural crest phenotypes: Schwann cells can differentiate into myofibroblasts. Int J Dev Biol 49:151–159 16. Motohashi T, Yamanaka K, Chiba K et al (2009) Unexpected multipotency of melanoblasts isolated from murine skin. Stem Cells 27:888–897 17. Lagasse E, Connors H, Al-Dhalimy M et al (2000) Purified hematopoietic stem cells can differentiate into hepatocytes in vivo. Nat Med 6:1229–1234 18. Motohashi T, Yamanaka K, Chiba K et al (2011) Neural crest cells retain their capability for multipotential differentiation even after lineage-restricted stages. Dev Dyn 240:1681–1693 19. Nagai T, Ibata K, Park ES et al (2002) A variant of yellow fluorescent protein with fast and efficient maturation for cell-biological applications. Nat Biotechnol 20:87–90 20. Motohashi T, Kunisada T (2018) Direct conversion of mouse embryonic fibroblasts into neural crest cells. Methods Mol Biol. https:// doi.org/10.1007/7651_2018_145 21. Rugh R (1990) The mouse. Oxford University Press, New York

Methods in Molecular Biology (2019) 1879: 267–284 DOI 10.1007/7651_2018_143 © Springer Science+Business Media New York 2018 Published online: 28 April 2018

Regeneration of Mouse Skin Melanocyte Stem Cells In Vivo and In Vitro Ke Yang, Weiming Qiu, Pei-Rong Gu, and Mingxing Lei Abstract Coordinated regeneration of melanocyte stem cells (McSCs) and hair follicle stem cells (HSCs) contributes to generation of pigmented hairs. Synchronous regeneration of McSCs with activation of HSCs occurs not only during initiation of a new hair cycle in vivo but also during reconstitution of hair follicles in vitro. The duration of the quiescent state of these stem cells becomes longer and longer in lifespan of mammals, leading to a decreased regenerative ability to form hair follicles. Here, we describe methods to activate McSCs during hair follicle regeneration in vivo, and isolate melanocytes from neonatal mouse skin to generate an immortalized cell line of melanocyte progenitors in vitro, aiming to use them for studying melanogenesis and future clinical application. Keywords Hair cycle, Hair follicle stem cells, Melanocyte stem cells, Melanogenesis, Regeneration, Skin

1

Introduction Adult stem cells reside in specialized niches which interact with the surrounding environment to orchestrate tissue homeostasis [1–3]. In mammals, skin melanocyte stem cells (McSCs) reside in hair follicle bulge, the niche where the epithelial hair follicle stem cells (HSCs) locate. McSCs originate from neural crest cells, exit from neural crest tube, and migrate through skin dermis to epidermis and hair follicle. In murine hair follicle, McSCs undergo cyclic activation, degradation, and quiescence in synchrony with HSCs during hair cycling [4] (Fig. 1). McSCs express dopachrome tautomerase (DCT) and paired box gene 3 (PAX3) [5]. McSCs are differentiated from melanoblasts and differentiate to generate melanocyte precursors, which express receptor tyrosine kinase (c-Kit), DCT, microphthalmia-associated transcription factor (MITF), PAX3, and tyrosinase related protein 1 (TRP1). Melanocyte precursors differentiate into mature melanocytes, which

Ke Yang and Weiming Qiu contributed equally to this work.

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Fig. 1 Melanocyte stem cells (McSCs) regeneration during hair cycling. At early anagen, receiving the stimulating signals from DP and HSCs, McSCs are activated and initiate to proliferate. At full anagen, McSCs exit hair bulge, migrate to hair bulb, and differentiate into melanocyte precursors and mature melanocytes. At telogen, McSCs reside in the bulge of hair follicle. Ana Anagen, DP dermal papilla, HG secondary hair germ, HB hair bulb

produce melanin to pigment the hairs. Melanocytes show Tyrosinase (TYR) positive in addition to the genes that melanoblasts express. Uncoupling of McSCs and epithelial HSCs behaviors occurs under certain transient conditions in skin, resulting in differentiation of McSCs not only to pigment hair follicle but also to pigment the skin epidermis. External stimuli such as exposure to ultraviolet radiation, chemical, and injury trigger activation and migration of McSCs into the skin epidermis, where the McSCs differentiate into melanocytes, leading to epidermis repigmentation [6, 7]. Hair graying-related diseases such as canities are common hypopigmentation problems that influence hundreds of thousands of people in the world [8]. Loss of hair follicle melanocytes, stalled activation of McSCs, or inhibition of melanogenesis causes hair graying. Interestingly, some of the gray hairs return pigmented

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upon clinical therapy, suggesting that the quiescent McSCs may still retain in the bulge of the gray hair follicles and can be reactivated to repigment the hairs. Thus, proper activation of McSCs is useful and effective for therapy of skin depigmentation diseases, such as canities and vitiligo [9]. Hair reconstitution assay indicates that melanoblasts can incorporate into the newly developing hair follicles and repopulate the McSCs population [10, 11]. However, isolation and expansion of melanocytes are difficult because they are normally rare in number and refractory to divide in vitro. Decades of studies have introduced the usage of mitogens like phorbol myristate acetate (PMA), cholera toxin, alpha-melanotropin, endothelin-1, or basic fibroblast growth factors in culture media for isolation and culture of melanocytes. An efficient culture system has been established to expand embryonic melanoblasts by adding a feeder layer of XB2 keratinocytes and several growth-stimulating factors [12]. Advances in the establishment of induced pluripotent stem (iPS) cell techniques enable reprogramming of somatic cells into the pluripotent state [13]. Combination of MITF, SOX10 (SRY-box 10), and PAX3 directly converts mouse and human fibroblasts to functional melanocytes [14]. Defined growth factors including Wnt3a, stem cell factors (SCF), and endothelin 3 (ET-3) induce differentiation of human and mouse pluripotent stem cells toward the melanocyte lineage [15]. These iPS cell-derived melanocytes successfully produce melanin pigment that can be delivered to surrounding keratinocytes during reconstitution assay. Collectively, it is significant to explore the McSCs regeneration for treating skin and hair hypopigmentation disorders. We recently discovered that exposure of hair-bearing skin to 12-O-tetradecanoylphorbol-13-acetate (TPA) induces regeneration, migration, and differentiation of McSCs in vivo [16]. We also established a step-by-step method to culture a conditionally immortalized cell line of mouse melanocyte progenitors in vitro [17]. In this chapter, we provide detailed protocols for regenerating McSCs in vivo and in vitro.

2 2.1

Materials Animals

2.2 12-OTetradecanoylphorbol13-Acetate Application In Vivo

Seven-week-old female C57BL/6 mice are raised in the individually ventilated cage system. 1. TPA,
99%. 2. Acetone (HPLC grade). 3. 1% Pentobarbital sodium. 4. Eye scissors.

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5. Electronic scale. 6. Canvas gloves. 7. Sterile disposable syringe (1 ml). 8. Electronic balance. 9. Beaker (1 L). 10. Pipette (100–1,000 μl, 20–200 μl, and 0.5–10 μl) and tips. 11. EP tubes (5.0 ml and 1.5 ml). 12. Straightedge. 13. Mark pen. 14. Cotton pad. 2.3 Fixation of Skin Tissues

1. 1% Pentobarbital sodium. 2. Eye scissors. 3. Tweezers. 4. Filter paper. 5. Ice-cold phosphate-buffered saline (PBS) buffers. 6. 4% Paraformaldehyde. 7. 50%, 75%, 85%, 95%, and 100% ethanol (HPLC grade). 8. Xylene (HPLC grade). 9. Paraffin. 10. Incubator at 60  C.

2.4

Immunostaining

1. Microtome. 2. Optical and fluorescence microscopes (Nikon N6000, Japan). 3. 50%, 75%, 85%, 95%, 100% ethanol and xylene. 4. PBS. 5. Slide and coverslip. 6. Citrate antigen retrieval solution (50 stock buffer, Beyotime Biotechnology, China). 7. Antigen retrieval box. 8. Microwave. 9. Bovine serum albumin (BSA, Beyotime). 10. Goat anti-DCT antibody (1:200; Santa Cruz, USA). 11. Rabbit anti-TYR antibody (1:300; Bioworld, USA). 12. Mouse anti-PCNA antibody (1:10,000; Abcam, USA). 13. Refrigerate at 4  C. 14. CY3-conjugated donkey anti-rabbit secondary antibody (1:300; Beyotime), CY3-conjugated donkey anti-goat

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secondary antibody (1:300; Beyotime), and DyLight 488-tagged donkey anti-mouse secondary antibody (1:300; Bioworld). 15. 40 ,6-Diamidino-2-phenylindole (DAPI, Beyotime). 16. Anti-fluorescence quenching reagent (Beyotime). 17. Double distilled water. 2.5 Isolation and Cell Culture Reagents

1. Dorsal skin from the neonatal C57BL/6 mouse. 2. Culture medium: Dulbecco’s modified eagle medium (DMEM) + Glutamax (Thermo Fisher Scientific) + 1% of 10 mg/ml penicillin/streptomycin. 3. Heat-inactivated fetal bovine serum (FBS, Gibco). 4. TPA. 5. Cholera toxin. 6. 0.25% Dispase II (Sigma). 7. Penicillin (10,000 U/ml); streptomycin 10,000 μg/ml. 8. D-Hank’s Balanced Salt Solution (HBSS, Ca2+ and Mg2+ free). 9. 0.25% Trypsin–EDTA.

2.6 Cell Culture Equipment

1. Fine tipped stainless-steel forceps. 2. 10 ml sterile pipettes. 3. Flask-25 cm2. 4. 15 ml conical tubes. 5. 10 cm tissue culture dishes. 6. Centrifuge fitted for 15 and 50 ml conical tubes.

3

Methods

3.1 12-OTetradecanoylphorbol13-Acetate Application to Induce Melanocyte Stem Cells Regeneration in Mice

In this section, we describe the key steps to inducing McSCs regeneration upon TPA application in vivo.

3.1.1 Preparation of Mice and 12-OTetradecanoylphorbol-13Acetate Working Solution

1. Obtain 7-week-old female mice with pink dorsal skin, which indicates that the hair follicles are at telogen, rather than anagen or catagen of the hair cycle. 2. Weigh the mouse with an electronic balance. Anesthetize mice by intraperitoneally injecting 1% pentobarbital sodium at room temperature, with 30–50 mg/kg per mouse.

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3. Mark 3  3 cm2 area of dorsal skin with a mark pen. Clip dorsal fur of this area with eye scissors. 4. Make a stock solution of 10 mg/ml TPA by dissolving 1 mg TPA in 0.1 ml acetone. Store the stock solution at 20  C (see Note 1). 5. Make the working solution of TPA at 0.01 mg/ml. For example, to make 4 ml working solution, dissolve 4 μl stock solution in 3.996 ml acetone. Aliquot the TPA working solution into 1.5 ml EP tubes, with 400 μl per tube. 3.1.2 Application of 12-OTetradecanoylphorbol-13Acetate to the Mice

1. Randomize the mice into four groups (Fig. 2a), including A2 group (acetone treatment for two doses, as the control group), T2 group (TPA treatment for two doses), A8 group (acetone treatment for eight doses, as the control group), and T8 group (TPA treatment for eight doses). Raise the mice in different cages. 2. Put a mouse on a cotton pad. In TPA treatment groups, pipette 200 μl TPA working solution from the 1.5 ml EP tubes, and apply it onto the 3  3 cm2 hair-clipped dorsal skin area from the left to the right, and from the anterior to the posterior (Fig. 2b) (see Note 2). 3. Apply another dose of 200 μl TPA working solution after the first applied acetone is fully volatilized. Apply 200 μl acetone onto the dorsal skin of control groups twice (see Note 3). 4. Put the mice back into the cages, and raise them under a standard housing condition (see Note 4). 5. Three days later, perform another application of TPA or acetone. 6. Continue twice weekly applications of TPA working solution or acetone onto the clipped area of the mice. For example, schedule TPA applications on Monday and Thursday each week. 7. Pigmented hairs grow out of the dorsal skin in T8 group after the eighth treatment with TPA, whereas the skin in acetonetreated groups (A8) remains pink.

3.2 Analyze Melanocyte Proliferation and Differentiation in Mice 3.2.1 Sampling

Perform immunofluorescence staining on paraffin sections to check the proliferation, differentiation, and migration of melanocytes after TPA treatment.

1. Twenty-four hours after the second application of TPA or acetone to the mice, sacrifice the mice in A2 and T2 groups, and collect the skin tissues (Fig. 2a). 2. Forty-eight hours after the eighth treatment with TPA or acetone to the mice, anesthetize the mice by intraperitoneally

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Fig. 2 Regeneration of McSCs in mouse dorsal skin after 12-O-tetradecanoylphorbol-13-acetate (TPA) treatment. (a) Schematic shows timing of TPA application. (b) Schematic shows method of TPA application. (c) Double immunostaining detects the proliferating McSCs (DCT- (dopachrome tautomerase) and PCNApositive cells) after two doses of TPA treatment. (d) Immunostaining for DCT shows that melanocytes are located in skin epidermis and hair infundibulum after two doses of TPA treatment. (e) Immunostaining for DCT shows that melanocytes are located in hair bulb of full anagen hair follicle. A acetone treatment, T TPA treatment. Red arrowhead indicates DCT-positive cells, and yellow arrowhead shows DCT- and PCNA-positive cells

injecting 1% pentobarbital sodium in A8 and T8 groups, shave the newly regenerated hairs, and collect the skin tissues. 3. Wash the skin samples with ice-cold PBS, and cut them into 1 cm2 pieces with eye scissors and tweezers. 4. Flatten the skin samples onto a filter paper, and fix them in 4% paraformaldehyde at 4  C for overnight. 5. Wash the skin samples in PBS for 10 min and dehydrate them with a gradient series of ethanol (50%, 75%, 85%, and 95% ethanol for 1.5 h, respectively, 100% ethanol twice for 1 h each).

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6. Transparentize the skin samples by treating them with xylene twice for 7 min each. 7. Put skin samples in melted paraffin (melting point: 56–58  C) for 4 h. Embed skin samples in 12-well culture plates to make paraffin blocks. 3.2.2 Immunostaining

1. Section the skin tissues at 5 μm thickness using microtome and heat the paraffin slides at 60  C for 2 h (see Note 5). 2. Dewax the slides with xylene twice for 15 min each. 3. Put the slides in a gradient series of ethanol (100% ethanol twice for 5 min each; 95%, 85%, 75%, and 50% ethanol for 5 min, respectively), and then wash the slides with PBS for 5 min (see Note 6). 4. Make 1 antigen retrieval working solution by diluting the 50 stock citrate antigen retrieval solution with double distilled water. Put the diluted solution into an antigen retrieval box and boil it by using microwave. Put the slides into the boiled antigen retrieval buffer and microwave them with low power for 10 min. Cool down the slides in the antigen retrieval solution to room temperature (see Note 7). 5. Rinse the slides in PBS twice for 5 min each. Add 50 μl 5% BSA in PBS to the skin samples to block nonspecific antibody binding at room temperature for 1 h. 6. Incubate the skin samples with primary antibody (goat antiDCT antibody, diluted with 5% BSA in PBS) at 4  C for overnight. For double-immunofluorescence staining, dilute goat anti-DCT antibody and mouse anti-PCNA antibody with 5% BSA in PBS, and add them to the skin samples, which are then incubated in a humidified chamber at 4  C for overnight. 7. Rinse the samples three times in PBS for 10 min each. 8. Incubate the samples with secondary antibody (CY3-conjugated donkey anti-goat secondary antibody) in a humidified chamber at room temperature for 2 h. For the double-immunofluorescence staining, mix the CY3-conjugated donkey anti-goat secondary antibody with the DyLight-conjugated donkey anti-mouse secondary antibody) and add them to the skin samples, which are then incubated in a humidified chamber at 37  C for 1 h. 9. Remove the secondary antibody on the samples. Stain the nuclei with DAPI solution for 5 min. 10. Rinse the samples in PBS three times for 10 min each, add antifluorescence quenching reagent to slides and cover the samples with a coverslip. 11. Obtain images by fluorescence microscope.

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1. After two doses of TPA treatment, the cell proliferation marker PCNA and melanocyte lineage marker DCT are co-expressed in several cells at hair bulge and secondary hair germ regions (Fig. 2c), indicating that McSCs are activated. 2. DCT-positive cells are also observed in hair infundibulum and interfollicular epidermis after two doses of TPA treatment, compared to the A2 control group in which DCT-positive cells are located at the bulge and secondary hair germ regions (Fig. 2d). 3. After eight doses of TPA treatment, pigmented hairs are regenerated. DCT-positive melanocytes reside in hair matrix (Fig. 2e), indicating that McSCs are differentiated into melanocytes to pigment the hairs. 4. McSCs are responsible for melanocyte regeneration in epidermis and hair follicles. In the following sections, we describe the method to establish immortalized melanocyte progenitors in vitro.

3.3 Isolation of Melanocyte Cells from Mouse Skin In Vitro

1. Complete culture medium: DMEM culture medium containing 10% FBS, 200 nM TPA, and 200 pM cholera toxin. 2. Plain culture medium: culture medium containing 10% FBS.

3.3.1 Medium Preparation 3.3.2 Isolation of Melanocyte from Mouse Skin

1. Fill several 10 cm plastic plates with cold sterile HBSS and place the plates on ice for use of skin collection (see Note 8). 2. Thaw aliquots of Dispase II. One milliter of Dispase II (0.25%) is used to digest one piece of the neonatal back skin sample. 3. Sacrifice newborn C57BL/6 mice by CO2. 4. Wash the mice with 70% ethanol for 30 s and then in sterile HBSS with penicillin (10,000 U/ml) and streptomycin 10,000 mg/ml for 10 min. 5. Dissect dorsal back skin using fine forceps and scissors. 6. Gently remove all visible subcutaneous adipose tissue, panniculus carnosus, and blood vessels as clean as possible. Wash the skin samples in HBSS and place them into 0.25% Dispase II in 15 ml conical tubes. Incubate the samples at 4  C for overnight (see Note 9). 7. Transfer the skin samples to a dry, sterile tissue culture plate with the epidermis side up. Peel off the epidermis from the dermis using sterile forceps and transfer the epidermis to another plate.

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8. Mince the epidermis tissues and transfer them into a 50-ml conical tube. Digest them in 0.25% trypsin–EDTA for 3–5 min at room temperature. 9. Dilute and neutralize trypsin by adding 10 ml plain culture medium containing 10% FBS. Mix them by gently pipetting up and down 3–5 times. 10. Centrifuge tissue samples using a tabletop centrifuge at 150  g for 5 min to collect the dissociated cells. Cell pellet is visible at the bottom of the tube. 11. Remove all supernatant. Resuspend the pellet in complete culture medium. Dilute the cells to an optimum density of 1–5  105 cells in 1 ml complete culture medium. Transfer 6 ml cell solution into a 25-cm2 flask and culture it in a 37  C, 5% CO2 tissue culture incubator (see Note 10). 3.4 Establishment of Immortalized Melanocyte Cell Lines 3.4.1 Additional Reagents

1. Plasmid pAmpho (Molecular Oncology Laboratory, Medical Center, The University of Chicago). 2. Plasmid SSR #41 (Molecular Oncology Laboratory, Medical Center, The University of Chicago). 3. Lipofectamine 2000 (Invitrogen). 4. 0.45 μm Cellulose acetate or polysulfonic (low protein binding) filters. 5. Hygromycin B (Invitrogen). 6. Polybrene (Sigma-Aldrich).

3.4.2 Generation of Retrovirus Supernatants

1. Seed 1  105 HEK293 cells in a 25-cm2 flask (usually one flask for making one stable line). 2. Four hours later, gently wash the cells with 3 ml serum-free DMEM medium, and add 2.5 ml serum-free DMEM medium into the flask. Transfect the cells with a mixture of 250 μl DMEM (serum-free), 5 μl pAmpho (~0.5–1.0 μg DNA), 10 μl SSR #41 (~1.0–2.0 μg DNA), and 15 μl Lipofectamine (Invitrogen) (see Note 11). 3. Change the medium with 4 ml complete culture medium 4 h after transfection. Change the culture medium at 36, 60, 84, and 108 h, collect the supernatant which contains retroviruses, and store them at 4  C. The total volume of retrovirus supernatant should be about 16 ml. 4. Centrifuge the retrovirus supernatant at 150  g for 5 min at 4  C to remove the cell debris. 5. Filter retrovirus supernatant through a 0.45-μm cellulose acetate or polysulfonic (low protein binding) filter. Add 80 μl of 2 mg/ml polybrene to 16 ml retrovirus supernatant to prepare polybrene-containing retrovirus supernatant.

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1. When melanocytes grow to confluency (~20–30%), change the medium with 4 ml polybrene-containing retrovirus supernatant (see Note 12). 2. 6–8 h later, add another 4 ml polybrene-containing retrovirus supernatant to the cells and culture them for overnight. The retrovirus supernatant medium is replaced every 8 h with 3–4 rounds (see Note 13). 3. Melanocytes are selected with hygromycin B. 0.3 mg/ml Hygromycin B is added to 4 ml polybrene-containing retrovirus supernatant at the second, third, and fourth infection. About 7–10 days later, the melanocytes grow stably in the hygromycin B selection culture medium (see Note 14). 4. Immortalized melanocytes can grow in the plain culture medium containing 10% FBS. 2 ml 0.25% trypsin–EDTA is added to the plate to digest the cells when passage. Centrifuge the cells for 5 min at 150  g, remove the supernatant, and resuspend the cell pellet with plain culture medium containing 10% FBS (see Note 15). 5. Adjust the cell density to 1  105/ml with plain culture medium. Transfer 10 μl cell suspension to 1 ml culture medium and mix them by gently pipetting. Repeat the 1:100 dilution to obtain a final cell density at 10 cells per 1 ml culture medium. 6. Add 100 μl of the final diluted cell suspension to each well of 96-well plates. Incubate the plates at 37  C in a humidified CO2 incubator. Change culture medium every 3 days (see Note 16). 7. 7–10 days later, check each well and mark the wells that contain only a single colony. Digest the colony with 100 μl 0.25% trypsin–EDTA, wash the cells with DMEM containing 10% FBS, and culture the cells in 24-well plates. The cells are passaged into 12-well plates, then into 6-well plates (Fig. 3a–d). 8. Each cell colony can be cryopreserved in liquid nitrogen.

3.5 Screening and Detection of Immortalized Melanocyte Cell Lines 3.5.1 Additional Reagents

Collect and analyze each colony for evaluating cell differentiation as follows.

1. Triton X-100. 2. BCA protein assay kit (Pierce Biotechnology, Inc., Rockford, IL, USA). 3. L-DOPA. 4. Gold chloride solution (Abcam).

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Fig. 3 Generation of melanocytes in vitro. (a) Primary melanocytes can be isolated from newborn mouse skin. (b) Immortalized melanocytes grow in the plain culture medium. Impure cell populations show different morphology and melanin production. (c) and (d) show the cells that are passaged from the different single colony picked from 96-well plate

5. Nuclear fast red solution. 6. Sodium thiosulfate solution. 7. Antibodies against c-Kit, tyrosinase, TRP1, and DCT (Santa Cruz). 8. Texas Red-labeled secondary antibody (Jackson). 9. Cell Counting Kit-8 (CCK-8, Biyotime). 10. Bromodeoxyuridine (BrdU, Sigma). 11. Wnt3a recombinant protein (R&D). 3.5.2 Cell Pellets

1. Resuspend and wash cells with plain culture medium twice, and centrifuge them at 150  g for 10 min. 2. Observe the color of cell pellets. The pellet of melanoblast-like cells (MB-like) displays colorless, whereas the pellet of melanocyte-like (MC-like) cells shows brown or black (Fig. 4c).

Fig. 4 Analysis of immortalized melanocytes. (a) Masson Fontana staining of MB-like and MC-like cells. (b) Relative tyrosinase activity of MB-like and MC-like cells. (c) Cell pellets of MB-like and MC-like cells. (d) Immunofluorescence staining for DCT and c-Kit in MB-like and MC-like cells. MB-like, melanoblast-like cells. MC-like, melanocyte-like cells

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3.5.3 Tyrosinase Enzyme Activity

1. Plate 5  104 cells into 24-well plate. 2. Wash the cells with 500 μl HBSS twice after the cells grow to 80–90% confluency. Add 200 μl 1% Triton X-100 to each well to lyse the cells, and put the plate in 80  C over 2 h, followed by thawing completely in 37  C for 10 min. The cell lysate is centrifuged at 10,000  g for 10 min and the supernatant is harvested. 3. The protein concentration is determined using a BCA protein assay kit. 4. Add 40 μg protein lysates, 2.0 mM L-DOPA, and 0.1 M PBS (pH 6.8) to each well of 96-well plate and incubate them at 37  C for 1 h. The absorbance is measured at 475 nm by using an enzyme-linked immunosorbent assay reader. 5. Tyrosinase activity was calculated using the following formula. Tyrosinase activity (%) ¼ (OD475 of sample/OD475 of control)  100%. 6. The tyrosinase activity is significantly lower in MB-like cells than in MC-like cells (Fig. 4b).

3.5.4 Masson Fontana Staining

1. Seed 5  104 cells onto the coverslip and culture them for 24 h. Wash the cells with HBSS and fix them in 4% paraformaldehyde at 4  C for 1 h. Rinse the cells with HBSS 1 min for three times. 2. Incubate the cells in 0.2% gold chloride solution at room temperature for 30 s. Rinse the cells with distilled water 1 min for three times. 3. Incubate the cells in 5% sodium thiosulfate solution at room temperature for 1–2 min. Rinse the cells with running tap water for 2 min twice each. 4. Incubate the cells in nuclear fast red solution for 5 min. Rinse the cells in running tap water for 2 min twice each. 5. Dehydrate the cells with 100% ethanol three times for 2 min each. Transparentize the cells with xylene for 5 min and mount them with resinene. 6. Observe staining of melanin granules in MC-like cells. No obvious staining is observed in MB-like cells under phasecontrast microscope (Fig. 4a).

3.5.5 Immunofluorescence Staining

1. Cells on coverslip are fixed with methanol, permeabilized with 1% Triton-X100, and blocked with 10% donkey serum, following by incubating with antibodies against c-Kit, tyrosinase, TRP1, and DCT at room temperature for 60 min. 2. Wash the cells with PBS three times for 10 min each and then incubate them with Texas Red-labeled secondary antibody at room temperature for 30 min.

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3. Scan the image under a fluorescence microscope. Samples without adding primary antibodies, or with IgG control, are used as negative controls. 4. Immunofluorescence staining shows that the cells express DCT and c-Kit (Fig. 4d), but not tyrosinase and TRP1, indicating that these colonies should be melanoblast-like cells. 3.6 Analyze Proliferation and Differentiation of Melanocyte Cell Lines 3.6.1 Examine Melanocytes Proliferation Using Cell Counting Kit-8

1. Seed cells in a 96-well plate at a density of 104–105 cells/well in 100 μl culture medium. Culture the cells in a CO2 incubator at 37  C for 24 h. 2. Add 10 μl of a certain concentration of treatment factor, such as 100 ng/ml Wnt3a recombinant protein, to the plate. 3. Incubate the plate for an appropriate length of time (e.g., 6, 12, 24, or 48 h) in the incubator. 4. Add 10 μl CCK-8 solution to each well of the 96-well plate. Avoid introducing bubbles to the wells. 5. Incubate the plate in the incubator at 37  C for 4 h. 6. It is important to gently mix the solution on an orbital shaker at room temperature for 1 min to ensure homogeneous distribution of color, before measuring the absorbance. 7. Measure the absorbance at 450 nm using a microplate reader.

3.6.2 Examine Melanocytes Proliferation Using Bromodeoxyuridine Incorporation

1. Plates 2500 cells/well in 24-well plate and incubate them with growth factors, such as 100 ng/ml Wnt3a recombinant protein. Incubate the cells at 37  C for 24 h. 2. Add 1 mM BrdU working solution in 500 μl DMEM culture medium to the plate, which is then incubated in the cell culture incubator for 4 h. 3. Remove the culture medium. Add 100 μl fixing/denaturing solution per well, and keep the plate at room temperature for 30 min. 4. Remove the fixing/denaturing solution. Add 100 μl primary antibody against BrdU to the plate, and keep it at room temperature for 1 h. 5. Wash the cells in PBS three times for 5 min each. Add 100 μl HRP-conjugated secondary antibody solution to the plate, and keep it at room temperature for 30 min. 6. Wash the cells in PBS three times for 5 min each. Add 100 μl four methyl diphenyl amine hydrochloride (TMB) substrate. Incubate it at room temperature for 30 min. 7. Add 100 μl STOP Solution. Read absorbance at 450 nm (for optimal readings, read the plate within 30 min after adding STOP Solution).

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3.6.3 Differentiation of Melanocyte

1. Plate 1  105 cells/well in 24-well plate. 2. Replace the culture medium with DMEM culture medium containing 100 ng/ml Wnt3a to induce melanocyte differentiation, when the cells grow to 30% confluency. 3. Culture the cells in the incubator. Change the medium every other day by removing 50% of the medium and replace it with the same volume of fresh differentiation medium. 4. The culture medium gradually turns to dark after the cells are cultured for 5–7 days. Detect the cell differentiation following the method shown in Subheadings 3.5.2, 3.5.3, 3.5.4, and 3.5.5.

4

Notes 1. Warning: TPA is a tumor-promoting agent. Avoid handling it without any protection. 2. Remember to put on the canvas gloves to avoid bite when catching the mouse. 3. The application of TPA to the dorsal skin should be performed at room temperature because the mice are in an anesthetic condition. Keep the mice warm after TPA application. 4. TPA should be applied to the dorsal skin. Return the mice back to the cages after the acetone is completely volatilized. 5. Cut the sections longitudinally along the anterior–posterior orientation to make sure the display of full hair follicle structure. 6. To reduce the auto-fluorescence background when performing immunofluorescence staining, dewax the slides thoroughly in xylene, block the slides with 5% BSA in PBS at room temperature for 1 h, avoid drying of samples, and rinse the samples with PBS sufficiently. 7. Microwave the samples in pH 6.0 citrate butter to expose the antigens to reduce the false-negative results. 8. Dissected skin from neonatal mice can be stored in HBSS at 4  C for 24 h and even shipped during this period without significantly reducing cell yield or health. 9. The skin tissue can be cut into 1–2 cm2 pieces. It will be difficult to separate epidermis from dermis when larger tissues are prepared. Digest the skin tissues at 4  C for less than 24 h. 10. During the primary cell culture, add freshly prepared TPA in complete culture medium twice a week because TPA is unstable.

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11. pAmpho is a packaging vector of retrovirus, and SSR#41 is a shuttle vector of retrovirus containing SV40T-antigen (SV40T). The immortalized transformants can be induced by the expression of SV40T. 12. Most of the skin keratinocytes differentiate 3 days after culture, remaining melanocytes in the culture plate after hygromycin B selection. The adherent cells are trypsinized and passaged to make stable melanocyte cell lines. 13. After the primarily cultured melanocytes are passaged from the primary 25-cm2 flask, the retrovirus supernatant should be added to the culture plate to select melanocytes. The retrovirus supernatant can be stored at 4  C at the maximum of 3 days. 14. The Streptomyces hygroscopicus hyg gene encoding a hygromycin B phosphotransferase has been inserted in the packaging retrovirus plasmid SSR #41. When the immortalized melanocytes are established, the cells express hygromycin B phosphotransferase and obtain the resistance to hygromycin B. The immortalized melanocytes can be selected by hygromycin B. 15. Once the immortalized cells grow stably in the selected medium, the cells should be replated into 96-well plate to pick the single colony. The more time the cells are cultured in 25-cm2 flask, the more number of colonies can be picked in 96-well plate. Many colonies can form from the same immortalized cells. 16. Once the immortalized melanocyte cell line is established, the cells can be cultured in plain culture medium without TPA and cholera toxin.

Acknowledgements Mingxing Lei is supported by the Higher Education Enhancement Program, China Medical University, Projects Funded by China Postdoctoral Science Foundation (2016M590866), Fundamental Research Funds for the Central Universities (106112015CDJRC 231206), Special Funding for Postdoctoral Research Projects in C hongqing (Xm2015093), and a fellowship from the China Scholarship Council (2011605042). Ke Yang was supported by National Natural Science Foundation of China (81371718). Weiming Qiu is supported by National Natural Science Foundation of China (81602782) and Natural Science Foundation of Hubei Province (2016CFB348). The authors thank Dr. T.-C. He at the University of Chicago for gifting retrovirus vector and technical assistance.

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References 1. Lei M, Guo H, Qiu W et al (2014) Modulating hair follicle size with wnt10b/dkk1 during hair regeneration. Exp Dermatol 23(6):407–413 2. Lei M, Chuong CM (2016) Stem cells. Aging, alopecia, and stem cells. Science 351 (6273):559–560 3. Lei M, Yang L, Chuong CM (2017) Getting to the core of the dermal papilla. J Invest Dermatol 137(11):2250–2253 4. Rabbani P, Takeo M, Chou W et al (2011) Coordinated activation of wnt in epithelial and melanocyte stem cells initiates pigmented hair regeneration. Cell 145(6):941–955 5. Osawa M, Egawa G, Mak SS et al (2005) Molecular characterization of melanocyte stem cells in their niche. Development 132 (24):5589–5599 6. Chou WC, Takeo M, Rabbani P et al (2013) Direct migration of follicular melanocyte stem cells to the epidermis after wounding or uvb irradiation is dependent on mc1r signaling. Nat Med 19(7):924–929 7. Qiu W, Yang K, Lei M et al (2015) Scf/c-kit signaling is required in 12-o-tetradecanoylphorbol-13-acetate-induced migration and differentiation of hair follicle melanocytes for epidermal pigmentation. Cell Tissue Res 360 (2):333–346 8. Zhang Z, Lei M, Xin H et al (2017) Wnt/betacatenin signaling promotes aging-associated hair graying in mice. Oncotarget 8 (41):69316–69327 9. Birlea SA, Costin GE, Roop DR et al (2017) Trends in regenerative medicine: Repigmentation in vitiligo through melanocyte stem cell mobilization. Med Res Rev 37(4):907–935

10. Yonetani S, Moriyama M, Nishigori C et al (2008) In vitro expansion of immature melanoblasts and their ability to repopulate melanocyte stem cells in the hair follicle. J Invest Dermatol 128(2):408–420 11. Lei M, Schumacher LJ, Lai YC et al (2017) Self-organization process in newborn skin organoid formation inspires strategy to restore hair regeneration of adult cells. Proc Natl Acad Sci U S A 114(34):E7101–E7110 12. Eisinger M, Marko O (1982) Selective proliferation of normal human melanocytes in vitro in the presence of phorbol ester and cholera toxin. Proc Natl Acad Sci U S A 79 (6):2018–2022 13. Takahashi K, Tanabe K, Ohnuki M et al (2007) Induction of pluripotent stem cells from adult human fibroblasts by defined factors. Cell 131 (5):861–872 14. Yang R, Zheng Y, Li L et al (2014) Direct conversion of mouse and human fibroblasts to functional melanocytes by defined factors. Nat Commun 5:5807 15. Yang R, Jiang M, Kumar SM et al (2011) Generation of melanocytes from induced pluripotent stem cells. J Invest Dermatol 131 (12):2458–2466 16. Qiu W, Tang H, Guo H et al (2016) 12-oTetradecanoylphorbol-13-acetate activates hair follicle melanocytes for hair pigmentation via wnt/beta-catenin signaling. Cell Tissue Res 366(2):329–340 17. Yang K, Chen J, Jiang W et al (2012) Conditional immortalization establishes a repertoire of mouse melanocyte progenitors with distinct melanogenic differentiation potential. J Invest Dermatol 132(10):2479–2483

Methods in Molecular Biology (2019) 1879: 285–297 DOI 10.1007/7651_2018_155 © Springer Science+Business Media New York 2018 Published online: 13 June 2018

Interactions Between Epidermal Keratinocytes, Dendritic Epidermal T-Cells, and Hair Follicle Stem Cells Krithika Badarinath, Abhik Dutta, Akshay Hegde, Neha Pincha, Rupali Gund, and Colin Jamora Abstract The interplay of immune cells and stem cells in maintaining skin homeostasis and repair is an exciting new frontier in cutaneous biology. With the growing appreciation of the importance of this new crosstalk comes the requirement of methods to interrogate the molecular underpinnings of these leukocyte–stem cell interactions. Here we describe how a combination of FACS, cellular coculture assays, and conditioned media treatments can be utilized to advance our understanding of this emerging area of intercellular communication between immune cells and stem cells. Keywords Coculture, Conditioned media, Explants, FACS, Immune cells, Skin, Stem cells

1

Introduction The skin is among the few organs in the mammalian body that undergoes cyclical regeneration throughout the lifetime of the organism. Moreover, as the protective barrier of the body, the skin is often subjected to assault from physical stress, pathogens, and environmental toxins. Any damage results in the mounting of a rapid repair response in order to restore tissue homeostasis. Underlying both skin regeneration and repair are the numerous populations of stem cells housed in various niches in the skin [1, 2]. These different pools of stem cells are regulated via a multitude of regulatory networks [3, 4]. Of these, a burgeoning new signaling axis impacting stem cell behavior is the role of adaptive and innate immune cells that both reside within and are recruited into the skin. In fact, the presence of a plethora of dendritic epidermal T cells (DETCs), Langerhans cells, mast cells, and macrophages in the skin has led to the historic moniker of the skin as the “immune organ” [5, 6]. The recent literature is revealing activities of these immune cells beyond their classical role of pathogen defense and is expanding their functional repertoire to important new roles in regulating skin homeostasis and wound healing [7–9]. Among the variety of cutaneous immune cells, DETCs are an important subset of skin

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resident T cells which play critical role in both tissue homeostasis and wound repair. For instance DETCs are required for normal wound healing kinetics [10] and the stimulation of new hair follicle formation [11]. Using FACS, cell coculture assays, and conditioned media treatments, we recently demonstrated that the release of interleukin-1α (IL-1α) from injured keratinocytes activates DETCs, which in turn activates hair follicle stem cell proliferation in a wound healing scenario [12]. The phenomenon of immune signals functioning in noninflammatory scenarios is not restricted to the skin. It has already been established that inflammatory cytokines play regulatory roles during hematopoiesis [13], as well as muscle [14] and lung repair [15]. Thus, recent advances suggest that the interplay of immune cells and stem cells in the skin and other organs play as yet undiscovered critical roles. Elucidation of this crosstalk and their functional ramifications requires the ability to recapitulate these processes in vitro by isolating and culturing these cells and reconstituting their proximal interactions. Variations on this approach can be used to investigate other immune cell–stem cell crosstalk in the skin and other organs.

2

Materials

2.1 Instruments/ Labware

1. Sterile 10 cm non-toothed dissecting forceps (Mistry medical supplies, Cat. No. MMS5012 or equivalent). 2. Sterile 10 cm dissecting scissors (Multigate medical products, Cat. No. 06-311 or equivalent). 3. Scalpel (Fisher Scientific, 14-840-01 or equivalent). 4. Bacterial grade Petri dishes (Fisher Scientific, Cat. No.08-757100D or equivalent). 5. 1.5 ml microcentrifuge tubes No. 0030125150 or equivalent).

(Eppendorf,

Cat.

6. 15 ml conical tubes (Falcon, Cat. No. 05-527-90 or equivalent). 7. 50 ml conical tubes (Falcon, Cat. No. 14-432-22 or equivalent). 8. 10 ml Serological pipette (Eppendorf, Cat. No. 0030127722 or equivalent). 9. S1 Pipette filler (Thermo Scientific). 10. Laminar flow hood. 11. 5 ml Round Bottom Polystyrene Tubes (Corning, Cat. No. 352058 or equivalent). 12. Vacuum aspirator.

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13. Refrigerated centrifuge for 50 ml, 15 ml conical tubes, and 1.5 ml microcentrifuge tubes. 14. Four color flow cytometer, such as Becton Dickinson FACS Aria fusion. 15. Micropipettes (Thermo Scientific, Cat. No. 4641020N or equivalent). 16. 1 ml, 200 μl, and 10 μl micropipette tips. 17. CO2 tissue culture incubator. 18. Kimwipes (Kimberly-Clark corporation, Cat. No. 06-666 or equivalent). 19. 96-well tissue culture dish (BD Falcon, Cat. No. 08-772-3B or equivalent). 20. Integra Miltex biopsy punches, 8 mm (Fischer Scientific, Cat. No. 12-460-413 or equivalent). 2.2 Buffers and Reagents 2.2.1 Isolation of DETC Antibodies and Isotype Controls

Solution A—(Staining Buffer): Add 1 ml of fetal bovine serum to 49 ml of sterile PBS. (See Note 1.) One aliquot of Solution A should be ice cold and the other aliquot should be at room temperature. Rat Anti-Mouse CD16/CD32 (Clone 2.4G2), (BD Pharmingen, Cat. No. 553141). FITC Hamster Anti-Mouse Vγ3 TCR (BD Pharmingen, Cat. No. 553229).

(Clone

536),

FITC Hamster IgG1 κ Isotype Control (Clone A19-3) 553971 (BD Pharmingen, Cat. No. 553971). PE Hamster Anti-Mouse γδ T-Cell Receptor (Clone GL3), (BD Pharmingen, Cat. No. 553178). PE

Hamster IgG2 κ Isotype Control (BD Pharmingen, Cat. No. 550085).

(Clone

B81-3),

PE-Cy™7 Hamster Anti-Mouse CD3e (Clone 145-2C11), (BD Pharmingen, Cat. No. 552774). PE-Cy™7 Hamster IgG1, κ Isotype Control (Clone A19-3), (BD Pharmingen, Cat. No. 552811). 7-AAD, (BD Pharmingen, Cat. No. 559925). 2.2.2 DETC Culture and Activation

7-17 DETC cell line (4) and primary DETCs [16]. RPMI 1640 (Sigma Aldrich, Cat. No. R8758). FBS (Gibco, 16000-044). Recombinant IL-2 (Sigma Aldrich, Cat. No. I0523). Ultra-low attachment culture dishes 100 mm, (Corning, Cat. No. CLS3262).

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Anti-JAML (eBioscience, clone eBio4E10). Anti-CD3 (e-bioscience, Cat. No. 14-0032-8). rIL-1β (R&D systems, Cat. No. 401-ML-025). rIL-23 (R&D systems, Cat. No. 1887-ML-010). 2.2.3 Mice

3

Pups (P1–P6), 21-day-old mice, or 8–9-week-old adult mice. Separation of the epidermis is easier during the morphogenesis phase (P1–P6) and during the quiescent stage of hair follicle cycle—first telogen phase (usually around P21) and second telogen phase (Usually 8–9 weeks postnatal).

Methods

3.1 Isolation and Maintenance of Dendritic Epidermal T Cells (DETCs) from Murine Skin

The preparation of epidermal stem cell suspensions [17] and FACS mediated analysis of DETCs [18] have been described.

3.1.1 Extraction of Epidermal Cells from the Dorsal Skin of Mice for Isolation of DETCs

Extraction of epidermal cells from the dorsal skin of mice has previously been reported [17]. Freshly extracted epidermal cells should be kept in suspension in Solution A.

3.1.2 Isolation of DETCs from Epidermal Cell Suspension by FACS [18]

1. From the epidermal cell suspension, aliquot 1  105 cells/tube from the epidermal cell suspension into three 5 ml FACS tubes labeled-unstained control, 7-AAD and isotype control for primary antibodies. 2. Aliquot 1  106 cells from the epidermal cell suspension for experimental into new 5 ml FACS tube (see Note 2). 3. Block the cells to prevent nonspecific binding of antibody by incubating the cells with anti-CD16/CD32 (1 μg/million cells) for 5 min at 4  C. 4. Label the sample with antibodies as shown in Table 1 and incubate on ice for 30 min in the dark or by covering them with aluminum foil. 5. Mix the cells by gently flicking the tubes every 10 min to prevent the cells from settling down and aggregating at the bottom of the tube. 6. Pellet the cells by centrifuging the tubes at 250  g for 5 min at 4  C. 7. Wash the pellet in 1 ml of ice cold solution A. Carefully decant the supernatant. Repeat this wash two times.

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Table 1 Outline of experimental/control samples and antibody dilutions for fluorescence activated cell sorting of DETCs Sl. No.

Sample

Dilution

Minimum number of cells (see Note 2)

Final volume (μl)

1

Unstained

–

1  105

100

1:10

1  10

5

100

2

FITC isotype control

3

PE-Cy7 isotype control 1:50

1  105

100

PE isotype control

1:200

1  10

5

100

7-AAD

0.5 μl

1  10

5

100

Experimental samples

FITC anti-Vγ3 TCR (1:10) 1  10 PE-Cy7 anti-CD3ε (1:50) PE anti-γδ TCR (1:200) 7-AAD (5 μl)

6

300

4 5 6

8. Resuspend the pellet from unstained control, 7-AAD and isotype controls, in solution A and resuspend the pellet from experimental samples in 1 ml of solution A containing 5 μl of 7-AAD (as indicated in Table 1). 9. Keep the FACS tubes on ice until FACS analysis/sorting. 3.1.3 Preparation of DETC Growth Medium

Prepare DETC growth medium as described in [19]. Briefly, RPMI 1640 was supplemented with 10% FBS 0.5 IU/L penicillin, 500 mg/L streptomycin, 1% L-glutamine, 0.63 mM HEPES, 1 mM Na pyruvate, 1 μM nonessential amino acids (Gibco), and 5 U/ml IL-2 (Invitrogen, USA).

3.1.4 Maintenance of DETCs

The 7-17 DETC cell line can be cultured as described in [18, 19]. Primary DETCs can be isolated (Subheading 3.1.2) and cultured using the protocol described [16]. Briefly, both the 7-17 DETC cell line and primary DETC cells can be propagated as suspension cultures on ultra-low attachment culture dishes in RPMI 1640 media with 10% FBS and 20 U/ml rIL-2 at 37  C, 5% CO2. Every other day half of the medium should be gently removed by tilting the dish, letting the cells settle and gently pipetting out the spent medium, followed by replenishing with fresh medium. The cells should be passaged when the culture dish reaches 70% confluency. Collect total cell suspension into a 15 ml tube and centrifuge cells at 300  g for 5 min at 25  C. Gently aspirate the media and resuspend in 2 ml of fresh growth media and add 400 μl to a new 10 cm dish containing 10 ml of fresh growth media.

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3.2 Activation of DETCs

To stimulate the DETCs, plate the cells in a culture dish coated with 0.1 μg/ml anti-CD3 and 10 μg/ml anti-JAML as described in [12]. They can also be activated by adding rIL-1β (10 ng/ml), rIL-23 (10 ng/ml), anti-CD3 (1 μg/ml), or combinations of these, which will affect the level of DETC activation. Treat the DETCs for a period of 24–48 h at 37  C. The activation of DETCs can be monitored by assessing their cytokine expression pattern. Activated DETCs secrete factors into the growth media such as IL-17, FGF7, TNFα, and IFN-γ which can be detected by ELISA after 48 h as described in [18] or by analyzing their levels at the transcript level by RT-PCR after extracting RNA from DETCs after 24 h of treatment [12].

3.3 Coculturing Hair Follicle Stem Cells with DETCs

The coculture technique is advantageous as it can provide an experimental platform to understand the direct interaction between different cell types present in the tissue. Wendy Havran’s laboratory has pioneered the coculture technique of epidermal cells and DETCs [10, 12, 18–20] to investigate the direct interaction between two different cell types. Described below is a protocol to examine the direct interaction between DETCs and hair follicle stem cells. 1. Culture the primary hair follicle stem cells as described in [17]. 2. 7-17 DETC cell line or primary DETCs can be cultured as described in [16]. 3. Activate the DETCs as described in Subheading 3.2 for 24–48 h prior to coculture. 4. Plate 3  104 primary hair follicle stem cells in one well of a 96-well plate and incubate for 24 h at 37  C, 7% CO2. 5. Collect the activated (or inactivated) DETCs in a 15 ml falcon tube and centrifuge at 300  g for 5 min at 25  C. Aspirate the supernatant and resuspend the pellet in fresh E-media. 6. Add 1  105 of activated (or inactivated) DETCs on the cultured hair follicle stem cells (in step 4) and co-incubate at 37  C, 7% CO2 (see Notes 3 and 4). 7. To monitor the effect of DETCs on hair follicle stem cell proliferation, incubate DETCs and hair follicle stem cells together for 24–48 h (see Note 4). Proliferation can be analyzed by counting the number of hair follicle stem cells at various time points such as 24, 48, and 72 h after coculture using trypan blue exclusion assay as described in [12] (see Note 5). Also, the cells can be used for RNA and protein analysis (see Notes 6 and 7).

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3.4 Preparation of Conditioned Media 3.4.1 Media Preparation

3.4.2 Preparation of Conditioned Media from Epidermal Explants

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E media—Prepare media as previously published [17]. P-media for conditioning epidermal explants—P-media is E media without serum and additives. This media contains Gibco Invitrogen customized DMEM:F12 (3:1), sodium bicarbonate, Lglutamine, penicillin–streptomycin solution, and 0.05 mM CaCl2. The following procedure should be carried out under sterile conditions. 1. Sterilize the pups (see Notes 8 and 9) by rinsing them twice with 70% ethanol and autoclaved PBS for 1 min each in a Petri dish. 2. Decapitate the pups and amputate the limbs and tail (see Note 10) as close to the core body as possible (Fig. 1a–c). 3. With a sterile 10 cm dissecting scissor, cut the skin longitudinally from head to tail as shown in Fig. 1d–f. Starting from the dorsal side (the belly skin is much thinner so might tear) roll out the skin while pushing the inner body mass away gently with the help of sterile 10 cm non-toothed dissecting forceps (Fig. 1g–i). 4. Place the skin in a Petri dish, dermis facing up on the bottom of the dish. Spread out the skin using a non-toothed dissecting forceps making sure not to stretch the tissue beyond its normal size. (Fig. 1j). 5. Use forceps to remove any attached fascia (Fig. 1k). Using a scalpel, scrape off the subcutaneous fat loosely attached to the dermis (which endows the tissue with a shiny appearance), until the dermis becomes dull in appearance. Be careful not to damage the architecture of the dermis by inducing tears or scratches while scraping (Fig. 1l). 6. Use biopsy punches (8 mm) (see Note 11) to cut the skin into uniform sized circular biopsies (Fig. 1m, n). Take 2–3 biopsies from each skin. 7. Add 1 ml of dispase (2.5 mg/ml) in a 6-well plate or a 35 mm dish. 8. Place the skin biopsies in the dispase solution dermis side down and make sure that the biopsies are not curled so that the exposure of the dermis to the dispase solution is maximized. Do not stretch the biopsies beyond their normal size. 9. Incubate the skin biopsies for 1–2 h at 37  C. 10. Remove the skin biopsies from the dispase solution and place them into a new 6-well dish so that the epidermis side down. The epidermis should adhere to the bottom of the well (similar set up as Fig. 1j).

Fig. 1 Preparation of epidermal explants from postnatal day 3 pup. Description of the steps has been detailed in the protocol

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11. Use a sterile 10 cm non-toothed dissecting forceps to peel off the dermis. The epidermis will stick to the bottom of the well and appear as a white thin sheet. 12. Dip the epidermis in a 1.5 ml microfuge tube containing 1 ml of 1 PBS. Remove the epidermis and touch it to a kimwipe to remove excess PBS. 13. Use the non-toothed dissecting forceps to place the separated epidermis into a well of a 96-well plate in 200 μl of P-media (described in Subheading 3.4.1) (see Note 12) for 12–16 h at 37  C followed by collection of conditioned medium in appropriately labeled microfuge tubes. 14. Centrifuge the tubes at 300  g for 5 min at 25  C to remove any contaminating cells. Transfer the supernatant into a freshly labeled tube and centrifuge at 2000  g for 5 min to remove cell debris. 50 μl aliquots of the supernatant can be snap frozen in liquid nitrogen and stored at 80  C until further use (see Note 13). 3.5 Conditioned Media Approach to Investigate the Intercellular Communication Between Epidermal Keratinocytes, DETCs, and Hair Follicle Stem Cells Through Soluble Factors (See Fig. 2) 3.5.1 Assaying the Effect of Epidermal Explants on DETCs

1. Prepare epidermal explants from wild type and mutant mouse skins and collect conditioned media as described in Subheading 3.4.2. 2. Maintain DETC culture as described in Subheading 3.1.4 (see Note 14). 3. When the DETCs are 60% confluent, collect the cells in a 15 ml falcon tube and centrifuge at 300  g for 5 min at 25  C. Carefully aspirate the supernatant. 4. Dilute the wild type and mutant epidermal explant conditioned media in fresh RPMI media in 1:3 ratio (explant conditioned media:fresh RPMI media). 5. Resuspend the pellet (from step 3) with diluted conditioned media (from step 4). Plate 3  104 DETCs into a well of a 96-well plate. 6. Incubate the DETCs with 200 μl of the diluted conditioned media (from step 4) at 37  C for 24–48 h. 7. To check the activation of DETCs upon wild type or mutant epidermal explant treatment, their cytokine expression levels can be determined as described briefly in Subheading 3.2. Also, the cells can be processed to score for various cellular assays.

3.5.2 To Investigate the Effect of DETCs on Hair Follicle Stem Cells

1. Treat the DETCs with mutant (or wild type) epidermal explant conditioned media as explained above (Subheading 3.5.1) for 16–24 h at 37  C, 5% CO2. (See Notes 15 and 16.)

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Fig. 2 Investigation of the intercellular communication between epidermal keratinocytes, dendritic epidermal T cells (DETCs), and hair follicle stem cells (HFSCs) through soluble factors

2. Follow a sequential conditioned media:

centrifugation

process

to

collect

(a) Collect the cells along with media in a 15 ml falcon tube and centrifuge at 300  g for 5 min at 25  C. Transfer the supernatant carefully into a fresh falcon tube without disturbing the pellet. (b) Centrifuge the falcon tubes containing the supernatant at 2000  g for 5 min at 25  C to remove the cell debris. 3. Collect the supernatant (conditioned media) and discard the pellet (see Note 17). Use the conditioned media fresh or snap freeze it in liquid nitrogen for later use (see Notes 18 and 19). 4. Dilute the conditioned media in fresh E media in the ratio of 1:5 (conditioned media:E media). 5. Incubate the hair follicle stem cells with the diluted conditioned media (from step 4) at 37  C, 7% CO2 for 24–48 h (see Note 20). 6. To examine the effect of soluble factors from activated DETCs on hair follicle stem cell proliferation, count the number of hair follicle stem cells at various time points such as 24, 48, and 72 h after treating with activated DETCs conditioned media by trypan blue exclusion assay as described in [12]. The cells can also be processed for RNA and protein analysis.

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4

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Notes 1. 1 Penstrep and/or 1 gentamicin must be added to Solution A to maintain sterility of the tissue if the sorted cells will be used for culture. 2. The number of cells to be aliquoted is given only as a suggestion. The optimal number of cells required for FACS analysis depends upon the number of viable cells obtained from the epidermal cell suspension. The cell yield from the skin depends on several factors such as age of the mice, amount of skin collected, efficiency of proteolytic digestion, and efficiency of dissociation of epidermal cells to make the cell suspension. Multiple mice belonging to the same genotype can be taken to increase the total cell yield for experimental samples. 3. The ratio of DETC to hair follicle stem cells in the coculture can be varied and optimized for a given assay. 4. Co-incubation time can be optimized based on the assay requirement depending on whether transcriptional or translational or posttranslational level of changes needs to be determined. 5. Before proceeding with counting the cells to assess hair follicle stem cell proliferation, DETCs from the culture should be removed by collecting the used growth media (DETCs will be present in the media as they grow in suspension culture). Wash the plate once with sterile 1 PBS. Trypsinize the hair follicle stem cells and then proceed with cell counting. The collected DETCs can also be used to score for certain cellular assays. 6. To check for transcript level changes, co-incubate activated/ inactivated DETCs and keratinocytes for 12–24 h. Remove the used growth media and wash the plate gently with sterile PBS for 1 min and collect the hair follicle stem cells for RNA extraction [18]. Since the DETCs are suspended cells, these cells will be removed in the growth medium and can be pelleted and used as required. 7. To check for protein level changes, co-incubate activated/ inactivated DETCs and keratinocytes for 24–48 h. Collect the samples for protein analysis as mentioned in [18, 19]. 8. We have seen that the epidermis can be separated easily from the dermis in pups from an embryonic stage, E17.5 to postnatal day 4. Both back skin and belly skin can be used to make epidermal explants. 9. Epidermal explants can be prepared from skins of wounded, unwounded, and genetically engineered mice with this protocol.

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10. The tail clips from pups can be used for genotyping of genetically engineered animals if required. 11. Using biopsy punches helps in maintaining uniform size of the explants between the samples and ensuring that the conditioned media is prepared from the same volume of the skin. 12. Use the same medium for conditioning in which the subsequent culture experiments will be performed. It is essential that the serum free media is used in order to prevent the effects of serum constituents in future experiments. 13. Aliquoting the supernatant will prevent multiple freeze-thaw cycles of the samples and help maintain the activity of the soluble components. 14. Briefly wash the plates twice with 1 PBS to remove the serum prior to the addition of serum free media. Even trace amounts of serum has the potential to increase basal levels of certain signalling pathways. 15. DETCs can also be activated as described in Subheading 3.2. This can be used as a positive control for the experiment. 16. Incubating cells for 16 h conditioning has been reported [12]. Depending upon the assay and requirement, this conditioning time of the media can be optimized. 17. Do not use the cell pellet obtained in step 3 as the DETCs will not remain active. 18. Make sure to limit the freeze thaw cycles of conditioned media to maintain the activity of soluble factors. 19. Collected conditioned media can be concentrated if a soluble factor is at a very low concentration using centrifugal filter units of desired molecular weight cut off. 20. Amount and time of conditioned media treatment to the cells need to be optimized based on the assay requirement. Conditioned media can be diluted in the fresh growth media and added to the cells. References 1. Blanpain C, Fuchs E (2006) Epidermal stem cells of the skin. Annu Rev Cell Dev Biol 22:339–373 2. Goodell MA, Nguyen H, Shroyer N (2015) Somatic stem cell heterogeneity: diversity in the blood, skin and intestinal stem cell compartments. Nat Rev Mol Cell Biol 16:299–309 3. Gonzales KAU, Fuchs E (2017) Skin and its regenerative powers: an alliance between stem cells and their niche. Dev Cell 43:387–401

4. Lee B, Dai X (2013) Transcriptional control of epidermal stem cells. Adv Exp Med Biol 786:157–173 5. Salmon JK, Armstrong CA, Ansel JC (1994) The skin as an immune organ. WJM J 160:146–152 6. Hsu Y-C, Li L, Fuchs E (2014) Emerging interactions between skin stem cells and their niches. Nat Med 20:847–856

Immune Cell–Stem Cell Interactions 7. Paus R, van der Veen C, Eichmu¨ller S et al (1998) Generation and cyclic remodeling of the hair follicle immune system in mice. J Invest Dermatol 111:7–18 8. Castellana D, Paus R, Perez-Moreno M (2014) Macrophages contribute to the cyclic activation of adult hair follicle stem cells. PLoS Biol 12: e1002002 9. Ali N, Zirak B, Rodriguez RS et al (2017) Regulatory T cells in skin facilitate epithelial stem cell differentiation. Cell 169:1119–1129.e11 10. Jameson J, Ugarte K, Chen N et al (2002) A role for skin gamma delta T cells in wound repair. Science 296:747–749 11. Gay D, Kwon O, Zhang Z et al (2013) Fgf9 from dermal γδ T cells induces hair follicle neogenesis after wounding. Nat Med 19:916–923 12. Lee P, Gund R, Dutta A et al (2017) Stimulation of hair follicle stem cell proliferation through an IL-1 dependent activation of γδTcells. Elife 6:e28875 13. Zhang Y, Harada A, Bluethmann H et al (1995) Tumor necrosis factor (TNF) is a physiologic regulator of hematopoietic progenitor cells: increase of early hematopoietic progenitor cells in TNF receptor p55-deficient mice in vivo and potent inhibition of progenitor

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cell proliferation by TNF alpha in vitro. Blood 86:2930–2937 14. Burzyn D, Kuswanto W, Kolodin D et al (2013) A special population of regulatory T cells potentiates muscle repair. Cell 155:1282–1295 15. Arpaia N, Green JA, Moltedo B et al (2015) A distinct function of regulatory T cells in tissue protection. Cell 162:1078–1089 16. Goodman T, Lefranc¸ois L (1988) Expression of the γ-δ T-cell receptor on intestinal CD8+ intraepithelial lymphocytes. Nature 333:855–858 17. Nowak JA, Fuchs E (2009) Isolation and culture of epithelial stem cells. Methods Mol Biol 482:215–232 18. Nielsen MM, Lovato P, MacLeod AS et al (2014) IL-1β-dependent activation of dendritic epidermal T cells in contact hypersensitivity. J Immunol 192:2975–2983 19. Nielsen MM, Dyring-Andersen B, Schmidt JD et al (2015) NKG2D-dependent activation of dendritic epidermal T cells in contact hypersensitivity. J Invest Dermatol 135:1311–1319 20. Janis EM, Kaufmann SH, Schwartz RH et al (1989) Activation of gamma delta T cells in the primary immune response to Mycobacterium tuberculosis. Science 244:713–716

Methods in Molecular Biology (2019) 1879: 299–305 DOI 10.1007/7651_2018_148 © Springer Science+Business Media New York 2018 Published online: 24 May 2018

Isolating Immune Cells from Mouse Embryonic Skin Ambika S. Kurbet and Srikala Raghavan Abstract Skin is the primary barrier against the external environment and develops a robust immune network for its surveillance. The origin of the resident immune cells of the skin has become a focus of interest over past a decade. Fate mapping studies have revealed that the macrophages home into the skin as early as E12.5 and are derived from the yolk sac and fetal liver. The resident γδT cells are born in the thymus and home to the skin by E16.5. Recent work from our lab has shown that the embryonic macrophages can actively remodel the extracellular matrix in skin suggesting that the skin immune system can be activated long before exposure to foreign antigens. In this chapter, we present a detailed protocol for isolating monocytes, macrophages, and epidermal dendritic T cell populations from embryonic skin. Keywords Embryonic skin, Isolation, Macrophages, Monocytes, T cells, Flow cytometry

1

Introduction The adult skin contains different immune cell types, which include macrophages, dendritic cells, Langerhans cells, mast cells, natural killer cells, CD8, CD4, and γδT cells [1, 2]. In embryonic skin, on the other hand, γδT cells, mast cells, and macrophages (both resident and monocyte derived) form the bulk of the immune cells [3–5]. These resident immune populations are maintained throughout the life and self-renew independently without contribution from the bone marrow in homeostasis [5]. The embryonic macrophages are derived from the major hematopoietic sites, the yolk sac and fetal liver, whereas in adults the bone marrow is the primary site of hematopoiesis [3, 9]. Immune cells can affect stem cell homeostasis and skin regeneration. Recent work has shown that the resident macrophages contribute to reactivation of the hair follicle stem cells by releasing Wnt ligand [6]. Likewise, upon injury, epidermal γδT cell secretes FGF-7, FGF-10, and IGF-1 that are important for survival, proliferation, and migration of epidermal stem cells [7, 8]. These and other data suggest that the crosstalk between immune cells and epithelial cells is crucial for maintaining skin homeostasis and data from our lab suggests that this is set up as early as E16.5 [9–11]. Therefore, isolating embryonic immune

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Propidium Idodide live cell gating SSC-A (⫻1,000)

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cells and identifying their transcriptome and secretome will allow us to interrogate the very earliest events that set up this crosstalk which could lead to the identification of targets for therapeutic intervention. In this chapter, we describe a protocol for isolating leukocytes, macrophages, monocytes, and T cells from E16.5 to E18.5 embryos with milder methods of homogenization to achieve around 90% live cells. We use CD45, F4/80, CD11b, and CD3 antibodies for sorting by flow cytometry to isolate pure populations of cells which can then be used for various applications (see Figs. 1 and 2).

2 2.1

Materials Tissue

2.2 Dissection Instruments 2.3 Reagents and Supplies

Six- to eight-week-old female mice should be used to set up timed pregnancies to recover skin tissue from embryos of the desired age (E16.5, E17.5, and E18.5). Scissors (Sigma, S3146-1EA) and fine forceps (Sigma, Z1687771EA). 1. Dispase: Activity of dispase (Thermo scientific, 17105-041) is between 0.6–2.4 U/ml. It is ideal to prepare a fresh filter sterilized (0.22 μ filter) working stock (1 mg/ml ~ 0.5 U) for every experiment (dispase loses its activity after a week if stored at 4  C). One can also make 10 mg/ml (5 U) stock solutions and store at 20  C for use for up to 2 years. 2. 35 mm dishes (BD Falcon, 353001). 3. 0.25% Trypsin EDTA (Sigma, T4049).

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Fig. 2 Flow cytometric analysis of P2 gated populations: (a) Analysis for CD45 (leukocyte marker), (b) F4/80 (macrophage marker), (c) CD11b (monocyte marker), and (d) CD3 (pan T cell marker). All the plots are accompanied by images of E18.5 whole skin showing the immunofluorescence distribution of the respective immune cell markers (green) with Laminin5 (red) (which marks the basement membrane). Isotype controls for the antibodies used are shown in (e–g). Scale bar 10 μm

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4. Phosphate buffered saline (PBS), pH 7.4. 5. Collagenase IV (Sigma, C5138) working concentration of 0.2 mg/ml in sterile 1 PBS from a stock solution of 1 mg/ml. 6. Dulbecco’s Modified Eagle Medium (DMEM), Sigma D5796, and FBS (Gibco, 16000044). 7. Cell strainers, 40 μm (BD Falcon, 08-771-1). 8. FACS buffer: 2% FBS (Gibco, 16000044) and 0.1% sodium azide (Sigma, 71289) in 1 PBS. 9. FACS tubes: round bottom polystyrene test tube, with snap cap, sterile (Corning, 352054). 10. Propidium iodide (Sigma, P4864). 2.4

3

Antibodies

CD45.1 PE (12-0451-81, eBiosciences), F4/80 FITC (11-480182, eBiosciences), CD11b APC (17-0112-81, eBiosciences), CD3 (14-0032-82, eBiosciences), PE isotype control (12-4031-81, eBiosciences), APC isotype control (17-4321-41, eBiosciences), anti-rat Alexa Fluor 647 (A21247, Invitrogen), and FITC isotype control (11-4031-81, eBiosciences).

Methods This procedure describes the isolation of live (~90% viable) leukocytes, which include macrophages, monocytes, and lymphocytes from embryonic skin. 1. Pregnant females should be anesthetized and sacrificed based on the procedures approved by Institutional Animal Ethics Committee Guidelines (IAEC). Embryos are recovered and placed in sterile 1 PBS to avoid drying of the skin. 2. Add 1 ml of dispase in a 35 mm dish (or 6 well plate) for the skin sample from each embryo. 3. Using sharp, sterile scissors, decapitate the embryos, and cut the tail and limbs (see Note 1). 4. Place the embryos on a sterile tissue paper and cut the skin from the belly (ventral portion) to head region. 5. Using pair of sterile forceps, peel the skin of the embryos gently from the ventral to the dorsal side, making sure not to tear it (see Note 2). 6. Place the skin in dispase and incubate for 1 h at 37  C (see Note 3). 7. After dispase treatment, gently peel the epidermal sheet with a pair of fine forceps. Dispase allows the separation of epidermis from dermis by acting on ECM proteins of basement membrane of the skin. Place the epidermis in 1 ml of 0.25% trypsin

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in a 6 well, or 35 mm dish. Cut it into small pieces with a fine forceps and incubate at 37  C for 10 min. 8. Place the dermis in a dish containing 1 ml collagenase carefully spreading it and incubate at 37  C for 1 h for efficient activity (see Note 4). 9. Using a 2 ml or 5 ml sterile pipette, pipette the epidermal tissue up and down several times. Add 1 ml of media (DMEM with 10% FBS) to stop the action of trypsin. Continue pipetting till a uniform cell suspension is seen. 10. Strain the epidermal cell suspension using a 40 μm cell strainer into a 50 ml falcon tube and spin at 309  g (1500 rpm) for 5 min at 4  C in a eppendorf 5810R centrifuge. Remove the supernatant and resuspend in FACS buffer, and place the falcon tube on ice (see Note 5). 11. After collagenase treatment of dermis is complete, repeat the process detailed in steps 9 and 10 to collect the dermal cell suspension. Mix the epidermal and dermal cell suspension in a 50 ml falcon tube. 12. Centrifuge the cells at 309  g (1500 rpm) for 10 min at 4  C. 13. Resuspend the cell pellet in 1 ml of FACS buffer. Count the total number of cells in 1 ml (see Note 6). 14. Divide the cells into FACS tubes, each containing 2–3 million cells for each antibody. Label the tubes. Set approximately 105 cells aside for unstained and secondary controls. See Table 1. 15. Incubate the samples with primary antibodies and isotype controls (dilutions as described in datasheet) for 1 h on ice (see Note 7). 16. Wash the cells two times with 1 cold PBS, invert it few times, and centrifuge at 309  g (1500 rpm) for 5 min at 4  C. 17. Decant the supernatant carefully (do not use vacuum suction) and resuspend the samples with 1 ml of FACS buffer. Table 1 Sample tubes and corresponding controls used for staining Samples

Controls

1

Unstained

–

2

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6

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3

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7

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4

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8

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9

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18. For unconjugated primary antibodies, add the secondary antibody (1:500 dilution) and incubate for 30 min at 4  C. Keep tapping gently every 5 min for good mixing (see Note 7). 19. After the secondary staining, wash the cells as described in steps 16 and 17. 20. Resuspend the cells with 1 ml of FACS buffer. 21. Add propidium iodide (PI) at a concentration of 1 μg/ml before acquiring samples to gate dead cells out of the population. Dead cells take up PI (see Note 8). 22. We use the BD FACSAria Fusion cytometer, which is equipped with blue (488 nm), red (633 nm), violet (405 nm), yellowgreen (561 nm), and UV lasers. We use the appropriate filters for each antibody tagged with dye (see Note 9) and data is analyzed with BD FACSDiva software which is compatible with the machine. Live cells are used for gating and isolating different immune cell populations. 23. The cells are collected in a tube containing media (DMEM + 10% FBS) (see Note 10).

4

Notes 1. If the limbs and tail are improperly cut, you will face difficulties in taking the intact skin. 2. If you wish to isolate immune cells from E16.5, after isolating whole skin (described in steps 3 and 4), directly place the skin in collagenase for 1 h at 37  C. 3. Place the skin dermis side down. 4. For embryonic skin, 1 h of collagenase is sufficient. If you are working with P0 or P1 pups, it is recommended to leave it for 2–3 h. 5. For increased viability, it is recommended that the cells be maintained at 4  C throughout the procedure. 6. One Whole E18.5 skin will have approximately 15–20 million cells. 7. Keep mixing the sample for uniform staining, as the cells tend to form clumps. 8. One can also use DAPI at 1 μg/ml for gating out dead cells. 9. We use anti-rat 647 as a secondary antibody for CD3. The signal for secondary control and CD3 is acquired in APC filter, as the emission maxima lies in the same range. 10. If you still see debris or any clumps, strain the samples again and use. One can also use FACS tubes with strainer caps BD# 352235.

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Acknowledgments We would like to thank H. Krishnamurthy and the Central Imaging and Flow Facility (CIFF) at NCBS for the use of the confocal microscopes and FACs facility. Animal work was partially supported by the National Mouse Research Resource (NaMoR) grant (BT/PR5981/MED/31/181/2012; 2013-2016) from the DBT. We thank members of the Raghavan lab for feedback. The SR lab is funded through core funds from inStem supported by the Department of Biotechnology (DBT), DBT grant (BT/PR8655/ AGR/36/759/2013), and DST-SERB grant (EMR/2016/ 003199). References 1. Nestle FO, Di Meglio P et al (2009) Skin immune sentinels in health and disease. Nat Rev Immunol 9(10):679–679 2. Heath WR, Carbone FR (2013) The skinresident and migratory immune system in steady state and memory: innate lymphocytes, dendritic cells and T cells. Nat Immunol 14 (10):978–985 3. Jiang X et al (2010) Embryonic trafficking of gammadelta T cells to skin is dependent on E/P selectin ligands and CCR4. Proc Natl Acad Sci U S A 107(16):7443–7448 4. Gurish MF, Austen KF (2012) Developmental origin and functional specialization of mast cell subsets. Immunity 37(1):25–33 5. Hoeffel G et al (2012) Adult Langerhans cells derive predominantly from embryonic fetal liver monocytes with a minor contribution of yolk sac-derived macrophages. J Exp Med 209 (6):1167–1181

6. Castellana D et al (2014) Macrophages contribute to the cyclic activation of adult hair follicle stem cells. PLoS Biol 12(12):e1002002 7. Tay SS et al (2013) The skin-resident immune network. Curr Dermatol Rep 3:13–22 8. Jameson J et al (2002) A role for skin gammadelta T cells in wound repair. Science 296 (5568):747–749 9. Kurbet AS et al (2016) Sterile inflammation enhances ECM degradation in integrin β1 KO embryonic skin. Cell Rep 16(12):3334–3347 10. Sharp LL, Jameson JM, Cauvi G, Havran WL (2005) Dendritic epidermal T cells regulate skin homeostasis through local production of insulin-like growth factor 1. Nat Immunol 6 (1):73–79 11. Epelman S, Lavine KJ, Randolph GJ (2014) Origin and functions of tissue macrophages. Immunity 41(1):21–35

Methods in Molecular Biology (2019) 1879: 307–321 DOI 10.1007/7651_2018_145 © Springer Science+Business Media New York 2018 Published online: 24 May 2018

Direct Conversion of Mouse Embryonic Fibroblasts into Neural Crest Cells Tsutomu Motohashi and Takahiro Kunisada Abstract Neural crest cells (NCCs) are multipotent cells that emerge from the edges of the neural folds and extensively migrate throughout developing embryos. Dorsolaterally migrating NCCs colonize skin, differentiate into skin melanocytes, and lose their multipotency. Multipotent NCCs or NCCs derived multipotent stem cells (MSCs) were recently detected in their migrated locations, including skin, despite restrictions in cell fate acquisition following migration. Since many features of NCCs have yet to be revealed, the novel properties of NCCs represent an important and interesting field in stem cell biology. We previously reported the direct conversion of mouse embryonic fibroblasts (MEFs) into NCCs by the forced expression of the transcription factors C-MYC, KLF4, and SOX10. We herein describe the methods employed for direct conversion: retrovirus infection for the forced expression of transcription factors, a flow cytometry-sorting method for the isolation of converted NCCs, and culture methods for the maintenance and differentiation of the converted NCCs. Keywords C-MYC, Direct conversion, Flow cytometer, KLF-4, Neural crest cells, P75, SOX10

1

Introduction Neural crest cells (NCCs) emerge from the dorsal region of the fusing neural tube and extensively migrate throughout the developing embryo. They differentiate into many cell types, including neurons and glial cells of peripheral sensory and autonomic ganglia, Schwann cells, melanocytes, endocrine cells, smooth muscle, and skeletal and connective tissues of the craniofacial complex; and thus embryonic NCCs undoubtedly are multipotent [1]. With development, NCCs become committed to particular lineages and simultaneously lose their ability to give rise to diverse phenotypes. For example, the trunk NCCs dorsolaterally migrate, colonize skin, and become committed to melanocytes [1–3]. Multipotent neural crest stem cells (NCSCs) or NCC-derived MSCs were recently detected in NCC-derived fetal and adult tissues, including skin, despite restrictions in cell fate acquisition after migration. Multipotent NCSCs were shown to be maintained, even in adult rodent tissues, and have been detected in the adult sciatic nerve, gut, and carotid bodies to which NCCs have migrated

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[4–7]. Previous studies have reported that cells isolated from the adult skin of mice and humans exhibited self-renewal features and differentiated into NCC derivatives such as neurons, glial cells, smooth muscle cells, and melanocytes [8, 9]. Others studies demonstrated that cells in adult hair follicles were also capable of differentiating into derivatives of NCCs [10–12]. Thus, NCCs have many novel properties that have not yet been identified and represent an important and interesting field in stem cell biology. However, studies on NCCs are hampered by the difficulties associated with isolating and manipulating these cells. NCCs emerge as a continuous cell population, progressively disperse, and invade neighboring tissues, which results in challenges with their separation and isolation. NCCs generated from ES cells or directly converted from cultured fibroblasts are alternative sources for NCC studies. NCCs have been generated from mouse and human ES or iPS cells in various methods mimicking the developmental process [13–17], and NCCs have been directly converted from human fibroblasts via the forced expression of SOX10, an NCC specifier [18]. We also previously reported that mouse embryonic fibroblasts (MEFs) were directly converted into NCCs by the expression of SOX10, C-MYC, and KLF4 [19]. Converted NCCs have similar properties to in vivo NCCs; they exhibit differentiation potency into neuronal cells, glial cells, adipocytes and osteocytes, selfrenewal ability, and migration potency [19]. Converted NCCs may be advantageous for NCC research because they are easy to isolate and manipulate in in vitro cultures. In this chapter, we describe methods for the direct conversion of MEFs into NCCs: retrovirus infection for the forced expression of transcription factors, flow cytometric isolation, the maintenance of converted NCCs, and the differentiation into NCCs derivatives. We use Sox10-IRES-Venus mouse fibroblasts for the conversion and isolation of converted NCCs as SOX10-positive cells (Fig. 1a, b). Regarding the use of wild-type fibroblasts, converted NCCs may be isolated as P75-positive cells by using a P75 antibody (Fig. 1c).

2

Materials

2.1 Preparation and Infection of Retrovirus

1. Medium for maintenance of Platinum-E (Plat-E) Retroviral Packaging Cell Line: DMEM (Gibco) supplemented with 10% fetal calf serum (FCS), 1/100 GlutaMax-1 Supplement (Gibco), 50 μg/ml streptomycin, and 50 U/ml penicillin. Store at 4
C and use within 2 months. 2. 0.5 mM EDTA PBS solution.

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Fig. 1 Sox10, c-Myc, and Klf4 convert mouse embryonic fibroblasts (MEFs) into SOX10+ cells. (a) Experimental scheme for the conversion of Sox10-IRES-Venus MEFs into SOX10+ neural crest cells (NCCs). (b) Representative flow cytometry plots of Sox10-IRES-Venus MEF infected with Sox10, c-Myc, and Klf4 after 10 days. The numbers show the average percentages of SOX10+ NCCs (red line gate). PI propidium iodide. (c) Representative flow cytometry plots of wild-type MEF infected with Sox10, c-Myc, and Klf4 after 10 days. The numbers show the average percentages of P75+ NCCs (red line gate)

3. Polyethyleneimine (PEI) solution: Dilute 100 mg polyethyleneimine max (Polysciences, Inc., USA, Mx40,000) with 100 ml of distilled water and sterilize by passage through a ϕ 0.22-μm membrane filter. Storage at 20
C. 4. Polybrene solution: Dilute 80 mg hexadimethrine bromide (Nacalai Tesque, Japan) with 10 ml of distilled water or Caand Mg-free phosphate buffer saline (PBS) and sterilize by passage through a ϕ 0.22-μm membrane filter. Storage at 4
C. 5. Retrovirus vector (such as pMXs-GW, pMYs-GW). 6. OPTI-MEM (Gibco).

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2.2 Culture of Retrovirus-Infected Mouse Embryonic Fibroblasts

1. 0.05% Trypsin/0.5 mM EDTA (Gibco). 2. Medium for maintenance of ST2 stromal cells: RPMI-1640 (Gibco) supplemented with 5% FCS, 50 μM 2-mercaptoethanol, 50 μg/ml streptomycin, and 50 U/ml penicillin. Store at 4
C and use within 2 months. 3. Dexamethasone (Dex; Sigma): Store original stock solution at 102 M in ethanol at 70
C. Prepare the working stock solution at 103 M in ethanol at 4
C and dilute from this stock solution for each use. Stable for at least 1 year (stock at 70
C) or 3 months (stock at 4
C). 4. Human recombinant fibroblast growth factor-2 (bFGF; R&D Systems): Store the stock solution at 200 nM in 0.1% bovine serum albumin (BSA)/PBS at 70
C. Store at 4
C after thawing. Stable for at least for 6 months (stock at 70
C) or 1 month (stock at 4
C). 5. Cholera toxin (CT; Sigma): Store the stock solution at 50 nM in distilled water at 70
C. Store at 4
C after thawing. Stable for at least 1 year (stock at 70
C) or 1 month (stock at 4
C). 6. Human recombinant endothelin-3 (EDN3; Peptide Institute, Inc., Japan): Store the stock solution at 100 μg/ml in 0.1% acetic acid solution at 70
C. Store at 4
C after thawing. Stable for at least 1 year (stock at 70
C) or 1 month (stock at 4
C). 7. Basal medium for conversion: αMEM (Gibco) supplemented with 10% FCS, 107 M Dex, 20 pM bFGF, 10 pM CT, 100 ng/ml EDN3, 50 μg/ml streptomycin, and 50 U/ml penicillin. Store at 4
C and use within 2 months.

2.3 Flow Cytometric Analysis and Isolation of Generated Neural Crest Cells

1. Dispase II solution: Dilute 10 Dispase II (Sanko Junyaku Co., Ltd., Japan) with Ca- and Mg-free PBS and sterilize by passage through a ϕ 0.22-μm membrane filter. 2. Staining medium (SM): PBS containing 3% FCS. 3. Rat anti-mouse Fc gamma receptor (2.4-G2; BD Bioscience). 4. Rabbit-anti mouse P75 (ab8875; Abcam). 5. DyLight 649-conjugated rat anti-rabbit IgG (Biolegend). 6. PI solution: SM containing 3 μg/ml propidium iodide (PI) (Calbiochem). 7. Flow cytometer (such as Becton-Dickinson, FACS Vantage, or FACS Aria).

2.4 Maintenance and Differentiation of Generated Neural Crest Cells

1. Incubator for culture in hypoxic conditions (such as Hitachi and MCO-5M). 2. Poly-D-lysine (PDL) solution: Dissolve poly-D-lysine (Biomedical Technologies Inc., USA) in distilled water to a concentration of 150 μg/ml. Store at 4
C.

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3. Human plasma fibronectin (HPF) solution: Suspend human plasma fibronectin (Gibco) in PBS to a concentration of 150 μg/ml. Store at 20
C until. 4. Medium for maintenance of generated NCCs: 5:3 mixture of DMEM–low (Gibco)—neurobasal medium (Gibco) supplemented with 1% chick embryo extract (CEE, USBiological, USA), 1% N2 (Gibco), 2% B27 (Gibco), 50 μM 2-mercaptoethanol (Sigma), 35 ng/ml all-trans retinoic acid (Sigma), 20 ng/ml IGF-1 (R&D systems), 100 ng/ml EDN3 (Peptide Institute Inc., Japan), and 20 ng/ml bFGF (R&D Systems). Store at 4
C and use within 2 months. 5. Basal medium for differentiation of generated NCCs: the same components as the medium for maintenance of generated NCCs except that it contains 0.1% CEE, and 10 ng/ml bFGF. 6. Medium for neural cell differentiation: basal medium for differentiation of generated NCCs supplemented with 50 ng/ml BMP-2 (R&D Systems). 7. Medium for glial cell differentiation: basal medium for differentiation of generated NCCs supplemented with 1 nM forskolin (Sigma) and 1 nM Nrg-1 (R&D Systems). 8. Adipogenic induction medium (LONZA). 9. Adipogenic maintenance medium (LONZA). 10. Medium for osteocyte differentiation: MSCGM medium (LONZA) supplemented with 0.1 μM Dex (Sigma), 50 μg/ ml ascorbate (Wako Pure Chemical Industries Ltd., Japan), and 0.1% β-glycerophosphate (Sigma). 2.5 Immunohistochemical Analysis of Cultured Neural Crest Cells

1. 4% Paraformaldehyde (PFA) in PBS: pH ¼ 7.0–7.5 (see Note 1). 2. 0.1% Triton-X100 (Gibco) in 0.5% BSA PBS. 3. Blocking solution: 3% goat serum or 5% BSA in PBS. 4. Primary antibodies: anti-mouse neuronal class III β-tubulin (TuJ-1, Covance), anti-mouse glial fibrillary acidic protein (GFAP) (Z0334, DakoCytomation), anti-mouse α smooth muscle actin (1A4, Sigma), anti-mouse peripherin (MAB1527, Chemicon), anti-nestin (Rat401, Chemicon), and anti-S100β (SH-B1, Sigma). 5. Secondary antibodies: Texas-Red conjugated anti-mouse IgG (1:500; Molecular Probes) and Alexa Fluor 488-conjugated anti-rabbit IgG (1:500; Molecular Probes). 6. Hoechst 33258 (Sigma) for nuclei stain. 7. Fluorescence microscope (such as Olympus and IX-71).

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2.6 Oil Red O Staining for Generated Adipocytes

1. 4% PFA in PBS. 2. Oil Red O stock solution: Dissolve 0.5 g of Oil Red O (Wako Pure Chemical Industries Ltd., Japan) in 100 ml isopropyl alcohol. Store at room temperature. 3. Oil Red O stain solution: Dilute 6 ml of Oil Red O stock solution with 4 ml water solution.

2.7 Alkaline Phosphatase and Alizarin Red Stain for Generated Osteocytes

1. Methanol. 2. Alkaline phosphatase (ALP) staining kit (Muto Pure Chemicals Co. Ltd., Japan). 3. 0.028% Ammonium hydroxide water solution. 4. Alizarin Red S (Kanto Chemical Co., Inc., Japan).

3

Methods

3.1 Maintenance of Plat-E Cells

Plat-E cells are a packaging cell line for the production of an ecotropic retrovirus, and may only infect mouse or rat cells [20]. Plat-E cells are maintained in medium for maintenance of Plat-E. 1. Obtain confluent Plat-E cells in a dish or flask and dissociate them with 0.5 mM EDTA at 37
C for 4 min. 2. Add medium, gently pipette, and centrifuge them at 200  g for 4 min, and split cells 1:4 onto 100-mm dishes. 3. Maintain the cells by regularly passing them every 2 or 3 days.

3.2 Maintenance of ST2 Stromal Cells and Preparation of Medium for Conversion

ST2 stromal cells are used to prepare culture supernatants for the direct conversion. ST2 stromal cells are derived from mouse bone marrow stroma cells. ST2 stromal cells are maintained in medium for maintenance of ST2 stromal cells (see Note 2). 1. Obtain confluent ST2 cells in a dish or flask and trypsinize them with 0.05% trypsin solution at 37
C for 4 min. 2. Add medium, dissociate the cells by pipetting, centrifuge them at 200  g for 4 min, and split cells 1:4 onto 100-mm dishes. 3. Maintain cells by regularly passing them every 3 or 4 days. 4. In order to prepare the medium for conversion, discard the culture medium from the confluent ST2 cells on the 100-mm dish, and add the 10–12 ml basal medium for conversion. Two or 3 days later, harvest the culture supernatant, centrifuge them at 1,400  g for 5 min, and use for conversion.

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3.3 Preparation of a Retrovirus and Infection for Direct Conversion

The method for preparation of a retrovirus using Plat-E cells on 6-well plate is described below. Regarding the use of other plate sizes, calculate by the area of the plate.

3.3.1 Preparation of the Retrovirus

Retrovirus vectors are transfected to Plat-E cells by using PEI solution. Prepare the retroviruses supernatant in every experiment. The use of frozen-stock virus solution markedly reduces infection efficiency. 1. Seed Plat-E cells at approximately 1.0–1.5  105 cells/well. Twenty hours later, transfect retrovirus vectors into the Plat-E cells. 2. Mix 50 μl OPTI-MEM and 4.5 μl PEI solution and then vortex. 3. Add 2 μg retrovirus vector to the OPTI-MEM/PEI mixture and vortex (see Note 3). 4. Let stand for 30 min at RT. 5. Drop the mixture onto seeded Plat-E cells and incubate at 37
C. 6. Twenty-four hours later, change the medium to fresh medium for the maintenance of Plat-E. 7. Harvest individual supernatants containing virus at 48 h posttransfection for infection (see Note 4). 8. Centrifuge virus supernatant at 200  g for 5 min, and then at 1,400  g for 5 min to eliminate the Plat-E cells (see Note 5).

3.3.2 Infection with the Retrovirus

In order to increase infection efficiency, use detached MEFs for retrovirus infection. The following is the method for using 6-well plate. Regarding the use of other plate sizes, calculate by the area of the plate. 1. Prepare 1.0–1.5  105 cells/well of detached MEFs. 2. Add polybrene solution to the virus supernatant to a concentration of 4 μg/ml. 3. Mix MEFs with the virus supernatant and seed on 6-well plates. In order to achieve infection with three transcription factors, an equal volume of the retrovirus supernatant is mixed before infection (see Note 6). The start of the virus infection period is termed “day 0” (see Note 7). 4. After 24 h of infection, the medium is changed to the previously prepared medium for conversion. 5. Change the medium to the fresh medium for conversion every 2 or 3 days.

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3.4 Isolation of Converted Neural Crest Cells

Infected MEFs are converted into NCCs at 10 days after the infection with Sox10, c-Myc, and Klf4. We use MEFs derived from Sox10-IRES-Venus mouse embryos for the conversion. In these mice, the VENUS GFP variant gene [21] under the control of the internal ribosomal entry site (IRES) has been placed downstream of the Sox10 stop codon (Fig. 1a). By using the mice, the NCCs generated are isolated as SOX10-positive (VENUS-positive) cells from infected MEFs with flow cytometry (Fig. 1b). In the case of the direct conversion of wild-type MEFs, the generated NCCs may be isolated as P75-positive cells immunostained with anti-P75 antibodies (Fig. 1c). The P75 molecule is an NCC marker. This section describes the methods used to isolate the generated NCCs derived from wild-type MEFs.

3.4.1 Cell Dissociation for Flow Cytometry

1. Ten days after the retrovirus infection, infected MEFs are incubated at 37
C for 10–15 min in Dispase II solution (see Note 8). 2. Gently dissociate the cells by pipetting with a P200 or P1000 pipette. 3. Add 10 volumes of SM to quench the digestion, and then centrifuge the cells at 200  g for 4 min.

3.4.2 Immunostaining of Cell Suspensions for Flow Cytometry

1. Dissociated cells are washed with SM twice. 2. Add the rat anti-mouse Fc gamma receptor and incubate cells on ice for 30–40 min (see Note 9). 3. Wash twice with SM. 4. Add the rabbit anti-mouse P75 (see Notes 10 and 11). 5. Incubate on ice for 30–40 min. 6. Wash the cells twice with SM. 7. Add DyLight 649-conjugated rat anti-rabbit IgG and incubate the cells on ice for 30–40 min (see Note 12). 8. Wash the cells twice with SM. 9. Resuspend the cells in PI solution (see Note 13).

3.4.3 Isolation of Converted Neural Crest Cells by Flow Cytometry and the Culture of the Cells

Generated NCCs are cultured based on the method reported by Morrison et al. [5]. 1. Isolate P75-positive and PI-negative cells (P75+ cells) from the cell suspension by performing flow cytometry (Fig. 1c). 2. Regarding sorted cells culture, directly inoculate the sorted cells into the wells of culture plates by using the flow cytometry system (see Note 14). The wells need to have previously been coated with PDL and HPF solution and have contained the medium for maintenance of generated NCCs (see Notes 15 and 16).

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3. Incubate the sorted cells under hypoxic conditions with 5% O2 and 5% CO2 at 37
C. Medium is changed every 2 days. The day on which cells are sorted is defined as day 0. 3.5 Culture and Immunohistochemical Analysis of Isolated Neural Crest Cells

After 6 days of culture using medium for maintenance of generated NCCs, the sorted cells form colonies or clusters (Fig. 2). Cells are differentiated into NCC derivatives by changing to differentiation media.

3.5.1 Differentiation into Neural, Glial, or Smooth Muscle Cells

1. Change the medium for maintenance of generated NCCs to the basal medium for differentiation of generated NCCs and incubate under hypoxic conditions at 37
C. The day on which the medium is changed is defined as day 0. 2. Change the medium to fresh medium every 2 or 3 days.

Fig. 2 NCCs generated by the forced expression of SOX10/C-MYC/KLF4 formed clusters. Isolated cells generated clusters after a 6-day culture under the medium for maintenance of generated NCCs. Upper photo: lower-power field. Lower 2 photos: higher-power field of “a” and “b” in the upper photo

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3. Approximately 21 days later, neural cells, glial cells, or smooth muscle cells are generated in the colonies (Fig. 3a). 4. Regarding specific neural cell differentiation, change the medium for maintenance of generated NCCs to medium for neural cell differentiation. Approximately 21 days later, neural cells are generated in colonies (Fig. 3b). 5. Regarding specific glial cell differentiation, change the medium for maintenance of generated NCCs to medium for glial cell differentiation. Approximately 21 days later, glial cells are generated in the colonies (Fig. 3c). 6. All differentiation cultures are performed under hypoxic conditions at 37
C.

Fig. 3 Converted NCCs differentiated into neural cells, glial cells, osteocytes, and adipocytes. (a) TuJ1+ neural cells, GFAP+ glial cells, and α smooth muscle actin (αSMA) + smooth muscle cells differentiated from converted NCCs. Nuclei were stained with Hoechst 33258 (Blue). (b, c) In specific differentiation medium, converted NCCs differentiated into peripherin+ and nestin+ neural cells (b); and GFAP+/S100β+ glial cells (c). (d, e) Converted NCCs differentiated into adipocytes with positive Oil Red O staining (d); and osteocytes with alkaline phosphatase (ALP) activity and positive staining with Alizarin Red (e). Scale bar ¼ 200 μm in a, 50 μm in b, c, 100 μm in d, e

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1. Aspirate the culture medium from cultures and wash them three times with PBS. 2. Add 4% PFA in PBS to fix cells (see Note 17). 3. Incubate at room temperature for 15 min. 4. Wash three times with PBS. 5. Add 500 μl of 0.1% Triton-X100 in 0.5% BSA PBS to render cells permeable (see Note 18). 6. Incubate at room temperature for 30 min. 7. Wash three times with PBS. 8. Add a mixture of 3% goat serum or 5% BSA in PBS to block the nonspecific binding of antibodies. 9. Incubate at room temperature for at least 30 min. 10. Wash three times with PBS. 11. Dilute the primary antibody in 0.5% BSA PBS (see Note 11), and then add it to cells at room temperature (see Note 19). 12. Wash three times with PBS. 13. Dilute the secondary antibody in 0.5% BSA PBS (see Note 11), and then add it to react with the primary antibody. 14. Incubate at room temperature for at least 30 min. 15. Wash stained cells three times with PBS. 16. Add Hoechst 33258 to stain nuclei. 17. Incubate at room temperature for 1–3 min (see Note 20). 18. Wash stained cells three times with PBS, and examine the colonies under the fluorescence microscope (Fig. 3a–c) (see Note 21).

3.5.3 Differentiation into Adipocytes

1. Change the medium for maintenance of generated NCCs to adipogenic induction medium. 2. The medium is changed to adipogenic maintenance medium after 3 days. 3. The medium is changed to adipogenic induction medium after 3 days. 4. Four cycles of induction/maintenance are repeated. After the culture, adipocytes are generated in colonies (Fig. 3d). 5. All differentiation cultures are performed under hypoxic conditions at 37
C.

3.5.4 Oil Red O Stain of Generated Adipocytes

1. Aspirate the culture medium from the cultures and wash once with PBS. 2. Add 4% PFA in PBS to fix cells (see Note 17). 3. Incubate at room temperature for 15 min.

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4. Wash once with PBS. 5. Add Oil Red O stain solution (see Note 22). 6. Incubate at room temperature for 30 min. 7. Wash twice with distilled water and examine the colonies under the microscope (Fig. 3d). 3.5.5 Differentiation into Osteocytes

1. Change the medium for maintenance of generated NCCs to medium for osteocyte differentiation. The day on which the medium is changed is defined as day 0. 2. The medium is changed to the fresh medium every 2–3 days. 3. Approximately 21 days later, osteocytes are generated in the colonies (Fig. 3e). 4. Differentiation cultures are performed under hypoxic conditions at 37
C.

3.5.6 Alkaline Phosphatase Staining of Generated Osteocytes

Use the ALP staining kit (Muto Pure Chemicals Co. Ltd., Japan). 1. Dilute 1 ml of fix-preparation solution with 9 ml of methanol and keep it at 4
C (Fixation solution). 2. Dilute the Fast Blue RR salt with 10 ml of Substrate stock solution (Substrate solution). 3. Aspirate the culture medium from the cultures and wash once with PBS. 4. Add the fixation solution to fix the cells (see Note 23). 5. After 10 s, aspirate the fixation solution from the cultures. 6. Add the substrate solution (see Note 23). 7. Incubate at room temperature for 0.5-1H. 8. Wash three times with distilled water and examine colonies under the microscope (Fig. 3e).

3.5.7 Alizarin Red Staining of Generated Osteocytes

1. Dissolve 1 g Alizarin Red S in 100 ml distilled water (Solution A). 2. Adjust solution A to pH 6.0 using 0.028% ammonium hydroxide water solution (Solution B). 3. Aspirate the culture medium from cultures and wash them three times with PBS. 4. Add the ice-cold methanol to fix the cells at 4
C for 20 min. 5. Aspirate ice-cold methanol from the cultures and wash them three times with distilled water. 6. Add solution B and incubate at room temperature for 5 min. 7. Wash three times with distilled water and examine the colonies under the microscope (Fig. 3e).

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Notes 1. Do not make a stock solution. Prepare the fresh solution when needed. 2. ST2 cultures sometimes exhibit a change in appearance (i.e., change to a more dendritic shape or senescent appearance) when cultured for long periods. Discard these cultures and use freshly thawed ST2 cells from the frozen stock. 3. We use the retrovirus vectors pMYs-EGFP, pMYs-Sox10, pMYs-Klf4, and pMYs-cMyc for fibroblast conversion. These retrovirus vectors may be purchased from the human proteome expression resource (HuPEX) library (HuPEX clones in HGPD, http://www.HGPD.jp/, the National Institute of Advanced Industrial Science and Technology, Japan). 4. Use appropriate controls in order to verify gene transfer efficiency. We use retrovirus vector-inserted EGFP as a transfection control in every experiment. Gene transfer efficiency is typically 60%. High gene transfer efficiency is necessary for direct conversion. 5. Plat-E cells may also be eliminated by passage through a ϕ 0.45-μm membrane filter. 6. For example, mix 600 μl of each retrovirus supernatant for the infection of Sox10-, cMyc-, and Klf4-retroviruses when using 6-well plates. 7. Use appropriate controls in order to verify infection efficiency. We use the EGFP retrovirus as a control infection in every experiment. Infection efficiency is typically 40–60% after 3–4 days. 8. We use 500 μl/well of Dispase II solution for dissociation of MEFs seeded on 6-well plates. 9. Antibodies against the Fc gamma receptor block the nonspecific cell-surface binding of antibodies. Add an appropriate volume of the diluted antibody solution according to the supplier’s recommendations. 10. Keep a small amount of the sample without the addition of antibodies. This sample is used as a negative control in the flow cytometric analysis. 11. Add an appropriate volume of the diluted antibody solution according to the supplier’s recommendations. 12. Other fluorescein dye-conjugated anti-rabbit IgG may also be used. 13. Only dead cells are stained by PI.

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14. One hundred to five hundred and single sorted cells are introduced into 6-well and 96-well plates, respectively. 15. The methods used to coat plates with PDL and HPF solution are as follows: (a) PDL solution is applied to tissue culture plates and immediately withdrawn. (b) The plates are allowed to dry at room temperature, rinsed with distilled water, and allowed to dry again. (c) HPF solution is then applied to the dishes and immediately withdrawn. 16. Medium volumes are 2 ml/well and 200 μl/well in 6-well and 96-well plates, respectively. 17. Fixative volumes are 1 ml/well and 200 μl/well of 4% PFA for 6-well and 96-well plates, respectively. 18. 0.1% Triton-X volumes are 500 μl/well and 200 μl/well for 6-well and 96-well plates, respectively. 19. Incubate at room temperature for more than 30 min or at 4
C overnight. 20. An incubation for 1–3 min is sufficient for nuclear staining. 21. We use an Olympus IX-71 fluorescence microscope. 22. The volume of Oil Red O stain solution is 500 μl/well for 6-well plates. 23. The volumes of fixation solution and substrate solution are 1 ml/well for 6-well plates.

Acknowledgements This study was supported by the Gifu University Graduate School of Medicine Research Grant Program, by a research grant from the Japan Science and Technology Agency CREST, and by a grant from the program Grants-in-Aid for Scientific Research (C) from the Japan Society for Promotion for Science. A part of the work was also supported by the Intramural Research Program of the National Institutes of Health, National Institute on Aging, USA. References 1. Le Douarin NM, Kalcheim C (1999) The neural crest, 2nd edn. Cambridge University Press, Cambridge 2. Luo R, Gao J, Wehrle-Haller B et al (2003) Molecular identification of distinct neurogenic and melanogenic neural crest sublineages. Development 130:321–330

3. Wilson YM, Richards KL, Ford-Perriss ML et al (2004) Neural crest cell lineage segregation in the mouse neural tube. Development 131:6153–6162 4. Stemple DL, Anderson DJ (1992) Isolation of a stem cell for neurons and glia from the mammalian neural crest. Cell 71:973–985

Direct Conversion of Mouse Embryonic Fibroblasts into Neural Crest Cells 5. Morrison SJ, White PM, Zock C et al (1999) Prospective identification, isolation by flow cytometry, and in vivo self-renewal of multipotent mammalian neural crest stem cells. Cell 96:737–749 6. Kruger GM, Mosher JT, Bixby S et al (2002) Neural crest stem cells persist in the adult gut but undergo changes in self-renewal, neuronal subtype potential, and factor responsiveness. Neuron 35:657–669 7. Pardal R, Ortega-Saenz P, Duran R et al (2007) Glia-like stem cells sustain physiologic neurogenesis in the adult mammalian carotid body. Cell 131:364–377 8. Fernandes KJ, McKenzie IA, Mill P et al (2004) A dermal niche for multipotent adult skin-derived precursor cells. Nat Cell Biol 6:1082–1093 9. Toma JG, McKenzie IA, Bagli D et al (2005) Isolation and characterization of multipotent skin-derived precursors from human skin. Stem Cells 23:727–737 10. Sieber-Blum M, Grim M, Hu YF et al (2004) Pluripotent neural crest stem cells in the adult hair follicle. Dev Dyn 231:258–269 11. Amoh Y, Li L, Katsuoka K et al (2005) Multipotent nestin-positive, keratin-negative hairfollicle bulge stem cells can form neurons. Proc Natl Acad Sci U S A 102:5530–5534 12. Yu H, Fang D, Kumar SM et al (2006) Isolation of a novel population of multipotent adult stem cells from human hair follicles. Am J Pathol 168:1879–1888 13. Mizuseki K, Sakamoto T, Watanabe K et al (2003) Generation of neural crest-derived peripheral neurons and floor plate cells from mouse and primate embryonic stem

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cells. Proc Natl Acad Sci U S A 100:5828–5833 14. Lee G, Kim H, Elkabetz Y et al (2007) Isolation and directed differentiation of neural crest stem cells derived from human embryonic stem cells. Nat Biotechnol 25:1468–1475 15. Motohashi T, Aoki H, Chiba K et al (2007) Multipotent cell fate of neural crest-like cells derived from embryonic stem cells. Stem Cells 25:402–410 16. Pomp O, Brokhman I, Ziegler L et al (2008) PA6-induced human embryonic stem cellderived neurospheres: a new source of human peripheral sensory neurons and neural crest cells. Brain Res 1230:50–60 17. Lee G, Papapetrou EP, Kim H et al (2009) Modelling pathogenesis and treatment of familial dysautonomia using patient-specific iPSCs. Nature 461:402–406 18. Kim YJ, Lim H, Li Z et al (2014) Generation of multipotent induced neural crest by direct reprogramming of human postnatal fibroblasts with a single transcription factor. Cell Stem Cell 15:497–506 19. Motohashi T, Watanabe N, Nishioka M et al (2016) Gene array analysis of neural crest cells identifies transcription factors necessary for direct conversion of embryonic fibroblasts into neural crest cells. Biol Open 5:311–322 20. Morita S, Konjima T, Kitamura T (2000) PlatE: an efficient and stable system for transient packaging of retroviruses. Gene Ther 7:1063–1066 21. Nagai T, Ibata K, Park ES et al (2002) A variant of yellow fluorescent protein with fast and efficient maturation for cell-biological applications. Nat Biotechnol 20:87–90

Methods in Molecular Biology (2019) 1879: 323–345 DOI 10.1007/7651_2018_150 © Springer Science+Business Media New York 2018 Published online: 24 May 2018

High-Titer Production of HIV-Based Lentiviral Vectors in Roller Bottles for Gene and Cell Therapy Hazal Banu Olgun, Hale M. Tasyurek, Ahter Dilsad Sanlioglu, and Salih Sanlioglu Abstract Lentiviral vectors are becoming preferred vectors of choice for clinical gene therapy trials due to their safety, efficacy, and the long-term gene expression they provide. Although the efficacy of lentiviral vectors is mainly predetermined by the therapeutic genes they carry, they must be produced at high titers to exert therapeutic benefit for in vivo applications. Thus, there is need for practical, robust, and scalable viral vector production methods applicable to any laboratory setting. Here, we describe a practical lentiviral production technique in roller bottles yielding high-titer third-generation lentiviral vectors useful for in vivo gene transfer applications. CaPO4-mediated transient transfection protocol involving the use of a transfer vector and three different packaging plasmids is employed to generate lentivectors in roller bottles. Following clearance of cellular debris via low-speed centrifugation and filtration, virus is concentrated by high-speed ultracentrifugation over sucrose cushion. Keywords Gene therapy, Lentivirus, Roller bottles

1

Introduction Among the numerous viral vectors that have been tested in clinical gene therapy studies so far, HIV-based lentiviral vectors (LVs) stand out particularly due to the long-term transgene expression they provide. LVs have gained widespread use in recent years owing to their favorable features as effective gene transfer vehicles particularly for in vivo applications. The progressively increasing use of LVs in gene transfer studies has demonstrated the necessity to develop methods that will allow high-titer virus production. Scalable, effective, and robust production methods along with highyield purification steps are critical in this regard. Thus, we aimed to improve the current approaches for high-quality production of LVs in high concentrations in roller bottles, for use in experimental applications.

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Lentiviral Vectors

Lentiviruses are enveloped RNA viruses of the Retroviridae (Retrovirus) family. The most studied lentivirus is the human immunodeficiency virus type 1 (HIV-1), which the best characterized lentiviral vectors are derived from. These viruses enter the cell via membrane fusion, after which the positive sense RNA genome is converted into double-stranded DNA by the viral reverse transcriptase to form the proviral DNA. The proviral DNA is then carried into the nucleus to be integrated into the host cell genome by the viral integrase (IN) enzyme [1]. This integration step in the virus’ natural life cycle is crucial in providing long-term stable gene expression. Following integration, transcription directed by the LTR (Long Terminal Repeat) regions on the terminal portions of the viral genome takes place and the host cell initiates production of both the lentiviral RNA genome and the lentiviral proteins. Viruses produced in this way are released out of the cell via budding, to infect new host cells [2, 3]. Lentiviruses have the capacity to carry up to 9-kb-long genetic material and can transduce dividing and nondividing cells. Various gene therapy strategies utilize these features of lentiviruses to deliver therapeutic sequences to target cells [4, 5]. Lentiviral genome consists of cis- and trans-acting components. Important cis elements are: Long Terminal Repeats (LTRs), Rev-Responsive Element (RRE), and the packaging signal (Ψ) [1, 3]. LTR sequences reside in the 50 and 30 ends of the genome and act as promoters for transcription. Production of the structural and enzymatic proteins occurs under the control of the Rev regulatory protein, which binds to the RRE sequence located within the env gene. The posttranscriptional effects provided by Rev include inhibition of viral RNA splicing, stimulation of the nuclear export of the unspliced and incompletely spliced viral mRNAs, and enhancement of the translation of the RRE-containing RNAs [6]. The signal sequence Ψ function in recognition of the viral genome to be inserted into the capsid [7, 8]. The trans-acting components of the LV genome include nine open reading frames (ORFs) that code for 15 proteins. Three of these contain conserved genes that encode Gag, Pol, and Env, that are common in all retroviruses as they are required for viral replication. These polyproteins undergo posttranslational cleavage to generate the structural and enzymatic proteins that are essential for the viral life cycle. The outer envelope protein Env is cleaved to yield the surface (SU; gp120) and the transmembrane (TM; gp41) structural proteins. Matrix (MA), capsid (CA), nucleocapsid (NC), and p6 proteins, on the other hand, are the cleavage products of the Gag polyprotein and form the virion core. Pol also undergoes cleavage to produce three vitally important enzymes: protease (PR), reverse transcriptase (RT), and integrase (IN). These enzymes function in proteolytic processing of the viral precursor polyproteins, conversion of the viral RNA genome into DNA, and integration of this DNA molecule into the host cell genome, respectively [9, 10].

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High amounts of full-length transcripts are produced from the integrated proviral genome via the action of the trans-activator protein Tat, which stimulates transcriptional elongation. Tat specifically associates with its cellular cofactor TAK, which is composed of CDK9 and cyclin T, and hyperphosphorylates the carboxyterminal domain of the cellular RNA polymerase II (RNA Pol II) large subunit [11]. This phosphorylation increases RNA Pol II activity and enhances viral transcription to a large extent [12, 13]. The Rev protein, on the other hand, binds to the RRE sequence to provide the nuclear export of the intron-containing viral transcripts into the cytoplasm [14]. This step is essential in the viral life cycle, as unspliced transcripts are needed to be packaged for production of new infectious viral particles. The “accessory proteins” encoded by the remaining genes (Vif, Nef, Vpu, Vpr, and/or Vpx) are not critical for in vitro replication of the virus, as shown in cell culture systems [15]. For LVs to be used as effective gene therapy vectors, the transgene carried by the vector should be integrated into the target cell’s genome following infection, and provide transcription and translation of the therapeutic transgene only. Furthermore, biosafety of these vectors should be ensured by rendering them replication-deficient unlike the wild-type virus. Thus, the basic principle for production of biosafe LVs is elimination of the replication competence and prevention of reestablishment of this ability by the virus; so the viral genome should be modified accordingly [16]. Naldini et al. have developed a strategy in 1996 where the lentiviral vector components that are required for viral replication were splitted into three separate plasmids to be expressed during transient co-transfection of producer 293T cells [17]. Vectors produced via this three-plasmid system (packaging, transfer, and envelope) that minimizes the production of replication competent lentiviruses (RCLs) are defined as first-generation LVs [18]. The transfer plasmid contains the transgene, the promoter sequence required for the transcription of the transgene (CMV), the packaging signal (Ψ), and the LTR sequences which function in both conversion of the RNA genome into DNA and integration of the viral genome. All other trans-factors required for vector production are included in the packaging plasmid. Accidental packaging of the sequences that would lead to RCL formation is inhibited by deletion of the Ψ sequence. Furthermore, the env gene encoding the envelope proteins of the virus is also removed, and a third plasmid coding a heterologous envelope, the vesicular stomatitis virus glycoprotein (VSV-G), is used for pseudotyping the newly generated particles [17]. Use of VSV-G instead of lentiviral gp160 confers high stability to the viral particles and increases resistance to mechanical force. Additionally, VSV-G pseudotyping provides broad tropism over a range of target cells.

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Although first-generation LVs contributed a great deal to the overall level of biosafety, recombination risk of the plasmids with each other or with other viruses to generate RCLs could not be eliminated completely. Subsequent studies aimed minimization of the recombination risk associated with the first-generation LV production. Thus, second-generation “multiply attenuated” LVs were produced via removal of the sequences encoding the accessory genes (vif, vpr, vpu, and nef) that were included in the packaging plasmids of the first-generation LVs. Products of these genes are essential for the viral life cycle, yet not required for vector production, thus their removal did not impair vector yield or transduction efficiency [19]. Although a minimal risk of RCL formation still existed after these modifications, any such viruses would be devoid of the virulence factors [18]. By removal of the tat and rev genes that were included in the packaging plasmid used in the secondgeneration vector production, third-generation LVs were produced, to even further eliminate the RCL formation risk. However, Tat being an essential protein for lentiviral replication, the thirdgeneration vectors had to carry out Tat-independent transcription. This problem was solved by establishment of chimeric LTR regions with Tat-binding sequences removed, through which efficient lentiviral transduction could still be achieved [20]. Furthermore, introducing the essential rev gene to the producer cell line on a different plasmid also decreased risk of recombination and RCL formation. Further modifications to improve vector performance and biosafety in third-generation LVs include a 133-bp deletion introduced into the 30 LTR region of the viral genome, comprising also the TATA box and the Sp1 and NFkB binding sites. This deletion is naturally transferred to the 50 LTR site following reverse transcription and results in transcriptional inactivation of the LTR in the integrated provirus, effectively inhibiting viral RNA genome formation. Vectors modified in this way are defined as selfinactivating (SIN) vectors [21, 22]. The transfer plasmid in the third-generation LVs is also designed to contain the woodchuck hepatitis virus posttranscriptional regulatory element (WPRE) sequence, which increases the efficiency of transgene expression several folds by a posttranscriptional regulatory effect [23]. Through the above-listed modifications, the idea of using LVs as gene transfer vectors turned into a successful generation of a vector framework that has high efficacy and safety, and also ability to transduce both dividing and nondividing cells. 1.2 Lentiviral Vector Production

Human embryonic kidney (HEK) 293 cells and its derivatives are frequently used in production of the third-generation LVs by co-transfection of four plasmids [17, 20, 24, 25]. Besides a high transfection success in vector production, these human-based packaging cell lines also provide human-type glycosylation patterns on

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the Env proteins. This is a significant issue to be considered, especially if the vector is to be used in in vivo applications [26]. In fact, vectors with nonhuman glycosylation patterns are known to be targeted by the human complement system rapidly, within 20 min following application. The most preferred variant of the 293 cells in LV production is the simian virus 40 (SV40) T antigen-expressing HEK293T cells, which are proved to be more efficient cell lines for vector production with increased cell growth and transfection efficiency [25]. SV40 replication origin (SV40ori) in the plasmid backbones is defined as essential for plasmid replication [27]. This context enhances the nuclear import of expression vectors, thus increasing plasmids available for transcription [28]. Besides transient co-transfection of several plasmids to producer cell lines (Fig. 1), an alternative current strategy in LV production involves the use of stable, inducible packaging cell lines that express all lentiviral vector components except for the transfer vector [29]. Among the advantages of the transient gene expression approach compared to the stable packaging cell lines are its flexibility and overall process time [30]. It is an easily applicable method, and various different transient transfection methods have been developed, as it allows modification of different parameters (see Note 1). The protocol described in this chapter is a thirdgeneration LV vector production method that can be utilized as an intermediate step particularly in transition to large-scale production and summarizes the methodological algorithm to be followed in optimization of LV production, via evaluation of many different parameters suitable for optimization.

2

Materials l

293T cell line (ATCC CRL-3216)

l

FBS (fetal bovine serum) (Biochrom, 50115)

l

DMEM (Dulbecco’s modified Eagle’s medium) (SigmaAldrich, D5648)

l

IMDM (Iscove’s modified Dulbecco’s medium), (Sigma Aldrich, I7633)

l

Opti-MEM (Gibco, 26600134)

l

Chloroquine (Sigma, C6628)

l

Petri dishes, 150 mm (CELLSTAR, Greiner)

l

Roller bottles (CELLMASTER, Greiner, Ribbed surface)

l

pMDLg/pRRE (HIV-1 pGag-Pol, Addgene 12251)

l

Rev plasmid (pRSV-Rev, Addgene 12253)

l

pMD2.G (pVSV-G, Addgene 12259)

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LV-RFP plasmid (Addgene, 26001)

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Fig. 1 Transient transfection method for LV vector production. In transient transfection, the transfer vector, packaging, and envelope-coding elements are introduced into the 293T cells via a transfection agent such as CaPO4. Cells produce the vectors in the following few days after transfection. Vectors released from the cells at the end of the process are isolated from the cell supernatant

2.1

Recipes

l

1 PBS (phosphate buffered saline): 137 mM NaCl 2.7 mM KCl 4.3 mM Na2HPO4 1.47 mM KH2PO4 Dissolve the reagents listed above in 800 ml dH2O Adjust the pH to 7.4 Add distilled water to a total volume of 1 L Sterilize solution by autoclaving at 121  C for 15 min on liquid cycle

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Store at +4  C l

DMEM: One bottle of powder DMEM (high glucose), Sigma D5648 Dissolve in 800 ml dH2O Add 3.7 g NaHCO3 Add distilled water to a total volume of 1 L dH2O Sterilize solution through a 0.22-μm bottle-top filter Add 10% (v/v) FBS, 1% (v/v) Na-pyruvate, and 1% (v/v) pen-strep to 1 L DMEM under aseptic conditions in a Class II Laminar Flow Cabin Store at +4  C

l

Opti-MEM: 13.59 g Opti-MEM Gibco 26600134 2.4 g NaHCO3 Dissolve in 800 ml dH2O Adjust the pH to 7.3 Add distilled water to a total volume of 1 L Sterilize solution through a 0.22-μm bottle-top filter Store at +4  C Add 10% (v/v) FBS and 1% (v/v) pen-strep to 1 L Opti-MEM under aseptic conditions in a Class II Laminar Flow Cabin prior to use

l

IMDM: IMDM, powder, Sigma I7633 Dissolve in 800 ml dH2O Adjust the pH to 7.2 Add distilled water to a total volume of 1 L Sterilize solution through 0.22 μm bottle-top filter Store at +4  C Add 10% (v/v) FBS, 1% (v/v) pen-strep, and 25 μM chloroquine to 1 L IMDM under aseptic conditions in a Class II Laminar Flow Cabin prior to use

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2 HBS (HEPES-buffered solution): 280 mM NaCl 10 mM KCl 1.5 mM Na2HPO4 (anhydrous) 50 mM HEPES 12 mM Glucose

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Dissolve in 800 ml dH2O Adjust the pH to 7.05 Add distilled water to a total volume of 1 L Sterilize solution by 0.22 μm bottle-top filter Aliquot solution and store at 20  C l

2 M CaCl2: 2 M CaCl2 (anhydrous) Dissolve in double distilled water (ddH2O) Sterilize solution through 0.22 μm bottle-top filter Aliquot solution and store at 20  C

l

1 Tris–EDTA (TE): 10 mM TRIS, pH 8.0 1 mM EDTA, pH 8.0 Dissolve in dH2O

l

0.1 Tris–EDTA (TE): dH2O solution: Dilute 1 TE tenfold to prepare 0.1 TE. Mix 2 volumes of 0.1 TE solution and 1 volume of dH2O. Sterilize solution through 0.22 μm bottle-top filter. Store at +4  C.

l

HBSS (Hanks’ balanced salt solution): Buffer 1: Dissolve 8 g NaCl, 0.4 g KCl, and 1 g glucose in 100 ml dH2O. Buffer 2: Dissolve 0.358 g Na2HPO4 (anhydrous) and 0.6 g KH2PO4, in 100 ml dH2O. Buffer 3: Dissolve 0.73 g CaCl2 in 50 ml dH2O. Buffer 4: Dissolve 1.23 g MgSO4·7H2O in 50 ml dH2O. Buffer 5: Dissolve 0.35 g NaHCO3 in 10 ml dH2O. PREMIX buffer: Mix 10 ml #1, 1 ml #2, 1 ml #3, 1 ml #4, and 86 ml dH2O. Add 0.1 ml #5 solution to 9.9 ml PREMIX. Sterilize solution by 0.22 μm bottle-top filter and store at +4  C.

2.2

Instruments

l

Thermo HeraCell240i CO2

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Beckman Coulter, Optima L-90K, 365670

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Beckman SW28, 342204

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Multiskan Spectrum Spectrophotometer

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Thermo Class II Laminar Flow Cabin

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Thermo Multi RF Centrifuge

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Methods Transfection via calcium phosphate (CaPO4) precipitation is the most widely used method for production of LVs, as a cost-effective, readily applicable, and reproducible method with easily obtained components [31–33]. Nevertheless, the transfection efficiency is directly associated with parameters such as DNA–CaPO4 precipitation, amount of plasmid DNA, calcium and phosphate concentrations, temperature, and duration of the procedure [34]. Thus, various factors should be taken into consideration for successful optimization of the process (see Note 5). Precipitate size substantially affects the success of transfection, where small precipitate volumes lead to higher transfection efficiency. As increased incubation times during transfection, on the other hand, will lead to larger precipitates, shorter incubation times should be preferred (see Note 4) [34]. Another variable affecting the precipitate size is the technique used in preparation of the transfection mixture. Two different methods known as the bubble and vortex techniques were tested to define the more efficient method in our protocol, taking into consideration that the amount of precipitate affects transfection efficiency in both cases. Another factor affecting the transfection yield is the plasmid DNA amounts to be introduced into the producer cell line. Many studies that aimed to define the optimum plasmid amounts for successful transfection in the presence of multiple plasmids conclude that exceeding amounts of transfer plasmid introduced compared to the packaging and VSV-G plasmids result in much higher production yields [27, 35]. Yet the optimum amounts change substantially depending on the structure of the vector used. After extensive literature search, plasmid amounts to be used for transfection into 80–90% confluent cells in a 150-mm petri dish were decided as: 14 μg for Gag/Pol; 6 μg for Rev; 7.5 μg for VSV-G; and 22.5 μg for the transfer vector (RFP) (see Note 2). For production of high-titer LVs, it is essential to maintain viability of the cell line and stability of the changing physicochemical conditions during the transfection procedure (see Note 3). Basic transfection methods are not sufficient particularly for large-scale processes, thus additional agents are required for increased efficiency. Most preferred of these are sodium butyrate, chloroquine, cholesterol, and lipids [35–37]. Chloroquine is an amine that raises endosomal and lysosomal pH levels; the increase in lysosomal pH in turn is believed to prevent degradation of the transfected DNA (see Note 6) [38, 39]. However, an optimized concentration and duration of exposure should be established to avoid its concentration-, time- and cell type-dependent toxic effects [39]. Two different chloroquine concentrations were used in this study, as 25 and 40 μM [40] (see Note 8).

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Production yield was evaluated by RFP fluorescence. Viral particle numbers following ultracentrifugation were calculated in terms of the integrated copy number via quantitative PCR. Following all optimization studies, 1011 transduction units (TU) of lentiviral vector production were established in each roller bottle, and similar concentrations of virus could be obtained from each roller bottle via application of the optimized protocol [41, 42]. 3.1 CaPO4 Transfection Method

CaPO4 transfection may be done via the bubble technique or the vortex method:

3.1.1 Bubble Technique

1. Isolate and determine the concentrations of Gag/Pol, Rev, VSV-G, and transfer plasmids (see Note 7). 2. Calculate the required amounts of plasmids. 3. Add required volumes of plasmids into a Falcon tube. 4. Add 0.1 TE:dH2O solution into the plasmid mixture to a total volume of 875 μl. 5. Add 125 μl 2 M CaCl2, to obtain a final concentration of 0.25 M in a total volume of 1 ml. 6. Add 1 ml 2 HBS solution in a separate Falcon tube. 7. Create air bubbles in HBS solution by the help of a pipette controller. 8. Meanwhile, add the DNA–CaCl2 mixture dropwise. 9. Incubate the final mixture for 5 min at room temperature.

3.1.2 Vortex Method

1. Isolate and determine the concentrations of Gag/Pol, Rev, VSV-G, and transfer plasmids (see Note 7). 2. Calculate the required amounts of plasmids. 3. Add required volumes of plasmids into a Falcon tube. 4. Add 0.1 TE:dH2O solution into the plasmid mixture to a total volume of 875 μl. 5. Add 125 μl 2 M CaCl2, to obtain a final concentration of 0.25 M in a total volume of 1 ml. 6. Add 1 ml 2 HBS solution in a separate Falcon tube. 7. Adjust vortex speed to medium level. 8. Create a constant vortex in 2 HBS solution. 9. Meanwhile, add the DNA–CaCl2 mixture dropwise. 10. Incubate the final mixture for 5 min at room temperature.

3.2 Lentivirus Production

1. Dissolve 293T cells in 10% FBS-containing DMEM. 2. Passage 293T cells 2 prior to transfection.

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3. Add transfection media to a 150-cm2-surface area petri dish at 80–90% of cell confluence ratio. 4. Add transfection cocktails (prepared by either the bubble or the vortex technique) dropwise, following addition of the transfection media. 5. Incubate transfected cells for 4 days at 37  C in a CO2 incubator. 6. Observe RFP signals at 24, 48, 72, and 96 h. 3.2.1 Optimization of the Production Conditions

Besides the particle sizes, amount of medium and FBS are also known to affect the transfection yield in LV production. Thus, the optimum production conditions should be set accordingly. In this protocol, different media were used for transfection and collection purposes, and optimization studies were performed under three different conditions (Table 1). RFP plasmid was used as transfer vector in all trials, for the follow-up of the efficiencies of the procedures tested. At the end of the protocols, use of 10% FBS-containing medium along with the bubble technique appeared as the most efficient method (Fig. 2). Further optimization studies under eight different conditions were also carried out, where two different chloroquine concentrations (25 and 40 μM) and media with different contents were used, including Opti-MEM, which was reported to enable higher viral titer yields. These conditions are summarized in Table 2. As evident from the RFP fluorescence signals in Fig. 3, panels a and b, optimization studies enabled determination of the optimum transfection and production media, and detection of the optimum chloroquine concentration [40, 41].

Table 1 Three different conditions for LV production optimization Transfection media

Collection media

Exp A

Opti-MEM 2% FBS 1% penicillin/streptomycin

DMEM 2% FBS 1% penicillin/streptomycin

Exp B

Opti-MEM 2% FBS 1% penicillin/streptomycin

DMEM 10% FBS 1% penicillin/streptomycin

Exp C

Opti-MEM 10% FBS 1% penicillin/streptomycin

DMEM 10% FBS 1% penicillin/streptomycin

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Fig. 2 Comparison of the LV production efficiencies of the bubble and vortex methods 72 h after transfection under three different conditions (a, b, and c panels refer to the different conditions given in Table 1) 3.2.2 Scaling of the Optimized LV Production

Besides optimization of the production conditions, formation of an upscalable protocol is also very significant in LV production. For this purpose, testing of the applicability of the protocol for production in roller bottles is important. A primary task is to define the optimum seeding density and culturing rates for the 293T cells to reach the desired confluency in roller bottles.

Cell Culture in Roller Bottles

The surface area where cells can adhere and grow in roller bottles is approximately 1700 cm2, whereas the surface area of a 150-mm petri dish is approximately 150 cm2. Thus, the number of cells

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Table 2 Eight different conditions for optimization of LV production in terms of the optimum chloroquine concentration and media combinations Transfection media

Collection media

Exp 1

IMDM 25 μM chloroquine 10% FBS 1% penicillin/streptomycin

Opti-MEM 10% FBS 1% penicillin/streptomycin

Exp 2

IMDM 40 μM chloroquine 10% FBS 1% penicillin/streptomycin

Opti-MEM 10% FBS 1% penicillin/streptomycin

Exp 3

Opti-MEM 25 μM chloroquine 10% FBS 1% penicillin/streptomycin

Opti-MEM 10% FBS 1% penicillin/streptomycin

Exp 4

Opti-MEM 25 μM chloroquine 10% FBS 1% penicillin/streptomycin

Opti-MEM 10% FBS 1% penicillin/streptomycin

Exp 5

IMDM 25 μM chloroquine 10% FBS 1% penicillin/streptomycin

OPTIMEM 2% FBS 1% penicillin/streptomycin

Exp 6

IMDM 40 μM chloroquine 10% FBS 1% penicillin/streptomycin

Opti-MEM 2% FBS 1% penicillin/streptomycin

Exp 7

Opti-MEM 25 μM chloroquine 10% FBS 1% penicillin/streptomycin

Opti-MEM 2% FBS 1% penicillin/streptomycin

Exp 8

Opti-MEM 25 μM chloroquine 10% FBS 1% penicillin/streptomycin

Opti-MEM 2% FBS 1% penicillin/streptomycin

grown in a single roller bottle culture system roughly corresponds to that grown in 12 petri dishes. 1. First, a single flask of cells that reach a suitable density for transfection are trypsinized and counted, to define the time it will take 293T cells to reach 80–90% confluency in roller bottles. 2. Cells are seeded into roller bottles with 200 ml DMEM medium and followed for confluency.

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Fig. 3 RFP fluorescence signals at 24, 48, and 72 h of LV production with (a) 2% fetal bovine serum (FBS), and (b) 10% FBS-containing Opti-MEM used as collection media, along with four different combinations of Iscove’s modified Dulbecco’s medium (IMDM) and Opti-MEM media and 25 or 40 μM chloroquine at the transfection stage

3. Cells trypsinized from 12 different petri dishes are seeded in roller cell culture bottles, followed by overnight incubation in two different rates as 0.3 and 1 rpm. 4. Following trypsinization the next day, the rate at which 90% of the cells are attached to the surface is detected. 5. After the optimum rate for high attachment is defined, bottles with different densities of cells seeded are subject to incubation at different rates and different durations.

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Fig. 3 (continued)

6. Following incubation, cells in the roller bottles are trypsinized and cells counted separately. 7. Optimum parameters for obtaining cell numbers corresponding to 12 petri dishes are defined (Table 3). According to our results, 293T cells were ready for transfection under the seeding density and revolution time specified in the fifth setup (Fig. 4) [40, 41].

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Table 3 Optimization parameters for roller bottle cell culture seeding density, revolution speed, and incubation time Seeding density

Speed/time

1. Setup

Two petri dishes (40  106 cells)

0.3 rpm/48 h

2. Setup

Three petri dishes (60  106 cells)

0.3 rpm/48 h

3. Setup

Four petri dishes (80  106 cells)

0.3 rpm/48 h

4. Setup

Four petri dishes (80  106 cells)

0.3 rpm/24 h 1 rpm/24 h

5. Setup

Four petri dishes (80  106 cells)

0.3 rpm/24 h 1 rpm/16 h

Fig. 4 Cells ready for transfection in roller bottles with ribbed surfaces: (a) cells ready for transfection, seeded in roller cell culture bottles, (b) closer view of cells attached to the ribbed surface, (c) view of a roller cell culture bottle that has a 1700 cm2 surface area, and (d) 293T cells that have reached 100% confluence in 150 mm petri dishes

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Table 4 Exemplary calculation of the required plasmid amounts Plasmid

Required amount

Concentration (μg/μl)

Volume required

Gag/Pol

176

3.02

58

Rev

68

2.9

24

VSV-G

95

5.4

18

270

3.1

87

RFP (transfer vector)

LV Production in Roller Bottles Via CaPO4 Transfection

Cell numbers used for transfection in a single petri dish were adapted to the roller bottles. Accordingly, as the surface area of a single roller bottle corresponds to that of a total of 12 petri dishes, amounts of all solutions were multiplied by 12. Sample volume calculations are given in Table 4.

3.3 Optimized LV Production Protocol in Roller Cell Culture Bottles

HIV-based third-generation LV production via CaPO4 transfection method was optimized in roller cell culture bottles following the steps specified above (Fig. 5) [40, 41]. 1. Culture 293T cells in petri dishes that have a 15-cm2 surface area. 2. Trypsinize four of the petri dishes when cells are 100% confluent. 3. Add trypsinized cells to 200 ml DMEM medium (10% FBS and 1% pen-strep) and transfer to a single roller bottle. 4. Incubate cells in a roller bottle incubator for 24 h at 0.3 rpm speed, allowing them to adhere. 5. Adjust speed to 1 rpm for 16 h for cell expansion (Fig. 6a). 6. Replace the culture media with 180 ml transfection media (IMDM containing 10% FBS and 25 μM chloroquine). 7. Incubate roller bottles for 30 min at a speed of 0.3 rpm. 8. Add required amounts of Gag/Pol, Rev, VSV-G, and transfer plasmids into a Falcon tube. 9. Add 0.1 TE: dH2O mixture to a total volume of 10,500 μl. 10. Add 1,5 ml 2 M CaCl2, to obtain a final concentration of 0.25 M in a total volume of 12 ml. 11. Add 12 ml 2 HBS in a separate Falcon tube. 12. Add DNA–CaCl2 mixture dropwise using the bubble technique. 13. Incubate the final mixture for 5 min at RT to obtain surface neutralization (Fig. 6b).

Fig. 5 Figure schematizing production of HIV-based third-generation LVs via CaPO4 transfection method. Image created by Yunus Emre Eksi, MSc

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Fig. 6 Figure showing LV production via CaPO4 transfection following cell culture, and transfer into chloroquine-containing medium. (a) Incubation of cells ready for transfection, (b) preparation of the transfection cocktail, and (c) transfer of the transfection cocktail onto the cells in chloroquine-containing IMDM

14. Transfer the prepared transfection cocktail dropwise to the roller bottle containing the transfection media (Fig. 6c). 15. Incubate roller bottles at the speed of 0.3 rpm for 8 h at 37  C in a CO2 incubator. 16. Change chloroquine-containing transfection media with 10% FBS-containing Opti-MEM. 17. Incubate roller bottles at 37  C in a CO2 incubator up until 72 h following transfection. 18. Harvest the virus-containing supernatants when the incubation time is completed. 19. Centrifuge viral supernatants at 2000  g for 15 min to cleanse the supernatant from cell debris. 20. Filter the viral supernatants through a 0.45-μm vacuum filter. 21. Meanwhile, sterilize the ultracentrifuge tubes by UV irradiation. 22. Dispense viral supernatants as 30 ml per tube. 23. Create a sucrose cushion at the bottom of the tube with 5 ml sucrose solution (10% (v/v)). 24. Concentrate viral supernatants by ultracentrifugation at  82,000  g , 4  C for 2.5 h (Fig. 7a).

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Fig. 7 Ultracentrifugation procedure following the production process. (a) Ultracentrifugation instrument, (b) viral pellets obtained following ultracentrifugation, and (c) schematic representation of the lentivirus. Image created by Yunus Emre Eksi, MSc

25. Discard the supernatants following ultracentrifugation and resuspend the viral pellets in HBSS (Fig. 7b). 26. Collect LVs after resuspension and store aliquoted viral particles at 80  C, available as ready-to-use.

4

Notes 1. In PEI-based transfections used as an alternative to CaPO4 transfection, parameters such as PEI:DNA ratio, and polyplex amounts per cell differ between experiments that are performed with different media, cell line, plasmid structure, etc. [43, 44]; thus, optimization of the PEI-based techniques should be evaluated accordingly [30, 45]. 2. The success of the LV production is directly related to many different parameters such as the cell line used, size of the expression vector, whether a transfection agent is used or not, concentration of the transfection agent, the action mechanism of the transfection agent, and even the nature of the protein expressed from the transgene [28]. 3. Although the CaPO4 method provides a high transfection efficiency, it is negatively affected from changes in experimental

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parameters such as the pH, precipitation kinetics, and impurities in solutions [27, 32]. 4. In formation of the DNA/CaPO4 precipitate, lengthened incubation times cause increased precipitation volumes, thus a decrease in transfection efficiency. 5. Precipitation in CaPO4 transfection is carried out at RT in the presence of HBS at pH 7.05, containing 125 mM Ca2+ and 0.75 mM Na2HPO4 [46]. 6. During CaPO4 transfection in impure solutions, calcium ions in the precipitate may be substituted with ions such as Mg2+, Pb2+, and Zn2+, while phosphate ions may be exchanged with molecules such as carbohydrates in the medium or cellular CO2. The resulting alterations in CO2 and CO3+ concentrations lead to pH changes, thus affect the efficiency of the transfection. Buffer solutions such as HEPES should be added into the medium to avoid sudden pH changes [47, 48]. 7. The quantity, concentration, and purity of the plasmid DNAs that are introduced into the producer cell line affect the transfection yield. The purity of the plasmids obtained after isolation should be very close to an A260/280 value of 1.8. Quite pure and high concentration yields are obtained in plasmid isolation via the widely used commercial kits, compared to the cesium chloride and ethidium bromide-based isolation protocols. For production of LVs via CaPO4 transfection, the required recombinant DNA amounts are given generally as 1–15 μg/ 1  106 cells [27, 30, 49]. Optimization studies should be carried out for determination of the optimum plasmid amounts to be used in transfections where multiple plasmids are used. Exceeding amounts of transfer plasmid used compared to the other plasmids provide much higher production yields [27, 35]. 8. The action mechanisms of the transfection agents to be used in production should be well characterized. If the reagents used are capable of affecting cell viability and transfection, the optimum application method should be defined. If chloroquine is used, cell exposure should not exceed 8–12 h, to avoid toxicity and the resulting decrease in cell viability and viral titers.

Acknowledgments This study is supported by grants from Akdeniz University Scientific Research Administration Division (TYL-2015-1027) and the Scientific and Technological Research Council of Turkey (TUBITAK-112S114).

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References 1. Sakuma T et al (2012) Lentiviral vectors: basic to translational. Biochem J 443(3):603–618 2. Goff S (2001) Retroviridae: the retroviruses and their replication. In: Howley P et al (eds) Fields Virology. Lippincott, Williams and Wilkins, Philadelphia, PA, pp 1871–1939 3. Vogt PK (1997) Historical introduction to the general properties of retroviruses. In: Coffin JM et al (eds) Retroviruses. CSHL Press, Cold Spring Harbor, NY, pp 1–26 4. Tasyurek MH et al (2014) GLP-1-mediated gene therapy approaches for diabetes treatment. Expert Rev Mol Med 16:e7 5. Templeton NS (2009) Gene and cell therapy : therapeutic mechanisms and strategies, 3rd edn. CRC Press, Boca Raton 6. Powell DM et al (1997) HIV Rev-dependent binding of SF2/ASF to the Rev response element: possible role in Rev-mediated inhibition of HIV RNA splicing. Proc Natl Acad Sci U S A 94(3):973–978 7. Lever A et al (1989) Identification of a sequence required for efficient packaging of human immunodeficiency virus type 1 RNA into virions. J Virol 63(9):4085–4087 8. McBride MS, Panganiban AT (1996) The human immunodeficiency virus type 1 encapsidation site is a multipartite RNA element composed of functional hairpin structures. J Virol 70(5):2963–2973 9. Ciuffi A (2016) The benefits of integration. Clin Microbiol Infect 22(4):324–332 10. Frankel AD, Young JA (1998) HIV-1: fifteen proteins and an RNA. Annu Rev Biochem 67:1–25 11. Herrmann CH, Rice AP (1995) Lentivirus Tat proteins specifically associate with a cellular protein kinase, TAK, that hyperphosphorylates the carboxyl-terminal domain of the large subunit of RNA polymerase II: candidate for a Tat cofactor. J Virol 69(3):1612–1620 12. Kim YK et al (2002) Phosphorylation of the RNA polymerase II carboxyl-terminal domain by CDK9 is directly responsible for human immunodeficiency virus type 1 Tat-activated transcriptional elongation. Mol Cell Biol 22 (13):4622–4637 13. Jones KA, Peterlin BM (1994) Control of RNA initiation and elongation at the HIV-1 promoter. Annu Rev Biochem 63:717–743 14. Pollard VW, Malim MH (1998) The HIV-1 Rev protein. Annu Rev Microbiol 52:491–532 15. Collins DR, Collins KL (2014) HIV-1 accessory proteins adapt cellular adaptors to

facilitate immune evasion. PLoS Pathog 10 (1):e1003851 16. Simon V, Ho DD (2003) HIV-1 dynamics in vivo: implications for therapy. Nat Rev Microbiol 1(3):181–190 17. Naldini L et al (1996) In vivo gene delivery and stable transduction of nondividing cells by a lentiviral vector. Science 272(5259):263–267 18. Naldini L, Verma IM (2000) Lentiviral vectors. Adv Virus Res 55:599–609 19. Zufferey R et al (1997) Multiply attenuated lentiviral vector achieves efficient gene delivery in vivo. Nat Biotechnol 15(9):871–875 20. Dull T et al (1998) A third-generation lentivirus vector with a conditional packaging system. J Virol 72(11):8463–8471 21. Miyoshi H et al (1998) Development of a selfinactivating lentivirus vector. J Virol 72 (10):8150–8157 22. Zufferey R et al (1998) Self-inactivating lentivirus vector for safe and efficient in vivo gene delivery. J Virol 72(12):9873–9880 23. Zufferey R et al (1999) Woodchuck hepatitis virus posttranscriptional regulatory element enhances expression of transgenes delivered by retroviral vectors. J Virol 73(4):2886–2892 24. Stacey GN, M.O.-W. (2011) Host cells and cell banking. In: Merten OW, Al-Rubai M (eds) Viral vectors for gene therapy. Humana Press, Totowa, NJ, pp 45–88 25. Merten OW et al (2016) Production of lentiviral vectors. Mol Ther Methods Clin Dev 3:16017 26. Warnock JN et al (2006) Cell culture processes for the production of viral vectors for gene therapy purposes. Cytotechnology 50 (1–3):141–162 27. Segura MM et al (2007) Production of lentiviral vectors by large-scale transient transfection of suspension cultures and affinity chromatography purification. Biotechnol Bioeng 98(4):789–799 28. Geisse S (2009) Reflections on more than 10 years of TGE approaches. Protein Expr Purif 64(2):99–107 29. Ansorge S et al (2010) Recent progress in lentiviral vector mass production. Biochem Eng J 48(3):362–377 30. Ansorge S et al (2009) Development of a scalable process for high-yield lentiviral vector production by transient transfection of HEK293 suspension cultures. J Gene Med 11 (10):868–876

Lentivirus Production in Roller Bottles 31. Follenzi A, Naldini L (2002) Generation of HIV-1 derived lentiviral vectors. Methods Enzymol 346:454–465 32. Toledo JR et al (2009) Polyethylenimine-based transfection method as a simple and effective way to produce recombinant lentiviral vectors. Appl Biochem Biotechnol 157(3):538–544 33. Pham PL et al (2006) Large-scale transfection of mammalian cells for the fast production of recombinant protein. Mol Biotechnol 34 (2):225–237 34. Jordan M et al (1996) Transfecting mammalian cells: optimization of critical parameters affecting calcium-phosphate precipitate formation. Nucleic Acids Res 24(4):596–601 35. Mitta B et al (2005) Detailed design and comparative analysis of protocols for optimized production of high-performance HIV-1derived lentiviral particles. Metab Eng 7 (5–6):426–436 36. al Yacoub N et al (2007) Optimized production and concentration of lentiviral vectors containing large inserts. J Gene Med 9 (7):579–584 37. Sena-Esteves M et al (2004) Optimized largescale production of high titer lentivirus vector pseudotypes. J Virol Methods 122 (2):131–139 38. Karolewski BA et al (2003) Comparison of transfection conditions for a lentivirus vector produced in large volumes. Hum Gene Ther 14(14):1287–1296 39. Luthman H, Magnusson G (1983) High efficiency polyoma DNA transfection of chloroquine treated cells. Nucleic Acids Res 11 (5):1295–1308 40. Olgun H (2017) Optimized production and purification methods of third generation HIV-based Lentiviral Vectors for in vivo applications. Master’s thesis. Retrieved from Turkey Council of Higher Education Thesis Center

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41. Tasyurek HM et al (2018) Therapeutic potential of lentivirus-mediated glucagon-like peptide-1 (GLP-1) gene therapy for diabetes. Hum Gene Ther. https://doi.org/10.1089/ hum.2017.180 42. Tasyurek MH, Eksi YE, Sanlioglu AD, Altunbas HA, Balci MK, Griffith TS, Sanlioglu S (2018) HIV-based lentivirus-mediated vasoactive intestinal peptide gene delivery protects against DIO animal model of type 2 diabetes. Gene Ther. https://doi.org/10.1038/ s41434-018-0011-1 43. Backliwal G et al (2008) High-density transfection with HEK-293 cells allows doubling of transient titers and removes need for a priori DNA complex formation with PEI. Biotechnol Bioeng 99(3):721–727 44. Derouazi M et al (2004) Serum-free large-scale transient transfection of CHO cells. Biotechnol Bioeng 87(4):537–545 45. Durocher Y et al (2002) High-level and highthroughput recombinant protein production by transient transfection of suspensiongrowing human 293-EBNA1 cells. Nucleic Acids Res 30(2):E9 46. Yang YW, Yang JC (1997) Calcium phosphate as a gene carrier: electron microscopy. Biomaterials 18(3):213–217 47. Jordan M (2000) Transient gene expression in mammalian cells based on the calcium phosphate transfection method. In: Al-Rubeai M (ed) Cell engineering. Springer, Dordrecht 48. Kingston RE et al (2003) Calcium phosphate transfection. Curr Protoc Mol Biol Chapter 9: Unit 9.1 49. Tom R et al (2008) Transfection of HEK293EBNA1 cells in suspension with 293fectin for production of recombinant proteins. CSH Protoc 2008:pdb prot4979

Methods in Molecular Biology (2019) 1879: 347–365 DOI 10.1007/7651_2018_154 © Springer Science+Business Media New York 2018 Published online: 05 June 2018

High-Grade Purification of Third-Generation HIV-Based Lentiviral Vectors by Anion Exchange Chromatography for Experimental Gene and Stem Cell Therapy Applications Hazal Banu Olgun, Hale M. Tasyurek, Ahter D. Sanlioglu, and Salih Sanlioglu Abstract Lentiviral vectors (LVs) have been increasingly used in clinical gene therapy applications particularly due to their efficient gene transfer ability, lack of interference from preexisting viral immunity, and long-term gene expression they provide. Purity of LVs is essential in in vivo applications, for a high therapeutic benefit with minimum toxicity. Accordingly, laboratory scale production of LVs frequently involves transient cotransfection of 293T cells with packaging and transfer plasmids in the presence of CaPO4. After clearance of the cellular debris by low-speed centrifugation and filtration, lentivectors are usually concentrated by highspeed ultracentrifugation in sucrose cushion. Concentrated viral samples are then purified by anion exchange chromatography (AEX) after benzonase treatment to remove the residual cellular DNA. Here, we describe an improved practical method for LV purification using AEX, useful for experimental studies concerning gene and stem cell therapy. Keywords Anion exchange chromatography, Gene and cell therapy, Lentivirus

1

Introduction Gene therapy studies involve the introduction of genetic material into cells, tissues, or organs, via gene transfer tools called vectors [1]. Vectors used in gene therapy are often broadly categorized as viral and nonviral [2]. The most important feature that a gene therapy vector is expected to have is efficient transfer of the therapeutic gene into the target cell. Various factors should be taken into consideration in selection of a suitable transfer system, such as the characteristics of the target cell and tissue, immunogenicity of the vector to be used, size of the transgene to be transferred, and intended duration of gene expression [1]. Systems where naked/plasmid DNA is transferred into the target tissue directly or via chemical/physical methods are defined as nonviral vectors. Although nonviral systems are easier to apply and more cost-effective compared to the viral vectors, they also have major disadvantages such as low gene transfer efficiency and

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inadequate transcription level and stability [3]. Viruses’ ability to achieve efficient gene transfer to suitable host cells opened the way to their widespread use in a high number of preclinical studies and clinical trials; today, nearly 70% of the clinical trials utilize viral vectors [4]. In this regard, more effective and safer applications of viral vectors require enhanced purity and biosafety levels. Long-term and stable gene expression provided by the lentiviral vectors (LVs), and their relatively large cargo capacity in particular, have accelerated their increased use in gene therapy studies [5, 6]. LVs in this respect should be produced in accordance with the required quality and biosafety standards, for the clinical applicability of the potential gene therapy approaches. Our study aimed to improve LV purification process with various parameters evaluated and optimized, via use of chromatography-based techniques, known as highly productive and quality methods. 1.1 Purification of Lentiviral Vectors

Mammalian cells are widely used as host cells in production of LVs. The viral supernatant acquired at the end of the LV production process contains process-related residual media components and chemicals, along with metabolic wastes. Plasmid DNA and other free nucleic acids, and serum and other proteins are among the main sources of impurity. High molecular weight proteoglycans and DNA contaminants are quite difficult to eliminate, which are large particles that hold strong negative charges similar to LVs. At this point, downstream processing facilitates elimination of impurities contained in the harvested supernatants in terms of biosafety and provides increased vector concentration [7]. The basic principle of the viral purification process is selection of a method that will provide maximum purity with minimum number of experimental steps involved. This is because each extra step to be added has the potential to weaken the transduction ability of the vector [8]. Although virus purification processes are based on basic techniques used in protein purification, viruses’ being large molecules that are difficult to distinguish makes their purification more complicated [9]. The order of the downstream processes applied in the purification of the LV vectors following cell culture procedures is as follows: prefiltration, concentration via ultracentrifugation, removal of contaminant nucleic acids, and chromatography [10].

1.1.1 Prefiltration

The viral supernatant containing the viral vectors should be clarified via prefiltration prior to the concentration and purification steps. Removal of the residual cells and cell debris from the supernatant is achieved by low-speed centrifugation and microfiltration [7]. The first centrifugation step provides elimination of large particles before filtration. For the filtration step, diafiltration is frequently used, which provides the concentration of viral particles and salt removal [11, 12]. One of the major challenges encountered during the filtration process is membrane clogging. Techniques such as tangential-flow filtration

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(TFF) aim to avoid such problems, providing tangential flow of the solution across the ultrafiltration membrane instead of a direct flow [13, 14]. However, such filtration techniques primarily aim increased viral concentration, rather than enhanced purity of the vectors, which are usually used together with ultracentrifugation and chromatographic methods [15]. 1.1.2 Ultracentrifugation

Ultracentrifugation is one of the most preferred methods for concentration of the viral vectors harvested from the supernatant subsequent to the filtration step. Although up to a 100-fold concentration can be acquired via ultracentrifugation at 20,000–90,000
g, it is not usually accompanied by an increase in the transduction yield. Despite the fact that VSV-G pseudotyping increases resistance against mechanical force, lack of endurance for such a high centrifugal force for long durations is still expected. Furthermore, ultracentrifugationbased methods may constitute a disadvantage for large-scale processes in terms of duration and work load [12, 16, 17]. Following ultracentrifugation, the total volume decreases while the concentration of the vector increases. Use of these concentrated vectors in in vivo applications requires removal of the impurities that could not be removed by filtration, but precipitated along with the vector during centrifugation [18]. The most important of these impurities in LV production is the SV40T antigen, derived from the producer 293T cell lines. Removal of this antigen must be assured for the vectors to be applicable to clinical trials in terms of biosafety. Because the ultracentrifugation process does not provide a complete purification even if the sucrose cushion method is used, such contaminant proteins can only be removed by techniques such as chromatographic purification [11].

1.1.3 Contaminant Nucleic Acid Degradation

Although the prefiltration and concentration steps provide removal of many cell- and media-derived impurities, nucleic acid remnants from the cells and plasmids utilized in the vector production process still remain in the LV solution. The quality of the viral supernatant is negatively affected with time, as cellular lysis during LV production increases the contaminating host DNA, RNA, and free nucleic acids as well as cell debris contents. Besides constituting a biosafety issue, contaminant nucleic acids may also cause an increase in viscosity, which may lead to difficulties in the purification steps [9, 19]. The decontamination process gets further complicated with the fact that the residual nucleic acids possess a similar electrical charge as the viral vector particles themselves. Benzonase application is suggested and applied as a solution to this problem [19]. Although it is possible for the residual nucleic acids to be degraded into small fragments with this application, additional effective purification steps are required afterwards, for removal of both the benzonase and the degraded fragments. Size distribution of residual

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contaminant DNA in the viral stock is given as