In Vitro Models for Stem Cell Therapy: Methods and Protocols (Methods in Molecular Biology, 2269) 1071612247, 9781071612248

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Methods in Molecular Biology 2269

Peggy Stock Bruno Christ Editors

In Vitro Models for Stem Cell Therapy Methods and Protocols

METHODS

IN

MOLECULAR BIOLOGY

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

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

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

In Vitro Models for Stem Cell Therapy Methods and Protocols

Edited by

Peggy Stock and Bruno Christ Department of Visceral, Transplant, Thoracic and Vascular Surgery, Applied Molecular Hepatology Lab, University of Leipzig Medical Center, Leipzig, Germany

Editors Peggy Stock Department of Visceral, Transplant, Thoracic and Vascular Surgery Applied Molecular Hepatology Lab, University of Leipzig Medical Center Leipzig, Germany

Bruno Christ Department of Visceral, Transplant, Thoracic and Vascular Surgery Applied Molecular Hepatology Lab, University of Leipzig Medical Center Leipzig, Germany

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-1224-8 ISBN 978-1-0716-1225-5 (eBook) https://doi.org/10.1007/978-1-0716-1225-5 © Springer Science+Business Media, LLC, part of Springer Nature 2021 This work is subject to copyright. All rights are reserved by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, expressed or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. Cover Caption: Co-culture of primary mouse hepatocytes and human mesenchymal stromal cells highlighting the formation of tunneling nanotubes with courtesy of Dr. Hsu from our Lab This Humana imprint is published by the registered company Springer Science+Business Media, LLC, part of Springer Nature. The registered company address is: 1 New York Plaza, New York, NY 10004, U.S.A.

Preface Initially, it was the biology of the embryonic stem cell and its unique feature to develop into multiple lineages, which gave us insight into the molecular nature of developmental processes leading to specified tissues and organs. Yet, soon we understood that there was a much broader spectrum of applicability of stem cells in basic and applied sciences but increasingly also in clinical use. Today, this is of high medicinal and socio-economic interest facing an increasing shortage of organs for transplantation, which is very often the only remaining therapeutic option. In recent years, the therapeutic potential of both embryonic and adult stem cells have been investigated intensively in animal models of human diseases giving hope of clinical feasibility of stem cell therapy in the near future. However, there is a mandatory need to understand the way stem cells work under specified disease conditions, which are as diverse as the diversity of the diseases itself. Therefore, it is evident that stem cells from their nature as multi- or pluripotent cells may respond in a different way depending on the microenvironment of a specific disease and the affected tissue. This way of response may comprise direct impact on tissue regeneration by differentiation of a stem cell into the healthy cell of the tissue targeted but also paracrine effects such as secretion of growth factors and/or cytokines affecting the diseased tissue to support its self-regenerative capacity just to mention two major routes of action. This implies that before clinical translation of stem cell therapy it is warranted to characterize molecular and cellular mechanisms of stem cell actions more closely. Both impact and potential side effects of stem cell applications such as their tumorigenic potential, cytokine productivity and efficiency, and tissue specificity in in vitro models have to be assessed. To reduce biological complexity, and thus increase specificity and comprehensibility of stem cell actions, in vitro assays mimicking disease conditions gain attraction in stem cell research and clinical translation. This book addresses exemplified in vitro disease models representing the respiratory, hepatobiliary, osteochondral, nervous, dermal, ocular, and immune system as well as pathological biological processes like tumorigenesis for stem cell research. The contents of the book are prepared to cover a range of diseases and application of different kinds of stem cells such as adult stem cells, but also reprogrammed tissue cells (iPS), which are the types of cells most frequently discussed in the context of applied sciences and medicine. In order to support assessment of molecular and cellular mechanisms of stem cell actions more closely, the book includes novel methods to characterize and manipulate stem cells with the aim to better understand and optimize their biological performance. Hence, the description of the in vitro disease models and their application in preclinical stem cell research in this book may address basic scientists and clinicians to tailor the biological as well as the therapeutic potential of stem cell therapy in clinical use.

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The standardization of protocols and assays is mandatory for future implementation into regulatory documents presenting the results from different individual cell types in different in vitro models in order to prepare a preclinical study in vivo as required for later approval by regulatory bodies. Leipzig, Germany

Peggy Stock Bruno Christ

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

PART I

INTRODUCTION

1 Consistent Inclusion of Mesenchymal Stem Cells into In Vitro Tumor Models . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ˜ o F. Mano Luı´s P. Ferreira, Vı´tor M. Gaspar, and Joa 2 Pluripotent Stem Cells for Cell Therapy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Insa S. Schroeder

PART II

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DISEASE MODELING

3 In Vitro Methods for the Study of Glioblastoma Stem-Like Cell Radiosensitivity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 37 Joseph H. McAbee, Charlotte Degorre-Kerbaul, and Philip J. Tofilon 4 Bioimaging of Mesenchymal Stem Cells Spatial Distribution and Interactions with 3D In Vitro Tumor Spheroids . . . . . . . . . . . . . . . . . . . . . . . . 49 ˜ o F. Mano Luı´s P. Ferreira, Vı´tor M. Gaspar, and Joa 5 Investigation of the MSC Paracrine Effects on Alveolar–Capillary Barrier Integrity in the In Vitro Models of ARDS . . . . . . . . . . . . . . . . . . . . . . . . . . . 63 Johnatas Dutra Silva and Anna D. Krasnodembskaya 6 In Vitro Methods to Evaluate the Effects of Mesenchymal Stem Cells on TGF-β1-Induced Pulmonary Fibrosis. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 83 Ying-Wei Lan, Chuan-Mu Chen, and Kowit-Yu Chong 7 Study of Mesenchymal Stem Cell-Mediated Mitochondrial Transfer in In Vitro Models of Oxidant-Mediated Airway Epithelial and Smooth Muscle Cell Injury. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 93 Charalambos Michaeloudes, Xiang Li, Judith C. W. Mak, and Pankaj K. Bhavsar 8 Co-Culture of Peripheral Blood Mononuclear Cells and Endothelial Colony Forming Cells from Cord Blood of Preterm Born Babies . . . . . . . . . . . . . 107 Jan Baier, Anchang Charles Gwellem, Roland Haase, Ines Volkmer, Babette Bartling, and Martin S. Staege 9 Ex Vivo Model of Spontaneous Neuroretinal Degeneration for Evaluating Stem Cells’ Paracrine Properties . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 125 Ivan Fernandez-Bueno and Ricardo Usategui-Martin 10 Ex Vivo Normothermic Hypoxic Rat Liver Perfusion Model: An Experimental Setting for Organ Recondition and Pharmacological Intervention . . . . . . . . . . . . 139 Federica Rigo, Victor Navarro-Tableros, Nicola De Stefano, Alberto Calleri, and Renato Romagnoli

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Contents

Co-Culture of Human Mesenchymal Stromal Cells and Primary Mouse Hepatocytes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 151 Mei-Ju Hsu, Madlen Christ, and Bruno Christ A 3D Dynamic In Vitro Model of Inflammatory Tendon Disease . . . . . . . . . . . . . 167 Susanna Schubert, Luisa Brandt, and Janina Burk Generation of Epidermal Equivalents from Hair Follicle Melanocytes, Keratinocytes, and Dermal Fibroblasts. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 175 Vuk Savkovic, Marie Schneider, Hanluo Li, Jan-Christoph Simon, Mirjana Ziemer, and Bernd Lethaus

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CHARACTERIZATION AND PRE-CONDITIONING

Using Gene Expression Music Algorithms (GEMusicA) for the Characterization of Human Stem Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Martin S. Staege Evaluation of Extracellular Vesicles from Adipose Tissue-Derived Mesenchymal Stem Cells in Primary Human Chondrocytes from Patients with Osteoarthritis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ˜ , and Marı´a Jose´ Alcaraz Marı´a Isabel Guille´n, Alvaro Compan Generation of Neural Stem Cells from Pluripotent Stem Cells for Characterization of Early Neuronal Development. . . . . . . . . . . . . . . . . . . . . . . . Matthias Jung, Jovita Schiller, Carla Hartmann, Jenny Pfeifer, and Dan Rujescu Engineered Tissues Made from Human iPSC-Derived Schwann Cells for Investigating Peripheral Nerve Regeneration In Vitro . . . . . . . . . . . . . . . . . . . . Rebecca Powell and James B. Phillips Modulation of a Stem Cell Gene: LGR4 Knockout in a Human Cell Line by CRISPR/Cas Method . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Swetlana Rot and Matthias Kappler In Vitro Tool: 3D Cell Culture of Human Adipose Tissue-Derived Mesenchymal Stromal Cells on Low Stiffness Silicone Scaffolds. . . . . . . . . . . . . . . Peggy Stock

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

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Contributors MARI´A JOSE´ ALCARAZ • Interuniversity Research Institute for Molecular Recognition and Technological Development (IDM), Polytechnic University of Valencia, University of Valencia, Burjasot, Valencia, Spain JAN BAIER • Section for Neonatology and Pediatric Intensive Care, Department for Operative and Nonoperative Pediatric and Adolescent Medicine, University Hospital Halle (Saale), Medical Faculty of Martin Luther University Halle-Wittenberg, Halle (Saale), Germany BABETTE BARTLING • Institute of Agricultural and Nutritional Sciences Animal Health Management, Martin Luther University Halle-Wittenberg, Halle (Saale), Germany PANKAJ K. BHAVSAR • National Heart and Lung Institute, Imperial College London, London, UK; Respiratory & Critical Care Medicine, The University of Hong KongShenzhen Hospital, Shenzhen, Guangdong, People’s Republic of China LUISA BRANDT • Saxon Incubator for Clinical Translation (SIKT), University of Leipzig, Leipzig, Germany JANINA BURK • Saxon Incubator for Clinical Translation (SIKT), University of Leipzig, Leipzig, Germany; Institute of Veterinary Physiology, University of Leipzig, Leipzig, Germany; Equine Clinic (Surgery, Orthopedics), Justus Liebig University Giessen, Giessen, Germany ` della Salute ALBERTO CALLERI • General Surgery 2U, Liver Transplant Unit, A.O.U Citta e della Scienza di Torino, University of Turin, Turin, Italy CHUAN-MU CHEN • Department of Life Sciences, and Ph.D. Program in Translational Medicine, National Chung Hsing University, Taichung, Taiwan; The iEGG and Animal Biotechnology Center, and Rong Hsing Research Center for Translational Medicine, National Chung Hsing University, Taichung, Taiwan KOWIT-YU CHONG • Department of Medical Biotechnology and Laboratory Science, Graduate Institute of Biomedical Sciences, College of Medicine, Chang Gung University, Taoyuan, Taiwan; Division of Biotechnology, Graduate Institute of Biomedical Sciences, College of Medicine, Chang Gung University, Taoyuan, Taiwan; Department of Laboratory Medicine, and Hyperbaric Oxygen Medical Research Lab, Bone and Joint Research Center, Chang Gung Memorial Hospital-Linkou, Taoyuan, Taiwan; Centre for Stem Cell Research, Faculty of Medicine and Health Sciences, Universiti Tunku Abdul Rahman, Kajang, Selangor, Malaysia BRUNO CHRIST • Department of Visceral, Transplant, Thoracic and Vascular Surgery, Applied Molecular Hepatology Lab, University of Leipzig Medical Center, Leipzig, Germany MADLEN CHRIST • Department of Visceral, Transplant, Thoracic and Vascular Surgery, Applied Molecular Hepatology Lab, University of Leipzig Medical Center, Leipzig, Germany ALVARO COMPAN˜ • Interuniversity Research Institute for Molecular Recognition and Technological Development (IDM), Polytechnic University of Valencia, University of Valencia, Burjasot, Valencia, Spain CHARLOTTE DEGORRE-KERBAUL • Radiation Oncology Branch, National Cancer Institute, National Institutes of Health, Bethesda, MD, USA

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` della NICOLA DE STEFANO • General Surgery 2U, Liver Transplant Unit, A.O.U Citta Salute e della Scienza di Torino, University of Turin, Turin, Italy IVAN FERNANDEZ-BUENO • Instituto Universitario de Oftalmobiologı´a Aplicada (IOBA), Universidad de Valladolid, Campus Miguel Delibes, Valladolid, Spain; Centro en Red de Medicina Regenerativa y Terapia Celular de Castilla y Leon, Valladolid, Spain; Red Tema´tica de Investigacion Cooperativa en Salud (RETICS), Oftared, Instituto de Salud Carlos III, Valladolid, Spain LUI´S P. FERREIRA • Department of Chemistry, CICECO—Aveiro Institute of Materials, University of Aveiro, Campus Universita´rio de Santiago, Aveiro, Portugal VI´TOR M. GASPAR • Department of Chemistry, CICECO—Aveiro Institute of Materials, University of Aveiro, Campus Universita´rio de Santiago, Aveiro, Portugal MARI´A ISABEL GUILLE´N • Interuniversity Research Institute for Molecular Recognition and Technological Development (IDM), Polytechnic University of Valencia, University of Valencia, Burjasot, Valencia, Spain; Department of Pharmacy, Faculty of Health Sciences, Cardenal Herrera-CEU University, Alfara del Patriarca, Valencia, Spain ANCHANG CHARLES GWELLEM • Department for Operative and Nonoperative Pediatric and Adolescent Medicine, University Clinic and Outpatient Clinic for Pediatrics I, University Hospital Halle (Saale), Medical Faculty of Martin Luther University Halle-Wittenberg, Halle (Saale), Germany ROLAND HAASE • Section for Neonatology and Pediatric Intensive Care, Department for Operative and Nonoperative Pediatric and Adolescent Medicine, University Hospital Halle (Saale), Medical Faculty of Martin Luther University Halle-Wittenberg, Halle (Saale), Germany CARLA HARTMANN • Martin Luther University Halle-Wittenberg, University Clinic and Outpatient Clinic for Psychiatry, Psychotherapy, and Psychosomatic Medicine, Halle/Saale, Germany MEI-JU HSU • Department of Visceral, Transplant, Thoracic and Vascular Surgery, Applied Molecular Hepatology Lab, University of Leipzig Medical Center, Leipzig, Germany MATTHIAS JUNG • Martin Luther University Halle-Wittenberg, University Clinic and Outpatient Clinic for Psychiatry, Psychotherapy, and Psychosomatic Medicine, Halle/Saale, Germany MATTHIAS KAPPLER • Department of Oral and Maxillofacial Plastic Surgery, Martin Luther University Halle-Wittenberg, Halle, Germany ANNA D. KRASNODEMBSKAYA • Wellcome-Wolfson Institute for Experimental Medicine, Queen’s University of Belfast, Belfast, UK YING-WEI LAN • Department of Medical Biotechnology and Laboratory Science, Graduate Institute of Biomedical Sciences, College of Medicine, Chang Gung University, Taoyuan, Taiwan; Division of Biotechnology, Graduate Institute of Biomedical Sciences, College of Medicine, Chang Gung University, Taoyuan, Taiwan BERND LETHAUS • Department of Cranio Maxillofacial Surgery, University of Leipzig Medical Center, Leipzig, Germany HANLUO LI • Department of Cranio Maxillofacial Surgery, University of Leipzig Medical Center, Leipzig, Germany XIANG LI • Department of Medicine, The University of Hong Kong, Pok Fu Lam, Hong Kong SAR

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JUDITH C. W. MAK • Respiratory & Critical Care Medicine, The University of Hong KongShenzhen Hospital, Shenzhen, Guangdong, People’s Republic of China; Department of Medicine, The University of Hong Kong, Pok Fu Lam, Hong Kong SAR; Department of Pharmacology & Pharmacy, The University of Hong Kong, Pok Fu Lam, Hong Kong SAR JOA˜O F. MANO • Department of Chemistry, CICECO—Aveiro Institute of Materials, University of Aveiro, Campus Universita´rio de Santiago, Aveiro, Portugal JOSEPH H. MCABEE • Radiation Oncology Branch, National Cancer Institute, National Institutes of Health, Bethesda, MD, USA; Department of Clinical Neurosciences, University of Cambridge, Cambridge, UK; Wake Forest School of Medicine, WinstonSalem, NC, USA CHARALAMBOS MICHAELOUDES • National Heart and Lung Institute, Imperial College London, London, UK; Respiratory & Critical Care Medicine, The University of Hong Kong-Shenzhen Hospital, Shenzhen, Guangdong, People’s Republic of China ` VICTOR NAVARRO-TABLEROS • Scarl., Molecular Biotechnology Center (MBC), 2i3T, Societa per La Gestione Dell’incubatore Di Imprese e Per Il Trasferimento Tecnologico ` degli Studi di Torino, Turin, Italy Dell’Universita JENNY PFEIFFER • Martin Luther University Halle-Wittenberg, University Clinic and Outpatient Clinic for Psychiatry, Psychotherapy, and Psychosomatic Medicine, Halle/Saale, Germany JAMES B. PHILLIPS • UCL Centre for Nerve Engineering, Department of Pharmacology, UCL School of Pharmacy, University College London, London, UK REBECCA POWELL • UCL Centre for Nerve Engineering, Department of Pharmacology, UCL School of Pharmacy, University College London, London, UK ` della Salute FEDERICA RIGO • General Surgery 2U, Liver Transplant Unit, AOU Citta e della Scienza di Torino, University of Turin, Turin, Italy ` della RENATO ROMAGNOLI • General Surgery 2U, Liver Transplantation Center, AOU Citta Salute e della Scienza di Torino, University of Turin, Turin, Italy SWETLANA ROT • Department of Oral and Maxillofacial Plastic Surgery, Martin Luther University Halle-Wittenberg, Halle, Germany DAN RUJESCU • Martin Luther University Halle-Wittenberg, University Clinic and Outpatient Clinic for Psychiatry, Psychotherapy, and Psychosomatic Medicine, Halle/Saale, Germany VUK SAVKOVIC • Department of Cranio Maxillofacial Surgery, University of Leipzig Medical Center, Leipzig, Germany JOVITA SCHILLER • Martin Luther University Halle-Wittenberg, University Clinic and Outpatient Clinic for Psychiatry, Psychotherapy, and Psychosomatic Medicine, Halle/Saale, Germany MARIE SCHNEIDER • Department of Hematology, Cell Therapy and Hemostaseology, University of Leipzig Medical Center, Leipzig, Germany INSA S. SCHROEDER • Stem Cell Differentiation and Cytogenetics Group, Biophysics Department, GSI Helmholtz Center for Heavy Ion Research, Darmstadt, Germany SUSANNA SCHUBERT • Institute of Human Genetics, University of Leipzig Medical Center, Leipzig, Germany; Saxon Incubator for Clinical Translation (SIKT), University of Leipzig, Leipzig, Germany; Institute of Veterinary Physiology, University of Leipzig, Leipzig, Germany JOHNATAS DUTRA SILVA • Wellcome-Wolfson Institute for Experimental Medicine, Queen’s University of Belfast, Belfast, UK

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JAN-CHRISTOPH SIMON • Department of Dermatology, Venerology and Allergology, University of Leipzig Medical Center, Leipzig, Germany MARTIN S. STAEGE • Department for Operative and Nonoperative Pediatric and Adolescent Medicine, University Clinic and Outpatient Clinic for Pediatrics I, University Hospital Halle (Saale), Medical Faculty of Martin Luther University Halle-Wittenberg, Halle (Saale), Germany; Department of Surgical and Conservative Paediatrics and Adolescent Medicine, Martin Luther University Halle-Wittenberg, Halle, Germany PEGGY STOCK • Department of Visceral, Transplant, Thoracic and Vascular Surgery, Applied Molecular Hepatology Lab, University of Leipzig Medical Center, Leipzig, Germany PHILIP J. TOFILON • Radiation Oncology Branch, National Cancer Institute, National Institutes of Health, Bethesda, MD, USA RICARDO USATEGUI-MARTIN • Instituto Universitario de Oftalmobiologı´a Aplicada (IOBA), Universidad de Valladolid, Campus Miguel Delibes, Valladolid, Spain INES VOLKMER • Department for Operative and Nonoperative Pediatric and Adolescent Medicine, University Clinic and Outpatient Clinic for Pediatrics I, University Hospital Halle (Saale), Medical Faculty of Martin Luther University Halle-Wittenberg, Halle (Saale), Germany MIRJANA ZIEMER • Department of Dermatology, Venerology and Allergology, University of Leipzig Medical Center, Leipzig, Germany

Part I Introduction

Chapter 1 Consistent Inclusion of Mesenchymal Stem Cells into In Vitro Tumor Models Luı´s P. Ferreira, Vı´tor M. Gaspar, and Joa˜o F. Mano Abstract Over recent years, the role of distinct mesenchymal stem cell populations in cancer progression has become increasingly evident. In this regard, developing in vitro preclinical tumor models capable of portraying tumor-associated mesenchymal stem cells (TA-MSCs) interactions with the tumor microenvironment (TME), cellular and extracellular components, would allow to improve the predictive potential of these platforms and expedite preclinical drug screening. Although recent studies successfully developed in vitro tumor models in which the biomolecular and cellular behaviors of TA-MSCs were recapitulated in the context of their interactions with specific TME components, no consensus has yet been reached regarding distinct TA-MSCs influence in the evolution of solid tumors. The paradoxical observations regarding the roles of MSCs on in vitro tumor models can in part be associated to a lack of standardization in how MSCs integration is performed. Herein, we summarize some of the main parameters linked to phenotypic variations established upon MSCs inclusion and interaction within in vitro tumor models. A critical overview of recent studies and how standardization of key parameters could improve the reproducibility and predictability of current preclinical validation models containing MSCs is also provided. Key words Tumor-associated mesenchymal stem cells, In vitro tumor models, MSCs standardization, Tumor microenvironment, Therapeutics screening

1 Tumor-Associated Mesenchymal Stem Cells—A New Paradigm in Cancer-Stroma Cooperation Cancer progression is no longer grasped solely as a one-dimensional process driven by the accumulation of genetic mutations in malignant cells, being instead a multifactorial process involving the co-regulation of its surrounding tumor microenvironment (TME) [1]. At the initial stages of cancer development, cancer cells exert a disruptive influence in tissue homeostasis, altering the phenotype of surrounding stromal cells [2] and associated extracellular matrix (TME-ECM) [3]. Despite the growing knowledge concerning cancer progression and tumor microenvironment influence in disease progression, most preclinical drug discovery/screening is still performed in models that lack correct cellular composition and Peggy Stock and Bruno Christ (eds.), In Vitro Models for Stem Cell Therapy: Methods and Protocols, Methods in Molecular Biology, vol. 2269, https://doi.org/10.1007/978-1-0716-1225-5_1, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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homology, as well as ECM and/or three-dimensional representation [4]. The lack of correspondence between in vitro platforms in which preclinical validation of novel compounds is performed, and the highly complex in vivo scenario in which they are applied, is one of the major obstacles for the development of more effective treatments and their transition into the clinical setting [5]. Consequently, translation of candidate therapies that perform well in in vitro cancer models, specifically those targeting pathways that suffer TME-dependent modulation (e.g., anti-angiogenic drugs, immunotherapeutic treatments), commonly fail either by lack of efficacy, or due to the onset of unforeseen side-effects [5]. In this context, tumor-associated mesenchymal stem cells (TA-MSCs) capacity to influence cancer progression, invasion, and treatment response is now increasingly recognized as a new paradigm of cancer-stroma cooperation. Including this hallmark is therefore central to the development of in vitro tumor models for testing TME targeting or immunotherapeutic approaches. MSCs represent a diverse class of multipotent stromal cells with a fibroblast-like morphology, known to exert distinct regenerative, and immune-regulatory function in accordance to their tissue of origin and surrounding microenvironment [6]. Depending on their origin and tissue-specific subpopulation, MSCs can present distinct phenotypes with varying secretomes, metabolic activity, epigenetic profiles, and small variances in surface receptors [6]. These differences translate to distinct proficiencies in achieving lineage differentiation, and migration towards the microenvironment of solid tumors [7]. Nevertheless, given their generalized ability to migrate towards injury sites MSCs are attracted to the TME by the release of pro-inflammatory signals, of specific cytokines, and signaling mediators, (e.g., VEGF, TGF-β1 IL-1β, CXCL12, and SDF-1α [8, 9]). Once in the TME, MSCs migrate towards and interact with cancer cells, experiencing further phenotypical and metabolic alterations that promote the appearance of distinct pro- or anti-tumoral TA-MSCs phenotypes [10]. These TA-MSCs populations have been shown to exhibit anti- or pro-tumoral phenotypes both in vitro and in vivo [11] through the release of exosomes, mRNAs, cytokines, and growth factors, or by interacting directly with surrounding populations through the exchange of intracellular organelles and sections of their extracellular membrane [8]. Through these pathways of communication pro-tumoral TA-MSCs promote metabolic and regulatory alterations in both cancer cells and other key populations of the TME (e.g., cancer-associated fibroblasts (CAFs), macrophages, T regulatory cells (Treg), Natural Killer (NK) cells), by: (i) promoting the appearance of immune-suppressive and pro-angiogenic niches, (ii) increasing cancer cell migration and invasion, and (iii) leading to increased metastatic potential and resistance to therapy [12– 15]. Such interactions are regulated by the extremely complex

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setting of multifactorial paracrine and cell–cell interactions derived from communication with both cancer cells and other cellular and extracellular matrix (ECM) components of the TME [16, 17]. Therefore, given the distinct phenotypes presented by MSCs of diverse tissue-specific and subpopulation-specific origins, (e.g., MSCs derived from adipose tissue [18], lung [19], synovial membrane [20], muscle [21], peripheral blood [22], as well as from fetal/neonatal tissues [23]), several questions have arisen regarding the direct and indirect interactions of population-specific TA-MSCs with the surrounding TME [13]. Despite the accumulation of distinct findings portraying MSCs in pro- or anti-tumoral roles, thus far there is no consensus on how such an engagement occurs. Regarding this, an increasing number of reports is focusing on analyzing the molecular pathways of cell–cell and cell–ECM interactions, based mainly in in vitro observed exchanges. Such observations demonstrate that parameters such as the type/stage of cancer, and MSCs phenotype can act against cancer progression [14, 24, 25]. For example, bone marrow-derived MSCs (hBM-MSCs) have been found to stimulate TME immune cell recognition in colon cancer [26]. Furthermore, umbilical cord-derived MSCs (hUC-MSCs) can attenuate lung cancer and hepatocarcinoma in vitro proliferation by paracrine inhibition of Wnt and promotion of apoptosis [27]. However, the large number of experimental approaches, culture conditions, qualitative and quantitative methods, and in vitro or in vivo models used thus far to evaluate immune-regulatory properties of TA-MSCs lack standardized characterization of crucial parameters, such as the analysis of present subpopulations, and population ratios variation overtime. Consequently, leading to an excess of literature and raw data that is poorly comparable, and occasionally contradictory. Integrating such a highly variable class of cells in complex multi co-culture environments, either in 2D or 3D cultures, is not trivial. Variations in key factors such as: (i) tissue origin, (ii) tissue-specific subpopulations, (iii) patient age, (iv) culture medium, (v) culture substrate, (vi) pre-conditioning, and (vii) co-culture with other cells in varying ratios, has been found to promote non-concordant results [11, 28]. Therefore, developing novel standards and guidelines for the inclusion of MSCs into preclinical validation models of the TME could promote an increased level of efficacy, predictability, and reproducibility between obtained results. Herein, we present a critical overview of the diverse parameters which must be further characterized and standardized in order to allow the development of uniform protocols for both preparation and utilization of MSCs during preclinical disease models development.

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Mesenchymal Stem Cells—Origin and Characterization Currently, MSCs in vitro identification is based on the following minimal criteria: (i) MSCs must be capable of growing in adherent culture conditions, (ii) be able to differentiate into osteoblasts, adipocytes, and chondroblasts in vitro, and (iii) express a defined subset of cluster of differentiation (CD) markers established by the International Society for Cell and Gene Therapy. Mesenchymal stem cell molecular identification is mainly based in positive expression of surface makers such as CD105, CD90, CD73, and an overall lack of CD45, CD14, CD34 or CD11b, CD79a or CD19 markers [14, 29]. Using these parameters, several studies have been able to isolate mesenchymal stem cells from distinct tissue sources, such as bone marrow, adipose, breast, tissue, skin, lung, and vascular tissues [30, 31]. Moreover, through a combination of histology and immunohistochemistry analysis, of cell morphology and respective CD markers expression, TA-MSCs have been identified among cell populations of the TME of numerous cancers such as colorectal [32], pancreatic [33], glioma [34], breast [35], lung [19], prostate [17], gastric [36], and ovarian [37].

2.1 Selecting the Source of MSCs— Human Vs Animal-Derived Stem Cells

Considering either animal or human sources for MSCs used in the development of in vitro disease models is crucial when establishing cross comparisons. As recently reviewed elsewhere [13, 23, 38, 39], MSCs derived from either human or animal sources show distinct tendencies towards the acquisition of either pro-tumoral or antitumoral TA-MSCs phenotypes derived from the communications they establish with surrounding cells within in vitro tumor models. For example, as shown by Oloyo et al. 2017 [23], while in vitro studies implementing mice-derived MSCs showcase a tendency towards the establishment of pro-tumoral phenotypes, rat-derived MSCs exhibit a tendency towards displaying anti-tumoral phenotypes. Such variations in behavior can be directly correlated with distinct surface receptor presentation, and alternative signaling pathways resultant from variations in the host-immune system pathways of communication [38]. Therefore, for the development of predictive in vitro models capable of replicating protein/marker homology and the complex molecular pathways found in vivo, a continued focus on the utilization of human-derived MSCs must be considered. Furthermore, attention must be given towards possible heterogeneity in the genetic, secretory, and lineage-predisposition phenotypes arising from patient-to-patient variability in humanderived MSCs isolates [39, 40]. Albeit undesirable from a reproducibility standpoint, such variability if well registered and accounted for during study design, can in fact provide a valuable source of information [41]. Given that multi-lineage and distinct phenotypic presentations of human-derived MSCs (human-derived

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MSCs) are important for fundamental tumor biology studies and for drug screening, researchers must take into consideration both patient-to-patient variations and subpopulation variations [42]. 2.2 Tissue-Specific Phenotype Variations in Human-Derived MSCs

Most studies employing human-derived MSCs to analyze TA-MSCs role on the TME have mainly focused in analyzing the role of bone marrow-derived MSCs (hBM-MSCs), fewer studies have focused in adipose tissue-derived MSCs (hAT-MSCs), and umbilical cord-derived MSCs (hUC-MSCs). The majority of these reports has been performed in breast, colon, and lung cancer in vitro models [23, 43]. However, there is still a lack of knowledge regarding the specific roles that tissue-specific MSCs can play within distinct cancer microenvironments. Yet, a growing body of knowledge has emphasized that hBM-MSCs and hAT-MSCs, respectively, mostly demonstrate pro-tumoral and ambivalent phenotypes, highly dependent on the cancer type and its microenvironment. Contrarily, hUC-MSCs have been shown to consistently exhibit anti-tumoral properties [44]. Extensive reports have shown hUC-MSCs exhibit both distinct immune-regulatory potential, promoting recognition of cancer cells by TME-associated immune cells and promoting pro-apoptotic pathways in mutated cells [27, 45, 46]. Furthermore, hUC-MSCs showcase an increased proliferative potential when compared to adult human-derived MSCs [47]. While such characteristics might render hUC-MSCs highly desirable for the development of TME targeting therapies they also pinpoint this class of MSCs as unideal candidates to model pro-tumoral TA-MSCs within the context of TME communication networks, thus limiting their utility in the development of in vitro tumor models [13, 39].

2.3 Distinguishing Distinct Tissue-Specific Subpopulations

The established minimal criteria parameters for MSCs identification have proven valuable, providing the capacity to fully isolate MSCs from a distinct plethora of tissues and integrate this crucial TME cellular component into in vitro cellular therapy and disease modeling systems. Nevertheless, simple characterization of conventional CD markers has been unable to provide a clear window for distinguishing between human-derived MSCs of distinct tissue origin and present subpopulations. Not all MSCs are created equal, with subpopulations of tissue-specific MSCs being associated with varying levels of integrin molecules expression (CD49a, CD106, and CD 146) [38]. These patterns of integrin expression can be correlated with adhesion to distinct ECM environments, to differences in both molecular signaling mechanisms and levels of activity, allowing to differentiate between MSCs with distinct topographical location [48]. For example, within the bone marrow, perivascular hBM-MSCs have been found to express CD146, an adhesion marker mostly absent in bone-lining hBM-MSCs [49]. Furthermore, CD146 expression has been found to be

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associated with conditions of hypoxia and normoxia, known to promote distinct immune-regulatory and proliferative phenotypes in hBM-MSCs [50]. Other CD surface markers increasingly associated with increased proliferative phenotype in MSCs are STRO-1 and platelet-derived growth factor receptor alpha which have been shown to be increasingly expressed in high-proliferating populations of adult human-derived MSCs [51]. Thus far, comparative studies reporting the influence of human-derived MSCs proliferation and multi-lineage differentiation potential on TME populations and cancer progression is still lacking, with new basic research on this field providing new ways to more efficiently design in vitro models. 2.4 Recognition versus Stimulation of Immunosupportive/ Immunosuppressive TA-MSCs Phenotypes

As previously mentioned, several reports have highlighted that TA-MSCs phenotype is dependent on their tissue of origin. In this regard, there is a lack of basic research pertaining to the identification of the tissue origin of TA-MSCs found within patients TME. Recently in a study by Van der Velden et al. 2018 [14], the levels of circulating MSCs attracted to prostate, gastric, colon, and sarcoma solid tumor microenvironments was analyzed. The authors found that distinct levels of circulating MSCs were found in accordance with the treatment regimen and type of cancer, with prostate cancer and sarcoma treated with radiation and chemotherapy regiments showcasing the highest fold increase (1.73 and 2.10, respectively) in the levels of circulating MSCs [14]. However, the lack of tissue-specific biomarkers currently prevents researchers to draw meaningful conclusions regarding TA-MSCs tissue of origin. Furthermore, divergences regarding theoretical TA-MSCs origin can be correlated with the lack of a consensus pertaining to circulating MSCs origin, with prevailing views stating that these can either be derived from migrating hBM-MSCs, from surrounding tissue MSCs, or from a perivascular origin [30, 52, 53]. Therefore, reports employing TME-derived TA-MSCs are increasingly crucial for the development of robust in vitro tumor models that allow researchers to better evaluate MSCs–cancer cells interactions. Moreover, current reports use isolated populations that can contain MSCs subpopulations with distinct immuneregulatory capacities, providing an increasing factor of variation towards the design of complex co-culture models. Having been established in 2006, the currently used minimal criteria for characterizing human-derived MSCs, provides no advantage for the unequivocal identification of specific immunological or secretory subtypes present within isolated populations. In a similar fashion to T cells and macrophages classification, some studies have attempted to divide TA-MSCs into stimulatory (MSC-1), and immunosuppressive (MSC-2) like populations [54, 55]. Notably, a study by Krampera et al. [54], highlighted some CD-biomarker differences between freshly isolated MSCs and

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IFNγ-primed MSCs known to exhibit a pro-inflammatory profile. The latter demonstrated an upregulation of markers such as major histocompatibility complex I (MHCI) and major histocompatibility complex II (MHCII), immune modulatory molecules (CD200, CD274), and adhesion molecules (CD54 and CD106) [54]. These MSC-1 cells were associated to the expression of IL-6, IL-8, TGF-β, and respective polarization of co-cultured macrophages towards an M1- or M2-like phenotype, as demonstrated through the analysis of macrophage expression of CD markers (e.g., CD14, HLA-DR, and CD206). The MSC-2 phenotype is correlated with the release of immunosuppressive factors, such as IL-4, IL1RA, PG1E2, which lead to macrophage polarization towards an M2-like phenotype promoting events such as immune-evasion and suppression of Treg and NK cells activity. Such a bipolar classification of TA-MSCs probably fails to fully represent the intricated setting of communications, and contradictory interactions TA-MSCs can exert on the TME. In this regard, studies providing clear markers for either MSC-1 or MSC-2 like phenotypes and possible intermediate states of activation are necessary to better characterize and predictively develop novel models around such a crucial axis of communication. Hindering the identification of TA-MSCs in vivo is the fact that these cells are present in extremely low frequencies within the TME and surrounding tissues [14, 56]. Furthermore, direct isolation of TA-MSCs is highly challenging owing to MSCs plasticity, exhibiting phenotypic variation overtime rendering them similar to other stromal cells present in advanced solid tumors [6]. Alike CAFs in tumors, certain TA-MSCs have been characterized by increased expression of HIF-1α, MMP-11, VEGF, CXCL12, PDGFα accompanying tumor progression [6]. Moreover, the expression of these specific minimal criteria CD-biomarkers is not fully restricted to MSCs populations within the TME, with for example CD105, CD90, and CD73 also being expressed by vascular populations [57], by mature stromal fibroblasts and some CAFs [58, 59]. Recently, LRRC5 has been demonstrated to be positively expressed in mesenchymal and stromal cells present in diverse advanced solid tumor niches, being overexpressed in CAFs and cancer cells of mesenchymal origin [60]. Curiously, no results regarding LRRC5 expression in TA-MSCs have thus far been reported, to the best of our knowledge. The biomarker in question could possibly be used to further separate these populations from other stromal cells if under-expressed or help distinguish tumor committed versus pristine MSCs. Overall, these observations raise the important question of MSCs specificity when designing in vitro tumor models, and the necessity of further implementing standardized protocols for mimicking tumor-specific microenvironment communication while assuring an increased reproducibility and predictability

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[61]. While the origin of TMA-MSCs and MSCs in general is still debated, their observed tissue-specific effects on tumor progression have led to further analysis of tissue-specific populations in this context, with their identifications and standardized inclusion into in vitro tumor models being of the outmost importance [43, 61]. Currently, MSCs characterization is based only on the analysis of the minimal criteria established in 2006 with few studies analyzing the expression on newly suggested markers now increasingly associated with distinct TA-MSCs phenotypes. Therefore, there is an urgent need to discover population-specific biomarkers and develop highly sensitive cell sorting assays capable of distinguishing immature/undifferentiated MSCs from committed TA-MSCs and surrounding stromal cell populations. Lastly, it is crucial to recognize that the levels of expression of a minimal set of surface biomarkers specifically chosen to distinguish them from other populations is not a guarantee of isolated MSCs homogeneity. Consequently, research is underway to perform extensive characterization of both the genomic and proteomic profiles of TME-specific MSCs. Providing a new pathway to analyze TA-MSCs established mechanisms that mediate cancer progression and tumor immune-evasion. In keeping with the trophic functions of MSCs derived from their role in tissue repair, their secretory and exosome profiles may also reveal unique biomarkers that reflect their biological properties and could potentially provide the basis for their standardized selection and inclusion into biomimetic in vitro tumor models [31]. 2.5 Characterizing MSCs Phenotype Through Secretome Analysis

TA-MSCs influence in the tumor mass extends beyond the direct contact established by these cells with surrounding components of the TME. In line with direct interactions established with adjacent cancer and stromal cells, TA-MSCs also act as a critical link of regulation and communication through the release of extracellular vesicles (e.g., exosomes) [62], cytokines, growth factors, and chemokines into their surrounding environment. For example, during tumor development TA-MSCs have been found to influence both tumor-associated macrophages and vascular cell populations, acting as a promoting force in processes related with angiogenesis through the release of pro-angiogenic growth factors and cytokines (e.g., VEGF, FGF, PDGF-α, IL-6, IL-8, TGF-β) [63]. Furthermore, it has been widely reported that TA-MSCs are capable of directly affecting the activation and proliferation of all immune system cell types via indirect communication based on the release of microvesicles, immunoregulatory growth factors, and cytokines [64]. Within the TME, TA-MSCs-derived extracellular vesicles and exosomes can affect immune cells activity in major phases of immune response by: (i) augmenting or preventing antigen recognition and presentation, for example, by endowing cancer cells with dormant phenotypes and increased expression of PD-1 [65, 66];

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(ii) interfering in immune cells communication, proliferation, activation, and effector activity, for example, in T effector cells (CD8+) to T regulator (CD4+) cell ratios within the TME of prostate and colon cancers [26, 67]. Thus far, to analyze the effect of TA-MSCs on tumor growth and in surrounding TME populations, cells were either directly co-cultured with MSCs or exposed to MSC-derived factors and/or conditioned media. While in direct co-culture bidirectional communication can occur, promoting a physiologically relevant profile in experiments where cancer cells are simply exposed to conditioned derived media, containing the bio-instructive milieu of cell-sourced secretome and vesicular elements released by MSCs. Furthermore, the composition of these conditioned media can serve as an important standpoint for the analysis of TME influence in TA-MSCs phenotype, as seen in previous studies in which the conditioned media from TME primed and non-primed MSCs was analyzed. Differences in TA-MSCs potential phenotype in accordance with MSC origin have been demonstrated [68, 69]. For example, conditioned media derived from hAT-MSCs, hUC-MSCs, and hBM-MSCs presents distinct levels of pro-angiogenic factors, resulting in distinct capacities in promoting or inhibiting such events in the TME. Moreover, as proposed by Visozo et al. 2017 [70], isolation and analysis of secretome soluble components present in conditioned media extracts and co-culture media may be performed through centrifugation, filtration, polymer precipitation-based methodologies, ion exchange chromatography, and size-exclusion chromatography, in order to better analyze and understand the composition of extracellular vesicles [70]. It is therefore of extreme importance to proceed towards the characterization of these complex secretomes, and their standardization in the context of in vitro tumor models development to assure an increased reproducibility and data cross comparison.

3

Standardization of In Vitro Culture Conditions Observations regarding MSCs plasticity and the lack of a specific set of biomarkers for their differentiation evidence the necessity of correctly examining both the diverse phenotypes and secretomes of TA-MSCs within in vitro tumor models, in an attempt to provide a physiological setting for their study. Given that MSCs are known to present diverse media requirements, as well as growth, proliferation, secretome, matrix deposition, and differentiation capabilities [13], when compared with other cellular components of the TME [71] standardization of models containing MSCs requires an extensive characterization of both MSCs phenotype and culture platforms. Furthermore, distinct variations in population ratio can lead to the release of dissimilar factors and cytokines leading to

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contrary effects [13]. As such, specific standardization of co-culture parameters must be considered when developing such models. The elevated plasticity of MSCs and their acquisition of TME-specific phenotypes is extensively affected by the previously referred factors, and by others such as the morphology and composition of the culture substrate or pre-existing scaffolds in which MSCs are laden [72]. Moreover, depending on the implemented culture ratios and co-cultured population-specific type of MSCs may present diversified TA-MSCs phenotypes [73, 74]. Such disparities introduce sources of variation in obtained results, complicating the establishment and standardization of culture procedures for complex heterotypic co-culture models. 3.1 Passaging and Variations in Proliferative and Differentiation Potential

MSCs are commonly derived from patient isolated cells, showcasing a distinct capacity for differentiation in accordance with passage and culture settings [75]. In vitro expanded MSC colonies display a progressive increase in lineage commitment as recently reviewed by Kimzcak et al. 2018 [70], demonstrating varying differentiation potential, for example, in accordance with the stiffness, and composition of the substrate in which they are seeded. Increasing evidence has been accumulated regarding the unequivocal role that variations in isolation procedures, and ex vivo manipulation conditions can exert on proliferation, differentiation, and MSCs communication with other co-cultured cells [76–79]. While MSCs are susceptible to the variations induced by parameters such as type of culture medium and associated growth-serum, initial plating density and posterior passaging frequency and density, as well as cell culture period and number of passages, they are not equally affected by them [70]. Given MSCs biological role and presence within a distinct range of in vivo microenvironments it cannot be expected that a standardized culture setting can promote the appearance of diversified specific phenotypes present in the TME [80]. For example, TA-MSCs found within hypoxic and nutrient starved microenvironments of chondrosarcoma [81, 82] will not be primed by the same environmental factors as those that influence TA-MSCs within highly irrigated pancreatic cancer microenvironments [83, 84]. As such, careful consideration must be placed on the growth conditions to which MSCs are subjected prior to inclusion within in vitro tumor models. Curiously, several reports pinpoint that growth of hBM-MSCs in low O2 environments or in serum starved xeno-free media enhances their proliferative capacity and promotes the acquisition of activated immune-regulatory phenotypes [85]. However, most experiments performed to date seem to perform MSCs culture and expansion in standard but nontissue-specific conditions, which can result in selective pressures that promote non-TME-specific phenotypes. Moreover, regarding the utilization of growth-serum, as remarked by Jon et al. 2015 [38], the same principle of selection through growth in specific

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conditions could be applied. For example, hBM-MSCs cultured in media supplemented only with platelet lysates, could promote the expansion of PDGF-β receptive MSCs expressing high levels of CD140b, normally found to be upregulated in hBM-MSCs derived from the marrow of patients with breast cancer [49]. 3.2 Influence of Culture Substrates on Mesenchymal Stem Cells Phenotype Acquisition

Within the TME cells find themselves closely associated with a surrounding extracellular matrix composed by meshes of diverse biomacromolecules associated in distinct ratios, density and alignments specific to the tissue or tumor stage in question [86– 88]. Tumor progression commonly characterized by alterations in the composition, microarchitecture and mechanical properties of tumor extracellular matrix, is now increasingly recognized to actively contribute to the proliferation, motility, epithelial-to-mesenchymal transition, and invasion of cancer cells [89–91]. Changes to tumor ECM act as a positive feed-back loop in tumor promotion, being triggered by altered regulatory and secretory profiles of the stromal cells present within the TME. These alterations act themselves as promoters of pro-tumoral phenotype acquisition in TME populations. MSCs have both been found to be regulators and regulated by the alterations that tumor microenvironment ECM suffers over the course of tumor progression. Thorough examination of TA-MSCs and ECM exchanges has provided a new outlook on the complex dynamics of the TME, with promise of yielding novel therapeutic targets [75, 92, 93]. Therefore, careful implementation of biomimetic ECMs and substrates in which both MSCs expansion and models’ development occurs is necessary. As previously reviewed, the development of novel in vitro preclinical validation models is shifting towards the implementation of complex 3D-based culture systems [94–96]. Translated from the field of Tissue Engineering and Regenerative Medicine (TERM), these platforms provide complex 3D microenvironments in which cells can fully communicate and interact in a spatially defined biomimetic environment [71]. 3D in vitro tumor models also serve as vehicles for the inclusion and study of TME-mimetic ECM components, providing valuable insights about the interactions that such extracellular components of the TME can have in tumor progression and in TA-MSCs phenotype acquisition. The diverse plethora of methodologies that can be applied for the production of such models has been recently reviewed elsewhere and extends beyond the scope of this chapter [97–104]. In regard to scaffold-based 3D co-cultured models, thorough examination of the interactions and stimuli these scaffolds can have on MSCs behavior is crucial for the interpretation and comparison of such models [13]. Naive mesenchymal stem cells have been reported to commit to specific lineages and phenotypes in accordance with the levels of elasticity and stiffness of surrounding tissue.

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[105] While in the initial stages of post-isolation culture these cells can readily be reprogramed towards a specific lineage phenotype through the addition of soluble induction factors, over extended periods of culture cells start to commit and differentiate gradually to a lineage specifically promoted by the stiffness and elasticity of the subtract in which they are cultured [106]. Commonly employed 2D-based methods of MSCs culture rely in polystyrene-based substrates coated with small amounts of bioactive proteins (e.g., fibronectin). These surfaces poorly mimic the visco-elastic properties and stiffness of a living tissue, presenting instead an elevated elastic modulus (500 MPa to 3 GPa), several orders of magnitude higher than those of soft-tissues [107]. Moreover, as demonstrated in recent studies using collagen coated substrates the alignment and stiffness of these bioactive proteins can lead to distinct shifts in the differentiation potential of MSCs. For example, hBM-MSCs have been found to express increased osteogenic markers when cultured in stiffer collagen matrix substrates (25–40 kPa), and neurogenic markers when cultured in low-stiffness collagen (0.1–1 kPa) [105]. Furthermore, in a study by Ishihara et al. (2018), hBM-MSCs were found to promote breast cancer cells proliferation when cultured in stiffer substrates (20–40 kPa), furthermore exhibiting CAF-associated markers of differentiation [108]. Diverse types of materials can now be implemented to fabricate scaffolds used for in vitro 3D tumor models development. Extending from synthetic derived materials such as polycaprolactone or poly-L-lactic acid to naturally derived materials such as collagen or ECM-derived hydrogels, to functionalized polysaccharides and biopolymers such as alginate, hyaluronic acid, or chitosan [35, 109–115]. The majority of studies directed at analyzing how MSCs can interact with other distinct TME populations towards the alteration of TME-ECM have focused mainly in co-culture platforms assembled based on biomimetic scaffolds such as collagen I, Matrigel™, silk-fibroin, alginate, and hyaluronic acid scaffolds. Such a plethora of biomaterials allows for the development of highly diverse biomimetic substrates with varying customizable stiffness, alignment, and visco-elastic properties. Moreover, the selection of the matrix in which MSCs are cultured must consider the bio-instructive and signaling capacities of certain ECM-derived substrates [116]. Studies have shown that culture of MSCs in soft collagen and Matrigel hydrogels can effectively increase the expression of markers associated with wound-healing mechanisms [117]. However, bioactive hydrogels such as Matrigel provide an added level of complexity, derived from the existence of bound-growth factors which like growth-serum can prime and severely influence MSCs behavior [116]. Moreover, as recently reported by Loebel et al. 2019, variations in hydrogels mechanobiological properties in which MSCs are cultured such as

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differences in adhesion moieties, alignment, material stressrelaxation profiles, and stiffness can not only promote distinct morphologies and internal signaling mechanisms but furthermore promote distinct ECM secretory profiles in MSCs [118]. In their study, Loebel et al. demonstrated that in stiffer hydrogel networks (~20 kPa) MSCs deposition of nascent ECM proteins, responsible for facilitating cell adhesion and promoting cell spreading, was lower when compared to softer hydrogels (~3 kPa). Furthermore, the authors observed that the inhibition of cellular adhesion to fibronectin domains of nascent proteins, conjugated with a decreased fibronectin deposition and altered fibril formation by MSCs, leading to a decreased osteogenic differentiation and a shift towards adipogenic differentiation [118]. The use of these bioactive substrates therefore requires careful control and analysis of obtained MSCs phenotypes, with associated batch-to-batch variability straining direct comparisons of obtained results [119]. 3.3 Defining Co-Culture Ratios and Analyzing Overtime Population Variations

As previously stated, our knowledge regarding the exact composition and levels of TA-MSCs present within the microenvironment of solid tumors is wanting [86, 120, 121]. The lack of basic research studies capable of answering fundamental questions, such as the point-of-migration in the TME, time, or period during which MSCs translocate to the TME, amount of TME-associated TA-MSCs in accordance to type of tumor and stage, and lastly phenotype of TA-MSCs, has prevented the in vitro implementation of physiologically validated cellular ratios in co-culture models [8, 13]. In this regard, a severe problem arises when considering that the amount of MSCs present within the model can lead to diverse, sometimes contrary, responses, and interactions with cancer cells and that co-cultured populations can exhibit distinct proliferation ratios [11, 109, 115, 122, 123]. Alterations on population ratios can lead to confounding results, derived for example from altered ECM degradation or deposition, differentiation of MSCs towards distinct cell phenotypes, and even from cellin-cell mechanisms that can lead to the abrogation of present MSCs populations. Regarding cell-in-cell activity MSCs are particularly susceptible to the action of cancer cells, with diverse reports showcasing fusion and cannibalism mechanisms in the TME of prostate and breast cancers, respectively [124, 125]. These phenomena lead to the establishment of chimeric cancer cells capable of expressing stem-like characteristics that allow them to more easily evade immune system recognition, and/or enter states of dormancy associated with cancer resurgence [124, 125]. Consequently, alterations in cultured cellular populations must be carefully monitored, creating the necessity of developing standardized high-throughput compatible cell-tracking methodologies capable of reporting cellular population variation and differentiation within direct co-cultured models [13]. As previously discussed

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in detail, immunostaining-based techniques have been used to a certain success in differentiating TA-MSCs from other TME populations within tumor models [126, 127]. However due to the appearance of overlapping receptors, in part from the membrane exchange mechanisms associated with TA-MSCs and cancer cells direct communication, current markers are unable to fully differentiate between distinct TA-MSCs subpopulations and TA-MSCsderived cells. Alternatively, the used cell-tracking techniques based in cellular transfection with non-integrative fluorescent protein reporter genes, (e.g., GFP, YFP, RFP) via viral or non-viral transfection vectors has been extensively employed [128, 129]. Moreover, development of multi-color fluorescence reporter sequences (dTomato, CFP, or YFP) correlated with the expression of specific proteins and other cellular lineage-specific reporting genes will undoubtedly provide new pathways to study TME TA-MSCs differentiation and integration [130, 131]. However, currently such methodologies still imply elevated associated costs and timeconsuming protocols, with the need for more affordable and easier to perform protocols such as stable membrane labeling probes [132], providing an alternative route for cell-tracking within co-culture TME models.

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Conclusion Establishing complex in vitro tumor models capable of replicating the role TA-MSCs exert in the TME is not a trivial pursuit. Given the elevated variability of this cell class, the lack of tissue and subpopulation-specific biomarkers, standardized growth conditions in accordance with intended application and tissue of origin, and other discussed factors, the comparison of MSCs containing in vitro tumor models must be made with careful reflection. The needed for implementation of higher characterization of used MSC populations and reproducible culture conditions becomes increasingly apparent through the analysis of the elevated number of conflicting reports regarding TA-MSCs role in the TME. Such conflicting views can derive not only from the inherent plasticity of MSCs but more importantly from the lack of GMP compliant methodologies, such as those implemented in the study and analysis of immune cell interactions in disease models. Moreover, the overall difficulty in tracking co-cultured cellular populations hinders the ability to use more complex models for the analysis of specific cellular interactions. The main points of variation associated with MSC inclusion are: (i) different initial seeding ratios lead to distinct phenotype and secretomes; (ii) the spatiotemporal manner in which cells are introduced to the model can lead to diverse morphological organizations; (iii) the diverse tissue-specific and subpopulation-specific MSCs presenting diverse proliferative

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rates; (iv) population phenotype varies in accordance to passages; and (v) the requirement of different types of culture media and associated supplements, which can increase variability. In summary, the establishment of standardized practices for the development of in vitro MSC-cancer co-culture testing platforms requires further study and optimization of analysis methodologies focused at tackling the several challenges unique to MSCs inclusion. References 1. Maman S, Witz IP (2018) A history of exploring cancer in context. Nat Rev Cancer 18:359–376. https://doi.org/10.1038/ s41568-018-0006-7 2. Bolm L, Cigolla S, Wittel UA et al (2017) The role of fibroblasts in pancreatic cancer: extracellular matrix versus paracrine factors. Transl Oncol 10:578–588. https://doi.org/10. 1016/j.tranon.2017.04.009 3. Karamanos NK, Theocharis AD, Neill T, Iozzo RV (2019) Matrix modeling and remodeling: a biological interplay regulating tissue homeostasis and diseases. Matrix Biol 75–76:1–11. https://doi.org/10.1016/j. matbio.2018.08.007 4. Breslin S, O’Driscoll L (2013) Threedimensional cell culture: the missing link in drug discovery. Drug Discov Today 18:240–249. https://doi.org/10.1016/j. drudis.2012.10.003 5. Begley CG, Ellis LM (2012) Drug development: raise standards for preclinical cancer research. Nature 483:531–533. https://doi. org/10.1038/483531a 6. Sacchetti B, Funari A, Remoli C et al (2016) No identical “mesenchymal stem cells” at different times and sites: human committed progenitors of distinct origin and differentiation potential are incorporated as adventitial cells in microvessels. Stem Cell Rep 6:897–913. https://doi.org/10.1016/j.stemcr.2016.05. 011 7. Naji A, Eitoku M, Favier B et al (2019) Biological functions of mesenchymal stem cells and clinical implications. Cell Mol Life Sci. https://doi.org/10.1007/s00018-01903125-1 8. Timaner M, Tsai KK, Shaked Y (2019) The multifaceted role of mesenchymal stem cells in cancer. Semin Cancer Biol:0–1. https://doi. org/10.1016/j.semcancer.2019.06.003 9. van Schaijik B, Wickremesekera AC, Mantamadiotis T et al (2019) Circulating tumor stem cells and glioblastoma: a review. J Clin

Neurosci 61:5–9. https://doi.org/10.1016/ j.jocn.2018.12.019 10. Norozi F, Ahmadzadeh A, Shahrabi S et al (2016) Mesenchymal stem cells as a doubleedged sword in suppression or progression of solid tumor cells. Tumor Biol 37:11679–11,689. https://doi.org/10. 1007/s13277-016-5187-7 11. Poggi A, Varesano S, Zocchi MR (2018) How to hit mesenchymal stromal cells and make the tumor microenvironment immunostimulant rather than immunosuppressive. Front Immunol 9:1–17. https://doi.org/10. 3389/fimmu.2018.00262 12. Eid JE, Garcia CB (2015) Reprogramming of mesenchymal stem cells by oncogenes. Semin Cancer Biol 32:18–31. https://doi.org/10. 1016/j.semcancer.2014.05.005 13. Ferreira LP, Gaspar VM, Henrique R et al (2017) Mesenchymal stem cells relevance in multicellular bioengineered 3D in vitro tumor models. Biotechnol J 12:1700079. https:// doi.org/10.1002/biot.201700079 14. van der Velden DL, Houthuijzen JM, Roodhart JML et al (2018) Detection of endogenously circulating mesenchymal stem cells in human cancer patients. Int J Cancer:1–18. https://doi.org/10.1002/ijc.31727 15. Ridge SM, Sullivan FJ, Glynn SA (2017) Mesenchymal stem cells: key players in cancer progression. Mol Cancer 16:31. https://doi. org/10.1186/s12943-017-0597-8 16. Baglio SR, Lagerweij T, Pe´rez-Lanzo´n M et al (2017) Blocking tumor-educated MSC paracrine activity halts osteosarcoma progression. Clin Cancer Res 23:3721–3733. https://doi. org/10.1158/1078-0432.CCR-16-2726 17. Ishida Y, Kido A, Akahane M et al (2018) Mesenchymal stem cells up-regulate the invasive potential of prostate cancer cells via the eotaxin-3/CCR3 axis. Pathol Res Pract 214:1297–1302. https://doi.org/10.1016/ j.prp.2018.06.012

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Chapter 2 Pluripotent Stem Cells for Cell Therapy Insa S. Schroeder Abstract In an increasingly geriatric population, in which elderly people frequently face chronic diseases and degenerative conditions, cell therapies as part of novel regenerative medicine approaches are of great interest. Even though today’s cell therapies mostly rely on adult stem cells like the mesenchymal stem cells or primary somatic cells, pluripotent stem cells represent an enormously versatile cell model to explore possible new avenues in the field of regenerative medicine due to their capacity to grow indefinitely and to differentiate into the desired cell types. The discovery of reprogramming somatic cells into induced pluripotent stem cells augmented the pool of applicable cell entities so that researchers nowadays can resort to embryonic stem cells, but also to a plethora of patient- and disease-specific induced pluripotent stem cells. The ease of targeted genome engineering is an additional benefit that allows using pluripotent stem cells for disease modeling, drug discovery, and the development of cell therapies. However, the task is still demanding as the generation of subpopulations and a sufficient cell maturation for some cell entities have yet to be achieved. Likewise, even though for some applications the cells of interest can be produced in the large-scale dimensions and purity that are required for clinical purposes, proper integration, and function in the host tissue remain challenging. Nonetheless, the immense progress that has been made over the last decades warrants the prominent role of pluripotent stem cells in regenerative medicine as in vitro models to broaden our knowledge of disease onset/progression and treatment as well as in vivo as a substitution of damaged/aged tissue. Key words Embryonic stem cells, Induced pluripotent stem cells, Differentiation, Cell therapy, Clinical translation

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Introduction Pluripotent stem cell-based (PSC) in vitro approaches may be complex, however, they offer a great advantage to primary cells/ tissues or adult stem cells due to their unlimited proliferation capacity and their ability to form all cell types of the body. Ever since the advent of reprogramming techniques and the generation of induced pluripotent stem cells (iPSC), this field has gained even more interest as now patient/disease-specific PSC allow mimicking the onset and progression of many diseases and will lead to the development of potent drugs for intervention. Disease modeling

Peggy Stock and Bruno Christ (eds.), In Vitro Models for Stem Cell Therapy: Methods and Protocols, Methods in Molecular Biology, vol. 2269, https://doi.org/10.1007/978-1-0716-1225-5_2, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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and subsequent treatment with cell replacement therapies requires that a couple of prerequisites are fulfilled. Firstly, the in vitro generated target cells have to resemble those in vivo in their characteristics and function. Secondly, pure populations of the target cells have to be generated as contaminating cells could mask the response of those target cells. Thirdly, especially those disorders with genetic backgrounds require the use of isogenic controls, meaning repairing mutations in patient-derived cells or generating the mutation in wild-type PSC or the use of several patient-specific iPSC to ensure that the observed phenotype results from the disease-specific genetic alteration and truthfully resembles the in vivo situation. Finally, all differentiation protocols and studies initiated as basic research have to be translated into clinical settings requiring upscaling, the use of clinical grade components, automation/standardization of laboratory procedures, rigid quality control, and many more. All of these issues are addressed by the research community and despite the increasing demands, there are promising pre-clinical and clinical PSC-based studies ongoing for major health issues such as cardiovascular and CNS-related diseases, macular degeneration and diabetes, which will be discussed.

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Use of PSC in Modeling and Treating Cardiovascular Disease Cardiovascular disease is one of the main health issues worldwide and amounts to approximately one third of all deaths globally [1]. This is due to the increasingly aging population but also because the heart is one of the organs with finite regenerative capacity. Treatment options are still suboptimal, delaying disease progression by treating the symptoms rather than offering a real cure. Taken together with the limited availability of transplantable donor tissue, the cardiologic field would benefit most by PSC-based drug discovery and cell therapies. Accordingly, a number of differentiation protocols have been established that allow generating various subtypes of cardiac cells such as ventricular [2, 3] and atrial cardiomyocytes [4–7], sinoatrial node cells [8– 10], and epicardium [11–13]. However, as is the case with most of the PSC-derived cells, the newly generated cardiac entities rather resemble fetal developmental stages and sufficient maturation remains a challenge. The issue, however, has been addressed by several groups taking into account the metabolic changes that occur upon birth and using electromechanical forces indicative for the heart. While the fetal heart relies on anaerobic glycolysis to meet its energy demand, this changes after birth, when hemodynamic shifts occur due to the expansion of the lungs and the cardiac workload increases dramatically [14]. Both incidences require the maturation and reorganization of the cardiac tissue and a change to oxidative

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metabolism to meet the high-energy demand of an adult heart. Consequently, the heart uses all energy sources, carbohydrates, lipids, proteins, and lactate, to assure proper function. As these metabolic alterations are linked to hormonal changes, several groups added thyroid and steroid hormones to their differentiation protocols [15], reduced the glucose concentration in the media [16], treated cardiac progenitor cells with fatty acids [17], or used a combination of all factors mentioned above [18]. A rather old approach is the use of lactate as a selecting measure to obtain mature cardiomyocytes [19]. With a variety of differentiation protocols at hands, cardiac disease modeling and drug discovery has become a possibility and alternative to mouse models that differ in heart rate and calcium and potassium currents contributing to cardiac action potential. Thus, ion channelopathies like the long QT syndrome were the first to be analyzed using patient-derived iPSC carrying the heterozygous missense mutation R190Q in the KCNQ1 gene [20]. Cardiomyocytes derived from these iPSC faithfully showed the symptoms of the long QT syndrome such as a prolonged action potential duration and arrhythmia. Many others including those, which model arrhythmogenic right ventricular cardiomyopathy, catecholaminergic polymorphic ventricular tachycardia, and dilated cardiomyopathy, have followed this first study. Additionally, multifactorial cardiac diseases such as atrial fibrillation can be modeled [21] and Shafaattalab et al. [22] were able to show the atrial-specific toxicity of ibrutinib, a tyrosine kinase inhibitor that is used to treat B cell cancers and frequently causes atrial fibrillation. First attempts to transplant human PSC-derived cardiomyocytes into guinea pigs resulted in the integration with the host myocardium in a myocard infarction model [23]. Later several groups used non-human primate models of myocard infarction. However, electrical coupling to the host myocardium resulted in ventricular tachyarrhythmia that was caused by ectopic pacemaker activity from the graft [24, 25]. The most similar study to the human cardiac physiology is one transplanting PSC-derived cardiomyocytes into a pig myocard infarction model [26]. Here as well the substantial engraftment of human cardiomyocytes within the infarcted scar tissue was accompanied by tachyarrhythmia for several weeks after transplantation. Such side effects were not seen in the only in-man trial, in which a small number of cardiac progenitor cells were transplanted into six patients with left ventricular systolic dysfunction [27]. However, the observed improved function was likely due to paracrine effects rather than repopulation and engraftment of the transplant.

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Use of PSC in Modeling and Treating CNS Diseases CNS-related cell therapies are envisioned for spinal cord injuries, stroke, traumatic brain injury, or degenerative diseases such as Parkinson and Alzheimer’s. Presently, most of the clinical trials use bone marrow or umbilical cord-derived mesenchymal stem cells. However, one Parkinson disease-related study using human embryonic stem cells (hESC)-derived neural precursor cells is performed at “The first affiliated hospital of Zhengzhou University,” Zhengzhou, Henan, China (“Safety and Efficacy Study of Human ESC-derived Neural Precursor Cells in the Treatment of Parkinson’s Disease,” ClinicalTrials.gov Identifier: NCT03119636). This Phase I/II, open-label, non-randomized clinical trial is supposed to enroll 50 patients for cell injection, administering a single dose of neural precursor cells by stereotaxic intra-striatal injection. However, no results have been published to date. Certainly, the most prominent clinical trial in the field of CNS diseases was conducted by the California-based company Geron attempting to treat spinal cord injury using hESC-derived oligodendrocyte progenitor cells (GRNOPC1 cells, ClinicalTrials.gov Identifier: NCT01217008). Five patients were treated with one injection of 2 million GRNOPC1 cells. Even though the trial was terminated in 2011 for economic reasons and a change in the company’s portfolio, a follow-up revealed that the HLA-mismatched cell transplants were well tolerated even after termination of an immunosuppressive regimen. However, there was no significant improvement of neurological function (see review [28]). The study, however, has been resumed by Asterias Biotherapeutics, Inc. (SCiStar study, ClinicalTrials.gov Identifier: NCT02302157) escalating the administered dose to up to 2 million cells in five cohorts of a total of 25 patients with subacute motor complete (AIS-A or AIS-B) cervical (C-4 to C-7) spinal cord injury. No adverse effects were reported. Instead, at 6 months, 15 out of 18 participants recovered at least 1 motor level on at least one side and 4 out of 18 subjects recovered two or more motor levels on at least one side (https:// www.globenewswire.com/news-release/2018/02/28/1401685 /0/en/Asterias-Provides-Update-for-its-AST-OPC1-Phase-1-2aClinical-Trial-in-Severe-Spinal-Cord-Injury.html, accessed November 27, 2019). The scarcity of CNS-related clinical trials involving PSC-derived cells is in opposition to the huge number of in vitro approaches to generate complex brain-like tissues and CNS cells from human PSC. Here, the generation of organoids, selfassembling 3D aggregates, have greatly improved our ability to model complex CNS disorders. In contrast to neurospheres, that also consist of various types of CNS cells, brain organoids mirror the cellular architecture and the developmental trajectories seen

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in vivo (reviewed in [29]). Guided differentiation that uses small molecules and growth/differentiation factors force PSC to generate brain region-specific organoids resembling the hippocampus [30], fore- or midbrain [31, 32], cerebellum [33], or the spinal cord [34] while unguided approaches or minimal guidance generally lead to whole brain/cerebral cortex organoids [35–38]. The region-specific organoids can be fused enabling researchers to examine interactions, e.g., cell migration, between those regions [39]. “Fusion-organoids” or hybrid organoids can also be used to introduce other cell types such as endothelial cells [40] making it a perfect modular system with the architecture and complexity of real tissue. Another benefit is the possibility of long-term culturing (>100d, [41]), which allows organoid studies that can be easily compared to those performed in rodents. Organoids have also been generated to model a broad range of brain disorders including microcephaly, Alzheimer’s disease, autism spectrum disorder, or even brain tumors and have been reviewed comprehensively elsewhere [42].

4

Use of PSC in Modeling and Treating Retinal Diseases PSC have been most successfully introduced in clinical trials as the source of retinal cells treating macular degeneration. As the eye is an immune privileged site and can be easily monitored noninvasively, it facilitates even allogeneic stem cell-based therapies as possible side effects such as immune rejection do not occur or can be diagnosed and counteracted immediately. Masayo Takahashi and co-workers from the RIKEN Center for Developmental Biology paved the way for the use of iPSC in this field addressing protocol improvements to generate retinal pigment epithelium (RPE) from allogeneic and autologous iPSC sources [43], the automation of the RPE generation for clinical use [44], tumorigenicity tests for iPSC and their derivatives [45], and critical handling procedures like storage and shipping [46, 47]. Two patients were initially enrolled in a study for autologous iPSC-derived RPE transplantation of whom one received the RPE sheet in 2014. Just recently, the 4-year followup was published [48] showing that the graft did survive and support photoreceptors and choroidal vessels. As vision acuity remained stable, this patient report proofs the concept of a feasible and safe procedure. The abandonment of the treatment of the second enrolled patient reflects the tight supervision and awareness of safety issues related with the use of iPSC-derived cells: In his case, one of the generated iPSC lines expressed copy number variations that were not found in the original patient’s fibroblasts used for the reprogramming [49]. Even though a tumorigenicity of the RPE sheets could not be detected, the genetic variations (three deletions, of which one was found on the X chromosome) might have

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had an effect on the affected genes and their flanking regions. Due to the fact, that one of the deletions affected the only X chromosome of the male patient and his ability to still respond moderately to the standard anti-VEGF therapy, the patient was not treated. As the generation of autologous iPSC and their derivatives is immensely time-consuming and cost-intensive, allogeneic HLA-matched cells are favorable and the first patient, who suffered from wet-type age-related macular degeneration, was treated with such iPSC-derived cells on March 28, 2017 (http://www.cdb. riken.jp/en/news/2017/ topics/0404_10343.html, accessed November 30, 2019). In addition to the above-mentioned iPSC-based studies, at present 19 clinical trials are listed at ClinicalTrials.gov using hESC-derived RPE for the same purpose of which 6 have been completed (https://clinicaltrials.gov using the search terms human embryonic stem cell and macular degeneration, accessed November 30, 2019).

5

Use of PSC in Modeling and Treating Diabetes Mellitus Certainly, one of the most serious health concerns is the number of patients with diabetes mellitus, type 1 and type 2. Even though there are many treatment options available, certain types of labile diabetes may require cell replacement therapies that cannot be fully met by providing and transplanting donor tissue. Thus, there has been numerous attempts in the last decades to generate insulinproducing beta-like cells from PSC. Even though such protocols are very elaborate [50], they lead to mono-hormonal insulinexpressing cells that can be distinguished from poly-hormonal cells and selected by the surface marker glycoprotein 2 [51]. Such protocols also have been established from type 1 diabetes patients [52, 53]. As encapsulation methods have also been improved, there are meanwhile four clinical trials sponsored by Viacyte to test the safety and efficacy of pancreatic islet progenitor cells (PEC-01 cells) in type 1 patients with extreme glycemic lability and/or recurrent severe hypoglycemic episodes (ClinicalTrials.gov identifiers NCT03162926, NCT02239354, NCT03163511, and NCT02939118).

6

Conclusion Clinical use of PSC-based products is still in an early phase and faces many challenges that makes it a high risk and cost-intensive research and development field. Collaborative/interdisciplinary effort from industry, academia as well as the national regulatory authorities such as the U. S. Food and Drug Administration and the

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European Medicines Agency will be required to meet the needs and to ensure a successful outcome for an increasing number of patients. However, the progress that has been made in differentiating PSC and identifying disease-related mechanisms/pathways using PSC and their progeny warrants this collaborative effort and will be beneficial to the medical field in due time.

Acknowledgments I.S.S. is funded by the German Federal Ministry of Education and Research (BMBF, support code 02NUK049A). References 1. Joseph P, Leong D, McKee M et al (2017) Reducing the global burden of cardiovascular disease, part 1: the epidemiology and risk factors. Circ Res 121:677–694 2. Lee JH, Protze SI, Laksman Z et al (2017) Human pluripotent stem cell-derived atrial and ventricular cardiomyocytes develop from distinct mesoderm populations. Cell Stem Cell 21:179–194 3. Zhang JZ, Termglinchan V, Shao NY et al (2019) A human iPSC double-reporter system enables purification of cardiac lineage subpopulations with distinct function and drug response profiles. Cell Stem Cell 24:802–811 4. Devalla HD, Schwach V, Ford JW (2015) Atrial-like cardiomyocytes from human pluripotent stem cells are a robust preclinical model for assessing atrial-selective pharmacology. EMBO Mol Med 7:394–410 5. Laksman Z, Wauchop M, Lin E et al (2017) Modeling atrial fibrillation using human embryonic stem cell-derived atrial tissue. Sci Rep 7:5268 6. Lemme M, Ulmer BM, Lemoine MD et al (2018) Atrial-like engineered heart tissue: an in vitro model of the human atrium. Stem Cell Reports 11:1378–1390 7. Lemme M, Ulmer BM, Lemoine MD et al (2018) Atrial-like engineered heart tissue: an in vitro model of the human atrium. Stem Cell Rep 11:1378–1390 8. Birket MJ, Ribeiro MC, Verkerk AO et al (2015) Expansion and patterning of cardiovascular progenitors derived from human pluripotent stem cells. Nat Biotechnol 33:970–979 9. Liang W, Han P, Kim EH et al (2019) Canonical Wnt signaling promotes pacemaker cell specification of cardiac mesodermal cells derived from mouse and human embryonic

stem cells. Stem Cells 38:352. https://doi. org/10.1002/stem.3106 10. Schweizer PA, Darche FF, Ullrich ND et al (2017) Subtype-specific differentiation of cardiac pacemaker cell clusters from human induced pluripotent stem cells. Stem Cell Res Ther 8:229 11. Gambardella L, McManus SA, Moignard V et al (2019) BNC1 regulates cell heterogeneity in human pluripotent stem cell derivedepicardium. Development 146:dev174441. https://doi.org/10.1242/dev.174441 12. Guadix JA, Orlova VV, Giacomelli E et al (2017) Human pluripotent stem cell differentiation into functional epicardial progenitor cells. Stem Cell Rep 9:1754–1764 13. Iyer D, Gambardella L, Bernard WG et al (2016) Robust derivation of epicardium and its differentiated smooth muscle cell progeny from human pluripotent stem cells. Development 143:904 14. Piquereau J, Ventura-Clapier R (2018) Maturation of cardiac energy metabolism during perinatal development. Front Physiol 9:959 15. Parikh SS, Blackwell DJ, Gomez-Hurtado N et al (2017) Thyroid and glucocorticoid hormones promote functional T-tubule development in human-induced pluripotent stem cellderived cardiomyocytes. Circ Res 121:1323–1330 16. Correia C, Koshkin A, Duarte P et al (2017) Distinct carbon sources affect structural and functional maturation of cardiomyocytes derived from human pluripotent stem cells. Sci Rep 7:8590 17. Horikoshi Y, Yan Y, Terashvili M et al (2019) Fatty acid-treated induced pluripotent stem cell-derived human cardiomyocytes exhibit adult cardiomyocyte-like energy metabolism

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phenotypes. Cell 8(9):1095. https://doi.org/ 10.3390/cells8091095 18. Lin B, Lin X, Stachel M et al (2017) Culture in glucose-depleted medium supplemented with fatty acid and 3,30 ,5-triiodo-l-thyronine facilitates purification and maturation of human pluripotent stem cell-derived cardiomyocytes. Front Endocrinol (Lausanne) 8:253 19. Kadari A, Mekala S, Wagner N et al (2015) Robust generation of cardiomyocytes from human iPS cells requires precise modulation of BMP and WNT signaling. Stem Cell Rev Rep 11:560–569 20. Moretti A, Bellin M, Welling A et al (2010) Patient-specific induced pluripotent stem-cell models for long-QT syndrome. N Engl J Med 363:1397–1409 21. Benzoni P, Campostrini G, Landi S et al (2019) Human iPSC modeling of a familial form of atrial fibrillation reveals a gain of function of if and ICaL in patient-derived cardiomyocytes. Cardiovasc Res 116:1147. https://doi.org/ 10.1093/cvr/cvz217 22. Shafaattalab S, Lin E, Christidi E (2019) Ibrutinib displays atrial-specific toxicity in human stem cell-derived cardiomyocytes. Stem Cell Rep 12:996–1006 23. Shiba Y, Fernandes S, Zhu WZ et al (2012) Human ES-cell-derived cardiomyocytes electrically couple and suppress arrhythmias in injured hearts. Nature 489:322–325 24. Liu YW, Chen B, Yang X et al (2018) Erratum: human embryonic stem cell-derived cardiomyocytes restore function in infarcted hearts of non-human primates. Nat Biotechnol 36:899 25. Liu YW, Chen B, Yang X et al (2018) Human embryonic stem cell-derived cardiomyocytes restore function in infarcted hearts of non-human primates. Nat Biotechnol 36:597–605 26. Romagnuolo R, Masoudpour H, PortaSanchez A et al (2019) Human embryonic stem cell-derived cardiomyocytes regenerate the infarcted pig heart but induce ventricular tachyarrhythmias. Stem Cell Rep 12:967–981 27. Menasche P, Vanneaux V, Hagege A et al (2018) Transplantation of human embryonic stem cell-derived cardiovascular progenitors for severe ischemic left ventricular dysfunction. J Am Coll Cardiol 71:429–438 28. Martin U (2017) Therapeutic application of pluripotent stem cells: challenges and risks. Front Med (Lausanne) 4:229 29. Benito-Kwiecinski S, Lancaster MA (2019) Brain organoids: human neurodevelopment in a dish. Cold Spring Harb Perspect Biol 12:

a035709. https://doi.org/10.1101/ cshperspect.a035709 30. Sakaguchi H, Kadoshima T, Soen M et al (2015) Generation of functional hippocampal neurons from self-organizing human embryonic stem cell-derived dorsomedial telencephalic tissue. Nat Commun 6:8896 31. Qian X, Nguyen HN, Song MM et al (2016) Brain-region-specific organoids using minibioreactors for modeling ZIKV exposure. Cell 165:1238–1254 32. Qian X, Jacob F, Song MM et al (2018) Generation of human brain region-specific organoids using a miniaturized spinning bioreactor. Nat Protoc 13:565–580 33. Muguruma K, Nishiyama A, Kawakami H et al (2015) Self-organization of polarized cerebellar tissue in 3D culture of human pluripotent stem cells. Cell Rep 10:537–550 34. Meinhardt A, Eberle D, Tazaki A et al (2014) 3D reconstitution of the patterned neural tube from embryonic stem cells. Stem Cell Rep 3:987–999 35. Lancaster MA, Renner M, Martin CA et al (2013) Cerebral organoids model human brain development and microcephaly. Nature 501:373–379 36. Lancaster MA, Knoblich JA (2014) Generation of cerebral organoids from human pluripotent stem cells. Nat Protoc 9:2329–2340 37. Mansour AA, Goncalves JT, Bloyd CW et al (2018) Erratum: an in vivo model of functional and vascularized human brain organoids. Nat Biotechnol 36:772 38. Mansour AA, Goncalves JT, Bloyd CW (2018) An in vivo model of functional and vascularized human brain organoids. Nat Biotechnol 36:432–441 39. Xiang Y, Yoshiaki T, Patterson B (2018) Generation and fusion of human cortical and medial ganglionic Eminence brain organoids. Curr Protoc Stem Cell Biol 47(1). https://doi. org/10.1002/cpsc.61 40. Song L, Yuan X, Jones Z et al (2019) Assembly of human stem cell-derived cortical spheroids and vascular spheroids to model 3-D brain-like tissues. Sci Rep 9:5977 41. Yoon SJ, Elahi LS, Pasca AM et al (2019) Reliability of human cortical organoid generation. Nat Methods 16:75–78 42. Amin ND, Pasca SP (2018) Building models of brain disorders with three-dimensional organoids. Neuron 100:389–405 43. Kamao H, Mandai M, Okamoto S et al (2014) Characterization of human induced pluripotent stem cell-derived retinal pigment

Pluripotent Stem Cells for Cell Therapy epithelium cell sheets aiming for clinical application. Stem Cell Rep 2:205–218 44. Matsumoto E, Koide N, Hanzawa H et al (2019) Fabricating retinal pigment epithelial cell sheets derived from human induced pluripotent stem cells in an automated closed culture system for regenerative medicine. PLoS One 14:e0212369 45. Kawamata S, Kanemura H, Sakai N et al (2015) Design of a tumorigenicity test for induced pluripotent stem cell (iPSC)-derived cell products. J Clin Med 4:159–171 46. Hori K, Kuwabara J, Tanaka Y (2019) A simple and static preservation system for shipping retinal pigment epithelium cell sheets. J Tissue Eng Regen Med 13:459–468 47. Kitahata S, Tanaka Y, Hori K et al (2019) Critical functionality effects from storage temperature on human induced pluripotent stem cellderived retinal pigment epithelium cell suspensions. Sci Rep 9:2891 48. Takagi S, Mandai M, Gocho K et al (2019) Evaluation of transplanted autologous induced

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pluripotent stem cell-derived retinal pigment epithelium in exudative age-related macular degeneration. Ophthalmol Retina 3:850–859 49. Mandai M, Watanabe A, Kurimoto Y et al (2017) Autologous induced stem-cell-derived retinal cells for macular degeneration. N Engl J Med 376:1038–1046 50. Pagliuca FW, Millman JR, Gurtler M et al (2014) Generation of functional human pancreatic beta cells in vitro. Cell 159:428–439 51. Cogger KF, Sinha A, Sarangi F et al (2017) Glycoprotein 2 is a specific cell surface marker of human pancreatic progenitors. Nat Commun 8:331 52. Millman JR, Xie C, Van DA et al (2016) Corrigendum: generation of stem cell-derived betacells from patients with type 1 diabetes. Nat Commun 7:12,379 53. Millman JR, Xie C, Van DA (2016) Generation of stem cell-derived beta-cells from patients with type 1 diabetes. Nat Commun 7:11,463

Part II Disease Modeling

Chapter 3 In Vitro Methods for the Study of Glioblastoma Stem-Like Cell Radiosensitivity Joseph H. McAbee, Charlotte Degorre-Kerbaul, and Philip J. Tofilon Abstract Ionizing radiation is a critical component of glioblastoma (GBM) therapy. Recent data have implicated glioblastoma stem-like cells (GSCs) as determinants of GBM development, maintenance, and treatment response. Understanding the response of GSCs to radiation should thus provide insight into the development of improved GBM treatment strategies. Towards this end, in vitro techniques for the analysis of GSC radiosensitivity are an essential starting point. One such method, the clonogenic survival assay has been adapted to assessing the intrinsic radiosensitivity of GSCs and is described here. As an alternative method, the limiting dilution assay is presented for defining the radiosensitivity of GSC lines that do not form colonies or only grow as neurospheres. In addition to these cellular strategies, we describe γH2AX foci analysis, which provides a surrogate marker for radiosensitivity at the molecular level. Taken together, the in vitro methods presented here provide tools for defining intrinsic radiosensitivity of GSCs and for testing agents that may enhance GBM radioresponse. Key words Glioblastoma, In vitro radiosensitivity, Clonogenic survival, Limiting dilution assay, γH2AX

1

Introduction Radiotherapy remains a primary treatment modality for glioblastomas (GBMs) significantly contributing to the prolongation of patient survival [1]. However, whereas many GBMs initially respond, they essentially all recur; even in combination with surgery and chemotherapy, the median survival of patients with GBM continues to be dismal with the majority succumbing to disease within 2 years of diagnosis [2]. Although GBM cells are generally considered to be highly migratory and invasive [3], local recurrence is overwhelmingly within the initial treatment volume, which indicates that GBM cells in situ are extremely radioresistant [4]. Defining the processes and molecules responsible for this radioresistance should provide a rational basis for designing target-based strategies that enhance GBM radiosensitivity and therapeutic response.

Peggy Stock and Bruno Christ (eds.), In Vitro Models for Stem Cell Therapy: Methods and Protocols, Methods in Molecular Biology, vol. 2269, https://doi.org/10.1007/978-1-0716-1225-5_3, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Towards this goal, a model system that accurately simulates the biology of GBMs is a critical requirement. Whereas the biology of long-established glioma cell lines has little in common with these brain tumors in situ [5], GBM stem-like cells (GSCs) are a clonogenic subpopulation thought to be critical to the development, maintenance, and treatment response of GBMs [6–8]. For in vitro experimentation, GSCs are initially isolated from human GBM surgical specimens as neurospheres and grown in neural basal medium containing epidermal growth factor and basic fibroblast growth factor, that is, stem cell growth medium. As a model system for investigating GBM radioresistance, we previously determined the in vitro radiosensitivity of GSCs using a clonogenic assay [9], the gold standard for defining intrinsic radiosensitivity. The in vitro clonogenic survival assay measures the consequences of radiation on the proliferative potential of individual cells; the survival curves generated from this assay reflect the two principal mechanisms of radiation-induced cell death: apoptosis and mitotic catastrophe [10]. Of note, as for cells isolated from most human solid tumors, irradiation of GSCs induces a minimal level of apoptosis with the majority of cells undergoing mitotic death [9]. Critical to the clonogenic assay is the ability of cells to grow as an attached monolayer and to form distinct colonies when seeded sparsely (i.e., at clonogenic density). Initial attempts at defining GSC radiosensitivity by clonogenic analysis used culture media containing 10% FBS [8]. However, although facilitating colony formation, the presence of FBS creates a differentiationinducing environment resulting in the loss of stem cell characteristics [5, 11], which complicates data interpretation. An alternative approach is to seed GSCs disaggregated from neurospheres onto standard tissue culture plates coated with poly-L-lysine (PLL) containing stem cell growth medium and maintained at 5% O2 [9, 12]. Under these conditions, the GSCs grow as adherent colonies and, in contrast to growth in medium containing FBS, maintain their stem-like cell characteristics [9, 11]. This method allows for the generation of radiation clonogenic survival curves that can be used to compare the radiosensitivity of GSC lines as well as to evaluate potential radiosensitizers. However, not all GSCs are amenable to the clonogenic assay. Some GSC lines only proliferate when grown in suspension as neurospheres. Other GSCs proliferate as monolayer cultures when tissue culture plates are coated with PLL or, as an alternative substrate that enhances attachment, polyornithine/laminin but do not form the distinct colonies necessary for clonogenic survival analysis. In these situations, the limiting dilution assay (LDA) may be applicable. Similar to clonogenic assays, the LDA determines the effects of radiation on cell proliferation and/or the ability to grow into neurospheres, which can be especially applicable to GSCs. The LDA typically involves seeding a serial dilution of cells into rows of

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a 96-well plate (see below), before or after irradiation [13, 14]. After allowing 2–3 weeks for proliferation, the wells are scored as positive or negative and survival curves constructed (see below). While it may be possible to determine surviving fraction based on neurospheres per well, it must be cautioned that at the time of irradiation wells do not contain single cells. In contrast to the clonogenic assay, which involves irradiation of single cells as they are attached in monolayer culture, irradiation of wells containing multiple cells growing in suspension can result in aggregation, which could influence neurosphere counts and complicate data interpretation. Finally, short-term cellular assays based on cell proliferation at 2–3 days after irradiation are not appropriate for assessing radiosensitivity of cells that undergo mitotic death (e.g., those isolated from solid tumors, normal fibroblasts). Whereas this type of assay may be applicable to chemotherapy, it does not account for the transient cell cycle delay that occurs after irradiation. In addition to the cellular assays described above, at the molecular level, analysis of γH2AX foci provides insight into GSC radiosensitivity. γH2AX foci correspond to radiation-induced DNA double-strand breaks (DSBs) and their dispersal correlates with DSB repair [15, 16]. Because DSBs are the critical lesion in radiation-induced cell death, γH2AX foci can serve as a surrogate measure of radiosensitivity [17, 18]. Moreover, given that a shared target of many radiosensitizing agents involves some aspect of the DSB repair process, quantifying γH2AX foci dispersal as a function of time after irradiation can be useful in identifying drugs that enhance radiation-induced cell death. Because foci can be evaluated in 1–2 days after irradiation and because their measure is amenable to high-throughput technology [19], γH2AX foci analysis provides an advantage as a screening approach for GSC radiosensitizers.

2 2.1

Materials Cell Culture

1. Stem cell medium: DMEM/F12, 2% B27 supplement without vitamin A, 50 ng/mL each of human EGF and FGF. 2. TrypLE Express. 3. Defined Trypsin Inhibitor. 4. Phosphate-buffered saline (PBS). 5. 40 μm cell strainers. 6. Beckman coulter cell counter and isotonic diluent (or similar cell counting platform). 7. 37
C, 5% CO2, 5% O2 Incubator (see Note 1). 8. Light microscope. 9. Centrifuge.

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2.2 Clonogenic Survival Assay

1. 0.1% Poly-L-Lysine in H2O. 2. 6-well plates. 3. 0.5% Crystal violet solution: 2% Crystal violet diluted in methanol. 4. Stereomicroscope.

2.3 Limiting Dilution Assay

1. Flat bottom 96-well plates. 2. Multichannel pipette. 3. Light microscope. 4. ELDA: Extreme Limiting Dilution Analysis online software (http://bioinf.wehi.edu.au/software/elda/).

2.4

γH2AX Staining

1. Poly-L-Ornithine: 1:500 in PBS. 2. Laminin: 1:500 in PBS. 3. 2-Chamber slides (see Note 2). 4. Fixative: 10% neutral buffered formalin. 5. PBS-T: PBS with 0.5% Tween 20. 6. Permeabilization buffer: PBS with 0.2% Triton. 7. Blocking buffer: PBS-T, 5% Goat serum, 1% BSA. 8. Primary antibody: anti-phospho-Histone H2AX Ser 139 (Millipore; 1:1000 concentration). 9. Secondary antibody: Alexa Fluor 488 goat anti-mouse IgG (Molecular Probes; 1:1000 concentration; or use secondary antibody of choice). 10. ProLong Gold Antifade with DAPI. 11. Fluorescent microscope. 12. Plate rocking device. 13. Image analysis software (e.g., ImageJ).

2.5 Ionizing Radiation Source

3

1. XRAD320 (Precision X-ray, Inc.) or similar source of ionizing radiation.

Methods

3.1 Clonogenic Survival Assay

1. All steps should be performed under sterile conditions until the final steps of staining and analysis. Once made or opened, reagents should be stored at 4
C unless otherwise specified by manufacturer. 2. Add 1 mL poly-L-lysine (PLL) to each well of a 6-well plate. Rock plates gently to ensure that all well surfaces are coated.

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Allow poly-L-lysine to stand for 1 h to overnight in 37
C incubator. 3. Remove PLL from wells and wash twice with sterile PBS. PLL can be utilized up to three times before being discarded. Aspirate final PBS wash and let wells dry in 37
C incubator for several hours. Drying overnight is preferred. 4. As glioblastoma stem-like cells are typically maintained as neurospheres, spin down neurospheres at 138  g for 3 min at room temperature. To limit the amount of cell damage, the level of centrifugation should be kept to a minimum, i.e., just enough to pellet the neurospheres, which will depend on the size of the spheres. Aspirate supernatant and resuspend in 1 mL TryplE express. After appropriate incubation time (30 s to 5 min) at room temperature, add 2 mL Defined Trypsin Inhibitor and 4 mL sterile PBS (see Note 3). 5. Pipette up and down until neurospheres have been disaggregated and are no longer visible in pipette (see Note 4). Filter cell suspension through a sterile 40 μm cell strainer; ensure a single cell suspension using microscope. 6. Spin cells down at 215  g for 5 min at room temperature. Aspirate supernatant and resuspend in 5 mL sterile PBS. Count cells with Coulter counter or hemocytometer. 7. Spin cells down for a final time at 215  g for 5 min, aspirate supernatant, and resuspend in stem cell media at 1  105 cells/ mL in a 50 mL conical tube. Perform serial 1:10 dilutions in stem cell media until desired dilution is reached. 8. Add appropriate number of cells to PLL-coated plates with a final volume of 2 mL of stem cell media per 9.5 cm2 well (see Note 5). Incubate plates overnight at 37
C, 5% CO2, and 5% O2 to allow cells to attach prior to radiation treatment (see Note 6). 9. After allowing time for cell attachment and recovery, plates are irradiated using an X-ray or other ionizing radiation source. Radiation should be delivered prior to the first doubling (for GSCs this is typically 24 h), which ensures that single cells and not doublets are irradiated. The radiation dose range typically delivered to GSCs is 1–6 Gy using at least 3 doses to allow for the generation of a survival curve. After irradiation, plates are returned to a 37
C, 5% CO2, and 5% O2 incubator and fed up to twice per week by adding 1 mL of stem cell medium (see Note 7). 10. In general, the maximum number of GSC colonies form in 14–21 days, which should be detectable under a stereomicroscope. Gently decant media into a waste container; 0.5% crystal violet is then added by dripping stain along the wall of the well,

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which should prevent colony detachment. After allowing crystal violet to stain cells for up to 5 min at room temperature, rinse plates by immersion in a large bowl of water (do not expose colonies directly to running water). Properly dispose of crystal violet. At this point, colonies should be clearly visible. 11. Count number of colonies per well with a stereomicroscope and colony counting pen and record. Colonies are defined as containing at least 25 cells. After irradiation cell death (permanent loss of proliferative activity) can occur after 1 or more divisions. For this reason and because the goal of the clonogenic assay is to define “surviving” cells, it is important to set the minimum number of cells per colony at 25, which corresponds to 4–5 divisions. 12. Using these results, a radiation survival curve can be constructed. Towards this end, the surviving fraction (SF) for each dose of radiation is calculated by determining the plating efficiency (PE) (number of colonies divided by the number of cells seeded  100), which is then divided by the PE determined for the untreated control sample. Survival curves are constructed by plotting the surviving fraction versus radiation dose. The control (0 Gy) SF is set at 1.0 on the y-axis with the remaining SF data points plotted on a log scale (decreasing from 1.0); the x-axis corresponds to the radiation dose and is plotted on a linear scale (a semi-log plot). 3.2 Limiting Dilution Assay (See Note 8)

1. Under a sterile hood, collect neurospheres in suspension by centrifugation at 138  g for 3 min at room temperature. As in the clonogenic assay, use the minimum amount of centrifugation that will pellet the neurospheres. 2. Perform steps 4–6 in Subheading 3.1 as outlined above to obtain single cell suspension. Perform serial dilutions to obtain a suspension with 200 cells/100 μL (see Note 9). 3. Fill every well of a 96-well plate with 100 μL of stem cell medium. It is important to avoid the production of bubbles during the entire experiment. 4. Add 100 μL of cell dilution made above (200 cells/100 μL) to all the wells in row A for a final volume of 200 μL. 5. With the multichannel pipette, thoroughly mix solutions in row A and remove 100 μL from row A to transfer to row B. For the purposes of this example, the highest cell concentration will be 100 cells/well (After removing 100 μL from row A, row A is left with 100 cells). 6. Thoroughly mix solutions in row B and remove 100 μL to transfer cells to row C. Continue in a similar fashion through row H (leave 200 μL media in row H, while all other wells will

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have 100 μL) to obtain the example range: Row A (100 cells), B (50), C (25), D (12), E (6), F (3), G (1), H (0). 7. With a light microscope visually verify that wells A-G contain cell solutions composed of single cells and that there are no more than three cells in row H. Once verified, return plates to 37
C incubator with 5% O2. 8. Twenty-four hours after seeding cells, irradiate individual plates with a range of radiation doses, typically 2–8 Gy, and return each to incubator. 9. At 14–21 days post-irradiation, remove plates from the incubator and examine under a light microscope. Determine and record the number of positive wells (wells that contain one or more spheres of approximately 30 cells) and/or the number of spheres per well. 10. With the recorded number of positive wells, use ELDA: Extreme Limiting Dilution Analysis online software to calculate the cancer cell initiating frequency and significance (http://bioinf.wehi.edu.au/software/elda/) [20]. 3.3

γH2AX Foci

1. Add 1 mL poly-L-ornithine (PO) to each chamber of 2-chambered slides. Rock slides gently to ensure that all chamber surfaces are coated. Allow PO to stand for at least 4 h to overnight in 37
C incubator. 2. Remove PO from chambers and wash twice with sterile PBS. PO can be utilized up to three times before being discarded. Aspirate final PBS wash and add laminin in PBS to chamber and incubate at 37
C for at least 4 h to overnight. 3. Perform steps 3–6 in Subheading 3.1 as outlined above to generate a single GSC suspension. The number of cells seeded per chamber should be sufficient to generate an evenly distributed monolayer without overcrowding. Incubate cells on chamber slides until cells reach approximately 70% confluency. If cultures become confluent, it will be difficult to accurately visualize individual cells. 4. When cells reach appropriate density, slides can be irradiated. The dose range is typically 1–3 Gy; for dispersal studies, which correlate with repair of radiation-induced DNA double-strand breaks (DSBs), typically, slides receive 2 Gy and are fixed at 1, 6, 16, and 24 h post-irradiation (see Note 10). However, IR dose and time points can vary based on cell line and study variables. 5. For collection, chambers are washed with PBS prior to fixing cells with 10% neutral buffered formalin for 10 min at room temperature.

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6. Remove fixative and wash cells three times with PBS. After fixation, slides can be stored in the last PBS wash at 4
C for 7–10 days prior to staining. 7. For staining, permeabilize cell membranes with 0.2% Triton/ PBS for 10 min at room temperature. After permeabilization, rinse with PBS-T once. 8. To prevent nonspecific binding of primary antibody, add blocking buffer for 1 h at room temperature with shaking. 9. After blocking, add the primary antibody, anti-phospho-histone H2AX Ser 129, at 1:1000 concentration in blocking buffer. Incubate for 2 h at room temperature or overnight at 4
C. If utilizing a small volume of buffer with antibody, place slides on a plate rocking device. 10. When primary antibody incubation is complete, wash cells three times with PBS-T for 5–10 min per wash at room temperature. 11. Immediately add secondary antibody, Alexa Fluor 488 goat anti-mouse IgG (or secondary of choice), at 1:1000 concentration in blocking buffer. Incubate for 2 h at room temperature with shaking. 12. Again, wash cells three times with PBS-T for 5–10 min per wash. Remove final PBS-T wash and remove chamber from slide. 13. Apply 1 drop of ProLong Gold Antifade with DAPI (stains nuclei) to each chamber area and cover with glass cover slip. Allow to dry for 1 h to overnight with applied weight on top of slide to remove air bubbles. Slides can be stored for several weeks at 4
C prior to imaging. 14. Image on fluorescent microscope at 40–63. 15. Utilize ImageJ to analyze TIF image files to count number of foci per nuclei. Alternatively, foci can be counted manually in each nuclei. Count at least 25–50 nuclei per condition and time point.

4

Notes 1. Standard incubators use ambient oxygen levels (air) of approximately 20%. However, in the GBM in situ environment oxygen levels are generally 5% or less. We have found that culturing GSCs in an incubator that maintains an oxygen level of 3–5%, which better simulates the GBM microenvironment, enhances their stem cell-like characteristics, including their clonogenicity, as compared to cells maintained at ambient oxygen levels [12].

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2. Poly-L-ornithine/laminin coating of tissue culture plates or slides enhances attachment of GSCs and induces a proliferation pattern of an evenly distributed monolayer. This is in contrast to PLL, which also enhances attachment but results in a colony forming growth pattern when cells are seeded at a sparse (clonogenic) density. The uniform distribution of cells over the surface of the slide (the result of poly-L-ornithine/laminin coating) allows for the straightforward visualization of nuclei using standard fluorescent microscopy. 3. Incubation times with TryplE will depend on the GSC line. To minimize cell injury, exposure to TrypIE should be kept as short as possible. In general, 30 s of TryplE exposure is sufficient for many lines when coupled with physical disaggregation techniques (see Note 4). If neurospheres remain intact after 30 s of TryplE and pipetting, neurospheres may be spun down again and incubated with fresh TryplE for a slightly longer duration. 4. An effective method of physical disaggregation is to attach a sterile, unfiltered 200 μL pipette tip to a sterile 10 mL pipette and pipette the entire cell suspension up and down through the 200 μL tip until neurospheres are no longer visible. 5. The number of cells plated per well depends on the PE of the GSC line. For example, if a GSC line has a PE of 0.20, then control plates seeded at 100 cells should yield approximately 20 colonies. The number of cells seeded for the irradiated plates should then be increased to account for the expected level of cell killing. That is, if 2 Gy is expected to result in an SF of 0.50, then increase the cells seeded to 200, which should yield approximately 20 colonies (Irradiated PE/Control PE ¼ SF of 0.50). 6. It is also possible to plate cells for clonogenic analysis after treating with radiation. In this case, individual samples (cells in monolayer or neurospheres in suspension) are irradiated at a designated dose and then disaggregated into single cell suspensions that are counted and subjected to the necessary dilutions to obtain the desired cell concentration, which are then seeded into PLL-coated plates. However, to reduce the variability that can result from the multiple individual cell counts and dilution procedures, we find that reproducibility is improved when cells are plated and then irradiated, a protocol that involves the disaggregation and counting of a single sample. Moreover, plating cells and then irradiating after a recovery period reduces the potential for the disaggregation process to artifactually influence cellular radioresponse. 7. The clonogenic assay can be used to identify potential radiation modifying agents. This typically involves an agent delivered

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alone and combined with each dose of radiation. The critical calculation is to account for the cell death (decreased PE compared to control) induced by treatment with the agent alone. By normalizing for agent-induced cell killing both the radiation only and the radiation + agent survival curves start at an SF of 1.0 allowing for a direct comparison. The radiation only and radiation + agent survival curves can be compared according to a dose modifying factor (DMF). This is defined as the ratio of the radiation dose that results in a given surviving fraction (typically 0.10) to the dose of radiation in the combined treatment that results in the same surviving fraction. A DMF of greater than 1 indicates radiosensitization and less than 1 indicates radioprotection. 8. This 96-well plate assay can be used for adherent cells that do not form colonies on standard or coated tissue culture plates, cells with a migratory phenotype that only form an evenly distributed monolayer, or cells that divide only when grown as neurospheres. 9. The range of cell concentrations used is dependent on the frequency of initiating cells in the fraction and on the intrinsic radiosensitivity of the cells. Therefore, LDA will need to be optimized for each individual cell line. An initial LDA using a wide range of cell concentrations can be used to find the best range of cell dilutions. The range should include concentrations with 100% positive wells or sphere formation down to cell concentrations with no positive wells or sphere formation. 10. γH2AX foci analysis can also be used to test potential radiosensitizing agents. Because DSB repair correlates with radiosensitivity, testing a drug or a genetic/epigenetic manipulation involves determining the time course of foci dispersal after irradiation. For GSCs, the radiation dose is typically 2 Gy and foci are determined at 1, 6, and 24 h post-irradiation with and without the agent. A significantly greater number of foci remaining at 24 h in the radiation + agent samples as compared to radiation only is suggestive of an inhibition of DSB repair and putatively radiosensitization. At each time point, it is important to have samples treated with agent alone to control for the possible induction of γH2AX foci by the agent. The detection of a radiosensitizing agents using γH2AX foci analysis should be verified using the clonogenic assay.

Acknowledgments JHM is supported by the NIH OxCam and Gates Cambridge Scholarships.

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References 1. Stupp R, Mason WP, van den Bent MJ et al (2005) Radiotherapy plus concomitant and adjuvant temozolomide for glioblastoma. N Engl J Med 352:987–996. https://doi.org/ 10.1056/NEJMoa043330 2. Ostrom QT, Gittleman H, Truitt G et al (2018) CBTRUS statistical report: primary brain and other central nervous system tumors diagnosed in the United States in 2011–2015. Neuro Oncol 20:iv1–iv86. https://doi.org/ 10.1093/neuonc/noy131 3. Giese A, Bjerkvig R, Berens ME, Westphal M (2003) Cost of migration: invasion of malignant gliomas and implications for treatment. J Clin Oncol 21:1624–1636. https://doi.org/ 10.1200/JCO.2003.05.063 4. Chan JL, Lee SW, Fraass BA et al (2002) Survival and failure patterns of high-grade gliomas after three-dimensional conformal radiotherapy. J Clin Oncol 20:1635–1642. https://doi. org/10.1200/JCO.2002.20.6.1635 5. Lee J, Kotliarova S, Kotliarov Y et al (2006) Tumor stem cells derived from glioblastomas cultured in bFGF and EGF more closely mirror the phenotype and genotype of primary tumors than do serum-cultured cell lines. Cancer Cell 9:391–403. https://doi.org/10.1016/j.ccr. 2006.03.030 6. Galli R, Binda E, Orfanelli U et al (2004) Isolation and characterization of tumorigenic, stem-like neural precursors from human glioblastoma. Cancer Res 64:7011–7021. https:// doi.org/10.1158/0008-5472.CAN-04-1364 7. Singh SK, Hawkins C, Clarke ID et al (2004) Identification of human brain tumour initiating cells. Nature 432:396–401. https://doi. org/10.1038/nature03128 8. Bao S, Wu Q, McLendon RE et al (2006) Glioma stem cells promote radioresistance by preferential activation of the DNA damage response. Nature 444:756–760. https://doi. org/10.1038/nature05236 9. McCord AM, Jamal M, Williams ES et al (2009) CD133+ glioblastoma stem-like cells are radiosensitive with a defective DNA damage response compared with established cell lines. Clin Cancer Res 15:5145–5153. https://doi.org/10.1158/1078-0432.CCR09-0263 10. Hall EJ, Giaccia AJ (2012) Radiobiology for the radiologist, 7th edn. Lippincott Williams and Wilkins, Philadelphia

11. Pollard SM, Yoshikawa K, Clarke ID et al (2009) Glioma stem cell lines expanded in adherent culture have tumor-specific phenotypes and are suitable for chemical and genetic screens. Cell Stem Cell 4:568–580. https:// doi.org/10.1016/j.stem.2009.03.014 12. McCord AM, Jamal M, Shankavaram UT et al (2009) Physiologic oxygen concentration enhances the stem-like properties of CD133+ human glioblastoma cells in vitro. Mol Cancer Res 7:489–497. https://doi.org/10.1158/ 1541-7786.MCR-08-0360 13. Grenman R, Burk D, Virolainen E et al (1989) Clonogenic cell assay for anchorage-dependent squamous carcinoma cell lines using limiting dilution. Int J Cancer 44:131–136 14. Venere M, Hamerlik P, Wu Q et al (2013) Therapeutic targeting of constitutive PARP activation compromises stem cell phenotype and survival of glioblastoma-initiating cells. Cell Death Differ 21:258–269. https://doi. org/10.1038/cdd.2013.136 15. Rogakou EP, Pilch DR, Orr AH et al (1998) DNA double-stranded breaks induce histone H2AX phosphorylation on serine 139. J Biol Chem 273:5858–5868. https://doi.org/10. 1074/jbc.273.10.5858 16. Sedelnikova OA, Rogakou EP, Panyutin IG, Bonner WM (2002) Quantitative detection of (125)IdU-induced DNA double-strand breaks with gamma-H2AX antibody. Radiat Res 158:486–492 17. Bana´th JP, MacPhail SH, Olive PL (2004) Radiation sensitivity, H2AX phosphorylation, and kinetics of repair of DNA strand breaks in irradiated cervical cancer cell lines. Cancer Res 64:7144–7149. https://doi.org/10.1158/ 0008-5472.CAN-04-1433 18. Olive PL, Bana´th JP (2004) Phosphorylation of histone H2AX as a measure of radiosensitivity. Int J Radiat Oncol Biol Phys 58:331–335 19. Avondoglio D, Scott T, Kil W et al (2009) High throughput evaluation of gammaH2AX. Radiat Oncol 4:31. https://doi.org/ 10.1186/1748-717X-4-31 20. Hu Y, Smyth GK (2009) ELDA: extreme limiting dilution analysis for comparing depleted and enriched populations in stem cell and other assays. J Immunol Methods 347:70–78. https://doi.org/10.1016/j.jim.2009.06.008

Chapter 4 Bioimaging of Mesenchymal Stem Cells Spatial Distribution and Interactions with 3D In Vitro Tumor Spheroids Luı´s P. Ferreira, Vı´tor M. Gaspar, and Joa˜o F. Mano Abstract In solid tumors, mesenchymal stem cells (MSCs) are recognized to establish complex intercommunication networks with cancer cells and to significantly influence their invasion and metastasis potential. Such bidirectional interplay occurs between both tissue resident/tumor-associated MSCs (TA-MSCs) and also tumor infiltrating MSCs (TM-MSCs) that migrate from distant sites such as the bone marrow. Interestingly, malignant cells interactions with MSCs in the tumor microenvironment extends beyond conventional exchanges of signaling factors and extracellular vesicles, including unconventional direct exchanges of intracellular components, or cancer cells cannibalism of MSCs. In the context of 3D in vitro tumor models, cell tracking assays making use of cell-labeling probes such as membrane penetrating dyes, can be leveraged to shed light on these events, and allow researchers to analyze overtime cell-to-cell spatial distribution, fusion, internal organization, and changes in co-cultured populations ratios. Herein, we describe a highthroughput compatible method through which MSCs positioning and permanence within in vitro 3D multicellular tumor spheroid models (3D-MCTS) can be tracked overtime. Although we have focused on the interactions of human bone marrow-derived MSCs (hBM-MSCs) within heterotypic lung cancer A549 3D-MCTS, these procedures can be implemented for other 3D tumor spheroid models and types of cells, taking into consideration that optimization steps are undertaken. Key words Tumor-associated/infiltrating mesenchymal stem cells, 3D Multicellular Tumor Spheroids, Bioimaging, Cell tracking, Cell labeling

1

Introduction Mesenchymal stem cells influence in the tumor microenvironment is increasingly known to represent a crucial factor in tumor progression and acquisition of key disease hallmarks. However, their exact role remains controversial and highly underexplored [1, 2]. In the diseased tissues, tumor-associated/resident (TA-MSCs) and tumor infiltrating (TI-MSCs) mesenchymal stem cells exhibit lineage and tumor population-dependent behaviors, that partially dictate the acquisition of TA-MSCs and TI-MSCs tumor permissive or inhibitory phenotypes which profoundly influence tumor progression and the overall therapeutic outcome [1, 3]. For example, bone

Peggy Stock and Bruno Christ (eds.), In Vitro Models for Stem Cell Therapy: Methods and Protocols, Methods in Molecular Biology, vol. 2269, https://doi.org/10.1007/978-1-0716-1225-5_4, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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marrow and adipose tissue-derived MSCs have been reported to promote lung cancer proliferation and resistance to therapeutics [4, 5]. Conversely, in the context of malignant pleural mesothelioma, a specific type of lung cancer, lung resident MSCs seem to exert anti-tumor activity through direct interactions with cancer cells [6]. Moreover, TA-MSCs of distinct origins (e.g., adipose and bone marrow derived) have also been found to directly engage with cancer cells via mitochondrial and cell membrane exchanges that directly augment cancer cell proliferation, checkpoint mechanisms escape, and immune system evasion [7–10]. Considering the increasingly established role of MSCs in tumor progression, the study of direct/indirect interactions between tissue-specific MSCs with cancer cells and other TME cellular components is therefore crucial during preclinical screening and validation stages [1]. Maximizing the potential of laboratory models to mimic cancer cell–MSCs interactions in a controlled and reproducible setting could in fact lead to the identification and validation of targeted therapies [1, 3]. From the available testing platforms, 3D tumor models outperform the conventional flat 2D in vitro tumor models for both fundamental biology studies and preclinical drug-screening studies. 3D multicellular tumor spherical \spheroid in vitro tumor models are highly reproducible and better recapitulate key TME hallmarks (e.g., nutrient/oxygen gradients, necrotic core formation, gene expression patterns), cell–cell contacts, and cellular 3D spatial distribution allowing researchers to better investigate MSCs-cancer cells direct/indirect interactions in vitro [11]. Moreover, the easy inclusion of these models in microfluidics and high-throughput compatible platforms can be leveraged to generate more relevant data [1, 12, 13]. 3D co-culture models also provide a more predictive preclinical validation platform and have shown, in certain cases, to correlate directly with patients clinical data [14]. Importantly, several in vitro models aimed at studying the direct interactions of cancer cells and MSCs upon co-culture have at times obtained contradicting results, in part, derived from non-standardized experimental designs. In this regard, variations in internal cellular organization in the 3D microtumor mass, MSCs tropism toward the tumor niche, MSCs permeance within the tumor microenvironment, and subsequent lineage differentiation require careful assessment during the development of 3D in vitro models [1]. For example, when analyzing the co-culture of MSCs and cancer cells, Bartosh and co-workers [9] demonstrated the effects of cancer cell cannibalism and fusion with MSCs on model progression should be considered. In this study, breast cancer cells co-cultured within 3D tumor spheroids with bone marrow-derived MSCs, fused, and degraded surrounding MSCs populations. Such cannibalization of MSCs lead to acquisition of pro-tumoral phenotypes by breast cancer cells. This triggered survival mechanisms of

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dormancy, due to the accumulation of pro-inflammatory and tumor-suppressor mediators that despite promoting a senescentlike state prevented cell death [9]. Besides direct internalization or degradation of MSCs populations within 3D in vitro models, initial seeding ratios of co-cultured cellular populations can also change during culture, as a result of either cell death or differentiation. Such alterations must be carefully monitored, creating the necessity of tracking and analyzing each diversified cellular population in direct contact on 3D co-culture models. Given the importance of such interactions in cancer progression and therapeutic outcome, it is therefore necessary to improve current methodologies for standardizing the study of MSCs interactions within the TME. Several methodologies for tracking cell organization and the presence in in vitro platforms, such as specific Cluster of Differentiation (CD) surface biomarkers staining [15], fluorescent reporter gene transfection protocols [10], and membrane labeling solutions [16] have been developed and optimized for 3D tumor spheroid models. While immunostaining techniques targeting CD surface biomarkers have been used with success in histological sections of dense 3D microtumors [15], real-time tracking of cells with antibody-based staining is limited due to diffusional barriers. Moreover, the differentiation of both cancer stem cells and MSC populations can be difficult to follow due to the overlapping expression of key receptors [17–19]. Adding to this, MSCs of distinct origins present distinct surface receptors, showing different expression levels as a result of their potential interactions with cancer cells, and posterior lineage differentiation or subpopulation development [20]. The use of cell tracking techniques, such as transfection with fluorescent protein expressing transgenes (e.g., GFP, YFP, RFP, mCherry, mTomatto) delivered via viral or non-viral vectors has been employed as a valid alternative [21]. Transfection-based labeling solutions, making use of fluorescent proteins offer reliable staining protocols that can provide lineage-specific staining up to several generations [22]. However, such protocols are generally expensive, involve time-consuming transfection and analysis steps. Most importantly, such approaches are associated with a transient transgene expression, variable transfection rates in the cell population and alterations in cell behavior, requiring careful optimization which makes their use in high-throughput platform rather cumbersome [23, 24]. Alternatively, the method herein described makes used of lipophilic Vybrant™ cell-labeling probes to study the spatiotemporal organization, direct cell-cell interactions, fusion, and permanence of distinct cell populations within heterotypic triple culture 3D tumor spheroids comprising non-small cell lung cancer (A549) cells, human dermal fibroblasts (HF), and human bone marrowderived mesenchymal stem cells (hBM-MSCs) [25, 26]. The used cell labeling dyes provide an affordable and easy-to-use approach

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for performing cell tracking assays. By using highly lipophilic carbocyanine dyes DiI, DiO, and DiD, which present low cytotoxicity and high resistance to intercellular exchange, one is able to differentially stain co-cultured cell populations and analyze their direct interactions and organization within heterotypic spheroid models [25]. These dyes can be added directly to serum-free cell culture media to perform uniform labeling of the extracellular membrane of suspended cells. Having been found to remain present up to 14 days post culture [25], the membrane bound dyes can be additionally used to: (i) analyze cell–cell fusion, (ii) exchanged membrane fragments, or (iii) to evaluate the assembly of stable cell clusters which depending on the resolution of the used microscopy system can be identified by the existence of double cell labeling [27, 28]. Moreover, although Vybrant™ cell labeling dyes have not yet been tested in the context of tissue-clearing protocols, previous reports have showcased that chemically similar compounds such as CM-DiI, were compatible with commonly used tissue-clearing methodologies (e.g., CLARITY) [29]. These features render cell-retainable labeling dyes as valuable tools for the analysis of both direct-cell interactions, and spatial distribution of MSCs within 3D tumor spheroids volume.

2

Materials

2.1 Cell Culture and 3D Spheroids Fabrication

1. Temperature and moisture-controlled CO2 incubator (Model C 170, Binder). 2. Human Primary Dermal Fibroblasts (ATCC®-PCS-201012™). 3. Non-small Cell Lung cancer cell line A549 (ATCC CRM-CCL-185™). 4. Bone Marrow-derived Mesenchymal Stem Cells (ATCC®PCS-500-012™). 5. Falcon tube refrigerated centrifuge (Sigma 3-16KL). 6. Cell Strainer (SPL Life Sciences—ø ¼ 40 μm). 7. Hemocytometer or automated cell counter. 8. Inverted phase-contrast microscope. 9. T-75 cm2 cell culture-treated T-flasks. 10. Dulbecco’s Phosphate-Buffered Saline (D-PBS). 11. Ultra-Low-Adhesion (ULA) round-bottom 96-well plates. 12. Trypan Blue. 13. Antibiotic/antimitotic (ATB) solution: 10,000 units/mL penicillin, 10,000 μg/mL streptomycin.

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14. Complete DMEM-HG: Dulbecco’s Modified Eagle MediumHigh Glucose Medium supplemented with L-glutamine and sodium bicarbonate, without HEPES, 10% (v/v) fetal bovine serum (FBS), 1% (v/v) antibiotic/antimitotic (ATB) solution. 15. Complete HAMs-F12: Ham’s F-12K Kaighn’s Medium, supplemented with L-glutamine and sodium bicarbonate, without HEPES, 10% (v/v) FBS, 1% (v/v) ATB. 16. Complete α-MEM: Alpha Modified Eagle’s Medium supplemented with L-glutamine and sodium bicarbonate, without HEPES, 10% (v/v) FBS, 1% (v/v) ATB. 17. Trypsin/EDTA solution: 0.05% Trypsin, 0.02% EDTA (see Note 1). 2.2 Cell Membrane Labeling

1. FBS-free cell culture medium (see Note 2). 2. Cell Culture Mediums supplemented with 10% (v/v) FBS and 1% (v/v) ATB solution (see Note 3). 3. 1 mM Vybrant™ DiO cell-labeling solution—contains dimethylformamide. 4. 1 mM Vybrant™ DiI cell-labeling solution—contains ethanol. 5. 1 mM Vybrant™ DiD cell-labeling solution—contains ethanol.

2.3 3D Cell Tracking in Internal Organization and Migration Assays

3

1. Time-lapse high-resolution confocal microscope, or similar fluorescence imaging apparatus. 2. Software: Image J/Fiji (open source), Huygens Suite (Scientific Volume Imaging) or Imaris (Bitplane).

Methods

3.1 Heterotypic 3D Tumor Spheroids Fabrication

1. Trypsinize the desired cell lines that are to constitute the unitary building blocks of 3D microtumors by following conventional protocols. First, wash the cells once with D-PBS and then treat with Trypsin/EDTA solution place the cells in an incubator at 37  C, 5% CO2, for 5 min (see Note 4). 2. Inactivate trypsin/EDTA solution by resuspending detached cells in trypsin neutralizing medium, at a 1:2 v/v ratio. Use the appropriate medium for each specific cell culture, namely HAM-F12 for A549 cells, DMEM-HG for HF cells, and α-MEM for hBM-MSCs. Pipet repeatedly to create a singlecell suspension (see Note 5). 3. Centrifuge the obtained cell suspensions at 300  g, for 5 min, room temperature (RT 22–25  C) (see Note 6). 4. Aspirate the supernatant and resuspend the cells in a minimal volume of cell culture medium.

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Fig. 1 (a) Size variation of cultured 3D spheroids can be observed over time by optical contrast micrographs of 3D spheroids. These images can then be used to monitor morphology, size, and circularity of obtained spheroids at different time points (1, 7, and 14 days after initial seeding). (b) High resolution CLSM confocal imaging of the cellular organization in 3D spheroids containing a triculture of cancer cells, fibroblast and MSCs at day 7. (b1) Micrograph with merged channels. (b2) Magnified section showcasing close cellular arrangement. (b3) DiO labeled A549 cancer cells – green channel. (b4) DiL labeled fibroblasts – yellow channel. (b5) DiD labeled MSCs – red channel. Mesenchymal stem cells acquire a cluster-like arrangement. Scale bars = 500 μm. Reproduced from Ferreira et al. 2018 [26], with permission from Elsevier

5. Count the cells using a hemocytometer or an automatic cell counter. 6. Prepare the necessary cell suspensions with a final concentration of 150,000 cells/mL using HAM-F12 medium. Adapted to the desired ratio for each cell populations in question for mono, and heterotypic 3D spheroids assembly (see Note 7). 7. Using a single or multichannel pipette (see Note 8) add 200 μL of the desired cell suspension to each well of the ULA roundbottom 96-well plates so that 30,000 cells are seeded per well. 8. Confirm that the desired wells were seeded by observation in an inverted phase-contrast microscope. 9. Carefully store the plates in a temperature and moisturecontrolled incubator and avoid any movement over the next 24 h to 72 h. 10. Exchange the culture medium every 3–4 days (see Note 9). 11. At the specific time points of 24 h, 3, 7, and 14 days inspect that 3D spheroids formation is occurring and acquire optical contrast microscopy micrographs (Fig. 1) for further quantification of size and circularity (see Note 10). 3.2 Cell Membrane Labeling for Labeled 3D Spheroids Generation

All steps involving the manipulation of labeling agents should be performed in a low-light or darkened room. Selection of the labeling agent for the respective cellular population should take into account the possibility of conflicting auto-fluorescence derived

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from deposited and present proteins within the spheroid matrix or included material. Furthermore, considering the excitation/emission of the selected fluorophore. 1. Recover the intended Vybrant™ labeling solution from storage and let stabilize at RT protected from light (see Note 11). 2. Retrieve the desired cell lines for differential staining, (e.g., A549 to be stained with DiO, HF to be stained with DiI, and hBM-MSCs to be stained with DiD) and trypsinize them (see Subheading 3.1 steps 1–3). 3. Resuspend each cell line in a minimal volume of respective FBS-free cell culture medium (see Note 2). 4. Count the cells using a hemocytometer or an automatic cell counter and prepare the necessary cell suspensions with a final concentration of 1  106  cells/mL in serum-free medium. 5. In light protected conditions, add in 5 μL of the selected tracking dye per 1 mL of cell suspension (1  106 cells/mL) (see Note 12). 6. Incubate the cells in suspension with the labeling dye for 25 min, at 37  C and 5% CO2. 7. Recover stained cells by centrifugation at 300  g, for 5 min, at 37  C (see Note 6). 8. Carefully remove the supernatant and gently resuspend the cells in the respective serum containing medium (i.e., complete HAM-F12 cell culture medium) (see Notes 13 and 14). 9. Homogenize gently with the pipette and allow a resting period of 10 min within the incubator at 37  C, and 5% CO2, before performing the next washing step (see Note 15). 10. Repeat steps 7–9 twice, until a total of three washes are performed. 11. Recover cells as in step 7 and count again to confirm cell numbers (see Note 16). 12. Assemble the required cell suspensions and proceed to labeled 3D spheroid formation (see Subheading 3.1 steps 6–9). Once again, protect labeled cells from strong light so as to avoid any occurrence of tracking dyes photo-bleaching. 13. Optional step: seed the remaining labeled cells in 2D adherent culture wells or petri-dishes in low concentration, for confirming the efficiency of the labeling procedure. 14. Upon seeding and confirmation through optical microscopy analysis, partially shield the ULA round-bottom 96-well plate with aluminum foil or any other opaque material to prevent photo-bleaching of stained cells (see Note 17). 15. Optional step: After 24 h, check the labeling efficiency in 2D cell culture controls under the fluorescence microscope.

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3.3 MSCs Internalization Cell Tracking Assays in 3D Spheroids

In order to access MSCs ability to migrate and integrate into tumor spheroids upon spheroid formation, a procedure, similar to the previously discussed one, can be applied. Therefore, spheroids are firstly cultured, and on the desired time point labeled MSCs are added in the required density to the culture plate wells. 1. Proceed to obtain spheroids with the desired physiological and biochemical characteristics. Herein, seventh day triple co-culture, co-culture, and mono-culture 3D spheroids of A549, HF, and in the case of tri-culture hBM-MSCs, were used (see Note 18). 2. Inspect by optical phase-contrast microscopy that 3D spheroid formation occurred in accordance with the desired standards. 3. Prepare DiD-stained hBM-MSCs in accordance with the labeling protocol (see Subheading 3.2 steps 1–11). 4. Resuspend the freshly labeled hBM-MSCs to the desired range of concentrations to test and store in dark until use, at 37  C and 5% CO2. 5. Carefully tilt the ULA round-bottom 96-well plate containing the unstained spheroids, or differentially stained spheroids, in a 30 –45 angle toward the user and remove the medium without aspirating the 3D spheroids. 6. Add the labeled hBM-MSCs in the desired seeding density to the respective wells containing the spheroids. In this protocol, a ratio of 1:10 ratio of hBM-MSCs to cancer cells was used for spheroids with an initial cell seeding density of 30,000 cells per well. 7. Acquire a fluorescence image of the 3D spheroids and labeled hBM-MSCs at day 0. 8. Incubate for the desired amount of time at 37  C and 5% CO2, performing either live-cell imaging or time-lapse imaging at specific time points (Fig. 2). 9. The obtained images can then be processed for analysis and possible sectioning using Imaris, Volocity, or ImageJ (see Note 19).

4

Notes 1. Other commercial cell detaching agents such as TrypLE™ Express can also be used. 2. To avoid undesired interactions between labeling agents and FBS proteins, cell culture medium without FBS is required for efficient labeling of the desired cells with Vybrant™ labeling solutions. Ideally, to avoid changes in cell culture medium composition between conditions the same medium used for 3D spheroids culture should be employed in the staining steps.

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Fig. 2 Example of widefield fluorescence micrographs of mono (a), dual (b and c), and triple co-culture (d) 3D spheroids with A549, HF, and hBM-MSC populations differentially stained for overtime hBM-MSCs cell migration and spheroid internalization tracking. hBM-MSCs stained with DiD were added to 3D spheroids previously cultured for 7 days (t ¼ 0 h) in ULA plates. A549—green channel (DiO), HF—Orange channel (DiL), spheroid resident hBM-MSCs (non-labeled in condition b and d); newly administered hBM-MSCs—red channel (DiD). Scale bar ¼ 500 μm. Reproduced from Ferreira et al. 2018 [26], with permission from Elsevier

3. Select the medium or medium combination in which the spheroids are going to be cultured, in the presented protocol HAM-F12 was used to perform 3D spheroids culture and thus used in an FBS-free format for the initial labeling. In order to perform the washing of unbound labeling agents, the same medium now supplemented with 10% FBS and 1% ATB was used. 4. Protocol optimization resulted in the selection of the following volumes for the trypsinization of adherent cell lines at 80–90% confluency when cultured on cell-treated T-Flasks, namely: (a) T-25 cm2—1 mL, incubation for 5 min, at 37  C. (b) T-75 cm2—2 mL, incubation for 5 min, at 37  C. (c) T-175 cm2—5 mL, incubation for 5 min, at 37  C. Cell lines with increased matrix deposition, or for which the culture time was extended can require either increase volume of trypsin (suggested), or increased incubation time (care must be taken when increasing incubation times in trypsin solutions so as to avoid toxicity). For example, trypsinization of MSCs cultures with significant matrix deposition can require an extended incubation time of ~6–8 min. 5. Take care to gently add and mix the cells within the minimal volume required for a translucid solution to form. Concentrated cells suspension requires vigorous pipetting for disassembly of cellular clumps, taking care however not to disrupt

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cell membrane integrity. Alternatively, cell clumps can be removed by using a cell strainer (SPL Life Sciences—Cell strainer ø ¼ 40 μm). 6. If using the specified centrifuge or similar, perform the centrifugation with an acceleration of 9 and a breakage of 5. 7. Utilization of distinct cellular populations leads to an increased complexity of the 3D model in question, and thus to the necessity of accounting for a higher number of possible sources of error considering the number of controls/control experiments. It is therefore important to ascertain the influence of each individual and combined cell populations in 3D spheroids assembly and their final characteristics. Hence, parameters such as 3D spheroids stability, necrotic core formation, spheroid morphology, circularity, compaction rate, reproducibility and hBM-MSCs migration and/or inclusion into co-culture (HF and hBM-MSCs), and tri-culture 3D spheroids must be evaluated for each cell-to-cell population ratio to be used. 8. Assure the cell suspension is perfectly homogenized. Ideally, gently resuspend 5 using a pipette at every 3-well seedings in order to avoid cell aggregation and deposition along time during the seeding procedure. 9. To avoid disrupting the 3D spheroid only remove 70–80% of the medium present in the culture well, taking care to add an additional volume of 10–20 μL to circumvent any losses derived from evaporation phenomena. 10. 3D models formation should evidence a visible contraction of the suspended cells into a compact spheroid-shaped aggregate over the initial course of culture. Depending on the types of cell lines used, such contraction can take a longer period of time to occur and require higher cell seeding densities. For example, in the developed model, A549 cell seeding densities of 30,000 cells/well are required for mono-culture spheroids to form uniformly and with spherical shape. Whereas, upon inclusion of hBM-MSCs co-culture spheroids of 15,000 cells/ well were equally capable of achieving similar organization and contraction profiles. Overall, the formed 3D spheroid should present a reproducible spheroidal shape with appearance of a dense core, observable by inverted phase-contrast microscopy as a darker central zone. 11. Unused portions of labeling agent that are not required for immediate use should be stored tightly sealed and protected from light at room temperature for short-term storage, or at 4  C for mid to long-term storage (DiI V-22885, DiO V-22886,DiD V-22887). In the case of Vybrant™ DiR celllabeling solutions, storage should always be performed at 20  C (DiR V-22888).

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12. DiI, DiO, and DiD solutions used were all at the same concentration in the stock solutions, namely 1 mM; hence, the volume of staining agent required for each staining procedure is equal if the volume of cells/media is equal. 13. CRITICAL: Take extreme care not to disturb the pellet of cells, and gently resuspend it in a defined amount of volume. Ideally, resuspend cells in double the volume of supernatant removed to assure a thorough wash of any remaining staining agent. 14. Carefully select the medium or medium combination in which 3D spheroids are cultured. In this protocol, HAM-F12 was used to perform 3D spheroids culture and thus used in an FBS-free format for the initial labeling. In order to perform the washing of unbound labeling agents, the same medium now supplemented with 10% FBS and 1% ATB was used. 15. Depending on the cell line, the 10 min stasis might be omitted; however, it was observed that the inclusion of such a step not only assured uniform distribution of the staining agent on the cell membrane, but also avoided loss of cell viability, or membrane disruption. CRITICAL: Necessary precautions must be undertaken to avoid cell aggregation, either gently resuspending after 5 min or maintaining the cell suspension under gentle agitation. 16. During protocol optimization, it was observable that significant loss of cells can occur with each centrifugation and resuspension step, being therefore important to perform cells counting prior to models’ assembly. Furthermore, a resting period should be afforded to the washed cells before they are subjected to further processing into 3D models. 17. Take great care not to reduce air flow or temperature exchange rates on the well-plate by using excessive amounts of shielding material. hBM-MSCs and other MSCs are extremely sensitive to temperature, humidity, and consequent osmolality variations within the well plates, with little variations producing highly distinct results. 18. Optimization of labeling agent’s longevity might be required given the utilization of distinct experimental settings. During optimization, it was verified that Vybrant™ labeling agents remained well preserved, with no unstained population being visible, over a period of up to 10–14 days. 19. Previous reports have made use of these software packages to effectively analyze in a quantitative manner cell dispersion, alignment, migration, variation in population ratios, and number of fusion events. Extended literature can be analyzed and adapted to the analysis of other parameters. A detailed discussion on this topic could be found in the literature [30–34].

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Acknowledgments This work was developed under the support of the European Research Council grant agreement ERC-2014-ADG-669858 for project “ATLAS,” and under the financial support by the Portuguese Foundation for Science and Technology (FCT) through a Doctoral Grant afforded to Luı´s Ferreira (SFRH/BD/141718/ 2018). This work was also supported by the Programa Operacional Competitividade e Internacionalizac¸˜ao (POCI), in the component FEDER, and by national funds (OE) through FCT/MCTES, in the scope of the project PANGEIA (PTDC/BTM-SAL/30503/ 2017). Furthermore, Vı´tor Gaspar acknowledges funding in the form of a Junior Researcher Contract under the scope of the project PANGEIA. This work was developed within the scope of the project CICECO-Aveiro Institute of Materials, UIDB/50011/2020 & UIDP/50011/2020, financed by national funds through the FCT/MEC and when appropriate co-financed by FEDER under the PT2020 Partnership Agreement. References 1. Ferreira LP, Gaspar VM, Henrique R et al (2017) Mesenchymal stem cells relevance in multicellular bioengineered 3D in vitro tumor models. Biotechnol J 12:1700079. https:// doi.org/10.1002/biot.201700079 2. Oloyo AK, Ambele MA, Pepper MS (2017) Contrasting views on the role of mesenchymal stromal/stem cells in tumour growth: a systematic review of experimental design. Advs Exp Med 1083:103–124 3. Shi Y, Du L, Lin L, Wang Y (2016) Tumourassociated mesenchymal stem/stromal cells: emerging therapeutic targets. Nat Rev Drug Discov 16:35–52. https://doi.org/10.1038/ nrd.2016.193 4. Park YM, Yoo SH, Kim S-H (2013) Adiposederived stem cells induced EMT-like changes in H358 lung cancer cells. Anticancer Res 33:4421–4430 5. Zhang Y, Zhang Z, Guan Q et al (2017) Co-culture with lung cancer A549 cells promotes the proliferation and migration of mesenchymal stem cells derived from bone marrow. Exp Ther Med 14:2983–2991. https://doi.org/10.3892/etm.2017.4909 6. Cortes-Dericks L, Froment L, Kocher G, Schmid RA (2016) Human lung-derived mesenchymal stem cell-conditioned medium exerts in vitro antitumor effects in malignant pleural mesothelioma cell lines. Stem Cell Res Ther 7:25. https://doi.org/10.1186/s13287-0160282-7

7. Pasquier J, Guerrouahen BS, Al Thawadi H et al (2013) Preferential transfer of mitochondria from endothelial to cancer cells through tunneling nanotubes modulates chemoresistance. J Transl Med 11:94. https://doi.org/ 10.1186/1479-5876-11-94 8. Fais S, Overholtzer M (2018) Cell-in-cell phenomena in cancer. Nat Rev Cancer 18:758–766. https://doi.org/10.1038/ s41568-018-0073-9 9. Bartosh TJ, Ullah M, Zeitouni S et al (2016) Cancer cells enter dormancy after cannibalizing mesenchymal stem/stromal cells (MSCs). Proc Natl Acad Sci 113:E6447–E6456. https://doi. org/10.1073/pnas.1612290113 10. Li C, Cheung MKH, Han S et al (2019) Mesenchymal stem cells and their mitochondrial transfer: a double-edged sword. Biosci Rep 39:1–9. https://doi.org/10.1042/ BSR20182417 11. Ferreira LP, Gaspar VM, Mano JF (2018) Design of spherically structured 3D in vitro tumor models—advances and prospects. Acta Biomater 75:11–34. https://doi.org/10. 1016/j.actbio.2018.05.034 12. Rodenhizer D, Dean T, D’Arcangelo E, McGuigan AP (2018) The current landscape of 3D in vitro tumor models: what cancer hallmarks are accessible for drug discovery? Adv Healthc Mater 1701174:1701174. https:// doi.org/10.1002/adhm.201701174

Bioimaging of MSCs Within 3D In Vitro Tumor Spheroids 13. Huang BW, Gao JQ (2018) Application of 3D cultured multicellular spheroid tumor models in tumor-targeted drug delivery system research. J Control Release 270:246–259. https://doi.org/10.1016/j.jconrel.2017.12. 005 14. Grassi L, Alfonsi R, Francescangeli F et al (2019) Organoids as a new model for improving regenerative medicine and cancer personalized therapy in renal diseases. Cell Death Dis 10. https://doi.org/10.1038/s41419-0191453-0 15. Bajetto A, Pattarozzi A, Corsaro A et al (2017) Different effects of human umbilical cord mesenchymal stem cells on glioblastoma stem cells by direct cell interaction or via released soluble factors. Front Cell Neurosci 11:312. https:// doi.org/10.3389/fncel.2017.00312 16. Yumoto K, Berry JE, Taichman RS, Shiozawa Y (2014) A novel method for monitoring tumor proliferation in vivo using fluorescent dye DiD. Cytom Part A 85:548–555. https://doi.org/ 10.1002/cyto.a.22434 17. Dominici M, Le Blanc K, Mueller I et al (2006) Minimal criteria for defining multipotent mesenchymal stromal cells. The International Society for Cellular Therapy position statement. Cytotherapy 8:315–317. https://doi.org/10. 1080/14653240600855905 18. Park SC, Nguyen NT, Eun JR et al (2015) Identification of cancer stem cell subpopulations of CD34+ PLC/PRF/5 that result in three types of human liver carcinomas. Stem Cells Dev 24:1008–1021. https://doi.org/ 10.1089/scd.2014.0405 19. Leon G, MacDonagh L, Finn SP et al (2016) Cancer stem cells in drug resistant lung cancer: targeting cell surface markers and signaling pathways. Pharmacol Ther 158:71–90. https://doi.org/10.1016/j.pharmthera.2015. 12.001 20. Wilson A, Hodgson-Garms M, Frith JE, Genever P (2019) Multiplicity of mesenchymal stromal cells: finding the right route to therapy. Front Immunol 10:1–8. https://doi.org/10. 3389/fimmu.2019.01112 21. Dittmer A, Hohlfeld K, Lu¨tzkendorf J et al (2009) Human mesenchymal stem cells induce E-cadherin degradation in breast carcinoma spheroids by activating ADAM10. Cell Mol Life Sci 66:3053–3065. https://doi.org/10. 1007/s00018-009-0089-0 22. Xie C, Yang Z, Suo Y et al (2017) Systemically infused mesenchymal stem cells show different homing profiles in healthy and tumor mouse

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models. Stem Cells Transl Med 6:1120–1131. https://doi.org/10.1002/sctm.16-0204 23. Mandel K, Yang Y, Schambach A et al (2013) Mesenchymal stem cells directly interact with breast cancer cells and promote tumor cell growth in vitro and in vivo. Stem Cells Dev 22:3114–3127. https://doi.org/10.1089/ scd.2013.0249 24. Karlsson H, Frykn€as M, Larsson R, Nygren P (2012) Loss of cancer drug activity in colon cancer HCT-116 cells during spheroid formation in a new 3-D spheroid cell culture system. Exp Cell Res 318:1577–1585. https://doi. org/10.1016/j.yexcr.2012.03.026 25. Probes M (2001) Vybrant cell-labeling solutions. Solutions:1–3 26. Ferreira LP, Gaspar VM, Mano JF (2018) Bioinstructive microparticles for self-assembly of mesenchymal stem cell-3D tumor spheroids. Biomaterials 185:155–173. https://doi.org/ 10.1016/j.biomaterials.2018.09.007 27. Lehmann TP, Juzwa W, Filipiak K et al (2016) Quantification of the asymmetric migration of the lipophilic dyes, DiO and DiD, in homotypic co-cultures of chondrosarcoma SW-1353 cells. Mol Med Rep 14:4529–4536. https:// doi.org/10.3892/mmr.2016.5793 28. Tario JD, Humphrey K, Bantly AD et al (2012) Optimized staining and proliferation modeling methods for cell division monitoring using cell tracking dyes. J Vis Exp:e4287. https://doi. org/10.3791/4287 29. Jensen KHR, Berg RW (2016) CLARITYcompatible lipophilic dyes for electrode marking and neuronal tracing. Sci Rep 6:1–10. https://doi.org/10.1038/srep32674 30. Helmchen F, Denk W (2005) Deep tissue two-photon microscopy. Nat Methods 2:932 31. Pawley JB (2006) Handbook of biological confocal microscopy, 3rd edn. Springer, New York 32. Owen DM, Williamson DJ, Boelen L et al (2013) Quantitative analysis of threedimensional fluorescence localization microscopy data. Biophys J 105:L05. https://doi. org/10.1016/j.bpj.2013.05.063 33. Ljosa V, Carpenter AE (2009) Introduction to the quantitative analysis of two-dimensional fluorescence microscopy images for cell-based screening. PLoS Comput Biol 5:e1000603 34. Panula PAJ (2003) Handbook of biological confocal microscopy. J Chem Neuroanat 25:228. https://doi.org/10.1016/s08910618(03)00007-3

Chapter 5 Investigation of the MSC Paracrine Effects on Alveolar–Capillary Barrier Integrity in the In Vitro Models of ARDS Johnatas Dutra Silva and Anna D. Krasnodembskaya Abstract Acute Respiratory Distress Syndrome (ARDS) is a devastating clinical disorder with high mortality rates and no specific pharmacological treatment available yet. It is characterized by excessive inflammation in the alveolar compartment resulting in edema of the airspaces due to loss of integrity in the alveolar epithelial–endothelial barrier leading to the development of hypoxemia and often severe respiratory failure. Changes in the permeability of the alveolar epithelial–endothelial barrier contribute to excessive inflammation, the formation of lung edema and impairment of the alveolar fluid clearance. In recent years, Mesenchymal Stromal Cells (MSCs) have attracted attention as a cell therapy for ARDS. MSCs are known to secrete a variety of biologically active factors (growth factors, cytokines, and extracellular vesicles). These paracrine factors have been shown to be major effectors of the anti-inflammatory and regenerative properties observed in multiple in vitro and in vivo studies. This chapter provides a simple protocol on how to investigate the paracrine effect of MSCs on the alveolar epithelial–endothelial barrier functions. Key words Mesenchymal stromal cells, Paracrine effects, Endothelial cells, Epithelial cells, Barrier permeability

1

Introduction Mesenchymal stromal cells (MSCs) have gained extensive attention as a promising cell therapy for the treatment of a variety of human disorders due to their immunomodulatory and regenerative properties [1]. A multitude of preclinical studies, including our own work [2–6] demonstrated strong potential for MSCs as a promising cell-based therapy for a variety of inflammatory lung diseases. It is widely accepted that the therapeutic effects of MSCs are in large mediated by the various components of MSC secretome. Accumulating evidence suggests that major functional components of the MSC secretome are extracellular vesicles (EVs). It is also known

Peggy Stock and Bruno Christ (eds.), In Vitro Models for Stem Cell Therapy: Methods and Protocols, Methods in Molecular Biology, vol. 2269, https://doi.org/10.1007/978-1-0716-1225-5_5, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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that MSCs can adequately respond to the specific inflammatory microenvironments and can alter their phenotype and secretory profile according to the exogenous stimuli [7–9]. Acute Respiratory Distress Syndrome (ARDS) can be triggered by a variety of insults, including direct pulmonary (pneumonia, aspiration) and indirect extrapulmonary (trauma, sepsis) injury; however, the final common pathway of ARDS involves disruption of the alveolar–capillary barrier permeability and accumulation of protein-rich edema fluid in the alveolar and interstitial spaces [10]. The alveolar–capillary barrier is composed of the layer of alveolar epithelium cells (Type II and Type I) and pulmonary microvasculature. In a normal functional lung, these cells provide a selectively permeable membrane for gas exchange. In ARDS, this membrane is disrupted, with many epithelial and endothelial cells damaged. Barrier permeability is strongly associated with the robustness of endothelial and epithelial adherens junctions. Adherens junctions are destabilized by pro-inflammatory cytokines excessively secreted in ARDS. The resultant loss of integrity of the alveolar–capillary barrier allows a protein-rich exudate from the blood to enter the airspaces thereby hindering gas exchange. This is further complicated by epithelial and endothelial cell death and denudation of the basement membranes. Survival and recovery from ARDS are dependent on returning the alveolus to a functional state. This must involve removal of the edema fluid, inflammatory cells, and repair of the epithelium and endothelium. Many studies have reported that MSC modulates lung epithelial and endothelial cells through paracrine effects resulting in improvement of their barrier functions through various mechanisms including secretion of growth factors (KGF, ANG-1) and mitochondrial transfer through EVs [2, 11–13]. To model alveolar–capillary barrier in vitro we use primary murine or human alveolar epithelial and pulmonary microvasculature endothelial cells and to model ARDS environment we stimulate cells with LPS, cytomix (mixture of pro-inflammatory cytokines characteristic of ARDS (TNF-a, IL-1b, and INFy)) or samples of bronchoalveolar lavage fluid or plasma from ARDS patients available from past and ongoing clinical trials, samples from healthy volunteers are used as a control. The most widely used methods to assess barrier integrity are measurements of the (1) levels of the barrier permeability to the high molecular weight FITC-Dextran, (2) levels of transepithelial (or endothelial) electrical resistance TEER, and (3) more recently developed technology based on the measurement of electrical impedance. In addition, to achieve a more complex evaluation of reparative capacities of MSCs it is essential to assess their effects on the capacity of endothelial and epithelial cells to migrate and proliferate, the functional properties which are necessary for the full restoration of the barrier integrity. These functional properties can be evaluated by wound-scratch

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assay with subsequent staining with Ki67 Ab. Ki67 is a transcription factor, expressed only in proliferating cells [14] and thus in one assay the impact of MSC secretome on both migration and proliferation of target cells could be examined. The general protocol described below is divided into three primary steps. First, we describe techniques for isolation of mesenchymal cells derived from bone marrow as well as isolation of primary murine pulmonary endothelial and epithelial cells from the lung tissue. In the second part, we describe our approach for generation of MSC Conditioned Medium (which represent full MSC secretome), isolation of MSC-derived extracellular vesicles (as the major functional component of MSC secretome), and Transwell-co-culture system which allows to account for contribution of intercellular communication for MSC effect (in this system, MSCs are able to receive and respond to paracrine signals from the target cells). Finally, in the third part, we describe methods to evaluate the paracrine effects of MSC on the barrier permeability using FITC-Dextran, TEER, and impedance-based techniques. Finally, we finish with wound-scratch assay allowing to investigate effects of MSC on reparative capacities of cells through the ability to close the wound via migration and proliferation.

2 2.1

Materials Animals

2.2 Bone Marrow Mesenchymal Stromal Cells (MSCs) Isolation

The procedures involving animals have to be in accordance with national law. BALB/c mice, C57BL/6 mice, or Wistar rats are the strains commonly used for cell extraction and isolation [15, 16]. Typically, young animals (6–8 weeks old) are used. 1. 15 mL Conical tube. 2. Iscove’s Modified Dulbecco’s Medium (IMDM-MSC) complete: IMDM, 15 mM HEPES in solution, 1% L-glutamine in solution of 0.85% NaCl, 15% heat-inactivated fetal bovine serum (FBS), 1% penicillin-streptomycin antibiotic in solution of 0.85% NaCl, pH 7.4. 3. T25 flasks. 4. T75 flasks. 5. T175 flasks. 6. Phosphate-buffered saline (PBS). 7. 0.05% Trypsin-EDTA. 8. Scissors. 9. Forceps.

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2.3 Lung Endothelial Cells Isolation

1. Sterile petri dish. 2. 1% Type I collagenase. 3. Fetal calf serum (FCS) (heat-inactivated). 4. 70 μm Cell strainer. 5. 50 mL Conical tube. 6. Dynabeads biotin binder. 7. Magnetic stand—DynaMag. 8. Biotinylated Anti-CD54 (Biolegend). 9. 2% Gelatine diluted in PBS. 10. Tissue culture coated 6-well plates. 11. PBS. 12. IMDM complete medium (IMDM-L Endothelial C): IMDM, 20% calf serum, 1% L-glutamine in solution of 0.85% NaCl, 1% penicillin-streptomycin antibiotic in solution of 0.85% NaCl, pH 7.4. 13. Scissors. 14. Forceps. 15. PECAM-1/CD31 (eBioscience). 16. ICAM-2 (eBioscience). 17. VE-cadherin (eBioscience).

2.4 Lung Epithelial Cells Isolation

1. PBS. 2. Heparin solution: 10 mL PBS with 1000 IU heparin. 3. 18G needle cannula. 4. 0.5 M PBS/EDTA (adjust to pH 8.0). 5. Elastase solution: 4 IU/mL of elastase in Ham’s F12 medium. 6. Digestion solution: 0.5 mg/mL DNAse I, 0.01 mg/mL bovine serum albumin, 1% penicillin-streptomycin in Ham’s F12 medium. 7. 50 mL conical tube. 8. Forceps. 9. Scissors. 10. Syringes. 11. HBSS solution: 12.5 μg/mL amphotericin B in solution, 0.1 M EDTA in solution, 10 mM HEPES in solution, 2% FBS, 1% penicillin-streptomycin in Hanks’ Balanced Salt Solution. 12. FBS.

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13. IMDM complete medium (IMDM-L Epithelial C): 10% FBS, 1% penicillin-streptomycin antibiotic in solution of 0.85% NaCl (pH of medium to 7.4). 14. 70 μm Cell strainer. 15. Red blood lyser: 4.15 g NH4Cl, 500 mg KHCO3, and 18.6 mg EDTA in 500 mL 1 N deionized water, pH 7.4. 16. Biotinylated anti-mouse CD45, clone 30-F11, 0.5 mg/mL (eBioscience). 17. Biotinylated anti-mouse CD16/32, clone 2.4G2, 0.5 mg/mL (BD, Pharmingen™). 18. Biotinylated anti-mouse CD31, clone MEC13.3, 0.5 mg/mL (Biolegend). 19. Biotinylated anti-mouse TER119, clone TER119, 0.5 mg/mL (eBioscience). 20. Collagen type IV; Matrigel; or Fibronectin. 21. anti-EPCAM. 22. anti-SP-C. 2.5 Investigation of the MSC Paracrine Effect

1. T175 flasks. 2. IMDM-MSC (see Subheading 2.2). 3. IMDM-conditioning: IMDM, 1% FBS, 1% L-glutamine in solution of 0.85% NaCl, 1% penicillin-streptomycin antibiotic in solution of 0.85% NaCl, pH 7.4. 4. IMDM-EV-depleted: IMDM, 15 mM HEPES in solution, 1% L-glutamine in solution of 0.85% NaCl, 1% penicillinstreptomycin antibiotic in solution of 0.85% NaCl. 5. Phosphate-buffered saline (PBS). 6. 1
Trypsin. 7. Trypan blue. 8. 15 mL conical tube. 9. Transwell inserts of 0.4 μm pores suitable for a 24-well plate (Greiner Bio-One, Germany). 10. Tissue culture coated 24-well plates.

2.6 Assays to Measure Barrier Permeability

1. Transwell inserts of 0.4 μm pores suitable for a 24-well plate. 2. Tissue culture coated 24-well plates. 3. Forceps. 4. 70 KDa FITC-Dextran. 5. E-plate (ACEA Bioscience, San Diego, CA, USA). 6. Black 96-well plate with a transparent bottom. 7. PBS.

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8. Fixative: 4% Paraformaldehyde (PFA) in PBS; pH 7.4. 9. 0.5% Triton-X-100 in PBS. 10. Ki67 monoclonal antibody (SolA15): 1:200 dilution in 10% NGS (eBioscience). 11. 1 mg/mL DAPI solution. 12. Hoechst nuclear stain (undiluted). 13. 10% Normal goat serum (NGS) in PBS. 14. Goat Anti-Mouse/Rat Alexa Fluor (AF) 594: 1:200 dilution in 10% NGS (Thermo Fisher Scientific). 2.7

Equipment

1. Water bath (37  C). 2. Centrifuge. 3. Tube rotator. 4. Laminar flow cabinet (safety cabinet class 2). 5. Hemocytometer and coverslips (Improved Neubauer). 6. Incubator. 7. Microscope. 8. Ultracentrifuge Optima XNP (Beckman coulter). 9. XCELLigence RTCA System (ACEA BIO, model: xCELLigence RTCA SP System). 10. Epithelial Voltohmmeter (World Precision Instruments). 11. ENDOHM Chamber (World Precision Instruments). 12. FLUOstar Omega microplate reader. 13. EVOS FL Auto epifluorescent microscope. 14. Leica SP8 confocal microscope. 15. Image analyzing software (ImageJ).

3

Methods

3.1 Mesenchymal Stromal Cells Isolation from Bone Marrow

This protocol can be used to isolate bone marrow mesenchymal stromal cells (MSCs) derived from different species (see Note 1). 1. Pre-warm all media solution in a water bath 1–2 h prior to isolation and pre-warm the centrifuge to 21  C prior to isolation. 2. Euthanize the animal and remove the femurs and tibias under aseptic conditions. 3. Cut the proximal and distal ends from bones and place inside 1 mL pipette tip in a 15 mL conical sterile tube. 4. Centrifuge at 450
g for 5 min at room temperature (RT). This step is performed to remove bone marrow contents from

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Fig. 1 Bone marrow Isolation scheme. The end of the bone is cut and discarded, the bone is placed into 1 mL pipette tip inside a 15 mL conical sterile tube and centrifuged. After centrifugation, the 1 mL pipette tip is removed and discarded, and the bone marrow extract resuspended in complete medium and the content place in T25 culture flasks. MSCs are incubated in the 37  C in a humidified atmosphere containing 5% CO2. Each 2–3 days, check cell density and remove dead cells by washing with pre-warmed PBS. Adherent cells exhibit similar proliferation rates, and upon reaching 80% confluence, cells should be expanded

each isolated bone, alternatively, the bone marrow can be flushed out with 1–2 mL of complete medium using a syringe and needle directly into a 15 mL conical sterile tube (see Fig. 1). 5. After centrifugation, remove the 1 mL pipette tip with the bone inside and resuspend the cell pellet in 3.5 mL of Iscove’s Modified Dulbecco’s Medium (IMDM-MSC, see Subheading 2.2) complete. 6. Adjust the volume to 5 mL of complete medium and then place the content in T25 culture flasks. MSCs are incubated in the 37  C in a humidified atmosphere containing 5% CO2. Each 2–3 days, check cell density and remove dead cells by washing with pre-warmed PBS. 7. Upon reaching 80% confluence, cells are expanded to T75 Flasks and maintained in IMDM-MSC complete medium. Usually, one T25 MSCs flask is expanded to three T75 flasks and T75 flask is expanded to two T175 flasks. 8. For expansion, trypsinize the cells with 0.05% Trypsin-EDTA solution for 3–5 min, inactivate trypsin with twice the volume of IMDM complete medium, centrifuge at 450
g for 5 min at RT, discard the supernatant, and resuspend the cells in IMDM complete medium. For each expansion, consider one passage of the cells (Extraction ¼ P1, T25 for T75 ¼ P2, T75 for T175 ¼ P3). For experimental use, cells are used from P3-P5. 3.2 Lung Endothelial Cells Isolation

1. Pre-warm all media solutions in the water bath 1–2 h prior to isolation and pre-warm the centrifuge to 21  C prior to isolation. 2. Euthanize the animal, remove the heart and lungs en bloc under aseptic conditions.

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3. Under a laminar flow cabinet, isolated from the heart the lung tissue, rinse with cold PBS, transfer to a sterile Petri dish, and cut into small pieces (approximately 0.2–0.8 cm2). 4. Collect the dissected pieces and put in a conical sterile tube with 2 mL of 1% type I collagenase for 30 min to 1 h at 37  C. Gently vortex the tube every 10 min. 5. Add 6 mL of IMDM-L Endothelial C (see Subheading 2.3) complete medium to inhibit the proteolysis and filter using 70 μm cell strainer into a 50 mL sterile conical tube. Centrifuge at 600
g for 5 min at room temperature (RT). 6. Discard the supernatant and resuspend the cells using warmed PBS. Repeat the centrifugation step using the same parameters (600
g, 5 min, RT) described. 7. Wash the Dynabeads twice (25 μL of magnetic beads in 3 mL of PBS) using a 15 mL sterile conical tube. Place the conical tube on the magnetic stand to separate the beads. Discard the supernatant and resuspend the Dynabeads in PBS. 8. Resuspend the cell pellet in PBS containing biotinylated antiCD54 according to the manufacturer’s instruction and incubate for 30 min in ice or fridge. After 30 min, wash with PBS and centrifuge. Incubate the cell pellet with Dynabeads biotin binder for 30 min in ice or fridge. 9. During the incubation time with a primary antibody, coat the 6-well plate with 2% gelatine and leave in the hood for 1 h. Before plating the isolated cells, remove the excess gelatine from the plates and allow to dry for 20 min in the laminar flow cabinet (see Note 2). After, re-sterilize the plate in UV light for 30 min. 10. After 30 min of incubation with Dynabeads, place the 15 mL conical tubes into a magnetic stand. The Dynabeads, along with differentiated cells, are attracted and attached to the magnet. Carefully, remove the cell suspension without disturbing the attached beads and transfer into a new 15 mL sterile conical tube. Repeat the process three times for the maximal removal of remaining beads from the cell suspension. 11. Resuspend the cell pellet in IMDM-L Endothelial C complete medium and plate the cells in pre-coated 6-well plate. Every 2–3 days, check cell density, remove dead cells by washing with pre-warmed PBS, and change the medium. 12. The purity of endothelial cells can be determined using FACS analysis for endothelial surface markers (PECAM-1/CD31, ICAM-2, and VE-cadherin).

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3.3 Lung Alveolar Epithelial Cells Isolation

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1. Pre-warm all media solution in a water bath 1–2 h prior to isolation and pre-warm the centrifuge to 21  C prior to isolation. 2. Euthanize the animal, insert into the trachea a 18G cannula, fix the cannula with a cotton string, and remove the heart and lungs en bloc under aseptic conditions. 3. Under a laminar flow cabinet, gently inject 10 mL heparin solution in the right ventricle. 4. Using a cannula, wash the lung three times with 1 mL of PBS/EDTA slowly (see Note 3). Remove the lungs, wipe all connective tissues and still with the cannulated trachea, place the lung on a petri dish on ice for 30 min. 5. With the trachea still cannulated, inject 1 mL of elastase solution into the lungs for 10 min. Collect and inject 0.5 mL for 5 min. Repeat the process four times. 6. After the elastase washes, cut the lungs into small pieces (approximately 0.2–0.8 cm2) and transfer to a 50 mL conical tube with 5 mL of digestion solution and incubate for 15 min at 37  C. 7. Dissociate the tissue by gentle agitation, wash and incubate with 10 mL of HBSS solution. Transfer in the 50 mL sterile conical tube and mix gently using 1 mL pipette tip. 8. The remaining solution was transferred to a 70 μm cell strainer into a new 50 mL sterile conical tube and the reaction stop with 5 mL of FBS and centrifuge at 600
g for 5 min at 4  C. 9. Resuspend the cells and add 1 mL of red blood lyser for 1 min on the ice, wash with HBSS solution, and repeat the centrifugation step (600
g, 5 min, 4  C). 10. Resuspend the cells using complete medium with 10% FBS (IMDM-L Epithelial C, see Subheading 2.4) and plate the cells in the pre-coated wells on collagen type IV, matrigel, or fibronectin-coated surfaces or in other ways depending on your experimental question. Every 2–3 days, check cell density, remove dead cells by washing with pre-warmed PBS, and change the medium. 11. The purity of cells can be enriched using biotinylated antibodies against lineage markers: anti-CD45—mark hematopoietic cells (CD45 positive), anti-CD16/32—alveolar macrophages (CD45 positive and CD16/32 positive), anti-CD31—endothelial cells (CD31 positive), anti-Ter119—erythroid cells (Ter119 positive). 12. Incubate for 60 min on ice with gentle intermittent mixing of the cells. At the end of the antibody incubation, wash the cells once in appropriated medium (centrifuge at 300
g for

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10 min at 4  C) and resuspend in 5 mL of complete medium. Transfer cells to 15 mL tube with an appropriate number of washed Dynabeads biotin binder. Incubate in the cold room for 30 min on the rotator to prevent the beads sitting to the bottom. 13. After incubation, place the tube in the magnetic separator. Lineage positive cells will get stuck to the tube wall. Gently pipette out unattached cells and place into a new tube. Place the fresh tube in the magnetic and repeat magnetic depletion for a total of three times. 14. The purity of epithelial cells can be determined using FACS analysis for epithelial markers (EPCAM, SP-C). 3.4 Investigation of MSC Paracrine Effect 3.4.1 Generation of Conditioned Medium

1. MSCs (see Subheading 3.1) are grown to around 80% confluence in T175 flask under standard conditions (IMDM complete medium, 37  C in a humidified atmosphere containing 5% CO2). 2. Remove the MSC medium (IMDM-MSC) and wash two times with PBS. MSCs are used in experiments at passage 3–5. 3. Add 5 mL of 1
trypsin and leave in the incubator for approximately 2–5 min. 4. Once MSCs cells are detached, add 5 mL of complete medium do inhibit trypsin activity and transfer the cells into a 15 mL sterile conical tube. 5. Centrifuge the cells at 450
g for 5 min at RT. 6. After centrifugation, aspirate the supernatant and resuspend the cell pellet in 1 mL of medium. 7. Count the cells using a hemocytometer, using trypan blue exclusion and seed corresponding cell densities on the plate using the complete medium. 8. The following day, check if the MSC cells are attached to the plate, wash with PBS and replace the medium with IMDMconditioning to generate culture conditioned medium (CM). 9. After 24 h collect the CM and centrifuge at 10,000
g/rcf to remove detached cells and cells debris.

3.4.2 Indirect Co-Culture

1. MSCs are grown to around 80% confluence in T175 flask under standard conditions (IMDM complete medium, 37 in a humidified atmosphere containing 5% CO2). 2. Remove the MSC medium and wash two times with PBS. MSCs are used in experiments at passage 3–5. 3. Add 5 mL of 1
trypsin and leave in the incubator for approximately 2–5 min.

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4. Once MSCs cells are detached, add 5 mL of complete medium to inhibit trypsin activity and transfer the cells into a 15 mL sterile conical tube. 5. Centrifuge the cells at 450
g for 5 min at RT. 6. After centrifugation, aspirate the supernatant and resuspend the cell pellet in 1 mL of medium. 7. Count the cells using a hemocytometer, using trypan blue exclusion and seed MSCs in the transwell insert (use 0.4 μm porous hanging cell culture transwell inserts). Pre-soak the inserts in the medium for at least 20 min prior to seeding the MSCs to facilitate cell adherence (see Note 4). 8. Determine corresponding cell densities and plate with the cell type of interest in the bottom chamber. For our experience, we co-culture MSCs at a 5:1 ratio (cell of interest to MSCs). 3.4.3 Isolation of Extracellular Vesicles

1. MSCs are grown to around 80% confluence under standard conditions (IMDM complete medium, 37 in a humidified atmosphere containing 5% CO2). 2. To generate extracellular vesicles (EVs), MSCs are washed 2
with PBS and their medium replaced with 15 mL of IMDM-EV-depleted medium and incubated for 48 h at 37 in a humidified atmosphere containing 5% CO2. 3. After 48 h, the medium is collected in 15 mL sterile conical tubes and centrifuged at 2000
g/rcf for 45 min at 4  C to remove cellular debris. 4. Transfer the supernatant into ultracentrifuge tubes ensuring that they are equally balanced (see Note 4). Place the tube inside to a 70Ti rotor and centrifuge at 100,000
g/rcf (acceleration ¼ 9, deceleration ¼ 5) for 3 h at 4  C to isolate the EVs. 5. After centrifugation, gently aspirate the supernatant and resuspend the EVs in an appropriate volume according to the final MSC cell count (see Notes 5 and 6).

3.5 Assays to Measure Barrier Permeability 3.5.1 Permeability to FITC-Dextran

1. In 24-well plate, seed the cells at a density of 1
105 per well on the pre-coated 0.4 μm transwell inserts (see Note 2). 2. Incubate the plate in a 37  C in a humidified atmosphere containing 5% CO2, with medium changes every 2–3 days. 3. Transfer the inserts to the wells with medium using forceps for another dry well and wait 5–10 min (see Note 7). Observe if medium from the upper chambers leaks into the lower chamber. If there is no leak, proceed to the next step. If there is a leak, wait for 2–3 days until cells become fully confluent to ensure the formation of cell junctions and a good barrier.

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4. Following monolayer formation, replace the medium with the desired stimulation (cytokines, molecules which induced inflammatory responses such as lipopolysaccharide or recombinant proteins, as thrombin) or your treatment of interest (MSC CM, EVs, MSC co-culture) and incubate the plate in the 37  C in a humidified atmosphere containing 5% CO2 for desired time period. 5. Remove the stimulation and replace with fresh medium using 100 μL of 70 KDa FITC-Dextran in the upper chamber and 500 μL to the lower chamber. Add a positive control in which FITC-Dextran also is added to insert without cells. Add a blank well which contains PBS added to the upper chambers of inserts without cells. 6. Incubate the cells in the presence of FITC-Dextran in the 37  C in a humidified atmosphere containing 5% CO2 for 30 min. 7. After 30 min, 100 μL samples from the lower chamber are collected and added to a black 96-well plate with a transparent bottom. Every condition should be done at least in triplicate. 8. Fluorescence intensity should be read immediately using a microplate reader at 485 nm excitation and 520 nm emission. 9. Results are presented as % of positive control. A schematic outline procedure is shown in Fig. 2. 3.5.2 XCELLigence® Real-Time Cell Analysis (RTCA)

Stimulation

1. Pre-coat the E-plate as previously described (see Note 2). Add 50 μL of a pre-warmed medium into each well and leave the E-plate in the incubator for at least 15 min before starting the experiment to ensure that the culture medium and E-plate surface achieve equilibrium. FITC-Dextran 70KDa

Time incubation

30 min Collect lower chamber medium

Cells monolayer

Aliquot triplicate Read the microplate

Fig. 2 Schematic outline procedure of FITC-Dextran. Following monolayer formation, replace the medium with the desired stimulation and incubate the plate in the 37  C in a humidified atmosphere containing 5% CO2 for the desired time period. Remove the stimulation and replace with fresh medium using 100 μL of 70 kDa FITCDextran in the upper chamber and 500 μL to the lower chamber. Incubate cells in the presence of FITCDextran in the 37  C in a humidified atmosphere containing 5% CO2 for 30 min. After 30 min, 100 μL samples from the lower chamber are collected and added to a black 96-well plate with a transparent bottom. Fluorescence intensity should be read immediately using a microplate reader at 485 nm excitation and 520 nm emission

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2. Fill the well of E-plate with 100 μL of cell medium and place the plate in the XCELLigence RTCA System, which is set up in a 37  C in a humidified atmosphere containing 5% CO2. 3. Set the program to detect the resistance at the designated time points (every hour, every 15 min) and measure the background of E-plate with medium (see Note 8). 4. After background measurements, seed 2
105 cells in each well and maintain the plate in the incubator for at least 2–3 h until the cells adhere to the E-plate (see Note 9). 5. Connect the E-plate at the RTCA system and monitor the cells until they reach confluency. 6. After the cell index curve stabilizes, remove the E-plate of the RTCA system and add the desired stimulation/treatment into each well. Every condition should be done in triplicate. 7. Put the plate back into the XCELLigence RTCA system in the 37  C in a humidified atmosphere containing 5% CO2. 8. Set up the desired detection period and time interval and start the recording. 9. The cell index of every time point should be normalized over time zero to determine the relative cell index. The average is calculated and plotted versus time. 3.5.3 Transepithelial/ Endothelial Electrical Resistance (TEER)

1. TEER measurements are routinely used to characterize monolayer integrity in the context of cell monolayer permeability experiments. 2. Seed the 1
105 cells/cm2 in transwell inserts (0.4 μm) pre-coated as previously described. Cells are maintained in culture for up 2 weeks, with media changes every 2–3 days. 3. Transfer the inserts to the wells with medium using forceps for another dry well and wait 5–10 min (see Note 7). Observe if medium from the upper chambers leaks into the lower chamber. If there is no leak, proceed to the next step. If there is a leak, wait for 2–3 days until cells become fully confluent to ensure the formation of cell junctions and a good barrier. 4. Following monolayer formation, replace the medium with the desired stimulation (cytokines, molecules which induced inflammatory responses such as lipopolysaccharide or recombinant proteins, as thrombin), your treatment of interest (MSC CM, EVs, MSC co-culture on Transwells) and incubate the plate in the 37  C in a humidified atmosphere containing 5% CO2 for desired time period. 5. Add 2 mL of medium to the Epithelial Voltohmmeter (EVOM). This medium should be the same medium used to culture the cells on the inserts. Ensure the mode switch is turned to R.

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6. Turn on the power switch and press the Test R button. The meter display should be read 1000 Ω  10 Ω. 7. Change the medium from the Transwell inserts and add 500 μL of cell culture medium to the apical surface of each Transwell. Transfer each Transwell, one at a time, into the chamber. The chamber’s cap contains a pair of concentric electrodes. Current flows between these symmetrically opposing circular disc electrodes and the resistance imparted by the cell layer are measured. 8. Always measure the resistance on a blank insert (no cells). This should then be subtracted from the resistance reading across cells obtain the true resistance reading of the cell layer. 9. Gently place the cap back on the chamber. Measure the TEER by holding the measure button down until the values stabilize. 10. The value obtained on the screen is the TEER in Ohms (Ω). This should be transformed into Ω.cm2 by multiplying the obtained value by the growth area of the Transwell membrane. Repeat this measurement twice for each insert (3 readings in total). A schematic outline procedure is shown in Fig. 3.

Fig. 3 Transepithelial/endothelial electrical resistance. Following monolayer formation, transfer each well one at a time into the chamber. The transwell insert in which the cells of interest are transferred into an Endohm chamber. Current flows between these symmetrically opposing circular disc electrodes and the resistance imparted by the cell layer are measured (a). Alternatively, if the experiments are not performed on transwell inserts, resistance can be assessed by inserting the electrode into the well of the plate and performing the same sequence of measurements (b)

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3.6 Assays to Investigate Effects of MSC on Reparative Capacities

1. Seed and grow cells with an appropriate medium at the desired density and incubate at 37  C in a humidified atmosphere of 5% CO2 until the formation of a monolayer, confirm by daily microscopical analysis.

3.6.1 Scratch Wound Healing Assay

2. A horizontal line should be drawn on the reverse of the plate across the middle of the wells to orientate the direction of scratch (vertical). 3. Once cells have formed a monolayer, gently and slowly scratch the monolayer with a new and sterile 1 mL pipette tip across the center of well. Scratch a straight line in one direction (see Note 10). 4. Remove the debris by washing the cells once with 1 mL of PBS or HBSS and then add 1 mL of appropriate medium. The medium may contain stimulus of interest that you want to test, e.g., lipopolysaccharide, cytokines, conditioned medium, extracellular vesicles, chemicals that inhibit/promote cell proliferation. 5. The area of the wound is microscopically analyzed at 0, 6, 18, 24 h or at later time points according to the research question. 6. Plates are imaged using an inverted microscope at 10
magnification. 7. Take two images of each wound at 0 h (above and below a horizontal line drawn across the well). Compare these to the two images (above and below the horizontal line) of the same wound taken at later time points. 8. The wound area is then quantitatively evaluated using software such as Photoshop or Image J. For each well, calculate the percentage of wound closure using the equation adapted from Buachan et al. [17]. A schematic procedure of scratch wound assay is shown in Fig. 4.

3.6.2 Immunofluorescence Staining of Scratch Wound Assays with Ki67

1. Ki67 is a transcriptional factor which is expressed in the nuclei of proliferating cells. After the experiments, wash the cells three times with PBS and fix using 4% PFA for 15 min at RT. 2. After fixation, excess of PFA should be removed and cells washed twice with PBS prior to permeabilization using 0.5% Triton X-100 for 20 min at RT. 3. Wash the cells three times using PBS and after, block with normal goat serum (NGS) in PBS (200 μL for 2 h in 10%). 4. Dilute the primary antibody Ki67 (dilution of 1:200) in 10% NGS in PBS and apply 100 μL per well and incubate at 4  C overnight. For a secondary-only control, add 100 μL of 10% NGS.

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B

(A+B/2 – C+D/2) * 100

= % wound closure

(A+B/2) Image “A” at 0h or “C” thereafter

Image “B” at 0h or “D” thereafter

Edge of the wound

Horizontal line across well

Fig. 4 Schematic procedure of Scratch Assay. (a) Representative images showing wound repair in A549s cell line in different conditions: C (control group), LPS (E. coli Lipopolysaccharide) mimetizing inflammatory environment and LPS-Evs (A549s incubated with Extracellular Vesicles-EVs). Images were taken by Zeiss Axiovert 25 CFL microscope. Original magnification
20. (b) Diagram showing the location of wound imaging and the equation to calculate the percentage of wound closure. The blue box is image “A” at 0 h or “C” thereafter, the red box is image “B” at 0 h or “D” thereafter

5. After overnight incubation, wash the wells three times with 1
PBS. Add Goat Anti-Mouse/Rat Alexa Fluor (AF) 594 (dilution of 1:200) in 10% NGS (100 μL) in the dark for 1 h at RT. For a primary-only control, add 100 μL of 10% NGS. 6. Wash the cells three more times with PBS and counter-stain with DAPI diluted in PBS or Hoechst (undiluted) for 5 min in the dark at RT. 7. Wash the cells three more times with PBS and add PBS or mounting medium in the wells. The cell can be stored in the dark at 4  C until imaging. 8. Cells can be imaged using an EVOS FL auto epifluorescent microscope or confocal microscope. Images of the well should be taken at 4
magnification (see Fig. 5). Quantitative analysis can be performed using ImageJ. 9. To quantify, a grid of individual squares each of 100,000 pixels2 in size can be drawn over each image using ImageJ. Determine the average number of Ki67-positive cells per 100,000 pixels2.

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Fig. 5 Immunofluorescence staining for Ki67. Representative images of scratch wound assay stained with Ki67 using a Life Technologies EVOS FL Auto epifluorescent microscope. In the upper panels, representative images of cells at 0 h stained with DAPI (left) and Ki67 (middle). In the bottom panel, images are taken 24 h demonstrating cell proliferation and Ki67-positive cells (middle). Quantitative image analysis can be performed on ImageJ by creating a grid of 100,000 pixels2. Four squares of the grid for each well and cells displaying positive Ki67 staining should be counted

4

Notes 1. After isolation, MSCs must be characterized according to the guidelines of the International Society for Cellular Therapy (ISCT). First, MSC must be plastic-adherent in standard culture conditions. Second, must express specific markers like CD105, CD73, and CD90 and lack expression of CD45, CD34, CD14 or CD11b, CD79 alpha or CD19, and HLA-DR surface molecules. Third, must differentiate to osteoblasts, adipocytes, and chondroblasts in vitro [18]. 2. To prepare the gelatine to coat the plates, make up a 0.1% gelatine solution (100 mg gelatine powder in 100 mL of sterilized distilled water), sterilize by autoclaving and set aside until use. Under the sterile hood, add 1 mL of 0.1% gelatine solution in 6-well plates and leave for 1 h in the hood. Remove the excess of gelatine and let the plate open in a laminar flow hood to dry (15–30 min). Re-sterilize by UV light under the hood for 30 min and the plate are ready for experiments. The plate can be prepared 1 day before the experiment and can be sealed by parafilm and stored in the fridge. For epithelial cells, plates should be pre-coated wells on collagen type IV, matrigel, or fibronectin.

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3. The trachea of rats and mice can be cannulated with an 18G needle. To cannulate mice, ideally, the needle can be cut/broken in half to be cannulated, thereby avoiding and rupturing the trachea. For rats, it is not necessary to decrease the size of the needle. After cannulation, it is advisable to use a cotton thread to attach the cannula to the trachea. 4. Lung fibroblasts can be used as a cell control for the MSCs, to determine whether the effects observed were specific to MSCs. To measure effects on permeability, MSCs should be cultured in the bottom chamber; to measure the effects on wound closure, MSCs should be cultured on the insert. The above assays can be utilized with both murine and primary human cells, we use primary human small airway epithelial cells and pulmonary microvasculature endothelial cells commercially available from PromoCell. 5. It is established that the pellet of EVs derived from 1
106 MSCs is resuspended in 10 μL PBS or work-medium. Some protocols for EVs extraction add additional centrifugation in PBS using the same parameters to minimize contamination with protein aggregates. 6. After extraction, EVs must be characterized according to the guideline of the International Society for Extracellular Vesicles (ISEV) [19]. 7. Every medium change in this step should be done carefully and slowly as to not activate or detach the cells. Pipet tips should not touch the membrane. Refill the chambers with fresh medium in a dropwise manner to avoid shear flow. 8. Start the RTCA program and set up the schedule page for the plate running procedure. Experiments should be divided into multiple steps, which consist of one or several sweeps (one scan across all selected wells). Step 1 is considered the background (baseline) measurement. Step 1 should be scan prior to the addition of cells and is pre-programmed to be one sweep. Make a separate step for continuous scanning with different intervals between sweeps and shorter intervals between sweeps are useful for detecting changes and short events. 9. It is important to optimize the density of cells and the number of measurements and intervals for your cell line to avoid wasting E-plates. Some cells adhere very quickly and spread within 2 h. When starting the XCELLigence measures for the first time, you can use short intervals to monitor cell index and adjust the parameters accordingly. 10. The resulting distance of the gap, therefore, equals to the diameter of the end of the tip used. The gap distance can be adjusted by using different types of tips.

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References 1. de Castro LL, Lopes-Pacheco M, Weiss DJ et al (2019) Current understanding of the immunosuppressive properties of mesenchymal stromal cells. J Mol Med Ther Ber 97(5):605–618 2. Fergie N, Todd N, McClements L et al (2019) Hypercapnic acidosis induces mitochondrial dysfunction and impairs the ability of mesenchymal stem cells to promote distal lung epithelial repair. FASEB J 33(4):5585–5598 3. Morrison TJ, Jackson MV, Cunningham EK et al (2017) Mesenchymal stromal cells modulate macrophages in clinically relevant lung injury models by extracellular vesicle mitochondrial transfer. Am J Respir Crit Care Med 196(10):1275–1286 4. Jackson MV, Morrison TJ, Doherty DF et al (2016) Mitochondrial transfer via tunneling nanotubes is an important mechanism by which mesenchymal stem cells enhance macrophage phagocytosis in the in vitro and in vivo models of ARDS. Stem Cells 34 (8):2210–2223 5. Silva JD, Lopes-Pacheco M, Paz AHR et al (2018) Mesenchymal stem cells from bone marrow, adipose tissue, and lung tissue differentially mitigate lung and distal organ damage in experimental acute respiratory distress syndrome. Crit Care Med 46(2):e132–e140 6. Maron-Gutierrez T, Silva JD, Asensi KD et al (2013) Effects of mesenchymal stem cell therapy on the time course of pulmonary remodeling depend on the etiology of lung injury in mice. Crit Care Med 41(11):e319–e333 7. Brown C, McKee C, Bakshi S et al (2019) Mesenchymal stem cells: cell therapy and regeneration potential. J Tissue Eng Regen Med 13(9):1738–1755 8. Murray LM, Krasnodembskaya AD (2019) Concise review:intercellular communication via organelle transfer in biology and therapeutic applications of stem cells. Stem Cells 37 (1):14–25 9. Weiss DJ, English K, Krasnodembskaya A et al (2019) The necrobiology of mesenchymal stromal cells affects therapeutic efficacy. Front Immunol 10:1228. https://doi.org/10. 3389/fimmu.2019.01228 10. Ware LB, Bastarache JA, Bernard GR (2017) Acute respiratory distress syndrome. In:

Vincent J-L et al (eds) Textbook of critical care, 7th edn. Elsevier, Philadelphia, pp 413–424 11. Lee JW, Fang X, Gupta N et al (2009) Allogeneic human mesenchymal stem cells for the treatment of E. coli endotoxin-induced acute lung injury in the ex vivo perfused human lung. Proc Natl Acad Sci U S A 106 (38):16,357–16,362 12. X1 F, Neyrinck AP, Matthay MA, Lee JW (2010) Allogeneic human mesenchymal stem cells restore epithelial protein permeability in cultured human alveolar type II cells by secretion of angiopoietin-1. J Biol Chem 285 (34):26211–26222 13. Islam MN, Das SR, Emin MT et al (2012) Mitochondrial transfer from bone-marrowderived stromal cells to pulmonary alveoli protects against acute lung injury. Nat Med 18 (5):759–765 14. Miller I, Min M, Yang C et al (2018) Ki67 is a graded rather than a binary marker of proliferation versus quiescence. Cell Rep 24 (5):1105–1112.e5 15. Samary CS, Ramos AB, Maia LA et al (2018) Focal ischemic stroke leads to lung injury and reduces alveolar macrophage phagocytic capability in rats. Crit Care 22(1):249 16. Abreu SC, Antunes MA, Xisto DG et al (2017) Bone marrow, adipose, and lung tissue-derived murine mesenchymal stromal cells release different mediators and differentially affect airway and lung parenchyma in experimental asthma. Stem Cells Transl Med 6(6):1557–1567 17. Buachan P, Chularojmontri L, Wattanapitayakul S (2014) Selected activities of Citrus Maxima merr. Fruits on human endothelial cells: enhancing cell migration and delaying cellular aging. Nutrients 6:1618–1634 18. Dominici M, Le Blanc K, Mueller I et al (2006) Minimal criteria for defining multipotent mesenchymal stromal cells. The International Society for Cellular Therapy position statement. Cytotherapy 8(4):315–317 19. The´ry C, Witwer KW, Aikawa E et al (2018) Minimal information for studies of extracellular vesicles (MISEV2018): a position statement of the International Society for Extracellular Vesicles and update of the MISEV2014 guidelines. J Extracell Vesicles 7(1):1535750

Chapter 6 In Vitro Methods to Evaluate the Effects of Mesenchymal Stem Cells on TGF-β1-Induced Pulmonary Fibrosis Ying-Wei Lan, Chuan-Mu Chen, and Kowit-Yu Chong Abstract A co-culture model of mesenchymal stem cells (MSCs) and fibroblasts is an efficient and rapid method to evaluate the anti-fibrotic effects of MSCs-based cell therapy. Transforming growth factor (TGF)-β1 plays a key role in promotion of fibroblast activation and differentiation which can induce collagen deposition, increase ECM production in lung tissue, eventually resulted in pulmonary fibrosis. Here, we use this co-culture system and examine the ECM production in activated fibroblasts by western blot and quantitative real-time analysis to understand the therapeutic effects of MSCs. Key words Co-culture, TGF-β1, Pulmonary fibrosis, Extracellular matrix, Conditioned medium

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Introduction Pulmonary fibrosis (PF) is a progressive lung disorder affecting primarily elderly people with unknown etiology. PF carries a poor prognosis, high mortality rates and for which there are no effective treatment. Pulmonary fibrosis is characterized by excessive extracellular matrix (ECM) proteins production and accumulation which result in impaired pulmonary function [1]. Transforming growth factor (TGF)-β1 is considered to be the crucial factor involved in the progression of fibrotic changes [2]. TGF-β1 promotes fibroblast activation and differentiation which can induce collagen accumulation and increase ECM production in lung tissue, eventually lead to pulmonary fibrosis [3, 4]. Clinical trials of mesenchymal stem cells (MSCs)-based cell therapy are the novel approach with great therapeutic potentials for the treatment of lung diseases. MSCs can migrate to damaged sites and promote tissue repair through direct differentiation or secretion of multiple paracrine factors [5]. Mounting evidence suggested that MSCs can inhibit the activation of TGF-β signaling pathways in fibroblast through a paracrine mechanism, consequently, contribute to the development of PF [6–8].

Peggy Stock and Bruno Christ (eds.), In Vitro Models for Stem Cell Therapy: Methods and Protocols, Methods in Molecular Biology, vol. 2269, https://doi.org/10.1007/978-1-0716-1225-5_6, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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This chapter details the methods for setting up an in vitro experiment to evaluate the anti-fibrotic potential of MSCs on TGF-β1-mediated ECM production in fibroblast.

2 2.1

Materials Equipment

1. 10-cm culture petri dish. 2. Transwell inserts. 3. 6-well culture plates. 4. Hemocytometer. 5. Centrifuge. 6. Laminar biosafety hood. 7. CO2 incubator. 8. Spectrophotometer. 9. PCR machine. 10. LightCycler® 480 Instrument. 11. Amersham Imager 600 imaging system. 12. Shaker. 13. 0.22μm filter.

2.2

Cell Lines

1. Mouse bone marrow-mesenchymal stem cell (MSC). 2. MRC-5 cells.

2.3 Media, Buffers, and Solutions

1. Phosphate-buffered saline (PBS). 2. Minimal essential medium (MEM): 10% Fetal bovine serum (FBS), 1% Penicillin/streptomycin (P/S). Store at 4
C. 3. Dulbecco’s Modified Eagle’s Medium/Ham’s Nutrient Mixture F-12 (DMEM/F12): 10% FBS, 1% Penicillin/streptomycin (P/S). Store at 4
C. 4. 0.25% Trypsin-EDTA. Store at 20
C. 5. Opti-MEM medium. Store at 4
C. 6. 2.5 ng/mL TGF-β1 in Opti-MEM medium. 7. 2.5 ng/mL TGF-β1 in Opti-MEM:Conditioned Medium ¼ 1:1 (see Subheading 3.3). 8. UltraPure Water.

2.4 Collagen Detection

1. Sirius Red/Fast Green Collagen detection kit (Chondrex Inc., Redmond, WA, USA). 2. Kahle fixative buffer: 26.88% EtOH, 3.7% formaldehyde, 2% glacial acetic acid.

Evaluation of MSC on TGFβ1-Induced Lung Fibrosis in Vitro

2.5

Western Blot

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1. PRO-PREP™ Protein Extraction Solution. Store at 20
C. 2. BCA Protein Assay Kit. 3. 6 SDS sample buffer: 60% glycerol, 300 mM Tris–HCl, 12 mM EDTA; 12% SDS, 6% 2-mercaptoethanol, 0.05% Bromophenol Blue, adjust to pH 6.8. 4. SDS-PAGE and Bio-Rad’s western blotting systems. 5. PVDF transfer membrane. 6. TBS: 0.2 M Tris-Base, 1.5 M NaCl, adjust to pH 7.6. 7. TBS-Tween (TBS-T): 0.1% Tween-20 in TBS. 8. Blocking buffer: 5% skimmed milk powder in TBS-T. 9. Primary antibody dilution buffer: 2.5% Bovine serum albumin (BSA), 0.02% NaN3 in TBS-T. Store at 4
C. 10. 1:1,000 Anti-Fibronectin in primary antibody dilution buffer (Santa Cruz Biotechnology, Santa Cruz, CA, USA). Store at 4
C. 11. 1:2,000 Anti-Collagen type I in primary antibody dilution buffer (Proteintech Group Inc., Rosemont, IL, USA). Store at 20
C. 12. 1:10,000 Anti-β-actin in primary antibody dilution buffer (Novus Biologicals, Centennial, CO, USA). Store at 20
C. 13. 1:5,000 Horseradish peroxidase (HRP)-conjugated secondary antibodies. Store at 20
C. 14. Western Lightning ECL Plus. Store at 4
C.

2.6 RNA Isolation and Quantitative Real-Time RT-PCR

1. Direct-zol RNA MiniPrep kit. 2. MMLV Reverse Transcription Kit. Store at 20
C. 3. qPCRBIO SyGreen Mix. Store at 20
C. 4. LightCycler® 480 Multiwell Plate 384, white. 5. Primer: Fibronectin forward: 5’-CCCACCGTCTCAACATGCTTAG-3’ Fibronectin reverse: 5’-CTCGGCTTCCTCCATAACAAGTAC-3’ Collagen type I forward: 5’-TCGGCGAGAGCATGACCGAT GGAT-3’ Collagen type I reverse: 5’- GACGCTGTAGGTGAAGCGGC TGT-3’ β-actin forward: 5’-GCGAGAAGATGACCCAGATC-3’ β-actin reverse: 5’-CCAGTGGTACGGCCAGAGG-3’

2.7 Software to Represent Data

Data are represented in bar graphs displaying mean  SD. For multiple comparisons, one-way analysis of variance was used, followed by Dunnett’s post hoc test. All statistical analyses were

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performed with GraphPad Prism (GraphPad Software Inc., San Diego, CA, USA, https://www.graphpad.com). For all analyses, a p value 0.21 hyperoxia. 4. CRL-7449 fibroblast cell line. 5. Anaerobic bags. 6. Distilled water. 7. Inverted microscope.

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Table 2 Primers used for PCR of PMBC markers Gene name

Primer sequence (forward, reverse)

Product size (base pairs, bp)

β-Actin (ACTB)

F GGCGGCACCACCATGTACCCT R CGGACTCGTCATACTCCTGC

308

CD11b

F GCTTCTTCAAGCGGCAATAC R GTGCACACACTTGCACACAG

255

CD14

F GACCTAAAGATAACCGGCACC R GCAATGCTCAGTACCTTGAGG

161

CD31

F AACAGTGTTGACATGAAGAGCC R TGTAAAACAGCACGTCATCCTT

146

CD34

F AATCAGCACAGTGTTCACCAC R TGCCCTGAGTCAATTTCACTTC

161

CD45

F CTCCGCCGCCAATGCAAAACT R GAGCTGTGGTGTGCAAGGCTGAG

144

CD133

F GCCACCGCTCTAGATACTGC R TGTTGTGATGGGCTTGTCAT

123

CD146

F AACAGTGTTGACATGAAGAGCC R TGTAAAACAGCACGTCATCCTT

174

EMS1

F ACAGCAGTGAGTGCAAAAGCA R GCGGTAGCAAGTTTCTCCCC

104

2.5.2 Acetylated Low-Density Lipoprotein (LDL) Uptake and Ulex europaeus Lectin Binding

1. 10μg/mL of 3, 30 -dioctadecyloxacarbocyanine-acetylated low-density lipoprotein (DioAc-LDL) in cEGM. 2. 12-well plates. 3. PBS. 4. Fluorescence inverted microscope. 5. 10μg/mL of fluorescein-tagged Ulex europaeus lectin in PBS.

2.5.3 Single Cell Clonogenic Assay

1. FACS. 2. anti-CD34 conjugated to phycoerythrin (Becton-Dickinson, Heidelberg, Germany). 3. 96-well plates. 4. cEGM. 5. 2% porcine skin gelatine. 6. Hoechst stain. 7. Inverted microscope.

2.5.4 Wound Healing (or Scratch) Assay

1. 25 cm2 culture flask. 2. Pasteur pipette.

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3. PBS. 4. cEGM. 5. Incubator. 6. Live camera. 2.6 Co-Culture of Endothelial Colony-Forming Cells With Peripheral Blood Mononuclear Cells

1. 6-well plates coated with 2% gelatine (see Note 2). 2. cEGM. 3. Water bath. 4. 50 mL Tubes. 5. Centrifuge. 6. 0.4% Trypan blue solution. 7. Neubauer hemocytometer. 8. Normoxid cell culture incubator. 9. Incubator capable to provide FiO2 > 0.21 hyperoxia. 10. Anaerobic bags.

2.7 Microarray Analysis

1. RNA isolation kit of high quality. 2. Human Exon 1.0 ST microarrays (Affymetrix, Santa Clara, CA, USA). 3. Expression Console build 1.4.1.46 (Affymetrix, Santa Clara, CA, USA). 4. Microsoft Excel 2010. 5. Microarray scanner.

2.8 Validation by real-time PCR Analysis

3

1. TaqDNA Polymerase System. 2. Primer: see Table 3.

Methods

3.1 Isolation of PBMCs from Peripheral Cord Blood

1. Under aseptic conditions, collect at least 15 mL of CB samples in containers coated with sodium citrate anticoagulant by passive bleeding as follows with a second assisting person (see Note 1). 2. Compress the umbilical cord with your fingers or an elastic band approximately 5 cm from the distal clamp and make a clean cut between your compression and the distal clamp (see Note 3). 3. Disinfect the end of the cord and put the end into a container with sodium citrate anticoagulant.

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Table 3 Primers used for validation of microarray results using real-time PCR Gene name

Primer sequence (forward, reverse)

Refseq

ANGPTL4

F GGACCACAAGCACCTAGACC R GCGCCTCTGAATTACTGTCC

NM_001039667

ANPEP

F TCCCAGAAAACCTGATGGAC R TCATTGACCAGTGTGGCATT

NM_001150

CCL2

F CCCAATGAGTAGGCTGGAGA R AAAATGGATCCACACCTTGC

NM_002982

CXCL1

F AGGGAATTCACCCCAAGAAC R CACCAGTGAGCTTCCTCCTC

NM_001511

GAPDH

F GGTGGTGCTAAGCGTGTTAT R ACCTCTGTCATCTCTCCACA

NM_001256799

MLLT11

F GGACCCTGTGAGTAGCCAGTA R CAGCTCCGACAGATCCAGT

NM_006818

NEAT

F CCAGTTTTCCGAGAACCAAA R ATGCTGATCTGCTGCGTATG

NR_131012

SEC61A1

F TCAACGGAGCCCAAAAGTTATT R ACATCCCGGTCATCACATACA

NM_013336

SELE

F CAGGTGTCCACTCCCAGGTCCAAG R GGCAACTAGAAGGCACAGTCGAGG

NM_000450

TNFSF15

F GACCAAGTCTGTATGCGAAGTAG R CCATTAGCTTGTCCCCTTCT

NM_001204344

4. The assisting person elevates the placenta and squeezes it gently to milk out all CB. CB flow is regulated by gentle reduction of the compression (see Note 4). 5. Whenever the CB samples are not processed immediately, the samples are stored temporarily at 4  C for a maximum of 72 h. Transport to the laboratory right before isolation is performed at room temperature (RT). 6. Pipette a maximum of 15 mL of CB into sterile 50 mL tubes at RT and dilute with an equal volume of Dulbecco’s phosphatebuffered saline (PBS). 7. 15 mL Ficoll-Paque is pipetted into a separate 50 mL tube. 8. Layer aliquots of 10–15 mL of the diluted CB slowly over the Ficoll-Paque solution by gently pipetting the diluted blood down the side of the tube containing the Ficoll-Paque to avoid mixture with Ficoll-Paque. 9. Density centrifugation: for 30 min at 500  g at RT without brakes or with the lowest available brake settings.

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Lowest density Plasma layer Density gradient

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Mononuclear layer Ficoll layer

Erythrocyte layer Highest density

Fig. 1 Isolation of PBMCs by density gradient centrifugation of umbilical cord blood showing the layer of mononuclear cells to be used to derive PBMCs and ECFCs for an autologous co-culture

10. Coat 24-well culture plates as described in Subheading 2.1 step 9. 11. Using a sterile Pasteur pipette, the hazy layer of low-density PBMCs located at the interface between the Ficoll-Paque (bottom layer) and diluted plasma (upper layer, see Fig. 1) is removed and carefully dispensed into a sterile 15 mL tube containing 10 mL of PBS at RT. 12. Wash cell suspension for 10 min at 330  g at RT with a high deceleration brake setting. 13. After centrifugation, carefully aspirate and discard the supernatant. 14. Wash pelleted cells two more times with PBS for 10 min at 330  g at RT. 15. Suspend the PBMCs in 1 mL of complete endothelial growth medium (cEGM) prewarmed at 37  C for counting and subsequent freezing or culturing. 16. Count the cells by diluting the cell suspension with trypan blue solution 1:10 in one well of a 96 culture well plate.

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17. Place 10μL of the cell suspension-trypan blue mixture on a Neubauer hemocytometer chamber and cover with a cover slide. 18. Count the cells using an inverted microscope. 19. Determine cell concentration per mL (C) with the following formula: 20. C ¼ Number of cells  dilution factor  10,000 (correction factor)/Number of squares. 21. For cryopreservation resuspend 10 million PBMCs in DMSO/ FCS solution with a concentration of 1  106 cells per mL. 22. Freeze at 1 mL per tube in Mr. Frosty™ cryo 1  C freezing containers at
80  C. 23. Store cells at
80  C until being thawed at the time for co-culture with autologous ECFCs (see Subheading 3.6). 24. Separate 500,000 PBMCs for fluorescence-activated cell scanning (FACS) (see Subheading 3.4). 25. Resuspend the rest of the PBMCs not frozen in cEGM with a concentration of 3  106 PBMCs per mL and transfer 2 mL per well into a gelatine-coated 24-well plate at a seeding density of approximately 3  106 PBMCs per cm2. 3.2 Cultivation of PBMCs into ECFCs

1. Cultivate PBMCs under normal culture conditions of 37  C, 5% CO2, and 95% humidity. 2. Change medium 24 h after seeding to allow attachment and discard debris. 3. Afterwards change medium every 3 days. 4. Control wells routinely for appearance of ECFCs. Typically, ECFCs appear between day 5 and day 21 of culture exhibiting the typical cobblestone morphology of endothelial cells (see Fig. 2a). 5. Passage cells once they achieved 80–90% confluence: In order to passage the cultured ECFCs aspirate medium and wash the cells with 2 mL PBS. 6. Add 500μL of trypsin-EDTA solution to each well of 24-well plates and incubate for 3–5 min at 37  C, 5% CO2 until cells began to round up and detach as seen using the 10 phase contrast objective of an inverted microscope. 7. Tap the culture plate with both hands in order to detach all cells. 8. Add 1 mL of DMEM/FCS to each well to neutralize the trypsin. 9. Transfer the resulting suspension to a 50 mL tube. 10. Centrifuge suspension at 330  g for 7 min at RT.

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Fig. 2 Characterization of ECFC after third passage according to Ingram et al. [18]. (a) Example for a confluent ECFC-colony with typical cobblestone morphology. (b) FACS analysis using primary anti-human antibodies conjugated to fluorescein isothiocyanate (FITC; anti-CD45, anti-CD133/2, IgG1 isotype control) or conjugated to phycoerythrin (PE; anti-CD14, anti-CD31, anti-CD34, anti-CD105, antiCD-146, IgG2a isotype control) from Becton-Dickinson (Heidelberg, Germany). ECFCs showed 99–100% positivity for CD31, CD105, and CD146 and less than 10% positivity for CD34. ECFCs were negative for CD14, CD45, and CD133. (c) Uptake of DioAc-LDL by ECFC. (d) Tube formation by ECFC in Matrigel. (e) Binding of Ulex europaeus lectin by ECFC. Scale bars represent 100μm

11. Aspirate supernatant and resuspend the cell pellet in fresh cEGM prewarmed at 37  C. 12. Expand the cell culture by cultivating the harvested cell suspensions in 6-well plates and later 25 cm2 culture flasks. 13. Use ECFCs at passage 3–6 for further experiments (see Note 5). 3.3 Phenotypic Characterization of PBMCs and ECFCs

PBMCs after isolation are subjected to fluorescence-activated cell scanning in order to examine surface expression of hematopoietic markers such as clusters of differentiation (CD)14, CD45, CD133/2, and CD34 alongside lymphocyte markers including CD3, CD4, and CD8. Endothelial markers including CD31, CD34, CD146, and CD105 and hematopoietic markers such as CD14, CD45, and CD133 will be examined in ECFCs between passages 3 and 6.

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1. Suspend 500,000 isolated PBMCs or ECFCs in an FACS tube containing 50μL of PBS. 2. Add 20μL of fluorochrome-labeled human monoclonal antibodies to the respective samples, use primary anti-human antibodies conjugated to fluorescein isothiocyanate (FITC; antiCD3, anti-CD4, anti-CD45, anti-CD133, and the corresponding IgG1 isotype control) or conjugated to phycoerythrin (PE; anti-CD8, anti-CD14, anti-CD31, anti-CD34, anti-CD105, anti-CD-146, and the corresponding IgG2a isotype control). 3. Incubate at 4  C in a refrigerator for 20 min protected from light. 4. After incubation, resuspend each cell sample in 1 mL PBS. 5. Centrifuge cell samples at 330  g for 10 min at RT. 6. Remove the supernatant and resuspend the cell pellet in an FACS tube containing 1 mL PBS. 7. Analyze the samples in an FACScan flow cytometer using CellQuest Pro software to determine the percentage of cells that stain positively or negatively for each antigen. 8. For determining positivity, establish gates using isotype controls. An example for the results after flow cytometric analysis is given in Fig. 2b: PBMCs showed heterogeneous positive signals for CD3, CD4, CD8, CD14, less than 10% positivity for CD34 and >90% positivity for CD45. ECFCs showed 99–100% positivity for CD31, CD105, and CD146 and less than 10% positivity for CD34. ECFCs were negative for CD14, CD45, and CD133 (see Fig. 2b). 3.4 Analyzing the Expression of Molecular Markers for Peripheral Blood Mononuclear Cells and ECFCs

1. Extract total RNA from PBMCs after thawing from
80  C (see Subheading 3.6 steps 2–6) and ECFCs between passages 3 and 6 using the High Pure RNA Isolation Kit following the manufacturer’s recommendations. 2. Measure RNA concentration using a spectrophotometer and analyze purity by the 260/280 absorbance ratio. 3. Use RNA either directly or stored at
80  C. 4. Synthesize complementary DNA (cDNA) for subsequent PCR analyses from RNA using the protocol in Subheading 2.4 Table 1. 5. Incubate the mixture for 1 h at 40  C and stop reaction at 90  C for 5 min. 6. Perform all PCR analyses with a standard TaqDNA polymerase system with a final volume of 25μL.

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7. Test PBMCs for their expression of CD14 (hematopoietic marker), CD34 (endothelial cell marker), CD11b (a leucocyte marker), and β-actin (housekeeping gene) (see Subheading 2.4 Table 2). 8. Test ECFCs for the expression of CD31, CD146, CD45, CD133, CD34, CD14, and contractin (EMS1) (see Subheading 2.4 Table 2). 9. After initial denaturation use following conditions for amplification: 40 cycles: denaturation 94  C, 30 s; annealing 60  C, 30 s; elongation 72  C, 45 s. 10. To verify the length of PCR products, prepare an agarose/ ethidium bromide gel in TAE buffer (see Subheading 2.4). 11. Pour the solution in a gel electrophoresis cassette, containing combs to create 15 slots in the gel. 12. Allowed gel to solidify for 30 min. 13. Cover the cassette with TAE buffer and pipette PCR products into the slots of the gel. 14. Use one slot for PCR marker. 15. Gel is subjected to 110 V electrophoresis current for 45 min. 16. Photograph gel under ultraviolet light using imaging unit. 3.5 Functional Characterization of ECFC

Functional verification of ECFCs is performed according to Ingram et al. [18] with some modifications as follows:

3.5.1 In Vitro Tube Formation Assay on Matrigel and the Influence of Varying Oxygen Concentrations

1. To assess the tube forming ability of ECFCs, thaw Matrigel™ overnight at 4  C in a refrigerator. 2. Pipette in triplicates at 50μL/well in wells of a 96-well plate. 3. Incubate culture dishes for 40 min at 37  C, in a 5% CO2 atmosphere cell culture incubator to allow for polymerization and formation of a gel-like surface. 4. Cultivate harvested ECFCs on the Matrigel™ in cEGM, at a seeding density of 15,000 cells per well in an incubator at normoxia (21% oxygen) 37  C, 5% CO2, and 95% humidity. 5. Use CRL-7449 fibroblast cell line as negative control in the state of normoxia. 6. In order to assess the influence of oxygen concentration on the complexity of the vascular tubes formed on matrigel, repeat the assay in culture conditions of hyperoxia (40% oxygen) and anoxia (0% oxygen) at 37  C, 5% CO2, and 95% humidity. 7. Use an incubator capable to provide FiO2 > 0.21 to simulate hyperoxic condition.

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8. Provide the anoxic environment by placing the cell culture container in anaerobic bags moistened with distilled water and seal. 9. Following 18–20 h of incubation under hyperoxia, normoxia, and hypoxia, analyze cells under an inverted microscope (see Fig. 2d). 3.5.2 Acetylated Low-Density Lipoprotein (LDL) Uptake and Ulex europaeus Lectin Binding

1. Endothelial properties of ECFCs like lipid-uptake and binding of lectin is assessed as follows: 2. Incubate 200,000 ECFCs between passages 3 and 6 with DioAc-LDL in 12-well plates in cEGM for 4 h, under normal culture conditions of 37  C, 5% CO2, and 95% humidity. 3. After incubation, wash the cells three times with 2 mL of PBS and centrifuge cell suspension at RT at 330  g for 7 min. 4. Image cells in 50μL PBS, using the fluorescence setting of an inverted microscope for the uptake of Dio-Ac-LDL (see Fig. 2c). 5. Incubate 200,000 ECFCs with fluorescein-tagged Ulex europaeus lectin for 1 h at 37  C. 6. Image cells under fluorescence with an inverted microscope for the binding of Ulex europaeus lectin (see Fig. 2e).

3.5.3 Single Cell Clonogenic Assay

The single cell clonogenic assay was performed to assess the ability of ECFCs to form colonies. 1. Detach cells (see Subheading 3.2 steps 6–11) and sort ECFCs at passage 3 with FACS by CD34. 2. Seed as single cells in wells of a gelatine-coated 96-well plate in 150μL cEGM. 3. Change cEGM every 3 days. 4. At day 14, inspect wells under an inverted microscope. 5. Aspirate the culture media in the 96-well plates and add 100μL of Hoechst stain to the wells in order to visualize and categorize ECFC colonies. 6. Colonies with 2–50 cells will be classified as low proliferative and colonies with more than 100 cells will be classified as high proliferative.

3.5.4 Wound Healing (or Scratch) Assay

The in vitro scratch assay was performed to assess the ability of ECFCs to migrate in vitro. 1. To create a scratch scrape a straight line on the monolayer of cultured ECFCs at third passage, at 80% confluency in a 25 cm2 culture flask using a Pasteur pipette tip. 2. Remove debris by washing the cells once with 4 mL cEGM.

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3. Aspirate and replace 4 mL cEGM. 4. Place culture dish in a humidified incubator under normal culture conditions for 18 h and position a live camera in the incubator to take pictures of the cell migration at intervals of 5 min. 3.6 Co-Culture of Endothelial Colony Forming Cells with Peripheral Blood Mononuclear Cells

1. Seed ECFCs at third passage on gelatine-coated 6-well plates at a density of 50,000 cells/cm2 in 4 mL of cEGM for 24 h under normal culture conditions of 37  C, 5% CO2, and 95% humidity to allow cell attachment. 2. 24 h later, thaw autologous PBMCs from
80  C by swirling the sample in the cryotubes for 45 s in a warm water bath at 37  C. 3. Immediately, transfer the solution of PBMCs with a pipette into a 50 mL tube containing 10 mL of prewarmed 37  C cEGM. 4. Wash PBMCs by centrifugation at 330  g for 7 min at RT. 5. Resuspend washed PBMCs in 1 mL cEGM. 6. Determine the number of viable cells by trypan blue exclusion (see Subheading 3.1 steps 17–20). 7. To reach a ratio 1:4 of ECFC to PBMC in co-culture, seed approximately 2 million thawed PBMCs directly into the 6-well plates with ECFCs to allow direct cell–cell contact. 8. Prepare three parallel co-culture samples (ECFCs plus PBMCs) and three parallel control ECFC samples (without PBMCs) per CB sample. 9. Cultivate each co-cultured sample and its respective control for 16 h at 37  C, 5% CO2 either in hyperoxia (40% oxygen), normoxia (21% oxygen), or an anaerobic milieu (see Note 2).

3.7 Microarray Analysis

1. After 16 h collect the cells of co-cultures and controls by trypsinization (see Subheading 3.2 steps 6–11). 2. Extract RNA of each sample by using an RNA isolation kit of high quality according to the manufacturer’s recommendations. 3. Analyze RNA from all samples by using Human Exon 1.0 ST microarrays. 4. Hybridize, stain, and scan arrays according to manufacturer’s protocols. 5. Use Expression Console build 1.4.1.46 for primary analysis of cel files and probeset summarization. 6. Analyze cel files using the RMA algorithm at the extended gene level.

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7. Identify probesets with increased or decreased signal intensities by using Microsoft Excel 2010. 8. Further, analyze probesets above the 95th percentile of all probesets. 9. Calculate the ratios of the signal intensity for the following comparisons: (1) PBMC/ECFC co-cultures under hyperoxia (HIGH) versus PBMC/ECFC co-culture under normoxic conditions (NORM); (2) HIGH versus PBMC/ECFC co-cultures under hypoxia (LOW); (3) NORM versus LOW. 10. Consider probesets to be up-regulated under hyperoxia if the signal intensities from the hyperoxia sample are above the 95th percentile of all signal intensities, if the signal intensities from the hyperoxia sample are higher than the signal intensities from the normoxia sample, and if the signal intensities from the hyperoxia sample are at least 1.5 times higher in the hyperoxia sample than in the hypoxia sample. Vice versa, genes are considered to be up-regulated under hypoxia if the signal intensities from the hypoxia sample are above the 95th percentile of all signal intensities, if the signal intensities from the hypoxia sample are higher than the signal intensities from the normoxia sample, and if the signal intensities from the hypoxia sample are at least 1.5 times higher in the hypoxia sample than in the hyperoxia sample. 11. Identify up-regulated probesets in hyperoxia and up-regulated probesets in hypoxia assigned to known gen loci. 3.8 Real-Time PCR Analysis

1. For the second validation step based on PCR technique (see Subheading 3.4) randomly choose probesets from these two lists, which were within gen loci with known biological function related to inflammation or angiogenesis. 2. For example, use the following nine genes for further replication angiopoetin-like 4 (ANGPTL4), alanyl aminopeptidase (ANPEP), C-X-C motif chemokine ligand 1 (CXCL1), transcription factor 7 cofactor (MLLT11), matrix metallopeptidase 14 (MMP14), nuclear paraspeckle assembly transcript 1 (NEAT1), Sec61 translocon alpha 1 subunit (SEC61A1), selectin E (SELE), and tumor necrosis factor superfamily member 15 (TNFSF15) (see Note 6). 3. Perform replication with quantitative real-time PCR in co-cultures of at least two independent patient samples. 4. Normalize specific gene amplification to glyceraldehyde-3phosphate dehydrogenase (GAPDH). 5. Quantify by using the 2
ΔΔct method [19].

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Fig. 3 Validation of genes with oxygen-dependent differential expression in microarray experiments using quantitative real-time PCR in independent samples. Relative expression values were calculated based on the 2
ΔΔCt method using GAPDH as a housekeeping gene. High: Co-culture experiment with 40% oxygen, Norm: Co-culture experiment with 21% oxygen, Low: Co-culture experiment with almost 0% oxygen

In our study, the microarray results for CXCL1, MMP14, SEC61A1, and TNFSF15 could be reproduced, showing a pattern of up-regulation in hyperoxia, whereas ANGPTL4 showed up-regulation in hypoxia (see Fig. 3).

4

Notes 1. If the CB is not processed immediately after collection, use a container with sodium citrate anticoagulant plus dextrose-arginine-phosphate. For term babies, you can use a commercially available standard cord blood collecting system. 2. Depending on your scientific question, you may use a co-culture setting with cell culture inserts, especially when it is important to analyze the two cell lines separately after co-culture. 3. Do not use a clamp to compress the umbilical cord.

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4. Always keep a mild compression on the cord throughout the whole procedure and shake the container gently to avoid clotting. Collection should start as soon as possible after receiving the placenta in order to diminish loss due to coagulation processes. Collecting CB: If you do not have any assistance at hand put a supporting stand with a couple of open containers on the bottom of a sink. The faster you receive the placenta the higher are your chances to obtain enough CB. Starting the collection more than 10 min after birth is usually not successful. Do not waste your time and material with partially clotted samples. Collecting at least 15 mL CB of preterm babies is not easy, but the smallest preterm baby in our setting had a birth weight of 810 g. A sample with less than 15 mL CB will only rarely produce enough PBMCs for establishing an autologous co-culture but is useful for cultivation of ECFCs only especially at the beginning of a project. 5. The single cell clonogenic assay was performed using ECFCs at passage 3. 6. For the genes alanyl aminopeptidase (ANPEP), transcription factor 7 cofactor (MLLT11), nuclear paraspeckle assembly transcript 1 (NEAT1), and selectin E (SELE) the results from the microassay analysis could not be reproduced.

Acknowledgments This project was supported by Wilhelm Roux program at the Medical Faculty of Martin Luther University Halle-Wittenberg, Halle (Saale) (28/14). References 1. Gluckman E, Broxmeyer HA, Auerbach AD et al (1989) Hematopoietic reconstitution in a patient with Fanconi’s anemia by means of umbilical cord blood from an HLA-identical sibling. N Engl J Med 321:1174–1178 2. Koblas T, Harman SM, Saudek F (2005) The application of umbilical cord blood cells in the treatment of diabetes mellitus. Rev Diabet Stud 2:228–234 3. Jaing T-H (2014) Umbilical cord blood: a trustworthy source of multipotent stem cells for regenerative medicine. Cell Transplant 23:493–496 4. Almici C, Carlo-Stella C, Wagner JE et al (1995) Umbilical cord blood as a source of hematopoietic stem cells: from research to clinical application. Haematologica 80:473–479

5. Broxmeyer HE, Douglas GW, Hangoc G et al (1989) Human umbilical cord blood as a potential source of transplantable hematopoietic stem/ progenitor cells. PNAS 86(10):3828–3832 6. Bieback K, Kern S, Klu¨ter H et al (2004) Critical parameters for the isolation of mesenchymal stem cells from umbilical cord blood. Stem Cells 22:625–634 7. Murohara T, Ikeda H, Duan J et al (2000) Transplanted cord blood-derived endothelial precursor cells augment postnatal neovascularization. J Clin Invest 105:1527–1536 8. Zhao Y, Wang H, Mazzone T (2006) Identification of stem cells from human umbilical cord blood with embryonic and hematopoietic characteristics. Exp Cell Res 312:2454–2464

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9. Bertaina A, Bernardo ME, Caniglia M et al (2010) Cord blood transplantation in children with haematological malignancies. Best Pract Res Clin Haematol 23:189–196 10. Garbuzova-Davis S, Ehrhart J, Sanberg PR (2017) Cord blood as a potential therapeutic for amyotrophic lateral sclerosis. Expert Opin Biol Ther 17:837–851 11. Reddi AS, Kothari N, Kuppasani K et al (2015) Human umbilical cord blood cells and diabetes mellitus: recent advances. Curr Stem Cell Res Ther 10:266–270 12. Burri PH (2006) Structural aspects of postnatal lung development—alveolar formation and growth. Biol Neonate 89:313–322 13. Helenius K, Sjo¨rs G, Shah PS et al (2017) Survival in very preterm infants: an international comparison of 10 national neonatal networks. Pediatrics 140:e20171264 14. Gien J, Kinsella JP (2011) Pathogenesis and treatment of bronchopulmonary dysplasia. Curr Opin Pediatr 23:305–313

15. Wang J, Dong W (2018) Oxidative stress and bronchopulmonary dysplasia. Gene 678:177–183 16. Mokres LM, Parai K, Hilgendorff A et al (2010) Prolonged mechanical ventilation with air induces apoptosis and causes failure of alveolar septation and angiogenesis in lungs of newborn mice. Am J Physiol Lung Cell Mol Physiol 298:23–35 17. Chao CM, El Agha E, Tiozzo C et al (2015) A breath of fresh air on the mesenchyme: impact of impaired mesenchymal development on the pathogenesis of bronchopulmonary dysplasia. Front Med 2:27 18. Ingram DA, Mead LE, Tanaka H et al (2004) Identification of a novel hierarchy of endothelial progenitor cells using human peripheral and umbilical cord blood. Blood 104:2752–2760 19. Livak KJ, Schmittgen TD (2001) Analysis of relative gene expression data using real-time quantitative PCR and the 2(
Delta Delta C (T)) method. Methods 25:402–408

Chapter 9 Ex Vivo Model of Spontaneous Neuroretinal Degeneration for Evaluating Stem Cells’ Paracrine Properties Ivan Fernandez-Bueno and Ricardo Usategui-Martin Abstract Ex vivo neuroretina cultures closely resemble in vivo conditions, retaining the complex neuroretina cells dynamics, connections, and functionality, under controlled conditions. Therefore, these models have allowed advancing in the knowledge of retinal physiology and pathobiology over the years. Furthermore, the ex vivo neuroretina models represent an adequate tool for evaluating stem cell therapies over neuroretinal degeneration processes. Here, we describe a physically separated co-culture of neuroretina explants with stem cells to evaluate the effect of stem cells paracrine properties on spontaneous neuroretinal degeneration. Key words Neuroretina, Ex vivo neuroretina, Retinal degeneration, Neuroprotection, Advanced therapies, Cell therapy, Stem cells, Paracrine properties

1

Introduction Neuroretina organ cultures closely simulate in vivo retinal cellular and molecular dynamics and have been considered an adequate tool for improving the knowledge on retinal physiology and pathobiology [1–5]. There are some limitations of these ex vivo systems, such as the absence of choroidal and retinal blood flow, the lack of vitreous, and the axotomy of the ganglion cells, which may considerably limit the study of the inner retina modifications. However, several ex vivo neuroretina studies have shown that this organ model is comparable with in vivo conditions [6–9]. Furthermore, this model has some other advantages, it is inexpensive and easy to develop, and provide a better alternative to animal experimentation than do cellular cultures composed of only a single cell type that lose the cellular dynamics among the multiple cell types and intercellular matrices of the retina [10]. Neuroretina culture systems began to develop in the 1920s, using retinas of chicken embryos on plasma clots [2], in later years, this technique was applied with mammalian retinas [3]. In 1954,

Peggy Stock and Bruno Christ (eds.), In Vitro Models for Stem Cell Therapy: Methods and Protocols, Methods in Molecular Biology, vol. 2269, https://doi.org/10.1007/978-1-0716-1225-5_9, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Trowell developed the membrane culture method [4], in which the tissue was placed in a porous membrane on a metal grid, and maintained at an air-medium interface. This technique, placing the vitreous surface of the retina in contact with the supporting membrane, was used to study cell dynamics in retinal explants during the next three decades [11–14]. Finally, in 1989, Caffe´ et al. [1] published a method in which the neural retina was placed, with the photoreceptors layer facing downward, on rafts made by nitrocellulose filters and polyamide grids. Since then, ex vivo adult mammalian neuroretina cultures, from rat, mice, guinea pig, dog, pig, or cow, have been used to describe the differentiation processes of pre- and early-stage post-natal retinas [1, 15–17]; to provide valuable insights into retinal diseases processes [6, 18–31]; to test potential therapeutic substances [18, 24, 29, 32]; to examine the role of growth factors over retinal cells’ dynamics [29, 33–36]; and to assess nanostructured scaffolds [37] and the potential cytotoxicity of substances [38, 39]. Furthermore, the ex vivo neuroretina culture has been used for enhancing and, more recently, for evaluating stem cell therapies over neuroretinal degeneration [9, 40– 43]. Although mammal neuroretina is a good alternative over the dependency on human samples availability for research purposes, ex vivo adult-human neuroretina cultures have been also characterized [7] and have been used to improve the current knowledge about photoreceptors, ganglion cells, glial cells, and microglia dynamics [7, 8, 44–50]. Here, we describe ex vivo neuroretina culture over porous membrane inserts that physically separated them from stem cells cultured on the bottom of the culture inserts. Thus, the porous membrane insert prevented the stem cells from migrating and integrating into the retinal tissue, whilst allowing molecular exchange between the different cell types [9, 22]. Therefore, the factors potentially secreted by stem cells can diffuse through the pores of the membrane insert and influence neuroretina cells’ dynamics. This experimental approach resembles an intravitreal injection of stem cells, where the cells do not integrate into the neuroretina but are influenced by the factors secreted by the retinal cells [9]. It differs from other previous approaches where a droplet of stem cells suspension is deposited on the vitreous surface of the neuroretina explants [51, 52]. In these models, the stem cells come into direct contact with the retinal cells, being able to differentiate and integrate into the retinal tissue, and thus losing their paracrine properties.

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Materials

2.1 Ex Vivo Neuroretina and Stem Cells Co-Culture

1. Eyeballs. 2. Stem cells of choice. 3. Culture dishes: 90 mm. 4. Antibiotics and antimycotics mixture: 10,000 IU/mL Penicillin, 10 mg/mL Streptomycin, 25 μg/mL Amphotericin B. 5. Dulbecco’s Modified Eagle Medium (DMEM) with 10 antibiotics/antimycotics (DMEM-10AA): CO2-independent DMEM, without L-glutamine, 10% antibiotics, and antimycotics mixture. 6. DMEM: CO2-independent DMEM, without L-glutamine, 1% antibiotics, and antimycotics mixture. 7. Culture medium for stem cells of choice. 8. Complete Neurobasal A medium (NeuA): Neurobasal A medium, 2% B-27 supplement, 10% fetal bovine serum, 1% Lglutamine, 1% antibiotics, and antimycotics mixture. 9. 22G needle. 10. Sterile cotton swaps. 11. Paintbrushes. 12. Transwell® plate: 6-well plates, 24-mm diameter with 0.4-μm pore, polycarbonate membrane insert (Corning Inc., Corning, NY). 13. Straight-blade dissecting scrissors. 14. Straight Vannas scissors. 15. Westcott scissors. 16. 70% Ethanol. 17. Cell culture spatula. 18. Co-culture medium: 1:1 mixture of culture medium for stem cells of choice and Complete Neurobasal A medium. 19. Incubator for cell cultures. 20. Forceps.

2.2 Neuroretinal Explants Processing

1. 4% Paraformaldehyde saline (PBS)

2.2.1 Paraffin Wax Embedding

2. PBS.

(PFA)

in

phosphate-buffered

3. Cassettes for small specimen. 4. Molds. 5. Automated tissue processing machine for paraffin embedding. 6. 96%; 100% Ethanol.

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7. Xylene. 8. 1:1 mixture of 100% Ethanol/Xylene. 9. Paraffin wax. 10. Rotatory microtome. 11. Glass slides. 2.2.2 Cryopreservation

1. O.C.T. Tissue-Tek. 2. 4% PFA in PBS. 3. PBS. 4. 15%; 20%; 30% Sucrose in PBS. 5. Molds. 6. Cryostat. 7. Glass slides.

2.2.3 Resin Embedding

1. Epoxy resin: low-viscosity (Spurr, TAAB, Aldermaston, UK). 2. Fixing solution: 1% PFA, 1% glutaraldehyde in PBS. 3. PBS. 4. Postfix: 1% osmium tetroxide in PBS. 5. 50%; 75%; 90%; 96%; 100% Ethanol. 6. Propylene oxide. 7. 3:1 mixture of Propylene oxide/epoxy resin. 8. 1:1 mixture of Propylene oxide/epoxy resin. 9. 1:3 mixture of Propylene oxide/epoxy resin. 10. Molds. 11. Ultramicrotome. 12. Glass slides treated with (3-aminopropyl) triethoxy-silane. 13. Staining solution for ultrastructural study: 1% toluidine blue in 3% sodium tetraborate in PBS.

3

Methods

3.1 Ex Vivo Neuroretina and Stem Cells Co-Culture 3.1.1 Stem Cell Preparation

Carry out all procedures at room temperature and under aseptic conditions unless otherwise specified.

1. Determine stem cell viability and cell counts. 2. Seed 30,000 stem cells, with viability higher than 95%, in 1.5 mL of culture medium (see Note 1), on the bottom of Transwell® culture plates, and culture for 72 h.

Ex Vivo Neuroretina Culture for Evaluating Stem Cells Properties 3.1.2 Neuroretina Explants Preparation (See Note 2)

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1. Fully immerse each eyeball in 70% ethanol for 2 min (see Fig. 1a and see Note 3). Then, wash the eyeball three times in DMEM10AA, for 2 min every wash (see Fig. 1a). 2. Place the eyeball on a culture dish with DMEM and remove the remaining extraocular tissues using dissecting scissors. 3. Puncture the sclera with a sterile 22-gauge needle at the ora serrata until performing a macroscopically visible hole (see Fig. 1b). 4. Insert Westcott scissors through the scleral hole, previously performed, and dissect dividing the ocular globe into anterior and posterior eyecups (see Fig. 1c, d). 5. Discard the anterior eyecup with the iris and the lens. 6. Transfer the posterior eyecup to a new plate with NeuA medium. 7. Remove carefully the vitreous from the posterior eyecup with sterile cotton swabs and avoid touching the retinal tissue (see Fig. 1e). 8. Mechanically detach the neural retina as a whole from the retinal pigment epithelium (RPE) by gently brushing (see Fig. 1f) and cutting the optic nerve with Westcott scissors (see Note 4 and see Fig. 1g). 9. Deposit the detached neural retina into the NeuA medium by gently brushing (see Fig. 1h). 10. Unroll the neuroretina by brushing (see Fig. 1i) and cut into 5  5 mm explants with Vannas scissors (see Fig. 1j, k), in such a way to avoid the most peripheral retina and the presence of irregular edges (see Notes 5 and 6). 11. Transfer the neuroretinal explant on a cell culture spatula (Fig. 1l) and deposit it on the membrane insert of the Transwell® culture plates, with the aid of a paintbrush (see Note 7 and see Fig. 1m). 12. Unroll the neuroretinal explant and locate it in the middle of the membrane insert by gently brushing (see Note 8 and see Fig. 1n).

3.1.3 Neuroretinal Explants and Stem Cells Co-Culture

1. Add the appropriate co-culture medium to the Transwell® culture plates which contain the stem cells previously seeded (see Note 9 and see Fig. 1o). 2. Co-culture the neuroretinal explants on the insert membranes and the stem cells in the same Transwell® culture plate, but physically separated by the insert membrane (see Notes 10 and 11) (see Fig. 2).

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Fig. 1 Neuroretina explants preparation from eye globes. (a) Eyeball immersion in ethanol and washing in clean DMEM. (b) Scleral puncture at the ora serrata and (c, d) dissection of the ocular globe into anterior and posterior eyecups. (e) Vitreous removal from the posterior eyecup. (f) Neuroretina detachment from the retinal pigment epithelium by gently brushing and (g) cutting the optic nerve. (h) Detached-neuroretina deposition into culture media by brushing. (i) Neuroretina unrolls and (j, k) cut into explants. (l) Neuroretina explant transfer and (m) deposition on the membrane insert. (n) Neuroretina explant unrolls and location in the middle of the membrane insert by brushing. (o) Add appropriate culture medium to the culture plates which contain the insert membrane with the neuroretina explant

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Stem cells

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Neuroretinal explant

Paracrine factors Porous membrane

Culture medium

Fig. 2 Ex vivo neuroretina and stem cells co-culture. The neuroretina explants are cultured over porous membrane inserts that physically separate them from the stem cells cultured on the bottom of the culture inserts. Thus, the porous membrane insert prevents the stem cells from migrating and integrating into the retinal tissue, whilst allowing molecular exchange between the different cell types. Therefore, the factors potentially secreted by stem cells can diffuse through the pores of the membrane insert and influence neuroretina cells’ dynamics

3. Maintain the culture medium level in contact with the support membrane beneath the explant during the culture period (see Note 12). 4. Change culture medium every day during the culture period, with freshly prepared warmed medium (see Note 13). 3.2 Neuroretinal Explants Processing

3.2.1 Paraffin Wax Embedding (for Light or Epifluorescence Microscopy)

Cut the culture membrane insert surrounding the neuroretina explant, and cut the neuroretina specimen into pieces for further processing. Handle the neuroretina specimens with extreme care during processing and avoid tightening or breaking of the neuroretina tissue. 1. Fix the samples in 4% PFA in PBS for 2 h at 4  C. 2. Remove PFA and wash two times in PBS. 3. Embed the samples in paraffin using an automated tissue processing machine, by using a specific program for small specimens. In brief, the dehydration, clearing, and wax infiltration steps are programmed as follows: (a) 96% ethanol for 6 min at 36  C (b) 96% ethanol for 12 min at 36  C (c) 100% ethanol for 6 min at 36  C (d) 100% ethanol/xylene (1:1) for 6 min at 36  C (repeat two times)

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(e) Xylene for 6 min at 36  C (repeat two times). (f) Paraffin wax for 9 min at 60  C (repeat four times). 4. Fill a mold with molten wax and place the specimen (infiltrated with wax) into it, carefully orientate the specimen to determine the “plane of the section.” 5. Place a cassette on top of the mold, topped up with more wax, and place all the components on a cold plate to solidify. 6. Remove the paraffin block containing the tissue specimen from the mold. 7. Cut sections with a rotatory microtome and mounted it on glass slides. 8. Sections are ready to apply staining or immunochemistry protocols for light or epifluorescence microscopy. 3.2.2 Cryopreservation (for Epifluorescence or Confocal Microscopy)

1. Fix specimens in 4% PFA in PBS for 2 h at 4  C. 2. Remove PFA and wash two times in PBS. 3. Cryoprotect the specimens in increasing concentrations of sucrose, in brief: (a) 15% sucrose for 2 h at 4  C (b) 20% sucrose for 2 h at 4  C (c) 30% sucrose for 12 h at 4  C. 4. Fill with Tissue-Tek a previously cooled to 4  C mold and place the specimen into it, carefully orientate the specimen to determine the “plane of the section.” 5. Topped up with more Tissue-Tek and cooled all the components from 4  C to 80  C. 6. Remove the Tissue-Tek block containing the tissue specimen from the mold (see Note 14). 7. Cut sections with a cryostat and mount it on glass slides. 8. Sections are ready to apply immunochemistry protocols for epifluorescence or confocal microscopy.

3.2.3 Epoxy Resin Embedding (for Transmission Electron Microscopy)

1. Fix specimens overnight in 1% PFA and 1% glutaraldehyde in PBS. 2. Wash the samples in PB and postfix in 1% osmium tetroxide in PBS. 3. Dehydrate gradually in ethanol series: (a) 50% ethanol for 10 min (b) 75% ethanol for 10 min (c) 90% ethanol for 10 min (d) 96% ethanol for 10 min (e) 100% ethanol for 10 min (repeat two times).

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4. Infiltrate the specimens in graded concentrations of propylene oxide/epoxy resin, in brief: (a) Propylene oxide for 10 min. (b) Propylene oxide/epoxy resin (3:1) for 1 h. (c) Propylene oxide/epoxy resin (1:1) for 1 h. (d) Propylene oxide/epoxy resin (1:3) for 1 h. (e) Epoxy resin for 30 min at 60  C. 5. Fill a mold with epoxy resin and place the specimen into it, carefully orientate the specimen to determine the “plane of the section.” 6. Topped up with more epoxy resin and place all the components on a laboratory stove for 24 h at 60  C for epoxy resin polymerization. 7. Remove the epoxy resin block containing the tissue specimen from the mold. 8. Cut semithin sections with an ultramicrotome, mount it on glass slides treated with (3-aminopropyl) triethoxy-silane. 9. Stain it with 1% toluidine blue in 3% sodium tetraborate in PBS (see Note 15) to select specific areas for ultrastructural study. 10. Cut ultrathin sections with an ultramicrotome and mount it on glass slides treated with (3-aminopropyl) triethoxy-silane. 11. Specimens are ready for staining protocols for transmission electron microscopy.

4

Notes 1. Use the appropriate culture medium composition based on the type of stem cells to be used. 2. To minimize spontaneous retinal degeneration after death, the neuroretinal explants should be obtained as soon as possible. As a suggestion, the eyes from animals must be processed within 2 h after sacrifice and those from humans within 4 h after death. 3. If the eyes were obtained from a slaughterhouse or a nonpathogen-free source a first 2-min wash in povidone-iodine is highly recommended. 4. The procedure described here is commonly used for porcine eyes. However, detaching the neuroretina can be more or less difficult depending on the species. In human eyes, the retina (with the RPE) is easily detached from the choroid, and then the neuroretina must be separated from the RPE by gently brushing. In some rodents, strong adhesions can occur between the RPE and the neuroretina that makes it necessary using microsurgical scissors to separate them.

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5. To macroscopically determine how the neural retina is oriented in the culture dish, the neuroretinal tissue should be completely unrolled. If the neuroretinal edges appear folded up and the retinal vessels are visible, the photoreceptors were at the bottom and the inner limiting membrane was at the top, and vice versa. 6. If only the neuroretinal regions mostly constituted by cones will be used, it is necessary to identify the cone-enriched visual streak of each species before preparing the corresponding explants. 7. If changes in the outer layers of the neuroretina will be evaluated, it is better to place the explants with the photoreceptors facing the insert membrane. Therefore, the outer layers would be directly influenced by the factors present in the culture medium. On the other hand, if changes in the inner layers will be evaluated, it is better to place the explants with the inner limiting membrane facing the insert membrane. 8. Avoid damaging the neural retina tissue and the membrane insert. 9. Co-culture in complete Neurobasal A and the appropriate culture medium for the type of stem cells used (1:1). 10. Co-cultures are maintained at 37  C in an atmosphere of 5% CO2 with 95% humidity. 11. The porous cell culture membranes prevented the stem cells from migrating and integrating into the neuroretinal tissue, whilst allowing molecular exchange between the different cell types. Thus, factors secreted by stem cells can diffuse through the pores of the cell culture membrane and influence neuroretinal cell dynamics, and vice versa [9, 22]. 12. Normally 1.5 mL of the culture medium is enough to contact the insert membrane without exceeding it. 13. For the best cellular maintenance in these co-culture models, it is essential to change the culture medium every day. Since the volume of medium that can be provided (1.5 mL) is considerably small for the nutritional needs of the cells and quickly runs out nutrients. 14. Samples can be stored in an ultra-freezer or liquid nitrogen for long-term cryopreservation until use. 15. Staining commonly uses to analyze and select the areas for ultrastructural study at this step is 1% toluidine blue in 3% sodium tetraborate in PBS.

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effects on photoreceptors in the rd1 retina through the glial glutamate transporter GLAST? Mol Vis 11:677–687 34. Franke AG, Gubbe C, Beier M, Duenker N (2006) Transforming growth factor-beta and bone morphogenetic proteins: cooperative players in chick and murine programmed retinal cell death. J Comp Neurol 495:263–278. https://doi.org/10.1002/cne.20869 35. Lagre`ze WA, Pielen A, Steingart R et al (2005) The peptides ADNF-9 and NAP increase survival and neurite outgrowth of rat retinal ganglion cells in vitro. Investig Ophthalmol Vis Sci 46:933–938. https://doi.org/10.1167/iovs. 04-0766 36. Garcı´a M, Forster V, Hicks D, Vecino E (2002) Effects of mu¨ller glia on cell survival and neuritogenesis in adult porcine retina in vitro. Invest Ophthalmol Vis Sci 43:3735–3743 37. Mayazur Rahman S, Reichenbach A, Zink M, Mayr SG (2016) Mechanical spectroscopy of retina explants at the protein level employing nanostructured scaffolds. Soft Matter 12:3431–3441. https://doi.org/10.1039/ c6sm00293e 38. Saikia P, Maisch T, Kobuch K et al (2006) Safety testing of indocyanine green in an ex vivo porcine retina model. Invest Ophthalmol Vis Sci 47:4998–5003. https://doi.org/ 10.1167/iovs.05-1665 39. Pastor JC, Coco RM, Fernandez-Bueno I et al (2017) Acute retinal damage after using a toxic perfluoro-octane for vitreo-retinal surgery. Retina 37:1140. https://doi.org/10.1097/ IAE.0000000000001680 40. Johnson TV, Martin KR (2008) Development and characterization of an adult retinal explant organotypic tissue culture system as an in vitro intraocular stem cell transplantation model. Invest Ophthalmol Vis Sci 49:3503–3512. https://doi.org/10.1167/iovs.07-1601 41. Rodriguez-Crespo D, Di Lauro S, Singh AKAK et al (2014) Triple-layered mixed co-culture model of RPE cells with neuroretina for evaluating the neuroprotective effects of adipose-MSCs. Cell Tissue Res 358:705–716. https://doi.org/10.1007/s00441-014-19875 42. Mollick T, Mohlin C, Johansson K (2016) Human neural progenitor cells decrease photoreceptor degeneration, normalize opsin distribution and support synapse structure in cultured porcine retina. Brain Res 1646:522–534. https://doi.org/10.1016/J. BRAINRES.2016.06.039 43. Jones MK, Lu B, Chen DZ et al (2019) In vitro and in vivo proteomic comparison of human

Ex Vivo Neuroretina Culture for Evaluating Stem Cells Properties neural progenitor cell-induced photoreceptor survival. Proteomics 19:1800213. https:// doi.org/10.1002/pmic.201800213 44. Niyadurupola N, Sidaway P, Osborne A et al (2011) The development of human organotypic retinal cultures (HORCs) to study retinal neurodegeneration. Br J Ophthalmol 95:720–726. https://doi.org/10.1136/bjo. 2010.181404 45. Carter DA, Dick AD (2003) Lipopolysaccharide/interferon-gamma and not transforming growth factor beta inhibits retinal microglial migration from retinal explant. Br J Ophthalmol 87:481–487. https://doi.org/10.1136/bjo.87.4.481 46. Carter DA, Dick AD (2004) CD200 maintains microglial potential to migrate in adult human retinal explant model. Curr Eye Res 28:427–436. https://doi.org/10.1080/ 02713680490503778 47. Balasubramaniam B, Carter DA, Mayer EJ, Dick AD (2009) Microglia derived IL-6 suppresses neurosphere generation from adult human retinal cell suspensions. Exp Eye Res 89:757–766. https://doi.org/10.1016/j. exer.2009.06.019

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48. Busskamp V, Duebel J, Balya D et al (2010) Genetic reactivation of cone photoreceptors restores visual responses in retinitis pigmentosa. Science 329:413–417 49. Carr AJ, Vugler A, Lawrence J et al (2009) Molecular characterization and functional analysis of phagocytosis by human embryonic stem cell-derived RPE cells using a novel human retinal assay. Mol Vis 15:283–295 50. Murali A, Ramlogan-Steel CA, Andrzejewski S et al (2019) Retinal explant culture: a platform to investigate human neuro-retina. Clin Exp Ophthalmol 47:274–285. https://doi.org/ 10.1111/ceo.13434 51. Johnson TV, DeKorver NW, Levasseur VA et al (2014) Identification of retinal ganglion cell neuroprotection conferred by platelet-derived growth factor through analysis of the mesenchymal stem cell secretome. Brain 137:503–519. https://doi.org/10.1093/ brain/awt292 52. Osborne A, Sanderson J, Martin KR (2018) Neuroprotective effects of human mesenchymal stem cells and platelet-derived growth factor on human retinal ganglion cells. Stem Cells 36:65–78. https://doi.org/10.1002/stem. 2722

Chapter 10 Ex Vivo Normothermic Hypoxic Rat Liver Perfusion Model: An Experimental Setting for Organ Recondition and Pharmacological Intervention Federica Rigo, Victor Navarro-Tableros, Nicola De Stefano, Alberto Calleri, and Renato Romagnoli Abstract The gold standard for organ preservation before transplantation is static cold storage, which is unable to fully protect suboptimal livers from ischemia/reperfusion injury. An emerging alternative is normothermic machine perfusion (NMP), which permits organ reconditioning. The ex vivo NMP hypoxic Rat Liver Perfusion Model represents a feasible approach that allow pharmacological intervention on isolated rat livers by using a combination of NMP and infusion of a number of drugs and/or biological material (cells, microvesicles, etc.). The combination of these two techniques may not only be applied for tissue preservation purposes, but also to investigate the biological effects of molecules and treatment useful in tissue protection. The protocol describes an ex vivo murine model of NMP capable of maintaining liver function despite an ongoing hypoxic injury induced by hemodilution. Furthermore, with this NMP system it is possible to deliver cells treatment or pharmacological intervention to an ex vivo perfused liver and suggests that could represent an innovative approach to recondition organs. Key words Liver transplant, Normothermic machine perfusion, Ischemia reperfusion injury, Hypoxic damage, Pharmacological treatment, Organ reconditioning

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Introduction Currently, the gold standard for organ preservation is Static Cold Storage (SCS), however, along with donor pool expansion, it has emerged that SCS is imperfect in preserving suboptimal organs from the so-called extended criteria donors [1, 2]. Normothermic machine perfusion (NMP) is an innovative and valid alternative to SCS. This type of perfusion keeps the organ at physiological temperature while continuously providing oxygen and nutrients [3, 4], it permits real-time monitoring metabolic status and allow graft viability. Moreover, NMP offers the advantages to provide pharmacological interventions or other treatment during preservation [5]. Animal studies demonstrated the superiority of NMP

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compared with SCS [6–8], and a phase I study established safety and feasibility of this technique in humans [9]. In this chapter, we briefly describe our Ex Vivo Normothermic Hypoxic Rat Liver Perfusion Model. The model consists in 4-h ex vivo perfusion: the liver is perfused with William Medium solution. Isovolemic hemodilution (mean hematocrit of 9.67  0.66%) is performed to provide suboptimal oxygen delivery and to induce a limited but progressive hypoxic injury. During the perfusion, perfusate samples are hourly collected to evaluate the metabolic status (pH, pO2, pCO2), while the hepatic function is monitored by assessing cytolysis markers in the perfusate (AST, ALT, LDH). Furthermore, bile production is quantified. In particular, even in a short-duration model, pH self-regulation, and bile production confirm the organ function despite the ongoing injury. Nevertheless, the system can be used also to deliver drugs, cells, or cell by-products. The feasibility to use cells or their by-products has been demonstrated by us [10], where we explored the feasibility of pharmacological intervention on isolated rat livers by using a combination of NMP and human liver stem cells-derived extracellular vesicles (HLSC-EV). The results demonstrate the feasibility of this NMP model in basic and preclinical research.

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2.1

Animals

2.2

Surgery

The procedures involving animals have to be in accordance with the guidelines of the ethics committee. Male Wistar rats aged 8–12 weeks (200–250 g weight). 1. Polystyrene box with ice. 2. 0.2 mg/kg Tiletamine-zolazepam. 3. 16 mg/kg Xylazine. 4. 50 mL Celsior solution. 5. 25,000 IU/5 mL Heparin. 6. Petri dishes—TC Dish 100 Standard. 7. Cannulae 24 GA 0.7  19 mm flowrate 20 mL/min. 8. Cannulae 18 GA 1.3  45 mm flowrate 97 mL/min. 9. Cannulae 22 GA 0.9  25 mm flowrate 35 mL/min. 10. Sterile utility drapes. 11. 50 mL Falcon tube. 12. Infusion kit with 3 stopcocks manifold. 13. Disposable sterile infusion manifold three-way stopcock. 14. Sterile surgical gauzes. 15. 0.5 mL, 1 mL, 10 mL, 50 mL vol. Syringes.

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16. Cannula caps Luer Lock-red stopper. 17. Scissor: Blunt/Blunt, 11.5 cm. 18. Scissor: 3 mm cutting edge, 0.05 mm tip diameter. 19. Fine forceps angled 45 : smooth tip width 0.4 mm  9 cm. 20. Graefe forceps: serrated tip width 0.8 mm. 21. 0.9% NaCl solution. 22. 90% Ethyl alcohol. 23. Silk suture 3/0. 24. Shaver. 2.3 Normothermic Machine Perfusion System

1. Moist Chamber TYPE 834/8 with metal tube heat exchanger for ex vivo perfusion organ (Harvard Apparatus, Holliston, MA, United States). 2. Sets of mini ball joint holders and cannulae (Harvard Apparatus, Holliston, MA, United States). 3. Bubble trap for flow rate up to approx. 50 mL/min Vol 1.6 mL (Harvard Apparatus, Holliston, MA, United States). 4. Windkessel (Harvard Apparatus, Holliston, MA, United States). 5. Peristaltic device. 6. Peristaltic pump REGLO analog Peristaltic Pump—ISMATEC, Wertheim, Germany). 7. Oxygenator: Hollow Fiber Oxygenator Type D150, Hugo Sachs Elektronik, March-Hugstetten, Germany). 8. PLUGSYS Modular Measuring & Control System—Case Type 601(Harvard Apparatus, Hugo Sachs Elektronik, MarchHugstetten, Germany). 9. Transducer Amplifier Module (TAM-D) (Harvard Apparatus, Hugo Sachs Elektronik, March-Hugstetten, Germany). 10. Servo Controller (SCP Type 704) (Harvard Apparatus, Hugo Sachs Elektronik, March-Hugstetten, Germany). 11. Thermocirculator with temperature set at 37  C. 12. Pump tubing Tygon S3™ E-LFL i.d. 2.06 mm, wall 0.84 mm. 13. Silicone Tubing: Platinum L/S 16. 14. Oxygen: O2 3.0 99.9% pure, 40 L Cylinder. 15. Gas Cylinder Pressure Regulator.

2.4 Perfusion Solution

1. Complete Williams Medium: Phenol red-free Williams E Medium, supplemented with 11.6 mM glucose, 50 U/mL penicillin, 50 μg/mL streptomycin, 5 mM L-glutamine, 1 U/mL insulin, 1 U/mL heparin.

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2. Perfusion solution: Isovolemic hemodilution is performed by adding 20 mL of fresh rat blood to 50 mL of complete Williams Medium, thus obtaining a mean hematocrit of 9.67  0.66%, sparge with 99% oxygen using oxygenator and add 2 mEq of sodium bicarbonate as a pH buffer. 3. 500 U Heparin is added hourly during perfusion. 2.5

Analysis

1. Blood gas analyzer: with kit membrane for pH, CO2, O2; cleaning and calibration solution; gas cylinder (see Note 1). 2. Centrifuge. 3. 1.5 mL Micro tube. 4. Emogas analyzer. 5. Scale. 6. Thermometer.

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Methods

3.1 Normothermic Machine Perfusion System (See Fig. 1) 3.1.1 Setting of Open Perfusion Circuit

1. Connect tubing from pump to the oxygenator. 2. Connect tubing from oxygenator to the bubble trap. 3. Connect tubing from bubble trap to the liver portal vein (PV) through the cannula (moist chamber). 4. Connect tubing peristaltic pump.

from

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Fig. 1 Normothermic machine perfusion schematic overview: (A) Liver perfusion chamber. (B) Peristaltic pump. (C) Oxygenator. (D) Bubble trap. (E) Thermostatic circulator

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5. Insert and connect with tubing one infusion kit with 3 stopcocks manifold before the moist chamber (after the oxygenator). 6. Insert and connect with tubing one disposable sterile infusion manifold three-way stopcock after the chamber (see Note 2). 7. Connect the oxygenator to the O2 cylinder. 3.1.2 Setting of Warm Water Circuit

1. Connect tubing from the thermocirculator output to moist chamber input. 2. Connect tubing from the moist chamber output to the thermocirculator input. 3. Refill the thermocirculator container with distilled water (see Note 3). 4. Set the temperature at 37  C and switch on to start the water circulation.

Surgery

Two animals are necessary for each experiment: one rat is used for blood collection to complete the perfusion solution while the second rat is used for liver procurement.

3.2.1 Blood Collection for Perfusion Solution

1. Anesthetize animals through an intramuscular injection of 0.2 mg/kg Zolazepam and 16 mg/kg Xylazine.

3.2

2. Administrate 1250 U of heparin intraperitoneally. 3. Shave the abdominal surface and disinfect with ethanol. 4. Perform an intracardiac puncture, pricking just above the xiphoid appendix and directing the needle at 45 on the midline or slightly to the left. 5. Aspirate a satisfactory volume of blood (about 13–15 mL). 6. Open the abdomen and chest with a median sternotomy to expose the cardiac area. 7. Open the pericardial cavity, cut off the left ventricle, and complete the aspiration of any residual blood volume, approximately 5 mL to reach the final volume of 20 mL (see Note 4). 8. Perform (if necessary) a cervical dislocation to avoid any kind of suffering of the animal. 9. Collect blood in a 50 mL tube with 50 U heparin and maintain it in ice until you use for the perfusion solution. 3.2.2 Hepatectomy for Perfusion Procedure (See Fig. 2a, b)

1. Anesthetize animals through an intramuscular injection of 0.2 mg/kg Zolazepam and 16 mg/kg Xylazine. 2. Administrate 1250 U of heparin intraperitoneally. 3. Shave the abdominal surface and disinfect with ethanol.

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Fig. 2 (a) Surgery procedure: midline laparotomy (A), exposed common bile duct (B), cannula insertion through the common bile duct (C), cannula secured (D), portal vein cannula positioning (E). Cannula outputs secured with caps Luer Lock stoppers (F), flushing with cold celsior solution through the PV cannula (G–I). (b) Surgery procedure: hepatectomy (L), washing with NaCl (M), the liver graft placed in a petri dish on ice surrounded by celsior solution (N)

4. Proceed with midline laparotomy by pinching and retracting the skin with the forceps, thereby perform a small incision with the scissor tip. 5. Insert the scissor through the incision and slide the scissor gently until the sternum. 6. Separate the abdominal skin to the muscle by sliding the open scissor from up to down.

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7. Perform a midline laparotomy following the midline (Linea alba). First the skin, then the muscle. 8. Retract the borders with retractors to facilitate the overview (see Fig. 2a(A)). 9. Retract the bowel and expose the liver and the hepatic pedicle. 10. Place the first 5 cm 3/0 silk suture without needle around the common bile duct at 5–6 mm distal from the prehepatic bifurcation (see Fig. 2a(B)). 11. Place a second 5 cm 3/0 silk suture without needle around the common bile duct at 7–8 mm distal from previous one (see Fig. 2a(C)). 12. Insert the 22-G cannula through the common bile duct, making sure to position the cannula tip at 2–3 mm away from the prehepatic bifurcation. 13. Block the cannula by knotting the previously positioned silk sutures, first distal one and then proximal one (see Fig. 2a(D)). 14. Positioning the 18-G cannula similarly as described in steps 10–13 for portal vein (PV) (see Fig. 2a(E/F)). 15. Flush gently and slowly 10 mL of cold Celsior solution through the PV cannula (see Fig. 2a(G–I)). 16. After sternotomy, make an incision in the heart with the scissor to open allow the blood-Celsior solution efflux. 17. Restart the liver flushing with additional 30 mL of cold Celsior solution through the PV cannula. 18. After perfusion is completed, block all the cannulas outputs with the caps Luer Lock stoppers to avoid the Celsior outflux. 19. Proceed with the hepatectomy by transecting all liver ligaments, suprahepatic, and infrahepatic inferior vena cava and separating from all the adjacent organs (see Fig. 2b(L)). 20. Wash the liver with NaCl and place the liver in an ice precooled petri dish and weigh the liver (see Note 5 and see Fig. 2b (M/N)). 21. Fill the petri dish with around 20 mL ice-cold Celsior solution and transport on ice to the perfusion room and place it in the moist perfusion chamber (see Fig. 3). 22. Wash with around 10 mL complete Williams Medium through the PV cannula, connect to the NMP circuit, and perfuse at 37  C through PV cannula under controlled pressure and flow conditions during 4 h (see Subheading 3.3).

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Fig. 3 Liver graft in the perfusion moist chamber 3.3 Perfusion Procedure

1. Connect the liver through PV cannula and connect bile cannula to an external drainage to collect bile (see Note 6). 2. Set the pressure parameter in the controller at 8–10 mmHg (see Note 7). 3. Switch on the controller and peristaltic pumps to start the ex vivo perfusion.

3.4 Analysis and Results

1. Before starting the perfusion measure weight and temperature of the organ.

3.4.1 Preliminary Analysis and Operating Parameters

2. Record the warm ischemia time and the total ischemia time during the surgical procedure and until the beginning of the perfusion, respectively (see Fig. 4). 3. Monitor flow rates and venous resistances during perfusion at constant pressure (see Fig. 5).

3.4.2 Perfusate Analysis

1. Perform blood gas analysis hourly during perfusion on in-flow and out-flow perfusate samples, testing pO2, pCO2, and pH (see Fig. 6). 2. With a 1 mL syringe take two samples of 1 mL of blood each: one from the first sterile infusion manifold three-way stopcock (before the oxygenator) and the second from the other one manifold (after the oxygenator).

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Fig. 4 Parameters measured prior perfusion procedure

Fig. 5 Flow rates and venous resistances during normothermic machine perfusion

3. Put the syringe in the Emogas analyzer and wait for the analysis. 4. Take the first one sample (the first collected from the manifold before the oxygenator) for further biochemistry analysis and turn back the second sample in the circuit. 3.4.3 Biochemistry Analysis

1. Collect a sample for biochemistry analysis every hour (see Subheading 3.4.2 steps 2–4).

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Fig. 6 pH trend during normothermic machine perfusion

2. Take the 1 mL outflow samples and centrifuge at 2700  g for 10 min at 4  C to eliminate cells and cell debris. 3. Store the cell-free supernatants at 80  C until aspartate aminotransferase (AST), alanine aminotransferase (ALT), and lactate dehydrogenase (LDH) levels are assessed by the Biochemistry Laboratory (see Fig. 7). 4. After every sampling restore the volume with 1 mL of complete Williams Medium. 3.4.4 Bile Analysis

1. Collect bile during the perfusion by connecting the bile duct cannula to a reservoir to quantify bile production. 2. Quantify the volume at the end of each perfusion (see Fig. 8).

3.4.5 Histological Analysis (See Note 8)

1. At the end of the perfusion, two or more liver lobes can be collected for histological analysis. At the end of each perfusion, whole circuit must be cleaned by washing with distilled water. For moist chamber and oxygenator, follow the manual instructions.

4

Notes 1. If other parameters such as K, Na, and Lactate need to be investigated, specific electrodes (all from A.DeMori Group, Milan, Italy) should be acquired. 2. You can use optionally “Infusion kit with 3 stopcocks manifold” or “Disposable sterile infusion manifold three-way

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Fig. 7 Biochemical profile of hepatic cytolysis and function markers of rat livers assessed at different time points during ex vivo isolated perfusion. All values are normalized to the animal liver weight in grams. Data are represented as mean  SEM

stopcock” it is the same but with the first one it is probably more easy to take samples. 3. Refill the thermocirculator device with distillate water as appropriate. Do not use fresh tap water. For more details, follow the manual instructions. 4. If the blood collected from the first rat is not enough, the second animal must be used. In this case, 5 mL or the amount necessary to reach 20 mL of total blood must be collected in the same way, at the same time as cutting the left ventricle but just before perfusing with the Celsior solution (see Subheading 3.2.2 steps 15, 16). 5. Weight an empty petri dish plus cannulaes plus caps on a calibrated balance before assessing liver weight. In this way, you can have the real net weight of the liver at the end of the perfusion. 6. You can use centrifuge plastic tube to collect the bile and biochemistry sample in a 1.5 mL tube.

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Fig. 8 Amount of bile produced during normothermic machine perfusion

7. By setting the pressure controller at 8–10 mmHg, you can obtain a 1.1/1.3 mL/min/g flow. 8. At the end of the perfusion, two or more liver lobes can be collected for histological analysis. Standard histological procedures can be followed as properly. References ˜ o J, Marchal T, Padillo J et al (2002) 1. Bricen Influence of marginal donors on liver preservation injury. Transplantation 74:522–526 2. O’Callaghan JM, Morgan RD, Knight SR et al (2014) The effect of preservation solutions for storage of liver allografts on transplant outcomes: a systematic review and meta-analysis. Ann Surg 260:46–55 3. Ravikumar R, Leuvenink H, Friend PJ (2015) Normothermic liver preservation: a new paradigm? Transpl Int 28:690–699 4. Laing RW, Bhogal RH, Wallace L et al (2017) The use of an acellular oxygen carrier in a human liver model of normothermic machine perfusion. Transplantation 101:2746–2756 5. Goldaracena N, Spetzler VN, Echeverri J et al (2017) Inducing hepatitis C virus resistance after pig liver transplantation-a proof of concept of liver graft modification using warm ex vivo perfusion. Am J Transplant 17:970–978

6. Brockmann J, Reddy S, Coussios C et al (2009) Normothermic perfusion: a new paradigm for organ preservation. Ann Surg 250:1–6 7. Jamieson RW, Zilvetti M, Roy D et al (2011) Hepatic steatosis and normothermic perfusionpreliminary experiments in a porcine model. Transplantation 92:289–295 8. Tolboom H, Pouw RE, Izamis ML et al (2009) Recovery of warm ischemic rat liver grafts by normothermic extracorporeal perfusion. Transplantation 87:170–177 9. Ravikumar R, Jassem W, Mergetal H et al (2016) Liver transplantation after ex vivo normothermic machine preservation: a phase 1 (first-in-man) clinical trial. Am J Transplant 16:1779–1787 10. Rigo F, De Stefano N, Navarro-Tableros V et al (2018) Extracellular vesicles from human liver stem cells reduce injury in an ex vivo Normothermic hypoxic rat liver perfusion model. Transplantation 102:e205–e210. https://doi. org/10.1097/tp.0000000000002123

Chapter 11 Co-Culture of Human Mesenchymal Stromal Cells and Primary Mouse Hepatocytes Mei-Ju Hsu, Madlen Christ, and Bruno Christ Abstract Human mesenchymal stromal cells (MSC) are adult stem cells, which feature hepatotropism by supporting liver regeneration through amelioration of hepatic inflammation and lipid accumulation in a mouse model of non-alcoholic steatohepatitis (NASH), a more advanced stage of fatty liver. It remains open, how MSC impact on hepatocytic lipid metabolism. To study MSC actions on fatty liver mechanistically, we established an in vitro model of co-culture comprising MSC and isolated mouse hepatocytes at a ratio of 1:1. Lipid storage in hepatocytes was induced by the treatment with medium deficiency of methionine and choline (MCD). The protocol can be adapted for the use of other lipid storage-inducing agents such as palmitic acid and linoleic acid. This co-culture model allows to study, e.g., whether MSC act indirectly via MSC-born paracrine mechanisms or through direct physical interactions between cells beside others. The protocol allows us to detect the formation of extensions (filopodia) from MSC to contact the fatty hepatocytes or other MSC within 24 h of co-culture. These structures may represent tunneling nanotubes (TNT), allowing for long-range intercellular communication. Key words Co-culture, Mesenchymal stromal cells, Primary hepatocytes, Fatty liver

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Introduction Obesity is one of the most prevalent health problems worldwide, which has been attributed to changes in lifestyle by increasing food intake and/or reducing physical activity. It is associated with metabolic comorbidities such as non-alcoholic fatty liver diseases (NAFLD), which has a global prevalence of 24%, and is currently the leading cause of chronic liver disease in the USA and Europe [1]. NAFLD may progress from simple steatosis to chronically inflammatory diseases like non-alcoholic steatohepatitis (NASH), cirrhosis, and HCC [2, 3]. Liver transplantation in NASH patients has been shown to reduce the risk of progression into HCC [4, 5]. However, the shortage of cadaveric donor livers and the high costs make MSC an alternative to whole liver transplantation [6]. Indeed, transplantation of MSC has been performed clinically

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to improve liver diseases since 2010. Clinically, the application of MSC mitigated inborn errors of liver metabolism such as ornithine carbamoyltransferase deficiency [7], decreased the severe bleeding complications in a hemophilia A patient [8], and improved liver function in cirrhotic patients [9–11]. However, some studies revealed only transient or even negligible effects of MSC treatment on liver diseases [12, 13]. Thus, the identification of the mechanisms involved in MSC actions is crucial to establish and improve the therapeutic outcome of MSC therapy. Previous studies showed that MSC-derived conditioned medium may contain hepatotropic factors [14] and exosomes [15] that potentially support liver regeneration by paracrine or systemic effects. Indeed, infusion of soluble factors secreted by MSC was sufficient to support hepatocyte growth in a rat model of D-galactosamine-induced acute liver injury [16]. Besides, many pharmacological companies have successfully isolated exosomes and extracellular vesicles from MSC to treat other than liver diseases [17]. This chapter provides methods that can be used to study the molecular and cellular interactions between MSC and primary hepatocytes, allowing to understand the effects of MSC on the regulation of hepatocytic lipid homeostasis. The protocol describes the isolation of mouse hepatocytes and human bone marrowderived MSC, their growth and hepatogenic differentiation, as well as a direct co-culture system, in which MSC and mouse hepatocytes are seeded in the same culture dish and challenged with steatosis-inducing MCD medium. By using this co-culture system, we discovered that MSC reduced hepatocyte lipid load independent of paracrine mechanisms [18]. Instead, a specific way of intercellular communication was identified encompassing the extension of tubular structures from the MSC physically contacting other MSC or hepatocytes. In other disease models, these so-called tunneling nanotubes (TNT) may mediate MSC actions by bidirectional delivery of organelles (such as mitochondria) and cytoplasmic contents [19].

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Materials All solutions are prepared with deionized H2O. Cell culture media and buffers for mouse hepatocyte isolation are sterilized by filtration with a 0.2 μm filter. We recommend the usage of the cell culture media within 1 month after preparation. Surgical instruments, PBS, and glass culture wares should be sterilized by autoclaving. All consumables for cell culture were either autoclaved or purchased in sterile packages. Laminar flow cabinet should be kept sterile by disinfection spray prior to use.

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1. Maintenance Medium Containing 6 Factors (EM6F) MSC growth medium: D-MEM basic medium: add 3.7 g/L NaHCO3 to D-MEM containing 1 g/L glucose and 580 mg/L L-glutamine, adjust pH to 7.2. MCDB 201 basic medium: dissolve 17.7 g of powder with 1 L of distilled water, adjust pH to 7.2. Mix 225 mL D-MEM and 170 mL MCDB 201 for EM6F and add 5 ng/mL 98% selenious acid, 1 nM dexamethasone, 5 μg/mL insulin, 0.1 mM ascorbic acid, 5 μg/mL apotransferrin, 4.7 μg/mL 99% linoleic acid liquid, 15% (v/v) heatinactivated fetal calf serum (FCS), 100 IU/mL penicillinstreptomycin. Store at 4  C. 2. Hepatocyte Growth Medium (HGM): 500 mL D-MEM without glucose, 500 mL D-MEM with 4.5 g/L glucose, 2 g albumin, 11.1 mM galactose, 0.6 mM L-ornithine, 0.26 mM L-proline, 9.98 mM HEPES, 2.5 mM nicotinamide, 0.99 mM glutamine, 54.4 μg/L ZnCl2, 20 μg/L CuSO4  5 H2O, 75 μg/L ZnSO4  7 H2O, 25 μg/L MnSO4  H2O, 100 IU/mL penicillin-streptomycin, 0.1 μM dexamethasone, 0.1% (v/v) insulin-transferrin-sodium selenite supplement, 20 ng/mL epidermal growth factor (EGF), and 40 ng/mL hepatocyte growth factor (HGF). Store at 4  C (see Note 1). 3. MEM: MEM Earle’s, 500 ng/mL insulin, 50 μg/mL gentamycin, and store at 4  C. 4. HHMM: mix 250 mL HGM and 250 mL MEM, add 10 ng/ mL EGF, 20 ng/mL HGF, 2% (v/v) heat-inactivated FCS and store at 4  C (see Note 2).

2.2 Isolation of Human Bone Marrow-Derived Mesenchymal Stromal Cells (MSC)

Ethics Committee approval and written informed consent must be obtained from patients or their relatives prior to collection of human samples. Experiments must be performed strictly sticking to these permissions. 1. Human bone marrow tissue. 2. PBS. 3. Citrate buffer: 311 mg sodium citrate in 10 mL distilled water, adjust pH to 7.4. 4. Krebs-Ringer buffer: dissolve 700 mg NaCl, 36 mg KCl, 30 mg MgSO4  7 H2O, 16.7 mg KH2PO4, and 201.7 mg NaHCO3 in 100 mL of distilled water. 5. Collagenase NB4G: dissolve 15 mg in 20 ml D-MEM high glucose (see Note 3). 6. 125-μm nylon mesh. 7. Biocoll with the density of 1.077 g/mL.

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8. Heat-inactivated FCS. 9. Scale. 2.3 Culture Plate Coating

1. Cell culture plates. 2. Glass coverslips for imaging: coverslips must be sterilized by 70% ethanol or autoclave and air dry prior to coating. 3. Human fibronectin: 100 μg/mL stock, store at 20  C. Fibronectin working solution: 1:200 dilution with D-MEM or EM6F to 5 μg/mL. 4. Collagen R: 0.2% stock, store at 4  C. 0.01% collagen working solution: 1:20 dilution of stock with distilled water.

2.4

Culture of MSC

1. EM6F. 2. Uncoated flasks for growing fresh MSC after isolation; fibronectin-coated flasks for growing of thawed MSC.

2.5 Cryopreservation of MSC

1. 1 Trypsin-EDTA. 2. EM6F. 3. Prepare freezing medium prior to cell freezing: 7.5% DMSO in heat-inactivated FCS. We recommend preparing 1 mL for each cryotube. 4. Cryovials. 5. Mr. Frosty™ Freezing Container filled with isopropyl alcohol. 6. 0.4% Trypan blue solution.

2.6

Thawing of MSC

1. Water bath set at 37  C. 2. EM6F. 3. 15-mL tube. 4. Fibronectin-coated culture surfaces.

2.7 Hepatogenic Differentiation of MSC

1. MSC on fibronectin-coated plates grown to 90% of confluency. 2. 5-azacytidin (AZA) stock: 4 mM in D-MEM. Final concentration: 20 μM AZA in culture. 3. HHMM.

2.8

Isolation of HC

The protocol and approval from local legislation of the Animal Welfare Act must be obtained prior to any animal experiments. The animals are kept with a 12-h circadian rhythm with free access to standard diet and drinking water. Male Pfp/Rag2 / (C57BL/ 6N(B6.129S6-Rag2(tm1Fwa)Prf1(tm1Clrk))) mice aged around 14-week-old are used in the current study. However, the use of other mouse strains or another gender can be altered to fit the purpose of the study.

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1. Krebs-Ringer buffer: Dissolve 4.2 g NaCl, 0.22 g KCl, 0.18 g MgSO4  7 H2O, 0.1 g KH2PO4, and 1.21 g NaHCO3 in 600 mL of distilled water. 2. Pre-perfusion buffer: 300 mL of Krebs-Ringer buffer, 30 mg EGTA. Equilibrate with carbogen for 30 min. Adjust pH to 7.35. 3. Perfusion buffer: 50 mL of Krebs-Ringer buffer, 178.5 mg HEPES, 29.5 mg CaCl2  2H2O. Equilibrate with carbogen for 30 min. Adjust to pH 7.5. Add 250 μL collagenase NB4G and mix well prior to use. 4. Wash buffer: 1 L distilled water, 4.77 g HEPES, 7 g NaCl, 0.36 g KCl, 0.3 g MgSO4  7H2O, 0.16 g KH2PO4, 4 g bovine serum albumin. Equilibrated with carbogen for 30 min. Adjust to pH 7.4. Add 50 mg DNase I and mix well. 5. Isoflurane. 6. Narcoren: 16 g pentobarbital-natrium/100 mL. 7. 25,000 IU/1.25 mL Heparin. 8. 22-G catheter. 9. Glass funnel. 10. Gauze. 11. Low-rise beaker. 12. 50-mL tube. 2.9

Co-Culture

1. Trypsin-EDTA. 2. Co-Culture MEM: MEM, 2% FCS. 3. Collagen-coated culture dishes or well plates (optional: collagen-coated glass coverslips). 4. MCD medium: 1 L methionine-choline-deficient (MCD) medium add 2 g albumin, 11.1 mM galactose, 0.6 mM Lornithine, 0.26 mM L-proline, 9.98 mM HEPES, 2.5 mM nicotinamide, 0.99 mM glutamine, 54.4 μg/L ZnCl2, 20 μg/L CuSO4  5 H2O, 75 μg/L ZnSO4  7 H2O, 25 μg/L MnSO4  H2O, 100 IU/mL penicillinstreptomycin, 0.1 μM dexamethasone, 0.1% (v/v) insulintransferrin-sodium selenite supplement, 20 ng/mL epidermal growth factor, and 40 ng/mL hepatocyte growth factor. Store at 4  C (see Note 1).

2.10 Oil Red O (ORO) Staining of Lipids

1. Cells grown on collagen-coated glass coverslips. 2. 3.7% formalin in PBS. 3. ORO stock solution: 0.5 g in 100 mL isopropanol. Prepare working solution freshly by mixing three parts of ORO stock

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solution with two parts of distilled water. The solution is ready to use after filtration via filter paper. 4. ProLong™ Gold Antifade Mountant with DAPI. 5. Glass slides. 2.11 Staining of Co-Culture for F-Actin

1. Cells grown on collagen-coated glass coverslips. 2. Glass slides. 3. 3.7% formalin in PBS. 4. 0.1% Triton X-100 in PBS. 5. 0.1% Phalloidin-conjugated with fluorochrome (e.g., Phalloidin-iFluor 488 from Abcam) in PBS containing 1% BSA. 6. ProLong™ Gold Antifade Mountant with DAPI.

2.12 Equipment and Non-Disposable Materials

1. 2.5, 10, 20, 100, 1000 μL pipettes. 2. 4  C,

20  C, and

85  C fridges.

3. Autoclave. 4. Centrifuges for 1.5 mL, 15 mL, and 50 mL tubes. 5. Face shield. 6. Hemocytometer and coverslips. 7. Humidified incubator (37  C with 5% CO2). 8. Insulating gloves. 9. Laminar-flow cabinet. 10. Liquid nitrogen tank. 11. Mr. Frosty™ Freezing Container. 12. Phase contrast microscope. 13. Fluorescence microscope. 14. Safety goggles. 15. Surgical forceps. 16. Surgical scissors. 17. Vortexer. 18. Water bath. 19. Water purification system. 20. Rocking platform. 21. Perfusion pump with tubing. 22. Bubble trap. 23. Heat exchanger. 24. Fume hood. 25. Gloves for handling liquid nitrogen.

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2.13 Disposable Materials

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1. 2.5, 10, 20, 100, 1000 μL pipette and tips. 2. 3.5-, 6-, 10-, 15-cm culture dishes. 3. 10, 25, 50 mL serological pipets. 4. 12-well plates. 5. Filtermax rapid bottle-top or any filter with capacity of 500 mL. 6. Glass coverslips. 7. T-25 and T-75 sterile plastic flasks.

3

Methods For cell culture procedures, work in an aseptic tissue culture hood and warm all media at 37  C in a water bath before use. A schematic diagram of the experimental flow is shown in Fig. 1.

3.1

Isolation of MSC

1. After excision, store the bone marrow tissue in D-MEM at 4  C, and isolate MSC within a maximum of 16 h. 2. Record tissue weight and calculate the amount of collagenase NB4G required for tissue digestion (see Note 3). 3. Under a laminar flow cabinet, soak tissue in a solution consisting of 20 mL of D-MEM high glucose, 2 mL of citrate buffer and collagenase NB4G. 4. Incubate the tube for 25–30 min at 37  C on a rocking platform, but without vortexing.

Fig. 1 Schematic plan of experimental procedures and timetable. Mice aged between 10 and 14 weeks were used for hepatocyte (HC) isolation. The MSC were thawed and grown in EM6F for expansion until reaching 80–90% of confluency. 5-azacytidin (AZA) was added to prime the cells for differentiation. After 24 h, cells were cultured with hepatogenic differentiation medium HHMM for an additional 16 days. At the end of differentiation, MSC are ready for co-culture with HC. MSC were detached from the culture plates and HC isolated from liver tissue, respectively. Two to three hours after seeding, the unattached cells are removed, and co-culture was incubated in control (HGM) or stimuli/chemical of interest

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5. Stop the digestion of the tissue by adding 5 mL of heatinactivated FCS. 6. Remove the undigested remnant tissue by filtrating with a 125-μm nylon mesh. 7. Pellet down the cells by centrifugation for 7 min at 4  C, 228  g. 8. Carefully remove the supernatant by suction. 9. Resuspend pellet with 20–40 mL of cold PBS and centrifuge for 5 min at 4  C, 228  g. Repeat this wash step for 2–3 times. 10. Resuspend pellet in 1 mL of PBS and gently layer it onto 10 mL of Biocoll (1.077 g/mL) without disturbing the interface. 11. Centrifuge at 228  g, 4  C for 20 min without brake. 12. Slowly obtain the MSC-containing white band at the interface and transfer to a new 15 mL tube. 13. Wash the cell suspension with 10 mL of PBS and mix gently by inversion. 14. Centrifuge at 228  g, at 4  C for 5 min. Repeat this wash step once. 15. Resuspend cell pellet with EM6F and seed onto uncoated plastic tissue culture flasks (see Note 4). 16. Incubate the cells at 37  C, 5% CO2. 17. Change the medium 24 h after seeding to remove the non-adherent cells, then change medium every 4–5 days until reaching 90% of confluency. 3.2 Cryopreservation of MSC

1. After growth of cells to 90% of confluency, wash cells with PBS and dissociate cell monolayer by adding trypsin approx. 1.5 mL for a T75 flask. 2. Allow the trypsin digestion at 37  C in an incubator for around 3–5 min. Monitor cell dissociation under a microscope to adjust time of incubation. 3. Add 8.5 mL of EM6F to stop enzymatic activity. 4. Take 50 μL sample from cell suspension and determine the cell density and viability using trypan blue exclusion assay. 5. Mix cell suspension sample with trypan blue 1:1 (may require to be further diluted empirically). 6. Apply 10 μL of the solution to a hemocytometer. 7. Count viable, unstained cells and dead cells in blue under the microscope. 8. The number of cells is calculated according to the formula provided by the manufacturer of the hemocytometer.

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9. Calculate the viability of the cells: [total viable cells (unstained)  total cells (viable + dead)]  100%. 10. Centrifuge cell suspension at 228  g for 5 min and then remove the supernatant. 11. Resuspend cells in freezing medium. 12. Label the cryovial with date, cell number, and donor number (see Note 5). 13. Place the cryovials in Mr. Frosty™ Freezing Container filled with isopropyl alcohol and place it at a 80  C freezer overnight. 14. Transfer the tubes to the vapor phase in the liquid nitrogen for longer storage. 15. Record the date, cell number, donor number, number of tubes, and user name in the cryopreservation logbook. 3.3 Coating Well Plates or Coverslips

Depending on the experiments, cells are seeded onto different culture plates with different coatings. For the purpose of amplification of cell numbers for co-culture experiments and of harvesting protein or RNA, cells are grown on flasks or dishes; whereas cells are seeded onto well plates (with or without glass coverslips) for experimental purposes. For imaging assays, cells are seeded onto 12 and 18 mm glass coverslips in 24 and 12 well plates, respectively. 1. For the expansion of MSC for further differentiation, coat flasks or dishes with fibronectin. Rinse the culture surfaces with human fibronectin working solution without leaving rest of fluid. 2. After air drying at room temperature under a safety hood for around 30 min, the plates are ready to use. 3. Mouse hepatocytes, differentiated MSC and co-cultures are grown on collagen-coated surfaces (see Note 6). Prepare collagen working solution and add an appropriate volume of collagen solution to reach a final concentration of 6 μg/cm2:

Culture plates

Collagen working solution

10 cm dish

3600 μL

6 cm dish

1326 μL

3.5 cm dish

552 μL

6-well plate

542 μL/well

12-well plate

220 μL/well

24-well plate

115 μL/well

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4. Plates are left overnight under a safety hood to allow air drying. 5. The next day, wash the plates twice with sterile water. 6. After air drying, coated plates are ready to use, or can be carefully sealed to keep sterility and stored at 20  C. 3.4 Thawing and Growing of Frozen MSC

1. Set up a water bath at 37  C and prepare a 15-mL tube containing 9 mL of EM6F or D-MEM. 2. Wear a face shield or safety goggles and insulating gloves to ensure the safety of handling liquid nitrogen. 3. Take a cryovial with MSC from liquid nitrogen container, dip it in the water bath and gently swirl for 2 min until frozen content is thawed. 4. Pour the cells to the 15-mL centrifuge tube. 5. Centrifuge at 228  g for 5 min and remove the supernatant. 6. Resuspend the cells in 1 mL and determine the cell viability and density by the trypan blue exclusion assay (see Subheading 3.2 steps 4–9). 7. Seed the cells onto fibronectin-coated dishes or plates at a cell density of 400–500 cells/cm2 (see Note 7). 8. Change the cell growth medium every 3–4 days until reaching 90% of confluency.

3.5 Hepatogenic Differentiation of MSC

1. Add AZA stock solution directly to culture, dependent on the volume of medium to reach the final concentration (see Note 8). 2. After 24 h, replenish with HHMM every 3–4 days for a total of 16 days of differentiation.

3.6

Isolation of HC

1. Water bath set at 39.5  C and warm up the pre-perfusion buffer and perfusion buffer. 2. Anesthetize mice with inhalant isoflurane by using a 50-mL tube containing gauze soaked with isoflurane. 3. Record body weight. 4. Inject 50 μL of Narcoren and 50 μL of heparin intraperitoneally as supplementary material for anesthesia and prevention of blood clotting, respectively. 5. Inspect the anesthetic depth by toe pinch and monitoring the breathing. 6. Make ventral midline incision by surgical scissors and expose the inferior vena cava and portal vein. 7. Cannulate the inferior vena cava, remove the needle, press the heart, and let the blood fill the cannula. 8. Connect the cannula with the tube of the pump with tubes containing pre-perfusion buffer, cut portal vein.

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9. Adjust speed of the pump to 3.3 mL/min and perfuse the liver via the inferior vena cava for 1 min (see Note 9). Subsequently, perfuse the liver with perfusion buffer for around 15 min. 10. Remove the liver and record tissue weight. 11. Place liver in a 50 mL tube containing around 30 mL of wash buffer. 12. Transfer the liver to the safety hood in low-rise beaker; mince the liver tissue with scissors (see Note 10). 13. Layer the glass funnel with two layers of sterile gauze and place it on a 50-mL tube. 14. Collect the cell suspension into a 50-mL tube by pouring the liver digest through the gauze. 15. Centrifuge at 68  g for 5 min at 4  C. 16. Remove the supernatant and add 50 mL of wash buffer. 17. Repeat the steps 15 and 16 twice. 18. Add 50 mL of MEM and ascertain the cell density and viability using the trypan blue exclusion assay (see Subheading 3.2 steps 4–9 and see Note 11). 19. Adjust the viable cell density at 1.6  105 cells/mL. 3.7 Co-Culture of HC and Hepatogenic Differentiated MSC

1. Trypsinize the hepatogenic differentiated MSC for 5 min at 37  C (see Subheading 3.5). 2. Stop the digestion by adding co-culture MEM. 3. Centrifuge for 5 min at 4  C and remove the supernatant containing trypsin. 4. Resuspend cells in co-culture MEM. 5. Determine the cell density and viability with a hemocytometer under the microscope (see Subheading 3.2 steps 4–9). 6. Adjust the viable cell density at 1.6  105 cells/mL. 7. Seed the isolated HC together with hepatogenic differentiated MSC at different ratios onto collagen-coated coverslips or other culture surfaces in co-culture MEM at a total cell number of 4  104 cells/cm2. Ratios for seeding onto a 12-well plate (4 cm2/well): HC:MSC

HC

MSC

1:0

1000 μL

–

10:1

909.1 μL

90.9 μL

5:1

833.3 μL

166.7 μL

1:1

500 μL

500 μL

0:1

–

1000 μL

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8. Mix well by gently shaking. 9. Allow the cells to settle down for 2–3 h in the incubator. 10. Remove the supernatant. 11. Wash with PBS if too many floating cells are observed. 12. Replenish with medium containing the stimuli of interest, here HGM and MCD medium as control and steatosisinducing group, respectively.

3.8 Staining Lipids with ORO

13. Analyze co-cultures frequently for the formation of extensions (filopodia) from cells to detect tunneling nanotubes (TNT). 1. Remove supernatant and wash cells once with PBS. 2. Work in a fume hood, fix the cells with formalin for 15 min at RT. 3. Wash twice with distilled water (see Note 12). 4. Add 0.5 mL ORO working solution to cells for each well in a 12-well plate. 5. Allow the staining for 1 h at RT. 6. Wash the cells three times with distilled water. 7. Mount the coverslips onto glass slides with ProLong™ Gold Antifade Mountant with DAPI. An example of the lipid staining by ORO is shown in Fig. 2.

3.9 Staining of Co-Culture with Phallodin

1. Follow the same procedure as described in Subheading 3.8 steps 1–3. 2. Permeabilize the cell with 0.1% Triton X-100 in PBS for 5 min at RT (see Note 13). 3. Wash twice with PBS. 4. Stain cells with Phalloidin-iFluor 488 in 1% BSA for 90 min at RT.

Fig. 2 MCD medium promotes the accumulation of lipid droplets in mouse hepatocytes. Mouse hepatocytes were grown for 1 (a, c) and 3 days (b, d) either in hepatocyte growth medium (a, b, HGM), or in steatosisinducing MCD medium (c, d). Lipid droplets were stained with oil red O (ORO, red), and the nuclei were counterstained with DAPI (blue)

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Fig. 3 Morphological differences are sufficient to distinguish mouse hepatocytes and MSC. (a) Mouse hepatocytes (polygonal cells with enhanced nucleoli staining) and (b) human stem cells (spindle shape with abundant stress fibers) and oval nuclei. Nuclei and F-actin were stained with DAPI (blue) and phalloidin (green), respectively. (c) shows the presence of TNT (arrows) on day 1 after co-culture of HC and MSC

5. Wash twice with PBS. 6. Mount the coverslips onto glass slides with ProLong™ Gold Antifade Mountant with DAPI. An example of the cell morphology in culture and co-culture is given in Fig. 3.

4

Notes 1. Add EGF and HGF freshly prior to use. 2. Add EGF, HGF, and FCS freshly prior to use. 3. We recommend using 15 mg collagenase NB4G for digestion of tissue weighing 2 g for optimal results. 4. The cell population after isolation may contain other cell types in addition to MSC. Taking advantage of the plastic adherent property of MSC and removal of the supernatant on the next day would serve as a means to keep only MSC in culture. 5. We freeze 105–106 cells in each vial in our lab; this cell number is handy for downstream applications. However, the cell density can be adjusted by users according to their requirements. 6. According to our observation, the differentiated MSC after trypsinization showed better attachment onto collagen-coated than fibronectin-coated surfaces. 7. This cell density allows to achieve 90% of confluency (for further cell differentiation) after 6–10 days of culture. 8. AZA can be added either directly to the old cell supernatants or after changing to fresh media.

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9. The color of the liver turns to light yellow due to the replacement of blood with pre-perfusion buffer. 10. The buffer shall appear turbid and brown due to the release of HC from the liver. 11. A good manipulation would yield around 2.5  107 cells per mouse with the viability ranging from 70 to 90%. Do not use HC with viability less than 50%. 12. Collect the solution containing formalin and dispose the waste according to the local regulation. 13. This step is optional; it requires to be tested in different cell types. References 1. Younossi Z, Anstee QM, Marietti M et al (2018) Global burden of NAFLD and NASH: trends, predictions, risk factors and prevention. Nat Rev Gastroenterol Hepatol 15(1):11–20. https://doi.org/10.1038/nrgastro.2017.109 2. Tilg H, Moschen AR (2014) Mechanisms behind the link between obesity and gastrointestinal cancers. Best Pract Res Clin Gastroenterol 28(4):599–610. https://doi.org/10. 1016/j.bpg.2014.07.006 3. Calle EE, Rodriguez C, Walker-Thurmond K et al (2003) Overweight, obesity, and mortality from cancer in a prospectively studied cohort of U.S. adults. N Engl J Med 348 (17):1625–1638. https://doi.org/10.1056/ NEJMoa021423 4. Lewin SM, Mehta N, Kelley RK et al (2017) Liver transplantation recipients with nonalcoholic steatohepatitis have lower risk hepatocellular carcinoma. Liver Transpl 23 (8):1015–1022. https://doi.org/10.1002/lt. 24764 5. Cholankeril G, Wong RJ, Hu M et al (2017) Liver transplantation for nonalcoholic steatohepatitis in the US: temporal trends and outcomes. Dig Dis Sci 62(10):2915–2922. https://doi.org/10.1007/s10620-017-4684x 6. Aurich H, Sgodda M, Kaltwasser P et al (2009) Hepatocyte differentiation of mesenchymal stem cells from human adipose tissue in vitro promotes hepatic integration in vivo. Gut 58 (4):570–581. https://doi.org/10.1136/gut. 2008.154880 7. Sokal EM, Stephenne X, Ottolenghi C et al (2014) Liver engraftment and repopulation by in vitro expanded adult derived human liver stem cells in a child with ornithine carbamoyltransferase deficiency. JIMD Rep

13:65–72. https://doi.org/10.1007/8904_ 2013_257 8. Sokal EM, Lombard CA, Roelants V et al (2017) Biodistribution of liver-derived mesenchymal stem cells after peripheral injection in a hemophilia a patient. Transplantation 101 (8):1845–1851. https://doi.org/10.1097/ TP.0000000000001773 9. Kharaziha P, Hellstrom PM, Noorinayer B et al (2009) Improvement of liver function in liver cirrhosis patients after autologous mesenchymal stem cell injection: a phase I-II clinical trial. Eur J Gastroenterol Hepatol 21 (10):1199–1205. https://doi.org/10.1097/ MEG.0b013e32832a1f6c 10. Jang YO, Kim YJ, Baik SK et al (2014) Histological improvement following administration of autologous bone marrow-derived mesenchymal stem cells for alcoholic cirrhosis: a pilot study. Liver Int 34(1):33–41. https:// doi.org/10.1111/liv.12218 11. Suk KT, Yoon JH, Kim MY et al (2016) Transplantation with autologous bone marrowderived mesenchymal stem cells for alcoholic cirrhosis: phase 2 trial. Hepatology 64 (6):2185–2197. https://doi.org/10.1002/ hep.28693 12. Kanazawa Y, Verma IM (2003) Little evidence of bone marrow-derived hepatocytes in the replacement of injured liver. Proc Natl Acad Sci U S A 100(Suppl 1):11,850–11,853. https://doi.org/10.1073/pnas.1834198100 13. Higashiyama R, Moro T, Nakao S et al (2009) Negligible contribution of bone marrowderived cells to collagen production during hepatic fibrogenesis in mice. Gastroenterology 137(4):1459–1466. e1451. https://doi.org/ 10.1053/j.gastro.2009.07.006

Co-Culture of Stem Cells and Hepatocytes 14. Winkler S, Hempel M, Bruckner S et al (2016) Identification of pathways in liver repair potentially targeted by secretory proteins from human mesenchymal stem cells. Int J Mol Sci 17(7). https://doi.org/10.3390/ ijms17071099 15. Wang YH, Wu DB, Chen B et al (2018) Progress in mesenchymal stem cell-based therapy for acute liver failure. Stem Cell Res Ther 9 (1):227. https://doi.org/10.1186/s13287018-0972-4 16. van Poll D, Parekkadan B, Cho CH et al (2008) Mesenchymal stem cell-derived molecules directly modulate hepatocellular death and regeneration in vitro and in vivo. Hepatology 47(5):1634–1643. https://doi.org/10. 1002/hep.22236

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17. Gimona M, Pachler K, Laner-Plamberger S et al (2017) Manufacturing of human extracellular vesicle-based therapeutics for clinical use. Int J Mol Sci 18(6). https://doi.org/10. 3390/ijms18061190 18. Hsu M, Hempel M, Ku¨hne H et al (2019) Physical contact between mesenchymal stromal cells and hepatocytes via tunneling nanotubes favor the utilization of hepatocyte lipids. Paper presented at the German Association of the Study of the liver, Heidelberg, 22–23 Feb 2019 19. Mahrouf-Yorgov M, Augeul L, Da Silva CC et al (2017) Mesenchymal stem cells sense mitochondria released from damaged cells as danger signals to activate their rescue properties. Cell Death Differ 24(7):1224–1238. https://doi.org/10.1038/cdd.2017.51

Chapter 12 A 3D Dynamic In Vitro Model of Inflammatory Tendon Disease Susanna Schubert, Luisa Brandt, and Janina Burk Abstract Three-dimensional (3D) cell cultures combining multipotent mesenchymal stromal cells (MSC), tendon extracellular matrix scaffolds, and mechanical stimulation by a bioreactor have been used to induce tenogenic differentiation in vitro. Yet, these conditions alone do not mimic the environment of acute inflammatory tendon disease adequately, thus the results of such studies are not representatives for tendon regeneration after acute injury. In this chapter, we describe two different approaches to introduce inflammatory stimuli, comprising co-culture with leukocytes and supplementation with the cytokines IL-1β and TNF-α. The presented in vitro model of inflammatory tendon disease could be used to study musculoskeletal pathophysiology and regeneration in more depth. Key words Regenerative medicine, Tissue engineering, Horse, Mesenchymal stromal cells, Scaffold, Tenogenic Differentiation, Leukocytes, Interleukin-1 (IL-1), Tumor necrosis factor-α (TNF-α)

1

Introduction For tendon regeneration, multipotent mesenchymal stromal cells (MSC) are considered as potent therapeutic tools. In contrast to tenocytes as highly differentiated cells, MSC have higher selfrenewal and lineage differentiation potential. Harvesting of MSC is uncomplicated and MSC have a higher proliferation and synthetic activities compared to terminally differentiated cells [1]. Therefore, for treating tendon disease, the application of MSC is preferred over the use of tenocytes. Transplantation of MSC and their subsequent tenogenic differentiation may lead to cell replacement and matrix modulation. Yet their mode of action is still not fully understood. In particular, it is still unclear how the regenerative effects of MSC are mediated in the inflammatory environment of acute tendon disease [2]. The aim of the here presented approach is to create a threedimensional, dynamic co-culture model that facilitates the investigation of tendon regeneration in vitro by closely reflecting the

Peggy Stock and Bruno Christ (eds.), In Vitro Models for Stem Cell Therapy: Methods and Protocols, Methods in Molecular Biology, vol. 2269, https://doi.org/10.1007/978-1-0716-1225-5_12, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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environment of acute inflammatory tendon disease. The central components of this in vitro model are MSC, decellularized tendon scaffolds, mechanical stimulation by a cyclic strain reactor, and inflammatory stimuli such as co-cultured leukocytes. Decellularized scaffolds function as three-dimensional culture substrates in in vitro research and translational approaches [3– 5]. Furthermore, the application of decellularized tendon tissue is a possibility to reflect natural tissue characteristics through the tendon-specific extracellular matrix. Specific protocols are available to decellularize full-thickness large tendon samples suitable for scaffold production (e.g., the equine superficial digital flexor tendons) [3, 6, 7]. MSC cultured on such decellularized tendon scaffolds and subjected to mechanical stimulation in the form of moderate cyclic stretching have been reported to undergo tenogenic differentiation [8, 9], with detailed protocols described earlier [5]. Yet, this does not acknowledge that tendon disease is often accompanied by inflammation, with elevated levels of inflammatory mediators and leukocyte infiltration in acute disease. To simulate inflammation, different approaches can be used, such as supplementation with cytokines [10, 11] or co-culture with (activated) leukocytes [12–14]. With respect to tendon inflammation, important cytokines are interleukin-1 (IL-1) and tumor necrosis factor-α (TNF-α), which have been shown to impact on tenocytes and stimulate extracellular matrix degradation [15]. However, co-culture with leukocytes may reflect the naturally occurring pathology better than selective supplementation with highly concentrated cytokines. We consider both approaches as justified depending on the experimental question although using both approaches in parallel should yield the most conclusive results.

2

Materials This chapter describes an approach to investigate inflammatory conditions in a 3D dynamic in vitro tendon regeneration model. The protocol is adapted to using equine MSC and equine decellularized superficial digital flexor tendon scaffolds as starting material. The following list gives an overview of the otherwise required materials. In general, use laminar flow cabinet, sterile gloves, and 70% ethanol for sterile working conditions.

2.1

MSC Cultivation

1. Storage medium: Dulbecco’s Modified Eagle Medium low glucose, pyruvate supplemented with 10% fetal bovine serum (FBS), 1% penicillin-streptomycin, 0.1% gentamycin-solution, 10% DMSO.

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2. Cultivation medium: Dulbecco’s Modified Eagle Medium low glucose, pyruvate supplemented with 10% FBS, 1% penicillinstreptomycin, 0.1% gentamycin-solution. 3. Sterile phosphate-buffered saline (PBS): adjust to pH 7.4. 4. 1% Trypsin. 5. Sterile cell culture flasks/dishes. 6. Trypan blue. 7. Hemocytometer. 8. 50 mg/mL gentamycin-solution. 9. 10.000 U/mL penicillin-streptomycin. 2.2 Scaffold Preparation

1. Cryostat (CM 3050S, Leica Biosystems). 2. Tissue Freezing Medium (Leica Biosystems). 3. Microtome blades (Leica Biosystems). 4. Scalpel handle. 5. Scalpel blades no. 22. 6. Forceps. 7. Ultra-low attachment (ULA) cell culture plates/dishes.

2.3 Seeding of Tendon Scaffolds

1. 1% Trypsin. 2. PBS. 3. Trypan blue. 4. Hemocytometer. 5. Sterile cell culture dishes/well plates.

2.4 Dynamic Culture of Tendon Constructs

1. Cyclic strain bioreactor (Institute of Technical Chemistry, Leibniz University, Hannover, Germany): use parameters 1 Hz, 2% strain. 2. Sterile tools: wrench. 3. Sterile surgical tools: forceps.

2.5 Inflammatory Culture Conditions— Leukocyte Recovery and co-Culture

1. Heparinized peripheral blood from a donor horse. 2. 50 mL LeucoSep™ tube. 3. Leuko Spin Medium: 1.090 g/mL density. 4. 5,000 IE/mL Sodium-heparin. 5. Leukocyte washing buffer: Add 0.5% BSA, 2 mM SodiumEDTA to 500 mL PBS. 6. Trypan blue. 7. Hemocytometer. 8. 2.5 μg/mL Concanavalin A cultivation medium. 9. Centrifuge.

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2.6 Inflammatory Culture Conditions— IL-1β and TNF-α Supplementation

1. 10 ng/mL Recombinant Human IL-1β/IL-1F2 (R&D Systems).

2.7 Sample Harvesting

1. PBS adjust to pH 7.4.

2. 50 ng/mL Recombinant Human TNFα (R&D Systems).

2. 2 mL Tubes. 3. 15 mL Tubes.

3

Methods

3.1 MSC Cultivation (See Note 1)

1. Frozen MSC are thawed and cultivated up to 80% confluence.

3.2 Scaffold Preparation (See Note 2)

1. Incubate frozen decellularized tendons for 10 min at 20  C in the cryostat.

2. Culture medium is changed twice a week.

2. Using a scalpel, customize tendon length and width into dimensions suitable for your experiments; for cultivation in a cyclic strain bioreactor, leave the tendons long enough for fixing the scaffold ends with the bioreactor clamps. 3. Using tissue freezing medium, fix the tendon specimen on the object holder of the cryostat. 4. Section the tendon specimens longitudinally into 300 μm-thick tendon extracellular matrix scaffolds using the cryostat. 5. Transfer the frozen scaffolds into a culture dish and transport them to the laminar flow cabinet. 6. Under sterile conditions, place the dissected tendon scaffolds in ULA plates (well plates or dishes). 7. Supplement the scaffolds with a drop of PBS to prevent their drying-out. 8. During a short incubation at 37  C, start with the preparation of MSC.

3.3 Seeding of Tendon Scaffolds (See Note 3)

1. Detach the cells with trypsin and harvest the MSC. 2. Add the cells to the hemocytometer and calculate the cell count. 3. Resuspend the MSC in culture medium (104 cells per μL). 4. Seed the scaffolds using 3  105 MSC in 30 μL per 1 cm2 scaffold surface, by homogeneously distributing the cell suspension on the scaffold. 5. Allow MSC to attach for 6 h at 37  C.

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6. After the incubation period, carefully cover the seeded scaffolds with pre-warmed culture medium. 7. Incubate the seeded scaffolds in a humidified atmosphere at 37  C and 5% CO2 for 3 days before starting the experiments. 3.4 Dynamic Culture of Tendon Constructs (See Note 4)

1. Place and fix the MSC-seeded tendon scaffolds in a suitable cyclic strain bioreactor. 2. Fill the bioreactor chamber with an adequate volume of pre-warmed culture medium. 3. Establish inflammatory culture conditions as described below (see Subheadings 3.5 or 3.6), or continue directly for the controls without inflammatory stimulation. 4. Continue the incubation in a humidified atmosphere at 37  C and 5% CO2. 5. Daily subject the MSC-seeded scaffolds to mechanical stimulation, following a regime consisting of 15 min cyclic stretching, followed by 15 min relaxation and another period of 30 min cyclic stretching (1 Hz, 2% strain, without pre-loading).

3.5 Inflammatory Culture Conditions— Leukocyte Recovery and Co-Culture (See Note 5)

1. Heparinized peripheral blood is collected by venipuncture. 2. Dilute whole blood 1:2 with PBS for leukocyte isolation by density gradient centrifugation. 3. Carefully layer the diluted whole blood (35 mL) on top of Leuko Spin Medium (15 mL); avoid mixing of the phases. 4. Centrifuge at 800  g for 20 min without brakes. 5. Aspirate the upper layer and transfer the cell layer to a new conical tube. 6. Wash the cells with leukocyte washing buffer. 7. Centrifuge at 300  g for 10 min. 8. Determine the number of viable leukocytes using Trypan blue and a hemocytometer. 9. Add the leukocytes to the MSC culture by suspending them in the culture medium to obtain a direct co-culture (10:1). 10. Continue with dynamic culture of the tendon constructs (see Subheading 3.4).

3.6 Inflammatory Culture Conditions— IL-1β and TNF-α Supplementation (See Note 6)

1. Reconstitute the recombinant cytokines. 2. Add the required volume of reconstituted cytokines to the culture medium, e.g., to final concentrations of 10 ng/mL IL-1β and/ or 50 ng/mL TNF-α. 3. Continue with dynamic culture of the tendon constructs (see Subheading 3.4).

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3.7 Sample Harvesting (See Note 7)

4

1. At different time points (e.g., Day 1 and Day 3 after inflammatory conditions were established and cyclic stretching started), samples such as supernatants and seeded scaffolds are harvested, washed in PBS if appropriate and stored appropriately until their analysis.

Notes In general, be focused on sterile working conditions since the experimental setup includes the tissue transfer between its naturally sterile environment and the artificially sterile laboratory environment. 1. Nuclear cells were isolated by collagenase digestion and seeded in cultivation medium. Cells were incubated in a humidified atmosphere at 37  C and 5% CO2, passaged upon subconfluency and then cryopreserved in storage medium [16]. It is advisable to use MSC from an early passage (up to passage 3). 2. Equine distal limbs were collected at a local abattoir, and tendon tissue was resected from the metacarpal regions aseptically. After decellularization by combining freeze-thaw cycles with detergent treatment, decellularized tendons were stored at 80  C before further processing [6, 7]. Note that scaffolds should not thaw during the whole sectioning procedure described. 3. Seeding densities may have to be adapted depending on cell size, viability, and proliferation rate of the MSC used. MSC are allowed to attach for 6 h so they would not be washed off by the addition of culture medium. Cell attachment may occur faster, but 6 h should provide enough safety margin and drying-out was not observed within this period of time. If your experimental design includes monolayer cultures as controls, seed those at a density of 3  103 cells per cm2 at the same time point. 4. The custom-made cyclic strain bioreactor we use allows to fix three technical replicate constructs in the medium chamber, which can be subjected to variable stretching parameters via a 1 kN motor and an integrated software. We recommend to use only mild stretching regimes, such as described above, as we observed increased cell death after longer periods of stretching without adaptation periods, and as higher strains (e.g., 10%, as often reported) do not reflect the in vivo situation during rehabilitation from tendon injury. If your experimental design includes static monolayer or scaffold cultures as controls, perform a medium change and incubate accordingly without mechanical stimulation.

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5. We use allogeneic leukocytes for a mild immunostimulation of MSC and to mimic mild tendon inflammation, respectively. Stronger stimulation is achieved when leukocytes are activated before being used for co-culture, using concanavalin A or phorbol-myristate-acetate and ionomycin as described previously [13]. Yet, we have observed that even non-activated leukocytes impact on the tenogenic properties of MSC. However, leukocyte activation may be advisable when stronger effects are desired, and to activate the immunomodulatory potential of the MSC. For leukocyte recovery, other leukocyte separation media could be used accordingly, but leukocyte fractions obtained should be analyzed prior to the experiments. The separation medium given here is suitable to isolate all major leukocyte subpopulations from equine whole blood. 6. As an alternative to co-culture with leukocytes, direct supplementation with pro-inflammatory cytokines can be performed. The effects of cytokine supplementation should not be expected to be the same as those of leukocytes, but there are important parallels, such as both reduced scleraxis expression [12]. The concentrations of pro-inflammatory cytokines given here correspond to those frequently used in experimental studies and should be useful to observe distinct effects. However, it should be acknowledged that cytokine concentrations in vivo are expected to be much lower, such as 0.01 ng/mL IL-1β and 0.1 ng/mL TNF-α. A further experimental group using such low concentrations may be helpful if you aim to mimic the in vivo environment as closely as possible, yet based on our previous data, effects may vary between MSC from different donors [12]. 7. Short incubation times such as 1 or 3 days are preferable particularly for co-culture studies to mimic the acute phase of inflammation. Note that if cells are cultured for longer periods, the medium should be changed with the renewed supplementation with cytokines or leukocytes from the same donor. References 1. Lui PPY (2015) Stem cell technology for tendon regeneration: current status, challenges, and future research directions. Stem Cells Cloning 8:163–174. https://doi.org/10. 2147/SCCAA.S60832 2. Manning CN, Martel C, Sakiyama-Elbert SE et al (2015) Adipose-derived mesenchymal stromal cells modulate tendon fibroblast responses to macrophage-induced inflammation in vitro. Stem Cell Res Ther 6:74. https://doi.org/10. 1186/s13287-015-0059-4

3. Burk J, Erbe I, Berner D et al (2014) Freeze-thaw cycles enhance decellularization of large tendons. Tissue Eng Part C Methods 20(4):276–284. https://doi.org/10.1089/ten.TEC.2012.0760 4. Cheng CW, Solorio LD, Alsberg E (2014) Decellularized tissue and cell-derived extracellular matrices as scaffolds for orthopaedic tissue engineering. Biotechnol Adv 32(2):462–484. https://doi.org/10.1016/j.biotechadv.2013. 12.012

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5. Youngstrom DW, Barrett JG (2016) Tendon differentiation on decellularized extracellular matrix under cyclic loading. Methods Mol Biol 1502:195–202. https://doi.org/10. 1007/7651_2016_332 6. Roth SP, Glauche SM, Plenge A et al (2017) Automated freeze-thaw cycles for decellularization of tendon tissue—a pilot study. BMC Biotechnol 17(1):13. https://doi.org/10.1186/ s12896-017-0329-6 7. Roth SP, Erbe I, Burk J (2018) Decellularization of large tendon specimens: combination of manually performed freeze-thaw cycles and detergent treatment. Methods Mol Biol 1577:227–237. https://doi.org/10.1007/ 7651_2017_49 8. Youngstrom DW, Rajpar I, Kaplan DL et al (2015) A bioreactor system for in vitro tendon differentiation and tendon tissue engineering. J Orthop Res 33(6):911–918. https://doi.org/ 10.1002/jor.22848 9. Burk J, Plenge A, Brehm W et al (2016) Induction of Tenogenic differentiation mediated by extracellular tendon matrix and short-term cyclic stretching. Stem Cells Int 2016:7342379. https://doi.org/10.1155/ 2016/7342379 10. Han P, Cui Q, Yang S et al (2017) Tumor necrosis factor-α and transforming growth factor-β1 facilitate differentiation and proliferation of tendon-derived stem cells in vitro. Biotechnol Lett 39(5):711–719. https://doi. org/10.1007/s10529-017-2296-3 11. Zhang K, Asai S, Yu B et al (2015) IL-1β irreversibly inhibits tenogenic differentiation

and alters metabolism in injured tendonderived progenitor cells in vitro. Biochem Biophys Res Commun 463(4):667–672. https:// doi.org/10.1016/j.bbrc.2015.05.122 12. Brandt L, Schubert S, Scheibe P et al (2018) Tenogenic properties of Mesenchymal progenitor cells are compromised in an inflammatory environment. Int J Mol Sci 19(9). https://doi. org/10.3390/ijms19092549 13. Hillmann A, Paebst F, Brehm W et al (2019) A novel direct co-culture assay analyzed by multicolor flow cytometry reveals context- and cell type-specific immunomodulatory effects of equine mesenchymal stromal cells. PLoS One 14(6):e0218949. https://doi.org/10.1371/ journal.pone.0218949 14. Cassano JM, Schnabel LV, Goodale MB et al (2018) Inflammatory licensed equine MSCs are chondroprotective and exhibit enhanced immunomodulation in an inflammatory environment. Stem Cell Res Ther 9(1):82. https:// doi.org/10.1186/s13287-018-0840-2 15. Schulze-Tanzil G, Al-Sadi O, Wiegand E et al (2011) The role of pro-inflammatory and immunoregulatory cytokines in tendon healing and rupture: new insights. Scand J Med Sci Sports 21(3):337–351. https://doi.org/10. 1111/j.1600-0838.2010.01265.x 16. Burk J, Ribitsch I, Gittel C et al (2013) Growth and differentiation characteristics of equine mesenchymal stromal cells derived from different sources. Vet J 195(1):98–106. https://doi. org/10.1016/j.tvjl.2012.06.004

Chapter 13 Generation of Epidermal Equivalents from Hair Follicle Melanocytes, Keratinocytes, and Dermal Fibroblasts Vuk Savkovic, Marie Schneider, Hanluo Li, Jan-Christoph Simon, Mirjana Ziemer, and Bernd Lethaus Abstract Bench-to-bedside axis of therapeutic product development is currently being oriented towards minimum invasiveness on both ends—not only clinical application but harvesting of the starting biological material as well. This is particularly relevant for Advanced Therapy Medicinal Products and their specific legislative requirements, even more so in skin regeneration. It is precisely the skin equivalents and grafts that benefit from the minimum-to-noninvasive approach to a noteworthy extent, taking in account the sensitive nature of both skin harvesting and grafting. This chapter includes protocols for two separate steps of generating skin equivalent from the cells cultured from hair follicle outer root sheath. The first step is a non-pigmented epidermal equivalent generated from human keratinocytes from the outer root sheath named non-pigmented epidermal graft. The second step consists of co-cultivating human keratinocytes and human melanocytes from the outer root sheath, hereby producing a pigmented epidermal graft. Key words Epidermal equivalent, Skin equivalent, Hair follicle outer root sheath, Non-pigmented epidermal graft (NP-EPI), Pigmented epidermal graft (P-EPI), Human keratinocytes from the outer root sheath (HUKORS), Melanocytes from the outer root sheath (HUMORS)

1

Introduction We are about to describe the stepping stones for a noninvasively based ex vivo/in vitro production of an epidermal equivalent that can be used both as a skin graft and as an in vitro model. This particular approach of low-to-none invasive producing and applying of Advanced Therapy Medicinal Products (ATMP) or the in vitro toxicological tests has become a major prerequisite in their design and application [1, 2] . This concept applies to skin regeneration to a large extent [1, 3]. To this purpose, Human Keratinocytes from the Outer Root Sheath of human hair follicle (HUKORS) have been threedimensionally cultured to generate a Non-Pigmented Epidermal

Peggy Stock and Bruno Christ (eds.), In Vitro Models for Stem Cell Therapy: Methods and Protocols, Methods in Molecular Biology, vol. 2269, https://doi.org/10.1007/978-1-0716-1225-5_13, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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graft (NP-EPI) and co-cultured with Human Melanocytes of the same origin (HUMORS) to construct a Pigmented Epidermal graft (P-EPI). Both NP-EPI and P-EPI reproducibly display correct anatomic features of the epidermis and basic functions such as asymmetric division, differentiation, melanin production and distribution. Clear advantage of the NP-EPI and P-EPI as a test model is that they utilize stem cells from the hair follicle outer root sheath, which differentiate in the course of the applied procedures. Herewith, not only the direct impact of tested agents on melanocytes and keratinocytes is being detected, but also an impact on their development from the stage of an ectodermal stem cell or a precursor, over the stage of basal keratinocytes until the endpoint. The latter reflects the impact of tested agents on young or regenerative epidermis. Therefore, the epidermal equivalents described in this Chapter are both toxicological and developmental models. The protocol consists of four parts. Culturing HUKORS and HUMORS from the hair follicle is shown in two separate procedures, since both cultures have their particularities, even though they rely on the same principle of air-liquid-interface cultivation. Third part of the protocol displays how the HUKORS are engineered into a non-pigmented epidermal equivalent. Fourth part describes incorporation of HUMORS into a NP-EPI and shows generation of a pigmented epidermal equivalent P-EPI. At the end of the protocols the characterization procedures are supplied.

2 2.1

Materials Equipment

1. Forceps. 2. Scissors. 3. Ø 3.5; 6 cm Petri dish. 4. Parafilm. 5. 6- and 12-well culture plates. 6. Transwell inserts fitting 6- and 12-well culture plates. 7. Cryo vials. 8. T25; T175 culture flask. 9. Centrifuge. 10. Safety cabinet with vacuum unit. 11. Cell culture incubator, normoxic and hypoxic conditions. 12. Pasteur pipettes. 13. Cell counting chamber. 14. Inverted microscope with documentation unit. 15. Fluorescence microscope with software. 16. Binocular/Stereomicroscope.

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17. Scalpel. 18. Microtome including water bath unit for stretching. 19. Paraffin embedding unit. 20. 15 and 50 mL tubes. 21. Ø 70 μm, white cell strainer. 22.
80  C;
20  C Freezer. 23. Refrigerator. 24. Nunc Lab-Tek Chamber Slides. 25. Cover slips. 26. Pressure cooker. 27. Cuvettes. 28. Glass slides. 29. Water bath. 30. Thermomixer. 31. Isopropanol bath (Mr. Frosty™ Freezing Container). 32. Liquid nitrogen storage container. 33. Multiscan spectrum plate reader. 34. 96-well flat bottom plate. 2.2 Media and Buffers

1. Trypsin/EDTA (0.25, 0.04% (w/v)/ 0.03% (w/v)). 2. PBS wCa2+/Mg2+. 3. Stock Medium for K-, K0, and F-Medium: 3 parts of Dulbecco’s Modified Eagle’s Medium (DMEM) and 1 part of HAM’s F12 Medium (HAM’s F12), 2.73% 1 M HEPES, 0.0207 mg/ mL adenine, 0.0057 mg/mL insulin, 3.0  10
6 mg/mL triiodothyronine, 0.01 mg/mL hydrocortisone. The medium should be kept at +2 to +8  C for up to 28 days. 4. Keratinocyte-Medium with antibiotics for primary culture (K-Medium): for 250 mL of k-Medium add to 219.5 mL of Stock medium 25 mL of human serum, 2.5 mL Amphotericin B working solution, 2.5 mL of 100 Penicillin/Streptomycin, 250 μL cholera toxin working solution, and 25 μL of EGF working solution. The medium should be kept at +2 to +8  C for up to 21 days. 5. Keratinocyte-Medium without antibiotics for secondary culture (K0-Medium): for 250 mL K0-Medium add to 219.5 mL Stock medium 25 mL of human serum, 7.5 mL of DMEM, 2.5 mL of HAM’s F12, 250L of cholera toxin working solution and 25 μL of EGF working solution. The medium should be kept at +2 to +8  C for up to 21 days. 6. Fibroblast-Medium without antibiotics (F-Medium): for 100 mL F-Medium, add to 87.8 mL Stock medium 5 mL of

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human serum, 5.2 mL of DMEM, 1.8 mL HAM’s F12, 0.1 mL of cholera toxin working solution, and 10 μL EGF working solution. The medium should be kept at +2 to +8  C for up to 21 days. 7. Fibroblast-Freezing-Medium (FE-Medium): for 20 mL FE-Medium add to 17 mL F-Medium 2 mL of DMSO and 1 mL of human serum. The medium should be kept at +2 to +8  C for up to 21 days. 8. Keratinocyte-Freezing Medium (KE-Medium): for 25 mL KE-Medium add to 22.5 mL K0-Medium 2.5 mL of DMSO. The medium should be kept at +2 to +8  C for up to 21 days. 9. Washing Medium for hair follicles (W-Medium): for 50 mL W-Medium add to 44.75 mL DMEM 1.25 mL of 1 M HEPES, 2 mL of Amphotericin B working solution, and 2 mL of 100 Penicillin/Streptomycin. The medium should be kept at +2 to +8  C for up to 28 days. 10. Transport Medium for hair follicles with antibiotics (T-Medium): for 100 mL of T-Medium add to 85 mL DMEM 10 mL human serum, 3 mL 1 M HEPES, 1 mL Amphotericin B working solution, and 1 mL of 100 Penicillin/Streptomycin. The medium should be kept at +2 to +8  C for up to 28 days (see Note 1). 11. DermaLife-M-Medium (DLM(
)). 12. 0.1 mg/mL Mitomycin stock solution: 2 mg Mitomycin C add 20 mL PBS wCa2+/Mg2+. Aliquot 1.7 mL per cryo vial (sufficient for 11 times use), store at
20  C for up to 6 months. 13. 10 mM sodium citrate buffer. Adjust to pH 6.0. 14. 0.004237 mg/mL Cholera toxin working solution. 15. 0.1 mg/mL Epidermal growth factor (EGF) working solution. 16. 10 μg/mL Amphotericin B working solution. 17. 50 ng/μL Geneticin solution G--418 Sulfate in DLM. 18. 5 mg/mL Collagenase V in PBS. 19. HEPES buffer. 20. 0.1 M PBS: 0.1 M PO4-buffer adjust to pH 7.2, add 120 mM NaCl. 21. Trypsin neutralizing solution (TNS). 2.3 Cells, Reagents, and Antibodies

1. Cryopreserved fibroblasts: normal human dermal fibroblasts (NHDF). 2. Normal human epidermal keratinocytes (NHEK). 3. Normal human epidermal melanocytes (NHEM). 4. Mycoplasma PCR detection kit.

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5. 1:1 mix of Methanol and Acetone. 6. Tap water. 7. Gelatine. 8. 0.1% saponine-PBS. 9. 2% bovine serum albumin (BSA) in PBS. 10. 40 ,6-diamidino-2-phenylindole (DAPI). 11. 1 M Sodium hydroxide. 12. 0; 3.125; 6.25; 12.5; 25; 50; 100 μg/mL Synthetic melanin standard curve. 13. 10 mg/mL L-DOPA in PBS. 14. Nuclear Fast Red. 15. 4% buffered formalin. 16. Hematoxilin. 17. 1% Eosin. 18. 5% silver nitrate. 19. Xylol. 20. 100%; 96%; 80%; 70%; 50% Ethanol (EtOH) deparaffinization/rehydration gradient series. 21. Aqua dest. 22. Fluoromount-G™ Mounting Medium. 23. Leicamount mounting solution. 24. Rabbit polyclonal anti-KRT5 antibody. Dilute 1:100. 25. Mouse monoclonal anti-PMEL antibody. Dilute 1:100. 26. Goat anti-mouse Alexa Fluor 594 secondary antibody. Dilute 1:300. 27. Goat anti-rabbit R-Phycoerythrin secondary antibody. Dilute 1:50. 28. Rabbit polyclonal anti-KRT10 antibody. Dilute 1:100. 29. Mouse monoclonal anti-IVL. Dilute 1:200. 30. 5% Sodium-thiosulfate solution. 31. 0.2% Gold chloride solution. 32. Ammonia solution.

3

Methods

3.1 Protocol for the Feeder Layer

At least 12 days ahead of the follicle preparation d(
12), 11 days ahead of hair plucking 1. Thaw the cryopreserved fibroblasts. 2. Mix with 10 mL F-Medium.

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3. Centrifuge for 10 min, 190  g, at room temperature (RT). 4. Aspirate the supernatant. 5. Resuspend the pellet. 6. Count the cells by using a counting chamber. 7. Set the cell suspension to 1  106 cells/mL. 8. Seed the cells in T175 flask and add 25 mL F-Medium. 9. Cultivate for 1 week at 37  C, 5% CO2. Approximately 10  106 fibroblasts per T175 flask can be expected as a yield of the culture. 10. After a week of culture, inactivate the fibroblasts: 11. Harvest the fibroblasts by trypsin/EDTA for 5 min. Prolong to 10 min if needed. 12. Stop the trypsinization by adding 10 mL of F-Medium. 13. Collect and resuspend the cells. 14. Centrifuge for 10 min, 190  g, at RT. 15. Aspirate the supernatant and resuspend the pellet. 16. Count the fibroblasts. 17. Set the cell suspension to 3–4  106 cells /mL. 18. Seed the cells into a T175 flask and fill up with 8.4 mL of F-Medium. 19. Incubate for 2 days at 37  C, 5% CO2. 20. On day 3, add 1.7 mL of the 1 mg/mL Mitomycin stock solution (working conc. 0.1 mg/mL). 21. Incubate 3–5 h at 37  C, 5% CO2. 22. Wash the cells three times with PBS. 23. Add 20–25 mL F-Medium and culture for 1–2 days at 37  C, 5% CO2. 2 days ahead of follicle preparation d(
2), 1 day ahead of hair plucking 24. Detach the inactivated fibroblasts by using trypsin/EDTA for 5 min. Prolong to 10 min if needed. Either cryopreserve or use fresh by seeding fibroblasts on the lower side of the insert (see Fig. 1h). Once the inserts with a feeder layer are in place, you can begin with the primary culture of HUKORS. 3.2 Primary Culture of Human Keratinocytes from the Outer Root Sheath (HUKORS)

Hair follicles are plucked, washed, and cultivated in air-mediuminterface set up by the means of suspended inserts with a support network. As the cells migrate out of the follicle, they populate the support network and they are harvested at near confluence [2– 5]. From this point on, HUKORS can be either freshly used or

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Fig. 1 Primary HUKORS and HUMORS cultures from the outer root sheath of hair follicle. (a) Hair sampling. (b) Plucked anagen hair follicle. (c) Insert with support mesh. (d) follicle in the air-medium-interface setup. Scale bar corresponds to 500 μm. (e) Migration of cells from the follicle onto the mesh. Scale bar corresponds to 500 μm. (f) 2D culture of HUKORS. (g) 2D culture of HUMORS. (h) Feeder layer, Normal Human Dermal Fibroblasts (NHDF). (f–h) Scale bar corresponds to 100 μm

cryopreserved. Both cryopreserved and freshly cultured HUKORS can be used for generating epidermal equivalents with equal efficiency. 3.2.1 Preparations 2 Days Ahead of Sampling Follicles d(
2)

Feeder layer 1. Place the Transwell inserts upside down in the 20 cm Petri dish in laminar flow cabinet. 2. Seed 1.0 mL of the inactivated fibroblast cell suspension (0.2  106 cells/mL) to the back (lower) side of a Transwell insert by slowly pipetting. 3. Leave in the incubator for 24 h to adhere. 4. Prepare 1  6-well plate with 5–6 Transwell inserts with a feeder layer of inactivated fibroblasts per proband. HUKORS: primary culture

3.2.2 Hair Plucking (See Fig. 1)

2 days ahead of follicle preparation d(
2): written documentation Make sure that your written consent is prepared and that it contains the correct data of the particular donor. Presumably, you have already obtained a permission of the local Ethical Committee to sample hairs as human material. The template of the written donor’s consent would be an integral part of your permission proposal. 1 day ahead of follicle preparation d(
1): hair plucking

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Fig. 2 Work flow of HUKORS 3D cultures for generating NP-EPI

1. Pluck 30–40 anagen hairs from the temporal area of healthy donors of age 18–50 with unwashed hair (see Note 2 and Fig. 1a). 2. Firmly pinch a minimal number of hairs (1–3) as close to the skin as possible without pinching the skin (see Note 3 and Fig. 1b). 3. Store the plucked follicles for 24 h in T-Medium in an Ø6 cm Petri dish sealed with Parafilm. The follicles can be held in the T-Medium for up to 96 h if needed. 3.2.3 Follicle Preparation, HUKORS Cultivation and Harvest (See Fig. 2)

d0: Follicles on cell strainer, washing, placing onto the prepared inserts; photo-documentation. 1. Shorten the hairs at the distal side by the means of binoculars, scalpel, and forceps. 2. Place the shortened hairs with a follicle in a cell strainer. 3. Dip in an Ø3.5 cm Petri dish containing 3.5 mL K-Medium. 4. Transfer the Petri dish with the cell strainer and the follicles into the laminar flow hood. From this point on, you should be working sterile. 5. In order to remove the resident flora, a series of four washing steps is necessary: Prepare the four washing positions before plucking. 6. For each step, 3.5 mL freshly made W-Medium in an Ø3.5 cm Petri dish is used. 7. Submerge cell strainer into the W-Medium 15 times in a course of 60 s. 8. Remove the cell strainer from the W1 (washing point 1) to the W2 and repeat the procedure. 9. Repeat the washing procedure in W2, W3, and W4. 10. Place 3.5 mL of K-Medium to the bottom of a 6-well plate. 11. Submerge the cell strainer with washed follicles in the K-Medium. 12. Prepare the Transwell insert with a feeder layer by placing 1.5 mL F-Medium to the cavity of the 6-well, then placing the insert into the 6-well (see Note 4).

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13. Replace the F-Medium with 1.5 mL K-Medium. 14. Place 5–6 of the washed follicles onto the network of the insert (see Note 5 and Fig. 1c). 15. Avoid desiccation; the follicles should be kept wet at all times until the lift phase. 16. Document by taking images at 4 and 10 magnification (scale bar 500 and 200 μm, respectively, see Fig. 1d). 17. Cultivate at 37  C in normoxic conditions with 5% CO2 for the next 5 days. d5: photo-documentation (visible migration, see Fig. 1e). 18. Repeat photo-documentation at 4 and 10 magnification (scale bar 500 and 200 μm, respectively) d7: medium change, photo-documentation. 19. Change the K-Medium in the lower compartment. 20. Photo-document at 4 and 10 magnification (scale bar 500 and 200 μm, respectively). d10: medium change, flooding. 21. Change the K-Medium in the lower compartment. 22. Photo-document at 4 and 10 magnification (scale bar 500 and 200 μm, respectively). 23. Flood the follicles by adding 1 mL of K-Medium to the upper compartment of the Transwell insert. The follicles will now be covered with a layer of K-Medium. d12: medium change in both the upper and the lower cavity. 24. Aspirate the K-medium from the lower cavity by approaching the lower cavity with the Pasteur pipette through the openings in the sleeve of the Transwell insert. 25. Subsequently aspirate the K-Medium from the upper cavity. 26. Add 1.5 mL of fresh K-Medium to the lower cavity and to the upper cavity 1 mL. d14: medium change in both the upper and the lower cavity, photo-documentation. 27. Replace K-Medium in both cavities with fresh K-Medium. 28. Photo-document at 4 and 10 magnification (see Fig. 1f, scale bar 500 and 200 μm, respectively). d17: Endpoint of primary culture: Harvesting, sampling for mycoplasma test. HUKORS are now ready to be harvested and either freshly used, or cryopreserved (see Note 6).

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29. Sample the medium from both the upper and the lower cavity of all wells, pool and separate into 3  15 mL centrifuge falcons for Mycoplasma-Test according to the instruction sheet of the kit used. 30. Aspirate the rest of the medium. 31. Wash the insert on both sides twice with 2 mL PBS and subsequently aspirate. 32. Wash the upper side of the insert with 2 mL PBS. 33. Incubate for 5 min and aspirate. 34. Pipette 450 μL Trypsin/EDTA onto the insert. 35. Incubate at 37  C for 15 min. 36. Control the degree of detaching and prolong the incubation time up until 25 min if required. 37. Stop the trypsinization by adding 2 mL of K-Medium to the insert. 38. Prepare a 50 mL centrifuge tube with a cell strainer. 39. Collect cells from all inserts with a pipette. 40. Transfer the cells through the cell strainer into the collection tube. 41. Wash each insert with 2 mL K-Medium, resuspend by pipetting up and down and pass the residual cell suspension through the cell strainer into the collection tube. Repeat until the cells are removed from the insert. 42. Centrifuge the 50 mL tubes with cells two times, with a washing step in K-Medium in between, at 220  g, for 4 min at RT, set speed to 9 and brake to 1. 43. Aspirate the supernatant. 44. Resuspend the pellet in 5–7 mL K0-Medium, depending on the pellet size. 45. Count the cells. 46. Set the cell suspension to 1  106 cells/mL (see Note 7). 47. Centrifuge the cells at 220  g, for 4 min at RT, set speed to 9 and brake to 1. 48. Aspirate the supernatant. At this point, you can either directly use or store your HUKORS. Immediate use will save time needed to generate epidermal equivalents in a secondary 3D culture, whereas cryopreservation (see Subheading 3.2.4) will help synchronize subsequent steps, particularly those of co-culturing HUKORS with HUMORS, more easily.

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1. Add 1.8 mL of KE-Medium to the pelleted cells. 2. Resuspend the HUKORS and divide into 3–5 cryo vials to reach approximately 2  106 cells/vial. 3. Place the cryo vials into the isopropanol bath (Mr. Frosty™ Freezing Container), keep at
80  C for 24 h and store at
150  C subsequently.

3.3 Secondary 3D Culture: Non-Pigmented Epidermal Model Consisted of HUKORS (NP-EPI)

3.3.1 Preparation of Inserts with Feeder Layer d(
1)

HUKORS are in the state of basal keratinocyte at the point of harvesting. Freezing and thawing do not affect their ability to proliferate symmetrically and asymmetrically, forming the stratum basale and differentiating into the upper strata—stratum spinosum, stratum granulosum finalized with stratum corneum. This process is corroborated with nutrient and liquid supply from the lower side of the insert, mediation by fibroblasts from the feeder layer and exposure to air from the upper side by reducing the amount of liquid. The latter mimics the native conditions of the outer limit of epidermis (see Fig. 2). 1. 1 day before the 3D culture start prepare 1  12-well plate per proband with eight 12-well Transwell inserts coated with inactivated fibroblasts as a feeder layer: 2. Seed 0.3 mL of the cell suspension containing 0.4  106 cells/ mL inactivated fibroblasts onto the back/lower side of the middle size inserts. See Subheading 3.1 and 3.2.1 steps 1–3 for preparing the inserts with a feeder layer. 3. Prepare fresh media.

3.3.2 NP-EPI Culture

d0: thawing, seeding, photo-documenting. 1. Thaw HUKORS by using pre-warmed K0-Medium. 2. Warm 10 mL of K0-Medium in a 15 mL centrifuge tube. 3. Thaw the cryo vial by dropwise adding warm medium to the ice core. As soon as thawed, immediately transfer the entire content to the 15 mL tube and mix with the pre-warmed K0-Medium. 4. Centrifuge the thawed cell suspension at 220  g, for 4 min at RT, set speed to 9 and brake to 1. 5. Aspirate the supernatant. 6. Resuspend the pellet in 5–7 mL of K0-Medium, depending on the pellet size. 7. Count the cells. 8. Set the cell suspension concentration to 0.7  106 cells/mL. 9. Change the medium in the lower cavity of the prepared 12-well plate by aspirating the F-Medium and replacing it with 1 mL K0-Medium.

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10. Pipette 0.5 mL of the cell suspension into the prepared inserts with a feeder layer (see Subheading 3.1 and 3.2.1 steps 1–3). 11. Photo-document by taking images at 4 and 10 magnification (scale bar 500 and 200 μm, respectively). 12. Cultivate at 37  C, normoxic conditions with 5% CO2 for 48 h. d2: lift, change medium from the lower cavity, photo-document. 13. Lifting: carefully aspirate K0-Medium from the upper cavity. Epidermal equivalents are now exposed to air and should be kept this way as a prerequisite for keratinocyte differentiation and stratification. 14. Medium change: aspirate used medium from the lower cavity with a Pasteur pipette. 15. Add 1 mL K0-Medium to the lower cavity (see Note 8). 16. d5: Change the medium in the lower cavity. 17. Photo-document by taking images at 4 and 10 magnification (scale bar 500 and 200 μm, respectively) 18. d6: Aspirate the medium from the lower cavity and add 1 mL K0-Medium. If the epidermal equivalent is still covered by a wet film of liquid, reduce the amount of K-Medium to 0.7 mL. 19. Photo-document by taking images at 4 and 10 magnification (scale bar 500 and 200 μm, respectively) 20. d7: Change K-Medium in the lower cavity and replace it with fresh 1 mL K-Medium (see Note 9). 21. d8: Photo-document by taking images at 4 and 10 magnification (scale bar 500 μm and 200 μm, respectively) 22. d9: Change K-Medium in the lower cavity and replace it with fresh 1 mL K-Medium (see Note 9). 23. d12: Change K-Medium in the lower cavity and replace it with fresh 1 mL K-Medium (see Note 9). 24. d13: Photo-document by taking images at 4 and 10 magnification (scale bar 500 μm and 200 μm, respectively) 25. d14: Change K-Medium in the lower cavity and replace it with fresh 1 mL K-Medium (see Note 9). d15: endpoint non-pigmented epidermal equivalent. The NP-EPI should be fully stratified with a visible stratum corneum at top view as an indicator of the endpoint NP-EPI. See the characterization of NP-EPI 3D culture for detailed analysis (see Subheading 3.6).

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3.4 Primary Culture of Human Melanocytes from the Outer Root Sheath (HUMORS) 3.4.1 Procedure of Generating HUMORS

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The culturing of HUMORS is based on the same sampling procedure and air-medium-interface platform as HUKORS [3–5]. We will therefore refer to those steps only shortly and address the specific steps relevant to the HUMORS culture. For a more detailed protocol, please see [5] printed in an earlier edition of Methods in Molecular Biology 1210. A condense protocol for culture of HUMORS from plucked hair follicles is presented in further text. The time points in the protocol are the optimal ones and this time frame may deviate. d0 sample hair follicles (see Fig. 1a, b,c). 1. Pluck 30–60 hairs according to the procedure described in Subheading 3.2.2. 2. Cut off the proximal follicle end using stereomicroscope, fine forceps and a scalpel, in order to reduce a potential carry over of dermal fibroblasts. 3. Wash the follicles repeatedly in W-Medium. 4. Wash once with 2 mL PBS. 5. Incubate the washed follicles with Collagenase V for 10 min at 37  C. The remaining mid- and the semi-distal part of hair follicle are partially proteolized by digestion in Collagenase V. Proteolysis of collagen lowers contact inhibition within the Outer Root Sheath (ORS) and the ORS cells can migrate more easily. 6. Wash once with 2 mL PBS. 7. Place 10 follicles on the mesh of a 6-well Transwell insert and add 1 mL fresh DLM medium to the bottom. The shortened follicles are cultivated in the air–liquid interface. 8. Photo-document at 4 magnification. 9. Incubate at 37  C, 5% O2, 5% CO2 for 7 days. d7 change DLM medium, photo-document. 10. Observe and photo-document the growth of the ORS every 2 days. After no longer than 3 weeks incubation (4 weeks post-biopsy), migrating cells should be visible on the mesh outwards the ORS, forming a rough monolayer surrounding the hair follicle. 11. Change DLM medium every 2 days. d14 Expanding the primary culture of hair root ORS, flooding. 12. Aspirate the DLM medium from the lower cavity of Transwell insert. 13. Add 2 mL DLM medium to completely cover the cells.

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14. Add 0.5 mL of the DLM medium to the upper side of the mesh insert every 2 days. The cells should proliferate until near confluency. 15. Photo-document once a week. In case of rapid growth, document more frequently. 16. Further cultivate in DLM on polystyrene cell culture plastic will result in differentiated, partially melanotic melanocytes within the next five passages in DLM. d30 Starting the 2D primary culture of HUMORS, trypsinization. Confluent cells are trypsinized, transferred to cell culture flasks, and further cultivated as 2D culture in DLM in hypoxic conditions. The time point of subculture start is marked as the first passage (p1). The cells are further passaged upon reaching 70% confluence of the culture flask, roughly once a week. Doubling rates are donorspecific and may vary. 17. Photo-document at the point of near confluence (not later than 6 weeks post-biopsy). 18. Aspirate the medium. 19. Wash the cells once with HEPES buffer. 20. Aspirate the buffer. 21. Add 1 mL Trypsin. 22. Keep at 37  C for 4 min. 23. Add 1 mL of TNS. 24. Pipette up and down until the cells are detached and form a single-cell suspension. 25. Transfer to a 15 mL Falcon tube and add 5 mL of PBS. 26. Centrifuge at 220  g for 5 min. 27. Resuspend the pellet in 5 mL of DLM medium. 28. Transfer the suspension to the T25 cell culture flask. 29. Cultivate in DLM at 37  C with 5% O2; 5% CO2. 30. Change the medium every 2 days. Maintaining the primary 2D culture of HUMORS (see Fig. 1g). 31. Observe the cells every 2 days for adherence, morphology, and confluence. Photo-document all changes. 32. At the 80% confluence, passage the cells (see steps 17–28). 33. Change the DLM medium every 2 days. Selecting the HUMORS from the primary culture of ORS cells: Geneticin selection, differential trypsinization.

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Residual fibroblasts or keratinocytes should be removed by the means of Geneticin selection and differential trypsinization. The translation interference of the Geneticin affects more actively and quickly proliferating fibroblasts and keratinocytes. Short tripsinization does not affect keratinocytes and fibroblasts, and it releases melanocytes in the early fraction of detached cells. Upon an outgrowth of cells, the cultures are “flooded” and allowed to reach near confluence (90% of the Transwell surface). Near confluence occurs after up to 2 weeks of the flooded culture. 1. Observe and photo-document the cells every 2 days for adherence, morphology, and confluence. 2. At near confluence, treat with Geneticin in DLM medium for 24 h in case of rapid proliferation of all cells (doubling time less than 1 week) and for 48 h by normal proliferation rate (doubling time 1 week). 3. After the Geneticin treatment, leave the cells to recover not shorter than 24 h in DLM medium. The surviving cells should majorly be melanocytes. Cultivate until 80% confluence. 4. In case of persistent presence of keratinocytes or fibroblasts, apply differential trypsinization with a duration of 3 min instead of 4 min (see steps 17–28). The melanocytes are less adherent than fibroblasts and keratinocytes and greater part of melanocytes will detach during the shorter exposure. Collect this first fraction of detached cells and cultivate them further. 5. Apply 4 min trypsinization further by each passage. If fibroblasts or keratinocytes reappear in culture, repeat the 3 minlong differential trypsinization. 3.4.2 Characterizing the HUMORS in the ORS Cell Culture

The cells should be characterized for melanocyte morphology and marker expression characteristic for amelanotic and melanotic melanocytes. From the P3 on, the cells are being tested for purity and characterized on the basis of morphology, marker expression, enzymatic function (L-DOPA conversion), and melanin content. The cells that exhibit PMEL expression in cell soma and in the dendrites (see Subheading 3.6.4 and Fig. 5) as well as melanin presence are regarded as functional. This stage is usually reached by the eighth week post-biopsy. Ripe melanotic melanocyte status can be determined by the enzymatic activity of DOPA oxidase through quantifying L-DOPA conversion to melanin, and by quantifying melanin content of the cells. Melanin production quantification 1. Detach the cells with 1 mL Trypsin/EDTA solution (see Subheading 3.4.1 steps 17–25). 2. Incubate at 37  C for 4 min.

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3. Count the cells. 4. Pellet the cell suspension by centrifugation with 5 mL of PBS. 5. Resuspend the cell pellet in 150 μL of 1 M NaOH. 6. Keep at 60  C for 3 h. 7. Transfer 300 μL of lysate to a flat bottom 96-well plate. 8. Set up an array of dilutions for melanin standard curve (see Subheading 2.3 step 12). 9. Measure the optical density/extinction at 475 nm wavelength of all samples using plate reader. 10. Determine melanin concentration by comparing the extinction values with those of the melanin standard curve. 11. Express melanin content as melanin μg/culture, or ng/cell. L-DOPA

Reaction

1. Culture melanocytes for 72 h on glass slides coated with gelatin. 2. Wash cells twice with 0.1 M PBS (see Subheading 2.2 step 22). 3. Fix the cells for 30 s in ice-cold acetone/methanol (1:1) at
20  C. 4. Wash the cells twice with 0.1 M PBS. 5. Freshly prepare L-DOPA in PBS (see Subheading 2.3 step 13). 6. Incubate the cells for 6 h at 37  C. 7. Exchange DOPA solution three times, each after 1.5–2 h incubation. 8. Wash cells twice with 0.1 M PBS. 9. Incubate cells with Nuclear Fast Red for 10 min. 10. Rinse cells 2–3 min with water, air-dry the slide. 11. Mount with Fluoromount™. 3.5 Secondary 3D Culture Pigmented Epidermal Model Consisted of HUKORS and HUMORS (P-EPI)

To generate a pigmented epidermal equivalent, HUMORS are seeded and then co-cultured with HUKORS to form the stratum basale and further strata, much like in non-pigmented equivalents, until stratum corneum is formed. Within this 3D construct, HUKORS and HUMORS will interface, forming structures analogous to melanin epidermal units, enabling distribution of melanin within the epidermal equivalent [6, 7]. Pigmented epidermal equivalents can be engineered as autologous (HUMORS and HUKORS from the follicles of the same donor) or heterologous (from different donors) and both variants are functional in vitro (see Fig. 3).

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Fig. 3 Work flow of HUMORS and HUKORS 3D cultures for generating P-EPI 3.5.1 Preparation of Inserts with Feeder Layer

1. 1–2 days before the 3D culture starts prepare 1  12-well plate per donor with four pieces of 12-well Transwell inserts coated with inactivated fibroblasts as a feeder layer. 2. Seed 0.3 mL of the cell suspension containing 0.4  106 cells/ mL inactivated fibroblasts onto the lower side of the middle size inserts. See Subheadings 3.1 and 3.2.1 steps 1–3 for preparing the inserts with a feeder layer. 3. Prepare fresh media.

3.5.2 P-EPI Culture

d(
2): coating the inserts with HUMORS. 1. Thaw cryopreserved HUMORS in passage 9–12 or harvest them by trypsin from the 2D culture (see Fig. 1g). 2. Count the cells. 3. Set the HUMORS suspension to 0.07  106 cells/mL. 4. Add 0.5 mL of the cell suspension to the insert. 5. Pipette 1 mL DLM(
) to the lower cavity. 6. Cultivate at 37  C in hypoxic conditions 5% O2 and 5% CO2. d(
1): DLM medium change, photo-documentation inserts. 7. Aspirate DLM medium from the upper and the lower cavity. 8. Add 1 mL of 1:1 DLM:K0 Medium to the lower cavity. 9. Photo-document by taking images at 4 and 10 magnification (scale bar 500 and 200 μm, respectively). d0: thawing documentation.

HUKORS,

seeding

into

inserts;

photo-

10. Thaw HUKORS by using pre-warmed K0-Medium. Warm 10 mL of K0-Medium in a 15 mL centrifuge tube. 11. Thaw the cryo vial by dropwise adding warm medium to the ice core. As soon as thawed, immediately transfer the entire content to the 15 mL tube and mix with the pre-warmed K0-Medium. 12. Centrifuge the thawed cell suspension at 220  g, for 4 min at RT, set speed to 9 and brake to 1.

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13. Aspirate the supernatant. 14. Resuspend the pellet in 5–7 mL of K0-Medium, depending on the pellet size. 15. Count the cells. 16. Set HUKORS suspension to 0.7  106 cells/mL. 17. Change the medium in the lower cavity of the prepared 12-well plate by aspirating the 1:1 DLM:K0 Medium and replacing it with 1 mL K0-Medium. 18. Pipette 0.5 mL of the HUKORS cell suspension atop the pre-cultured HUMORS into the prepared inserts with a feeder layer (see Subheading 3.5.1 for feeder layer, Subheading 3.5.2 for seeding HUMORS). 19. Cultivate in K0-Medium at 37  C, normoxic conditions with 5% CO2. 20. Photo-document by taking images at 4x and 10x magnification (scale bar 500 and 200 μm, respectively). d2: Lift, change medium in the lower cavity, photo-document. 21. Lift: carefully aspirate the medium from the upper cavity in order to expose the upper side of the equivalent to atmospheric air. 22. Medium change: aspirate the used medium from the lower cavity with a Pasteur pipette, add 1 mL of 1:1 mixture of DLM:K0 Medium to the lower cavity. d5: change medium in the lower cavity (1 mL 1:1-Medium), photo-document. 23. Aspirate the used medium from the lower cavity with a Pasteur pipette. 24. Add 1 mL of 1:1 mixture of DLM:K0 Medium to the lower cavity. If the equivalent is wet at the upper side, add a reduced amount of 0.7 mL 1:1 mixture of DLM:K0 Medium to the lower cavity instead of 1 mL. 25. Photo-document by taking images at 4 and 10 magnification (scale bar 500 and 200 μm, respectively) (see Fig. 4a–c). d7: medium change. 26. Aspirate the Used Medium from the Lower Cavity with a Pasteur Pipette and Add 1 mL of 1:1 Mixture of DLM:K0 Medium to the Lower Cavity (see Note 10). d8: photo-documentation. 27. Photo-document by taking images at 4 and 10 magnification (scale ar 500 and 200 μm, respectively)

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Fig. 4 Melanin presence and distribution in the P-EPI. (a) Macro view, d14 of the P-EPI 3D culture. (b) Macro view, d14 of the P-EPI 3D culture. (c) HE staining of a d14 NP-EPI without visible pigmentation. (d) HE staining of a d14 P-EPI with visible pigmentation (dark dots). (a, b) Scale bar corresponds to 5 mm. (c, d) Scale bar corresponds to 100 μm

d9: medium change. 28. Aspirate the used medium from the lower cavity with a Pasteur pipette and add 1 mL of 1:1 mixture of DLM:K0 Medium to the lower cavity (see Note 10) d12: medium change. 29. Aspirate the used medium from the lower cavity with a Pasteur ipette, add 1 mL of 1:1 mixture of DLM:K0 Medium to the lower cavity (see Note 10) d13: photo-documentation. 30. Photo-document by taking images at 4 and 10 magnification (scale bar 500 and 200 μm, respectively) (see Fig. 4a–c) d14: medium change. 31. Aspirate the used medium from the lower cavity with a Pasteur pipette, dd 1 mL of 1:1 mixture of DLM:K0 Medium to the lower cavity (see Note 10). d15: Endpoint pigmented epidermal equivalent (see Fig. 4). The pigmented epidermal equivalent is now fully stratified, recognizable by the main landmark, stratum corneum. Presence of melanin should be observable in the 3D culture if you vary the focus. Melanin and melanosomes are visible as dark dots or zones.

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Fig. 5 Characterizaton of the 2D cell culture of HUKORS (a) and HUMORS (c). KRT5 is used as a marker of basal keratinocytes (a, b) and PMEL as a marker of melanosomes and herewith melanocytes (c, d). NHEK (b) and NHEM (d) are used as control cell lines. Scale bars correspond to 100 μm 3.6 Characterization of Epidermal Equivalents 3.6.1 Immunocytochemistry in 2D Cell Cultures (See Fig. 5)

To characterize HUKORS and HUMORS primary culture, Normal Human Epidermal Keratinocytes (NHEK) and Normal Human Epidermal Melanocytes (NHEM) can be used as control cell lines. 1. To characterize 2D cell culture of HUKORS or HUMORS, cultivate the cells for 2 days in chamber slides and then perform the staining. Stain for keratinocyte marker keratin 5 (KRT5) and melanocyte protein marker glycoprotein 100/pre-melanosome protein (PMEL) to localize melanosomes: 2. Fix cells with ice-cold methanol at
20  C for 10 min. 3. Permeabilize with 0.1% saponine-PBS for 10 min at RT. 4. Block with 2% BSA for 15 min. 5. Incubate separately with following primary antibodies: rabbit polyclonal anti-KRT5, mouse monoclonal anti-PMEL for 60 min at RT. 6. Wash for 2 min with PBS. 7. Incubate with following secondary antibodies: goat anti-mouse Alexa Fluor 594 and goat anti-rabbit R-Phycoerythrin, respectively, for 45 min at RT. 8. Wash two times 2 min with PBS. 9. Counter-stain the nuclei with DAPI. 10. Mount cover slips with Fluoromount™ cooled to 4  C. 11. Document protein microscopy.

3.6.2 Melanin Content of Epidermal Equivalents

expression

by

using

fluorescence

The most prominent function of melanocytes is melanin production and its further distribution to epidermal keratinocytes. This feature can be quantified by measuring the content of melanin from lysed cells of 2D culture or epidermal equivalents.

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1. Lyse cells by repeated freeze/thaw cycles (3–5 times) and subsequent incubation with 1 M sodium hydroxide for 5 h at 60  C. 2. Collect cell lysates. 3. Measure the extinction of the lysates at λ ¼ 475 nm using a photometer. 4. Perform the same measurement with a standard curve of synthetic melanin with known concentrations. 5. Calculate the melanin concentration in the lysate against the values of the curve (see Subheading 2.3 step 12). 6. Calibrate the concentration/amount to that of per cell for the 2D culture (see Subheading 3.4.2) and per cm2 for the equivalents. 3.6.3 Histological Characterization of Equivalents

For characterization purposes of NP-EPI, normal human epidermal keratinocytes (NHEK) can be used as control cell line but the HUKORS worked much better in our hands. In this case, additionally prepare a separate setup for the 3D protocol for generating non-pigmented epidermal equivalents with the NHEK. For characterization purposes of P-EPI, normal human epidermal keratinocytes (NHEK) and normal human epidermal melanocytes (NHEM) can be used as control cell lines in a separate additional setup for the 3D co-culture protocol for generating pigmented epidermal equivalents. Nevertheless, the 3D co-culture of HUKORS with HUMORS worked much better in our hands, whereas NHEK+NHEM P-EPI even developed aberrantly to a large extent. It is generally sufficient to characterize the NP-EPI or P-EPI endpoint at d14. If you are aiming at monitoring the growth before the endpoint NP-EPI/P-EPI, control points should be set at d5, d9, and d14. Include a separate setup for each of these points. For basic histological evaluation, perform cross-sectioning of the equivalents embedded in paraffin: 1. Fix the equivalents in 4% buffered formalin for 24 h. 2. Dehydrate in ethanol gradients, clear in xylene and embed in paraffin. 3. Perform sectioning with a mid-size blade, thickness 5–7 μm. 4. Keep in cold water bath. 5. Transfer to 48  C water bath. 6. Keep on the stretching table at 40  C for 1 h. 7. Store at RT. Stain sections routinely with hematoxylin and eosin (HE) (see Fig. 6).

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Fig. 6 Checkpoints for epidermal equivalents at d5, d9, and d14. Both the NP-EPI (a–c) and P-EPI (d–f) are exhibiting evident increase in thickness and in number of layers. The endpoint at d14 shows comparable structure to that of human skin (g). ba Stratum basale, sp Stratum spinosum, gr Stratum granulosum, co Stratum corneum. Scale bars correspond to 100 μm

HE staining 1. Keep the sections in 2  10 min in xylol. 2. Transfer to 100% EtOH, keep for 5 min, transfer to 96% EtOH, 5 min transfer to 80% EtOH, 5 min transfer to 70% EtOH, 5 min transfer to distilled water for 5 min. 3. 6 min in hematoxylin solution. 4. Wash with tap water. 5. Keep in warm water until the color changes to blue (10–15 min). 6. Wash with distilled water. 7. Briefly submerge in 1% eosin (2–5 min). 8. Briefly wash with distilled water. 9. To dehydrate, perform ascending EtOH gradient steps fast (70/80/96/100 EtOH). 10. Transfer to xylol (2 10 min) and keep in xylol until mounting. 11. Mount with Leicamount. 12. Measure section thickness of the equivalents manually directly from an HE-stained cross-section using the Keyence BZ-9000 Generation II Analyzer Software Version 2.2. Typically, the overall thickness will increase 4–6-fold between d5 and d14 (see Fig. 6). Masson-Fontana staining To detect the melanin presence and distribution in the pigmented equivalents perform Masson-Fontana staining (see Fig. 4). Preparing of ammoniacal silver solution

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1. Prepare the needed volume of 5% silver nitrate and keep stirring at all times. 2. Add ammonia solution dropwise. The solution will turn brown and then clear. Keep adding ammoniac drops until the solution is clear. 3. Now add silver nitrate dropwise until the solution turns cloudy. Staining 4. Deparaffinize the sections in xylol for 2–10 min, 5 min per step 100/96/80/70% EtOH. 5. Treat with freshly prepared ammoniacal silver solution at 60  C for 60 min. 6. Wash three times with distilled water. 7. Keep 5 min in distilled water. 8. Incubate with 0.2% gold chloride solution for 10 min at RT. 9. Wash three times with distilled water. 10. Wash three times with tap water. 11. To fix incubate in 5% sodium-thiosulfate solution for 2 min at RT. 12. Keep in Nuclear Fast Red solution for 2 min at RT. 13. Wash three times with tap water. 14. Instantly dehydrate in ascending EtOH steps (5 min 70/80/ 96/100% EtOH). 15. Incubate 2 10 min xylol with agitation. 16. Mount with Leicamount. 3.6.4 Immunohistochemistry of Equivalents

Staining of keratin (KRT) 5, KRT10, and involucrin (IVL) should be sufficient to characterize an NP-EPI. P-EPI should be additionally stained with PMEL. 1. De-parafinize paraffin histological sections in xylene and rehydrate by 4 min steps of the ethanol gradients: 100%, 96%, 80%, 70%, 50%, Aqua destilata. 2. Employ heat-mediated antigen retrieval using 10 mM sodium citrate buffer (pH ¼ 6.0) 12 min in a pressure cooker. 3. Stain sections with primary antibodies: rabbit polyclonal antiKRT10, mouse monoclonal anti-IVL, rabbit polyclonal antiPMEL: Incubate for 60 min at RT. 4. Wash two times for 2 min in PBS. 5. Incubate for 45 min with following secondary antibodies: goat anti-mouse Alexa Fluor 594, anti-streptavidin Alexa Fluor 594, and goat anti-rabbit R-Phycoerythrin at RT, respectively. 6. Counter-stain the nuclei with DAPI.

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Fig. 7 Immunohistochemical characterization of NP-EPI and P-EPI. KRT5 is used as a basal keratinocyte marker, present in basal and suprabasal strata, primarily in stratum basale (a–c). KRT10 is a marker of differentiated keratinocytes, expressed in the outer strata, concluded with stratum corneum (d–f). Involucrin, marker of terminal keratinocyte differentiation, is expressed primarily in stratum granulosum and stratum corneum (g–i). Scale bars correspond to 100 μm

7. For anti-KRT5 staining, see Subheading 3.6.1. 8. Mount slides and visualize by fluorescence microscopy. As basal keratinocyte marker, KRT5 should be visible in basal layer, KRT10 in suprabasal layers, whereas the terminal keratinocyte marker IVL should be detectable in the granular and corneal layer. PMEL will primarily be located in basal and only occasionally in suprabasal layers. PMEL signals may also be detected in keratinocyte membranes as well as intracellularly; these are the indicators of melanin distribution by melanocytes and internalization by keratinocytes (see Fig. 7). The amount of apoptotic, necrotic, and parakeratotic cells should be negligible. 3.7 Dermatological Scoring of Histological Sections (See Fig. 6)

The epidermal equivalents should exhibit regular histological and anatomical features of the epidermis. HE staining should be sufficient for this type of scoring. EPI and P-EPI should display

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following strata: organized stratum basale, stratum spinosum, stratum granulosum, and stratum corneum. The keratinocytes will assume progressive flattening towards the outer strata, starting with cuboidal shape keratinocytes with elongated nuclei at stratum basale, over a number of polygonal spinous keratinocyte layers in stratum spinosum, flattened, granula-containing keratinocytes in stratum granulosum, to assume a plate-like shape in the orthokeratotic stratum corneum. The keratinocytes in those layers should display monomorphic nuclei (DAPI), whereas the cells in stratum corneum typically lack the nucleus and will show signs of detachment at the outermost zone of the stratum corneum. Intensive deviations from the described layer structure or from the characteristic morphology of the cells in different strata indicate alterations in the keratinocyte differentiation. The scoring should be performed upon histological sections of equivalents sampled on d14, which is the endpoint of the 3D cultures. Alternatively, d5 and d9 can be taken into analysis. The parameters to follow are: percentage of tested equivalents displaying organized stratum basale, number of layers in stratum spinosum, percentage of tested equivalents with present stratum granulosum, percentage of tested equivalents with present stratum corneum, percentage of tested equivalents with present desmosomes, percentage of monomorphic keratinocyte nuclei, and percentage of tested equivalents with present melanin. Basal layer will consist of cuboidal keratinocytes and melanocytes at ratio 10:1. In some cases, stratum basale may appear less organized. Stratum spinosum will be multilayered, consisting of up to 13 layers. Stratum granulosum should be exhibited in more than half of tested equivalents. Stratum corneum should be present in majority of tested equivalents (not less than 80%). Desmosomes should be present in all equivalents. Majority of tested equivalents should display monomorph keratinocyte nuclei (not less than 61%). Presence of melanin (MF) should be detected in melanocytes and in all P-EPIs. The signal will look like dark zones in the cell soma of melanocytes. Higher magnification will reveal the presence of melanin within the finer structures, melanosome vesicles, which will be distributed along the secretory path, starting from the nuclear vicinity, throughout the soma and along the dendrites. Upon the delivery of melanosomes, they should be visible in keratinocytes as well. Alternative focusing depth may reveal several dendrites reaching from the melanocyte soma connected to the surrounding keratinocytes, hereby mimicking melanin epidermal units. With a bit of luck, the distribution of melanosomes from melanocytes to keratinocytes can be detected.

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Erratically, melanocytes will be visible in the layers other than stratum basale. This is an in vitro artifact, mostly due to the keratinocyte proliferation that can place a new keratinocyte beneath a melanocyte and result in its displacement atop the basal layer. Also, traces of distributed melanin will be visible in keratinocytes as dark dots and some of them will end up in the strata atop the stratum basale. The quantifiable dermatological scoring parameters will increase over time, reaching the highest values at d14.

4

Notes 1. Transport of hair follicles at RT for up to 96 h in T-Medium. 2. Always use unwashed hair: in order to prevent the loss of the natural oils and herewith the skin elasticity, since dry skin tends to retain much of the follicle material and this reduces the harvest. Anagen follicles are visible as whitish, glossy, and compact. Such follicles should be included into the selection of N ¼ 40 for primary culture. 3. Make a swift single wrist-guided movement in the direction of hair growth in order to pluck the hairs. Proximal end of the plucked hair should carry a follicle. Pay attention to the quality of the plucked follicles and collect only intact anagen follicles for the primary culture. 4. The amount of 1.5 mL should be sufficient for contact between the lower insert side and the medium in order to keep it wet at the upper side by the means of surface tension forces. One insert is sufficient for 5–6 follicles, meaning a single 12-well plate should do for a single donor. 5. Keep washed follicles away from one another and from the insert walls. The follicles should be covered with a wet film. 6. Do not be tempted to mechanically separate the cells from the insert. Use trypsin (see Subheading 3.2.3 steps 30–48). 7. Optional: set aside 1 mL of suspension for purposes of quality control. 8. The medium should contact the lower side of the insert. Herewith, the epidermal equivalents are supplied with nutrients and kept humid. 9. If the epidermal equivalent is still covered by a wet film of liquid, reduce the amount of K-Medium to 0.7 mL. 10. If the equivalent is wet at the upper side, add 0.7 mL of 1:1 mixture of DLM:K0 Medium to the lower cavity instead of 1 mL.

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References 1. Dieckmann C, Renner R, Milkova L et al (2010) Regenerative medicine in dermatology: biomaterials, tissue engineering, stem cells, gene transfer and beyond. Exp Dermatol 19(8):697–706. https://doi.org/10.1111/j.1600-0625.2010. 01087.x 2. Limat A, Hunziker T (2002) Use of epidermal equivalents generated from follicular outer root sheath cells in vitro and for autologous grafting of chronic wounds. Cells Tissues Organs 172 (2):79–85. https://doi.org/10.1159/ 000065615 3. Anderer U, Savkovic V (2015) Zelltherapien in der Regenerativen Medizin. In: Sack U (ed) Zellul€are Diagnostik und Therapie. De Gruyter, Berlin, pp 291–341 4. Savkovic V, Dieckmann C, Milkova L et al (2012) Improved method of differentiation,

selection and amplification of human melanocytes from the hair follicle cell pool. Exp Dermatol 21(12):948–950. https://doi.org/10. 1111/exd.12038 5. Schneider M, Dieckmann C, Rabe K et al (2014) Differentiating the stem cell pool of human hair follicle outer root sheath into functional melanocytes. Methods Mol Biol 1210:203–227. https://doi.org/10.1007/978-1-4939-14357_16 6. Fitzpatrick TB, Breathnach AS (1963) The epidermal melanin unit system. Dermatol Wochenschr 147:481–489 7. Schneider M, Ziemer M, Simon JC et al (2020) Generation of pigmented and dermoepidermal skin grafts from human hair follicles. Tiss. Eng. A (in Revision)

Part III Characterization and Pre-Conditioning

Chapter 14 Using Gene Expression Music Algorithms (GEMusicA) for the Characterization of Human Stem Cells Martin S. Staege Abstract Gene Expression Music Algorithms (GEMusicA) use the transformation of gene expression data into melodies for the representation of sample-specific gene expression patterns. Quantitative analysis of similarities between melodies can be used for sample classification. The same algorithm can be used as simple and efficient encryption method. Here, we describe the usage of GEMusicA for the analyses of gene expression data from different stem cell types and stem cell-like tumor cells. Key words Gene expression, Germ cell tumors, Embryonic stem cells, Differentiation, Encryption

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Introduction The biological behavior of cells depends on the genes that are expressed in these cells. Modern gene expression analysis techniques like DNA microarrays or next-generation sequencing (NGS)based methods allow the genome-wide characterization of expression signatures in different cell types. Expression signatures of stem cells differ markedly from the expression signatures of more differentiated cells of the same cell linage. The characterization of stem cell-specific gene expression features can lead to a better understanding of stem cell biology, and this knowledge is necessary for optimization of treatment strategies for human diseases using stem cells as therapeutic tools (for instance, in the field of regenerative medicine) or targets (as in the case of cancer treatment). With the developments of microarrays and NGS techniques, the number of primary data points obtained from a single experiment has increased dramatically. Processing of the primary data will usually shrink the data set to a few thousands or millions of

Electronic supplementary material: The online version of this chapter (https://doi.org/10.1007/978-1-07161225-5_14) contains supplementary material, which is available to authorized users. Peggy Stock and Bruno Christ (eds.), In Vitro Models for Stem Cell Therapy: Methods and Protocols, Methods in Molecular Biology, vol. 2269, https://doi.org/10.1007/978-1-0716-1225-5_14, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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sequence-specific expression values. Several methods (heat maps, MA plots, volcano plots, t-SNE plots, and so on and so forth) have been developed in order to further reduce the dimensionality of such data and to visualize gene expression. An alternative and complementary approach for the presentation and intuitive apperception of gene expression data is based on the transformation of these data into melodies [1, 2]. Such “Gene Expression Music Algorithms” (GEMusicA) have been used for the characterization of stem cell features of Ewing sarcoma cells [2]. Here, I describe usage of GEMusicA for the analysis of germ cell tumor cells with variable differentiation status [3] in comparison to embryonic stem cells and other stem cell types. I describe how to prepare and analyze melodies using a GEMusicA implementation in R (see the Supplementary R code). This code can be modified, extended, and used as a starting point for own variants of the algorithm.

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Materials

2.1 Software and Hardware Requirements

There are no special hardware requirements for the methods described in this chapter. The examples have been tested on an Intel Core i5–3470 CPU @ 3.20 GHz with 16.0 GB RAM and 64 bit operating system and >10 GB-free space on hard drive. An internet connection is only required for the analysis of raw data from public data bases. If processed gene expression data are already available as text file, these files can be directly used for further analysis. If raw data have to be processed, the hardware and software requirements depend on the primary data format. As an example, I describe analysis of Affymetrix HG_U133A DNA microarray data from the Gene Expression Omnibus (GEO) data base [4]. I used the R software environment for implementation of GEMusicA (see Note 1). R is available for different platforms from The Comprehensive R Archive Network (https://cran.r-project.org/; latest accessed date: 20 Nov 2019). We recommend the usage of R in conjunction with RStudio (https://rstudio.com/products/rstudio/; latest accessed date: 20 Nov 2019). Several R packages are available that can be used for processing of raw data or for transformation of calculated melodies into audio files (see Note 2). For visualization of melodies, music scores can be produced in a TeX/LaTeX environment and R can be used for automatic generation of the TeX code. We used MiKTeX (available at https:// miktex.org/; latest accessed date: 20 Nov 2019) together with the MusiXTeX package (available at the Comprehensive TeX Archive Network (https://ctan.org/pkg/musixtex?lang¼de; latest accessed date: 20 Nov 2019). We recommend usage of a LaTeX writing environment, e.g., TeXstudio (http://texstudio.

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sourceforge.net/; latest accessed date: 20 Nov 2019). Alternatively, simple music scores can be prepared with other music notation editors, e.g., GNU Denemo (available at http://www.denemo. org/; latest accessed date: 20 Nov 2019). For post-processing of audio files varying free software tools are available, e.g., Audacity (https://www.audacity.de/downloads/; latest accessed date: 04 Nov 2019). 2.2 Gene Expression Data

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Any type of quantitative (gene expression) data can be used for GEMusicA (see Note 3). Raw data can be generated in the own lab or downloaded from public data bases. A large collection of (microarray-based) gene expression data is available from the GEO database [4] (https://www.ncbi.nlm.nih.gov/gds; latest accessed date: 20 Nov 2019). Similarly, NGS-based raw data are available from the Sequence Read Archive (SRA; https://www. ncbi.nlm.nih.gov/sra/; latest accessed date: 20 Nov 2019). In this chapter, we demonstrate the methods by using Affymetrix HG_U133A microarray-based gene expression data from germ cell tumor cells [3], undifferentiated embryonic stem cells [5], hematopoietic stem cells [6], and bone marrow-derived mesenchymal stroma cells [7]. The supplementary R script “Experiment1.r” can be used for automatic download of the required .cel files from the GEO data base (see Supplementary R code).

Methods A typical GEMusicA analyse includes the following steps: (A) Selecting and pre-processing of gene expression data, (B) calculation of tone frequencies and tone lengths, (C) generation of audio files on the basis of the calculated frequencies and tone lengths. Finally, (D) the result can be visualized and quantified.

3.1 Preparing the Environment

3.2 Step A: Selection and Pre-Processing of Gene Expression Data

This section describes a GEMusicA-based analysis using the R code provided as supplementary material (see Note 4). Note that GEMusicA uses current computer technology and, therefore, floating point rounding errors might lead to slightly different results on different hardware/software combinations (see Note 5). 1. Select the gene expression data set. In the example presented here, we use publicly available DNA microarray data from the GEO data base. If processed gene expression data are already present on your hard drive, proceed to step 4. The supplementary R file “Experiment1.r” includes the information necessary for analysis of the stem cell/germ cell tumor set described below (see Note 6). For the analysis of other data sets, the accession numbers in the supplementary R code have to be changed accordingly.

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2. Download the raw data and process the data. For download and processing of .cel files the Init function (see Supplementary R code) can be used (see Note 7). You can also manually download raw data from public databases. Pre-processing of gene expression data includes the transformation of raw data into a table with gene/transcript-specific expression values for each sample. The final table will contain normalized log-2 transformed data for usage in the next steps (see Note 8). 3.3 Step B.1: Calculation, Filtering, and Sorting of Frequencies and Tone Length

GEMusicA uses a coding system for the transformation of gene expression data into melodies. Genes with high expression are encoded by tones with high frequency and genes with low expression are encoded by tones with low frequency (see Note 9). The calculations can be performed by the GEMusicA18 function in R (see Supplementary R code). It is possible to pre-define the set of possible frequencies. This can decrease the computation time because it is not necessary to calculate these values from scratch. However, it is more flexible to allow calculation of frequencies according to the user demands. If equal temperament is used for tuning, the logarithms of the distance between neighboring frequencies are always the same. Conventionally, the octave (two tones with a frequency ratio of 1:2) is divided into 12 equal half-tone steps. If this system is used, it is possible to visualize the resulting melodies by music scores. Other divisions are also possible (see Note 10). 1. Define the number of different tone pitches and define the frequencies of these pitches (see Note 10). 2. If you use pre-defined frequencies, proceed to step 5. Otherwise, define the lowest possible frequency (see Note 10). 3. Define the number of different tone steps that will be used (see Note 10). 4. Define the number of tones per octave. The default value used by the supplementary R code is 12. 5. Define the number of different tone lengths that will be used or define a standard length for all tones (see Note 11). 6. Set the number of tones that will be generated (see Note 12). 7. Run GEMusicA (e.g., by calling the GEMusicA18 function from the supplementary R code) with the defined parameters. After calculation of frequencies and tone length, the order of the tones will be determined. It is possible to sort the tones according to biological features of the corresponding genes/ proteins (e.g., on the basis of Gene Ontology information). However, the knowledge about the biology of genes/proteins is unsteady and genes with multiple functions can be pigeonholed in different ways. Sorting the tones on the basis of the median of expression samples is independent from additional knowledge.

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Adjusting the melodies to a reference melody or a single reference tone can increase the discriminatory power of the melodies [1]. The deviation of the gene expression of a given sample from a reference sample (see Note 13) can be transformed into the deviation of the sample-specific melody from the reference melody. 1. Define a reference melody or single reference frequency (see Note 14). 2. Adjust the melody to the reference melody or reference tone (see Note 14).

3.5 Step C: Generate Audio Files

1. Prepare audio files from the calculated frequencies and ton lengths (see Note 15). 2. Play audio files with available audio software (see Note 16).

3.6 Step D: (Optional) Visualization and Quantitative Analysis of Frequencies

GEMusicA was developed for auditory display of gene expression data. However, for documentation purposes it is often desirable to present the results visually (e.g., in printed publications). For this end, melodies can be transcribed into musical scores. In addition, it is possible to apply different strategies for the quantitative analysis of the generated frequencies (see Note 17). Here, we describe only one of these possibilities which is based on an interesting encryption/decryption feature of GEMusicA. 1. Music scores can be prepared by using the supplementary R code. Prepare TeX templates for generation of music scores, e.g., by calling the function GenerateTeXFiles form the supplementary R code (see Note 18). Example music scores from stem cells and germ cell tumor cells are presented in Fig. 1. 2. Use the calculated gene-specific frequencies for a t-distributed Stochastic Neighbor Embedding (t-SNE) analysis [8] or other graphical analysis methods (see Note 19). A typical result for the stem cell/germ cell tumor cell data set is shown in Fig. 2. 3. Define a text that will be used for calculation of similarity scores and calculate similarity scores for the melodies by running the function DecScore (see Supplementary R code). This function uses a single sample for encryption of a text by an audio file. The sample number is one part of the key for decryption. Thereafter, all other samples are tested for their ability to be used as key for recovering the original text. The percentage of correctly recalculated letters from the original text is used as score for similarity between the sample used for encryption and the sample used for decryption (see Note 20). The example from Fig. 3 has been generated by using a short text from Hugo the St. Victor [9] which is included in the supplementary R file “Experiment1.r”.

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Fig. 1 Melody examples from the germ cell tumor cell/stem cell data set. Presented are representative melodies (only the first 12 bars) for all analyzed cell types. Melodies were adjusted to the reference melody “Reference_1” (see supplementary R code). The embryonic stem cell-specific genes LIN28A, NANOG, and POU5F1 are highlighted. High frequencies of all three tones representing these genes are only present in undifferentiated germ cell tumor (GCT) cells (H12.1 cells) and embryonic stem cells (ESC), but not in mesenchymal stromal cells (MSC), hematopoietic stem cells (HSC), somatically differentiated GCT cells (1411HP), or extra-embryonically differentiated GCT cells (1777NRpmet). Somatically differentiated cells have lost expression of all three genes whereas in extra-embryonally differentiated cells expression of LIN28A is still high (see also reference [3]). Accidentals only apply to the immediate note. The complete scores of all samples from the germ cell tumor cell/stem cell data set (before and after adjusting to the reference melody) are available as supplementary pdf files. The corresponding audio files are presented as supplementary wavesound files

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Fig. 2 t-SNE analysis of GEMusicA frequencies. Melodies have been produced with the supplementary R code. The t-SNE analysis of the calculated frequencies was performed with the supplementary R code. The high similarity between undifferentiated germ cell tumor (GCT) cells and embryonic stem cells but not with hematopoietic stem cells or mesenchymal stroma cells is clearly visible. Differentiated GCT cells (somatically or extra-embryonically) have lost this similarity

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Notes 1. The calculation of frequencies and tone lengths from gene expression data can be performed with any standard spreadsheet calculation software [1]. However, this is only a makeshift and even for medium-sized data sets the required computation times are unacceptable. Moreover, it is necessary to use additional tools for processing of raw data and for rendering the calculated frequencies audible. All necessary procedures are available in R (see Supplementary R code). The code has been tested with varying data sets with stable and reproducible results. For the analysis of DNA microarray raw data, all necessary steps are included in this code. Alternatively, array-specific software solutions can be used. For example, for Affymetrix microarrays, the Transcriptome Analysis Console (Thermo Fisher Scientific, Waltham, Massachusetts, USA) allows export of tab delimited text files that can be used for further analysis.

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Fig. 3 Encryption/decryption-based quantification of similarities between melodies. Samples from the stem cell/germ cell tumor cell data set were used individually for encoding a text fragment from the Didascalicon of H. de St. Victor as wavesound file. Thereafter, the melodies from all samples were used successively for decoding the melody and recovering the original text. Percent identity between the letters of the original text and the recovered text were used as similarity score. As expected, ESC, MSC, and HSC have highest scores for their own class members. Undifferentiated germ cell tumor cells (H12.1, H12.5) have high scores for ESC whereas somatically differentiated germ cell tumor cells (H12.1.D, 1777NRpmet) have lower similarities to ESC and increased scores for MSC. Extra-embryonically differentiated germ cell tumor cells (1411HP, GCT72) show an intermediate behavior (see also reference [3])

2. After installing R (and optionally RStudio), additional packages for processing of gene expression data and generation of sound files are required. The following packages should be available for the example analyses described in this chapter: GEOquery, oligo, pd.hg.u133a, audio, t-SNE, seewave, tuneR, Biostrings. RStudio can be used for installing the required packages from the internet. For other gene expression (microarray) platforms, the corresponding packages have to be installed. 3. Alternatively to transcriptomics data, other data can also be analyzed with the algorithm. The only requirement is that the data can be stored in a table where columns are samples and rows are individual features, e.g., gene-specific signal intensities from microarray experiments, normalized reads from RNA-seq experiments, antigen-specific mean fluorescence intensities from flow cytometry experiments, and so on and so forth. 4. You can start a new R project (e.g., in RStudio) and run the supplementary GEMusicALight.r script. This will define all

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necessary functions that are required for GEMusicA. Running the supplementary Experiment1.r script will perform all analyses discussed in this manuscript. 5. During the different steps of the algorithm, multiple calculations with floating point numbers have to be performed. Unfortunately, these calculations are imprecise as a consequence of the necessity for internal transformation of floating point numbers into binary numbers. You can simply test this by entering the following short code into your R console: (1101579-579+57.9-0.9)¼¼(1101-579-579-0.9+57.9). R will tell you that the answer is FALSE but you can see that the result is true (a  2b + c  d ¼ a  2b  d + c). In Microsoft Excel the same problem occurs with the following formula: ¼(1101579-579-0.9+57.9)¼(1101-579-579+57.9-0.9). Depending on the software and operating system, such errors will occur more or less frequently. In GEMusicA applications, such rounding errors might become relevant for sorting and filtering steps as well as during decryption of audio files (see below). 6. In principle, GEMusicA can be used for the comparative analysis of two single samples. In this case, sample-specific gene expression features will lead to sample-specific melodies. Increasing the number of samples will shape the melodies into class-specific melodies if such classes exist. In our example, the data set includes six classes: three types of human stem cells (hematopoietic stem cells, embryonic stem cells, and mesenchymal stem/stroma cells) and three types of germ cell tumor cells (undifferentiated, somatically differentiated and extraembryonically differentiated). The differential gene expression signatures of these classes are strong enough to allow the generation of characteristic melodies for the stem cell types. If the signatures are only weakly present, secondary classes can come out from behind. One class with a strong impact on gene expression signatures is the gender. Transcripts from the Y chromosome as well as transcripts related to dosage compensation and X-inactivation in female cells are often included in gene lists with differentially expressed genes, especially if the numbers of male and female donors in the compared groups are not equal. 7. The Init function (see supplemental R code) uses the Robust Multi-array Average (RMA) algorithm for processing of microarray .cel files. The function has three arguments: “ExperimentName” is the name of the experiment which will be used for storage of the data. “ExpressionData” is a list with .cel file names. The file names must match the corresponding GEO accession numbers if the files have to be downloaded from the GEO archive. “GEO” indicates whether the .cel files have to be downloaded from the GEO archive or not. Use GEO¼0 if .cel files are already present in the corresponding Cel_Files folder.

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8. The data should be normalized and the Init function uses RMA normalization. For DNA microarray-based gene expression data, scaling algorithms (e.g., the Microarray Suite 5.0 (MAS5.0) algorithm) are also sufficient, especially if all samples have been processed with the same protocol in the same lab. For a single gene or transcript, multiple values might be present in the data set (for instance, in the case of DNA microarray data which contain more than one probeset per gene). It is possible to agglomerate all expression values from a gene into a single value. However, this is not necessary and I suggest starting the analysis without agglomeration. Agglomeration might become interesting if a single differentially expressed gene (or few genes) with multiple expression values (probesets) dominate the final calculated melodies. In this case, the analysis can be repeated with agglomerated data. If filtering of gene sets is performed (see below) the method must be compatible with the gene set filter (e.g., usage of the same probeset IDs). If all (un-filtered) genes are used, it is also possible to select for a single gene only the values with the highest variance in the complete data set. The disadvantage of this method is the fact that in different data sets different probesets might be selected. Alternatively, the median value of all values from a single gene can be used. In principle, all genes from a data set can be used for GEMusicA. A disadvantage of this approach can be that genes with high expression in individual samples can be overrepresented in the final melodies. This problem can be circumvented if it is known which subset of genes (e.g., a stem cellspecific gene set) is relevant for the current investigation. In this case, it is possible to use only this gene set (or few gene sets) for GEMusicA [1]. 9. Obviously, this is only one possible implementation of the algorithm. For example, it is possible to encode genes with high expression by loud (or long) tones and genes with low expression with soft (or short) tones. If the melodies are presented only as audio files (or played on true music instruments), other methods might be interesting. For example, from the tones of a single octave, the frequency of the octave has the largest difference to the frequency of the fundamental tone but the dissonance between fundamental tone and octave is very low. Therefore, increasing differences in the gene expression between samples might also be encoded by increasing dissonance. A disadvantage of such dissonance scoring system lies in the lower objectifiability. The definition of dissonance and concord has changed from time to time whereas the frequency of a tone is an objective physical parameter.

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10. The GEMusicA18 function in R (see Supplementary R code) can be used with pre-defined frequencies. If you use such pre-defined frequencies, set the variable DefinedFrequencies to the corresponding value. Otherwise, use the default value 0 for this variable. The GenerateFrequencyList function (see Supplementary R code) will be used by GEMusicA for selection or calculation of frequencies. These frequencies will be stored in the variable Frequencies. DefinedFrequencies¼0 will calculate the frequencies on the basis of the minimal frequency (MinFreq), the number of tone steps per octave (ToneSteps) and the total number of tone steps (NumKeys). In this case, equal temperament will be used. With DefinedFrequencies¼1 the function will use pre-defined frequencies which are stored in the variable TetraFreqs[1,]. An advantage is the fact that it is easy to define frequencies from tuning models that differ from equal temperament. For instance, the quart interval c–f can be divided not only into one whole-tone, one whole-tone and one semi-tone interval (c–d–e–f; which is equivalent to the major scale) or into one whole-tone, one semi-tone and one wholetone interval (c–d–e flat–f; which is equivalent to the minor scale), but also into other intervals, e.g., the tone steps 0.5, 0.75, and 1.25 or the tone steps 0.375, 0.375, and 1.75. Such intervals are unusual in today’s Western music but have been found, e.g., in ancient Greek music [10]. The supplementary R code includes three pre-defined sets of frequencies which can be used (note that the number of different tones is limited to the number of pre-defined frequencies). You can define your own set of frequencies. In this case, set the variable DefinedFrequencies to 1 and store the frequencies as variable Frequencies. If the octave is considered to be the master interval, this interval can be divided into 12 semi-tone steps, 24 quartertone steps or any other number of tone steps. It is even possible to calculate the corresponding frequencies if the octave is divided into a non-natural number of tone steps. For example, ToneSteps¼10.5 in the GenerateFrequencyList function leads to frequencies where after 21 tone steps the super-octave is reached. With such tuning, the octave is not included in the set of possible frequencies. 11. In the supplementary R code, the variables FixedDur and maxNdots are used for defining the tone length. The default values used by the supplementary R code are 4 and 2, respectively. Possible values for the variable FixedDur in the supplementary R code are 1, 2, 4, 8, and 16. With these values and the default values from the GenerateTeXFiles function (see below), the shortest tone length will be the crotchet, quaver, semiquaver, demisemiquaver, and hemidemisemiquaver,

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respectively. If negative values are used for FixedDur (e.g., FixedDur