Epithelial Cell Culture: Methods and Protocols (Methods in Molecular Biology, 2749) [2 ed.] 1071636081, 9781071636084

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Methods in Molecular Biology 2749

Mario Baratta  Editor

Epithelial Cell Culture Methods and Protocols Second Edition

METHODS

IN

MOLECULAR BIOLOGY

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

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

For over 35 years, biological scientists have come to rely on the research protocols and methodologies in the critically acclaimed Methods in Molecular Biology series. The series was the first to introduce the step-by-step protocols approach that has become the standard in all biomedical protocol publishing. Each protocol is provided in readily-reproducible step-by step 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.

Epithelial Cell Culture Methods and Protocols Second Edition

Edited by

Mario Baratta Department of Chemistry, Life Sciences and Environmental Sustainability, University of Parma, Parma, Italy

Editor Mario Baratta Department of Chemistry, Life Sciences and Environmental Sustainability University of Parma Parma, Italy

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-3608-4 ISBN 978-1-0716-3609-1 (eBook) https://doi.org/10.1007/978-1-0716-3609-1 © The Editor(s) (if applicable) and The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024 This work is subject to copyright. All rights are solely and exclusively licensed by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, expressed or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. This Humana imprint is published by the registered company Springer Science+Business Media, LLC, part of Springer Nature. The registered company address is: 1 New York Plaza, New York, NY 10004, U.S.A. Paper in this product is recyclable.

Preface Epithelial cell culture has emerged as a cornerstone technique in the field of cell biology, providing invaluable insights into the fundamental mechanisms underlying various physiological processes. It is with great pleasure and excitement that we present the second edition of Epithelial Cell Culture: Methods and Protocols, which builds upon the success of the first edition and further delves into the dynamic world of epithelial cell research. In this edition, we have taken a meticulous approach to ensure the integration of chapters from the previous edition (Chaps. 2, 3, 5, 9, 10) while also incorporating new issues that have surfaced since the first edition. The rapid advancements in cell culture methodologies and the ever-evolving understanding of epithelial cell biology have paved the way for novel techniques and applications. As such, this edition reflects the latest cuttingedge research, ensuring that our readers are equipped with the most up-to-date knowledge in the field. An essential hallmark of this volume is the comprehensive representation of epithelial cell culture models from different mammalian species. As our understanding of cellular biology becomes more intricate, the need to explore diverse organisms becomes imperative. Hence, in this book, you will find detailed methodologies applicable to epithelial cells derived from primates, pigs, bovines, and laboratory animals. This expansion in scope allows researchers and students alike to grasp the nuances of comparative biology, fostering a deeper appreciation of species-specific variations in cellular behavior. Epithelial Cell Culture: Methods and Protocols, Second Edition is not limited to conventional cell culture techniques; rather, it transcends boundaries by delving into the manipulation and differentiation of epithelial cells. As cellular engineering and regenerative medicine hold the promise of transformative medical therapies, understanding cellular behavior at this level of granularity becomes paramount. Therefore, the inclusion of these advanced methodologies is a reflection of our commitment to equipping researchers with the knowledge and tools needed to push the boundaries of scientific exploration. Recognizing the growing significance of the gastroenteric system in human medicine and nutrition, we have dedicated special attention to epithelial cell models in this vital system. Epithelial cells lining the gastrointestinal tract play a crucial role in nutrient absorption, maintaining barrier integrity, and orchestrating interactions with the complex gut microbiome. As emerging research continues to reveal the intricate interplay between the gut epithelium and human health, we felt compelled to offer a dedicated section that addresses the unique challenges and opportunities presented by this essential system. This second edition volume aspires to be more than just a reference guide; it aims to be a source of inspiration and knowledge that empowers researchers to push the boundaries of cellular science. We believe that the collective efforts of scientists and students in this field hold the key to unlocking groundbreaking medical discoveries, improving human health, and advancing our understanding of life itself. We extend our heartfelt gratitude to all the contributors and researchers who have generously shared their expertise, making this second edition possible. Their collective dedication has enriched this volume and enabled it to become a cornerstone resource in the realm of epithelial cell culture.

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Finally, we hope that the scientific community, educators, and students will find this edition to be a valuable companion in their journey to unravel the complexities of epithelial cell biology, cultivate curiosity, and inspire the next generation of groundbreaking research. Parma, Italy

Mario Baratta

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

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1 Culture of Mouse Thymic Epithelial Cells in Serum-Free Medium . . . . . . . . . . . . 1 Yasuhiro Adachi 2 Cultured Pig Thyroid Follicular Cells: Electrical Evaluation of Epithelial Integrity. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7 Peying Fong 3 Human Minor Salivary Glands: A Readily Available Source of Salivary Stem/Progenitor Cells for Regenerative Applications . . . . . . . . . . . . . . 25 Caitlynn M. L. Barrows, Danielle Wu, Simon Young, and Mary C. Farach-Carson 4 Salivary Organotypic Tissue Culture: An Ex-vivo 3D Model for Studying Radiation-Induced Injury of Human Salivary Glands . . . . . . . . . . . . 39 Akshaya Upadhyay, Migmar Tsamchoe, and Simon D. Tran 5 Differentiation of Pig Gastric Primary Cells into Mucus Producing Epithelial Cells. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 55 Macarena P. Quintana-Hayashi, Sinan Sharba, and Sara K. Linde´n 6 Isolation, Culture, and Microscopic Imaging of Guinea Pig Primary Gastric Tissue Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 65 Weronika Gonciarz and Magdalena Chmiela 7 Method for Two-Dimensional Epithelial Monolayer Formation Derived from Mouse Three-Dimensional Small Intestinal Organoids . . . . . . . . . . 73 Yuta Takase and Toshio Takahashi 8 Human Hepatic Spheroid Coculture Model for the Assessment of Drug-Induced Liver Injury . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 85 Linhao Li and Hongbing Wang 9 In Vitro Porcine (Explant) Colon Culture . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 91 Matheus de Oliveira Costa and Michael K. Dame 10 Bioencapsulation of Oocytes and Granulosa Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . 103 Massimo Faustini, Stella Agradi, Daniele Vigo, Maria L. Torre, and Giulio Curone 11 Culturing and Differentiation of Patient-Derived Ectocervical Epithelial Stem Cells Using Air-Liquid Interphase and Matrigel Scaffold . . . . . . 109 Rajendra Kumar Gurumurthy, Naveen Kumar, and Cindrilla Chumduri 12 Ovine Trophoblast Cells: Cell Isolation and Culturing from the Placenta at the Early Stage of Pregnancy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 123 Paola Toschi, Irene Viola, Isabella Manenti, Silvia Miretti, Elisabetta Macchi, Eugenio Martignani, Paolo Accornero, and Mario Baratta

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Amniotic Membrane and Amniotic Epithelial Cell Culture. . . . . . . . . . . . . . . . . . . Angelo Canciello, Adrian Cervero`-Varona, Maura Turriani, Valentina Russo, and Barbara Barboni 14 Evaluation of the Epithelial Barrier Integrity in Primary Cultures of Pig Mammary Epithelial Cells. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Chiara Bernardini, Debora La Mantia, and Monica Forni 15 Bovine Skeletal Muscle Satellite Cells: Isolation, Growth, and Differentiation. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Silvia Miretti, Isabella Manenti, Paola Toschi, Elisabetta Macchi, Eugenio Martignani, Paolo Accornero, and Mario Baratta 16 Equine Induced Pluripotent Stem Cell Culture . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Julia Falk and F. Xavier Donadeu

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

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Contributors PAOLO ACCORNERO • Department of Veterinary Sciences, University of Torino, Turin, Italy YASUHIRO ADACHI • Department of Anatomy, School of Medicine, University of Occupational and Environmental Health, Fukuoka, Japan STELLA AGRADI • Department of Veterinary Medicine and Animal Sciences, University of Milan, Milan, Italy MARIO BARATTA • Department of Chemistry, Life Sciences and Environmental Sustainability, University of Parma, Parma, Italy BARBARA BARBONI • Faculty of Bioscience and Technology for Food, Agriculture and Environment, University of Teramo, Teramo, Italy CAITLYNN M. L. BARROWS • Department of Diagnostic and Biomedical Sciences, The University of Texas Health Science Center at Houston, Houston, TX, USA; Katz Department of Oral and Maxillofacial Surgery, The University of Texas Health Science Center at Houston, Houston, TX, USA CHIARA BERNARDINI • Department of Veterinary Medical Sciences, University of Bologna, Bologna, Italy; Health Sciences and Technologies-Interdepartmental Center for Industrial Research (CIRI-SDV), Alma Mater Studiorum – University of Bologna, Bologna, Italy ANGELO CANCIELLO • Faculty of Bioscience and Technology for Food, Agriculture and Environment, University of Teramo, Teramo, Italy ADRIAN CERVERO`-VARONA • Faculty of Bioscience and Technology for Food, Agriculture and Environment, University of Teramo, Teramo, Italy MAGDALENA CHMIELA • Department of Immunology and Infectious Biology, Institute of Microbiology, Biotechnology and Immunology, Faculty of Biology and Environmental Protection, University of Lodz, Lodz, Poland CINDRILLA CHUMDURI • Department of Molecular Biology, Max Planck Institute for Infection Biology, Berlin, Germany; Laboratory of Infections, Carcinogenesis and Regeneration, Medical Biotechnology Section, Department of Biological and Chemical Engineering, Aarhus University, Aarhus, Denmark; Chair of Microbiology, University of Wu¨rzburg, Wu¨rzburg, Germany GIULIO CURONE • Department of Veterinary Medicine and Animal Sciences, University of Milan, Milan, Italy MICHAEL K. DAME • Department of Internal Medicine, Division of Gastroenterology, University of Michigan Medical School, University of Michigan, Ann Arbor, MI, USA MATHEUS DE OLIVEIRA COSTA • Large Animal Clinical Sciences, Western College of Veterinary Medicine, University of Saskatchewan, Saskatoon, SK, Canada; Population Health Sciences, Faculty of Veterinary Medicine, Utrecht University, Utrecht, The Netherlands F. XAVIER DONADEU • Division of Translational Bioscience, The Roslin Institute and Royal (Dick) School of Veterinary Studies, University of Edinburgh, Edinburgh, UK JULIA FALK • Division of Translational Bioscience, The Roslin Institute and Royal (Dick) School of Veterinary Studies, University of Edinburgh, Edinburgh, UK

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MARY C. FARACH-CARSON • Department of Diagnostic and Biomedical Sciences, The University of Texas Health Science Center at Houston, Houston, TX, USA; Department of Bioengineering, Rice University, Houston, TX, USA; Department of Biosciences, Rice University, Houston, TX, USA MASSIMO FAUSTINI • Department of Veterinary Medicine and Animal Sciences, University of Milan, Milan, Italy PEYING FONG • Department of Anatomy and Physiology, Kansas State University College of Veterinary Medicine, Manhattan, KS, USA MONICA FORNI • Health Sciences and Technologies-Interdepartmental Center for Industrial Research (CIRI-SDV), Alma Mater Studiorum – University of Bologna, Bologna, Italy; Department of Medical and Surgical Sciences, University of Bologna, Bologna, Italy WERONIKA GONCIARZ • Department of Immunology and Infectious Biology, Institute of Microbiology, Biotechnology and Immunology, Faculty of Biology and Environmental Protection, University of Lodz, Lodz, Poland RAJENDRA KUMAR GURUMURTHY • Department of Molecular Biology, Max Planck Institute for Infection Biology, Berlin, Germany NAVEEN KUMAR • Chair of Microbiology, University of Wu¨rzburg, Wu¨rzburg, Germany DEBORA LA MANTIA • Department of Veterinary Medical Sciences, University of Bologna, Bologna, Italy LINHAO LI • Department of Pharmaceutical Sciences, University of Maryland School of Pharmacy, Baltimore, MD, USA SARA K. LINDE´N • Department of Medical Biochemistry and Cell biology, Institute of Biomedicine, Sahlgrenska Academy, University of Gothenburg, Gothenburg, Sweden ELISABETTA MACCHI • Department of Veterinary Sciences, University of Torino, Turin, Italy ISABELLA MANENTI • Department of Veterinary Sciences, University of Torino, Turin, Italy EUGENIO MARTIGNANI • Department of Veterinary Sciences, University of Torino, Turin, Italy SILVIA MIRETTI • Department of Veterinary Sciences, University of Torino, Turin, Italy MACARENA P. QUINTANA-HAYASHI • Department of Medical Biochemistry and Cell biology, Institute of Biomedicine, Sahlgrenska Academy, University of Gothenburg, Gothenburg, Sweden VALENTINA RUSSO • Faculty of Bioscience and Technology for Food, Agriculture and Environment, University of Teramo, Teramo, Italy SINAN SHARBA • Department of Medical Biochemistry and Cell biology, Institute of Biomedicine, Sahlgrenska Academy, University of Gothenburg, Gothenburg, Sweden TOSHIO TAKAHASHI • Suntory Foundation for Life Sciences, Bioorganic Research Institute, Kyoto, Japan YUTA TAKASE • Suntory Foundation for Life Sciences, Bioorganic Research Institute, Kyoto, Japan ` degli Studi del MARIA L. TORRE • Dipartimento di Scienze del farmaco , Universita Piemonte Orientale, Vercelli, Italy PAOLA TOSCHI • Department of Veterinary Sciences, University of Torino, Turin, Italy SIMON D. TRAN • McGill Laboratory of Craniofacial Tissue Engineering and Stem Cells, Faculty of Dental Medicine and Oral Health Sciences, McGill University, Montreal, QC, Canada MIGMAR TSAMCHOE • Department of Anatomy and Cell Biology, McGill University, Montreal, QC, Canada; Research Institute of the McGill University Health Center, McGill University, Montreal, QC, Canada

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MAURA TURRIANI • Faculty of Bioscience and Technology for Food, Agriculture and Environment, University of Teramo, Teramo, Italy AKSHAYA UPADHYAY • McGill Laboratory of Craniofacial Tissue Engineering and Stem Cells, Faculty of Dental Medicine and Oral Health Sciences, McGill University, Montreal, QC, Canada DANIELE VIGO • Department of Veterinary Medicine and Animal Sciences, University of Milan, Milan, Italy IRENE VIOLA • Department of Veterinary Sciences, University of Torino, Turin, Italy HONGBING WANG • Department of Pharmaceutical Sciences, University of Maryland School of Pharmacy, Baltimore, MD, USA DANIELLE WU • Department of Diagnostic and Biomedical Sciences, The University of Texas Health Science Center at Houston, Houston, TX, USA; Department of Bioengineering, Rice University, Houston, TX, USA SIMON YOUNG • Katz Department of Oral and Maxillofacial Surgery, The University of Texas Health Science Center at Houston, Houston, TX, USA

Chapter 1 Culture of Mouse Thymic Epithelial Cells in Serum-Free Medium Yasuhiro Adachi Abstract Primary cell culture systems are widely used as a valuable method for analyzing the biological functions of specific cells in vitro. Recently, various serum-free primary cell culture methods have been developed that do not involve the use of animal serums. Since the thymus is comprised of many cell types, such as thymocytes, thymic epithelial cells, macrophages, and fibroblasts, thymic epithelial cells must be isolated for their functional analysis in vitro. This chapter describes the detailed protocol for the selective primary culture of thymic epithelial cells using defined serum-free medium. Key words Thymic epithelial cells, TECs, Collagenase, Primary cell culture, Serum-free medium

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Introduction The thymus is the primary lymphoid organ responsible for the differentiation and maturation of T lymphocytes that function in the periphery. Thymic epithelial cells (TECs) are divided into two subpopulations (cortical TEC and medullary TEC), both of which play a role in inducing the differentiation and maturation of bone marrow-derived T lymphocyte progenitor cells (thymocytes) [1]. In early TEC analysis using cell culture systems [2, 3], the use of animal serums as an essential component of the culture medium caused several problems, including the proliferation of fibroblasts, the promotion of TEC differentiation by calcium ions, the effects of unknown factors in serum, and reproducibility problems due to serum quality. Subsequently, various improvements have been made, and serum-free culture methods have been established [3–7]. In recent studies on TECs, accurate analysis [8] and isolation of TEC subpopulations have been performed, alongside the investigation of differentiation stage-specific marker molecules [9] using flow cytometry (FCM) and fluorescence-activated cell sorting (FACS). However, the serum-free culture system is useful

Mario Baratta (ed.), Epithelial Cell Culture: Methods and Protocols, Methods in Molecular Biology, vol. 2749, https://doi.org/10.1007/978-1-0716-3609-1_1, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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for analyzing the effects of specific molecules such as cytokines, growth factors, and nutrients, since it has a defined medium composition and other serum components do not influence the experiment. This chapter describes the basic culture protocol for growing thymic epithelial cells (TECs), including the dissection of thymus from newborn mice, pretreatment for culture, and culture methods using serum-free media.

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Materials Newborn Mice

2.2 Preparation of Culture Wells

Mice should be systematically mated according to institutional guidelines (see Note 1). 1. Sterile culture wells: 35 and 60 mm plastic dishes or fourchamber slides. 2. Sterile Ca2+/Mg2+-free phosphate-buffered saline [1 × PBS (-)]. 3. 0.1% (w/v) gelatin: Cell culture grade gelatin should be used. Add 0.5 g of gelatin powder in a bottle containing 500 mL of purified water. Autoclave to dissolve gelatin completely and sterilize (see Note 2). Add 5 mL of 0.1% gelatin solution to a 60 mm plastic dish and incubate at 4 °C overnight. The recommended gelatin solution volumes are 10 mL for 100 mm dishes, 2 mL for 35 mm dishes, and 0.5 mL for 24-well plates and 4-chamber slides. Wash the culture wells with the same volume of 1× PBS (-) just prior to use. 4. Basic medium: 1:1 mixture of Ca2+-free DMEM and Ham’s F-12 nutrient mix supplemented with 2 mM L-glutamine, 1 mM sodium pyruvate, 100 U/mL penicillin, and 100 μg/ mL streptomycin. 5. Culture medium: Basic medium supplemented with 3 μg/mL recombinant human insulin (INS), 20 ng/mL recombinant human epidermal growth factor (EGF), 0.5 μg/mL hydrocortisone (HC), and 10 ng/mL cholera toxin (CT). For the preparation of 1000× stock solutions, mix together 3 mg/mL INS in basic medium, 20 μg/mL EGF in basic medium, 0.5 mg/mL HC in ethanol, and 10 μg/mL CT in purified water, and store at -20 °C. To make 1 mL of culture medium, add 1 μL each of 1000× growth supplement to 1 mL of basic medium just prior to use.

2.3 Dissection of Thymus from Newborn Mice

1. 70% (v/v) ethanol. 2. 60 mm plastic dishes. 3. Ice-cold, sterile Ca2+/Mg2+-free Hank’s balanced salt solution (HBSS). 4. Ice-cold, sterile 1× PBS (-).

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5. Two sets of sterile, regular, and microdissecting scissors and forceps: one for dissection and trimming the thymus and one for sterile use. 6. Stereomicroscope. 2.4 Enzymatic Dissociation of Thymus Fragments

1. Sterile 14 mL polypropylene tubes. 2. Micropipettes with 200 μL and 1000 μL tip: The tips should be cut at 2 mm. 3. Basic media supplemented 1 mg/mL DNase I and 1 mg/mL collagenase with low protease activity. 4. A water bath preheated at 37 °C.

2.5 Culture in Serum-Free Medium

1. 0.1% gelatin-coated sterile culture dishes: 35, 60 mm, chamber slides. 2. Sterile micro-forceps. 3. Micropipettes with 200 and1000 μL tips (see item 2, Subheading 2.4). 4. Culture medium (see item 5, Subheading 2.2). 5. Trypsin/ethylenediaminetetraacetic acid (EDTA). 6. 1 mg/mL trypsin inhibitor. 7. Incubator set at 37 °C and 5% CO2. 8. Refrigerated centrifuge precooled at 4 °C.

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Methods

3.1 Dissection of Thymus from Newborn Mice

The following operations should be performed under conditions as sterile as possible. 1. Prepare two sets of two 60 mm plastic dishes on ice with 5 mL of 1× PBS (-) and HBSS, respectively. 2. Sacrifice newborn mice (usually six to ten pups) by CO2 asphyxiation following institutional ethical regulations, and sterilize by immersion in 70% ethanol (see Note 3). 3. Remove the anterior thoracic wall with sterile scissors and forceps, and dissect the thymus under a stereomicroscope. Place the dissected thymus in a plastic dish containing ice-cold 1× PBS (-) to wash out the blood (Fig. 1). 4. Transfer the washed thymus to the next dish containing 1× PBS (-), and thoroughly remove connective tissue and capsule around the thymus with micro-forceps. Store the thymus in the next dish containing HBSS on ice. 5. Mince the trimmed thymus into 1–2 mm squares with sterile micro scissors in the dish containing HBSS.

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Fig. 1 Dissection of the thymus obtained from newborn mice. Panel A: front view of thoracic organs. Anterior thoracic wall has been removed. T: thymus; H: heart; L: lung; d: diaphragm; and li: liver. Panel B: dissected thymus (“T” in panel A) before trimming in 1× PBS (-). Bar = 3 mm

6. Collect the thymic fragments in a sterile 14 mL tube containing 5 mL of HBSS, and suspend by 15-gentle pipetting with a 1000 μL micro tip with the tip cut about 2 mm to release the bulk of thymocytes. 7. Stand the tube on ice for 5 min to allow the thymic fragments to settle. 3.2 Enzymatic Dissociation of Thymus Fragments

1. Transfer the supernatant containing thymocytes to another sterile tube. 2. Add 5 mL of 37 °C pre-warmed basic medium containing DNase I and collagenase to settled thymus fragments, and incubate for 30 min at 37 °C. Stir by ten-gentle pipetting every 5 min. 3. Transfer the supernatant to the tube of Step 1, and add 5 mL of new pre-warmed basic medium containing enzymes. Repeat three rounds of enzymatic dissociation (see Note 4). 4. Resuspend the resultant thymus fragments that are smaller in size in the 2 mL of culture medium.

3.3 Culture in Serum-Free Medium

1. Add growth supplements (EGF, INS, HC, and CT) to the basic medium stored on ice to prepare the required volume of culture medium. 2. Aspirate 0.1% gelatin in culture wells, wash wells with same volume of 1 × PBS (-) once, and add culture medium to each well. 3. Resuspend the enzymatically dissociated thymus fragments in culture medium, and add to the well in equal volumes. Place and arrange the thymus fragments evenly in a culture well with sterile micro-forceps (see Note 5).

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4. Start culture in 5% CO2 incubator at 37 °C. The initial medium change should be performed after 3 days, and half the medium volume should be exchanged every 2 days after that (see Note 6). 5. In the second or third week of culture, cells may be transferred to new matrix-coated culture wells by trypsinization. Aspirate the culture medium off, and add 0.5 mL of trypsin-EDTA to a 60 mm dish. Incubate the culture at 37 °C for 10–20 min. At this time, check whether the cells have detached from the bottom of the culture wells; if not, incubate the culture for a few more minutes. To stop the enzymatic reaction, add 0.2 mL of 1 mg/mL trypsin inhibitor to the culture. Serum-containing medium should not be used as an inhibitor of trypsin here. 6. Collect the detached cells by pipetting, and centrifuge at 1000–1200 rpm for 5 min at 4 °C. Resuspend cells in fresh culture medium and seed onto new matrix-coated culture wells at a density of 105 cells/mL.

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Notes 1. There is no significant difference in the proliferation of thymic epithelial cell from the thymus of mice up to 5 days old, but the authors use 3-day old mice for more efficient cell growth. In addition, pregnant mice can be purchased from animal suppliers. 2. Gelatin is used as a coating matrix for the adhesion of thymus fragments and cells to the dish in this protocol; type-I collagen can also be used [10]. Type-I collagen-coated culture wells can be purchased from a variety of suppliers. 3. Sacrificed mice should be wrapped in aluminum foil wiped with 70% ethanol and cooled in ice. Attention should be paid to the operator’s processing capacity and the number of mice to be processed. 4. The thymus fragments after the second enzymatic dissociation would be smaller because of the release of thymocytes. These should not be removed with supernatant containing thymocytes. Furthermore, the supernatant containing thymocytes which has been collected in a tube so far can be used for other analyses, such as flow cytometry. 5. An appropriate number of thymic fragments to be placed in culture wells may be 2 or 3 fragments in 4-chamber slides, 10–15 fragments in a 35 mm dish, and 20–30 fragments in a 60 mm dish. Depending on the number of placed thymus fragments, if the culture is successful, TECs may start expanding by day 2 or 3 and be confluent in 1 or 2 weeks (Fig. 2).

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Fig. 2 Proliferation of thymic epithelial cells on day 3 and 7 after the start of serum-free primary culture. Asterisks indicate enzymatically dissociated thymus fragments. Bar = 500 μm

6. Cultures should not be moved from incubator at least for the initial 48 h. The thymus fragments could detach if the medium shakes when it is removed from the incubator for observation. The reattachment of the thymus fragments is difficult, and detached fragments soon die. If the thymic fragments have not adhered and expanded at the bottom of the well 3 days after the start of the culture, the experiment failed. References 1. Abramson J, Anderson G (2017) Thymic epithelial cells. Annu Rev Immunol 35:85–118. https://doi.org/10.1146/annurev-immunol051116-052320 2. Sun TT, Bonitz P, Burns WH (1984) Cell culture of mammalian thymic epithelial cells: growth, structural, and antigenic properties. Cell Immunol 83(1):1–13. https://doi.org/ 10.1016/0008-8749(84)90219-3 3. Farr AG, Eisenhardt DJ, Anderson SK (1986) Isolation of murine thymic epithelium and an improved method for its propagation in vitro. Anat Rec 216(1):85–94. https://doi.org/10. 1002/ar.1092160115 4. Nieburgs AC, Picciano PT, Korn JH, McCalister T, Allred C, Cohen S (1985) In vitro growth and maintenance of two morphologically distinct populations of thymic epithelial cells. Cell Immunol 90(2):439–450. https://doi.org/10.1016/0008-8749(85) 90208-4 5. Ropke C, Elbroend J (1992) Human thymic epithelial cells in serum-free culture: nature and effects on thymocyte cell lines. Dev Immunol 2(2):111–121. https://doi.org/10.1155/ 1992/95098

6. Ro¨pke C (1997) Thymic epithelial cell culture. Microsc Res Tech 38(3):276–286 7. Sands SS, Meek WD, Hayashi J, Ketchum RJ (2005) Medium calcium concentration determines keratin intermediate filament density and distribution in immortalized cultured thymic epithelial cells (TECs). Microsc Microanal 11(4):283–292. https://doi.org/10.1017/ S1431927605050282 8. Gray DH, Chidgey AP, Boyd RL (2002) Analysis of thymic stromal cell populations using flow cytometry. J Immunol Methods 260(1–2):15–28. https://doi.org/10.1016/ s0022-1759(01)00493-8 9. Wang HX, Pan W, Zheng L, Zhong XP, Tan L, Liang Z, He J, Feng P, Zhao Y, Qiu YR (2019) Thymic epithelial cells contribute to thymopoiesis and T cell development. Front Immunol 10:3099. https://doi.org/10.3389/fimmu. 2019.03099 10. Ropke C, van Deurs B, Petersen OW (1990) Short-term cultivation of murine thymic epithelial cells in a serum-free medium. In Vitro Cell Dev Biol 26(7):671–681. https://doi. org/10.1007/BF02624423

Chapter 2 Cultured Pig Thyroid Follicular Cells: Electrical Evaluation of Epithelial Integrity Peying Fong Abstract Thyroid epithelial cells organize as enclosed follicles containing thyroid hormone precursor, iodinated thyroglobulin, with lumina bordered by the cellular apices. Transepithelial transport determines composition of compartmental milieu essential for both prohormone formation and its downstream conversion to thyroxine. Hence, not only do follicular lumina function as storage vessels but also as physiological reaction chambers into which reactive components, together with the proper salts and water, are secreted. Polarized, two-dimensional cultures of pig thyroid epithelia, prepared using established protocols, provide a convenient system for assessing transport processes subserving hormone production. This chapter details established methods for growing and evaluating integrity of primary pig thyroid cultures for downstream analysis of transport and other key physiological functions. Key words Thyrocyte, Apical, Basolateral, Transport, Resistance, Voltage, Current

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Introduction Epithelia organize as polarized arrays of cells joined by tight junctional complexes defining distinct apical and basolateral domains. This structural organization confers the ability to move solutes and water directionally – by either secretion or absorption – between their blood-facing (or interstitial) compartments and luminal compartments. In the case of epithelia that can be isolated from tissues as two-dimensional sheets, such as the intestine, colon, and bladder, both apical and basolateral sides can be accessed, enabling simple, yet powerful, experimental interrogation of secretory and absorptive processes. Studies of transport processes germane to thyroid function are limited by the three-dimensional, follicular organization of thyrocytes. In situ voltage- and ion-sensing microelectrode measurements can provide insights [1] but also are technically challenging and moreover limited by inaccessibility of the luminal compartment

Mario Baratta (ed.), Epithelial Cell Culture: Methods and Protocols, Methods in Molecular Biology, vol. 2749, https://doi.org/10.1007/978-1-0716-3609-1_2, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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to perfusion of experimental solutions. Thyrocytes grown on solid surfaces do develop polarity [2] but present the opposite problem of limiting accessibility to the basolateral aspect of the epithelium. Moreover, many epithelial cell types do not fully express transport proteins and polarize when grown on plastic but do when grown on permeable membrane supports [3]. Indeed, thyroid epithelial cells grown on permeable supports respond to thyroid-stimulating hormone (TSH) with iodide uptake and release [4, 5], permitting measurement of directional transport function [6, 7]. Early monolayer cultures of adult pig thyroids utilized fully dissociated cell suspensions [4, 6, 7]. Taking into account the greater proliferative capacity of neonatal and juvenile tissue and their potential for infiltration by fibroblasts introduced by complete cell dissociation procedures [8], we implemented isolated follicles for cultivating monolayer cultures of neonatal pig thyroids [9]. We based our procedures on the follicle isolation methods for culture of human thyroid [10], as well as adult pig [5]. This approach therefore favored development of high electrical resistance suitable for transport measurements [5]. Indeed, polarized thyrocyte epithelial monolayers derived from genetically engineered neonatal pigs proved indispensible for investigations seeking to understand the impact of these genetic manipulations on directional transport [11, 12]. Pig thyrocytes grown on permeable supports render thyroid epithelia suitable for measurements of transepithelial voltage, resistance, and short-circuit current. They also lend themselves well to tracer-based measurements of transport [5, 13], as well as studies of membrane polarization and subcellular trafficking using microscopic, cell biological and biochemical assays [6, 7, 11, 12]. In addition to reviewing our published protocol for culturing thyrocytes [9], the present chapter details a method for evaluating epithelial integrity necessary for terminal transport measurements. It serves as a functional tool for verifying development of monolayer polarity, regardless of the downstream application.

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Materials Review the following standard procedures for handling of reagents and materials. • All surgically collected tissues are placed into sterile, ice-cold media and transported to the laboratory on ice. • Solutions and growth media are stored at 4 °C and warmed to 37 °C before use. • Serum and antibiotic stocks are aliquoted and stored at -20 °C to avoid multiple freeze-thaw cycles. Aliquots of stocks are thawed and mixed immediately before preparation of media.

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• Store powdered enzymes at 4 °C in tightly lidded secondary containers filled with desiccant. • Media for tissue culture, as well as for solutions for measuring monolayer electrical parameters, are sterilized by passing through disposable vacuum filtration units fitted with 0.22 μm pore size polyethersulfone membranes. • For cell culture procedures, all plasticware and dissection instruments are sterile. • Comply with all approved local safety procedures for disposal of biological waste, plasticware, and sharps. 2.1 Solutions for Thyroid Follicle Isolation and Culture

Life Technologies supplies both liquid cell culture media and sera for our cell culture protocol. The compositions listed below correspond to their formulations. • Hank’s buffered saline solution (HBSS): 137 mM NaCl, 4.17 mM NaHCO3, 5.33 mM KCl, 0.44 mM KH2PO4, 0.34 mM Na2HPO4, 0.41 mM MgSO4.7H2O, 0.49 mM MgCl2.6H2O, 1.26 mM CaCl2, 5.56 mM D-glucose. If necessary, filter-sterilize the solution. Store at 4 °C. • Ca2+/Mg2+-free HBSS: 137 mM NaCl, 4.17 mM NaHCO3, 5.33 mM KCl, 0.44 mM KH2PO4, 0.34 mM Na2HPO4, 5.56 mM D-glucose. Filter-sterilize if necessary and store at 4 °C. • Tissue wash solution (TWS): HBSS containing penicillinstreptomycin (50 U/mL and 50 μg/mL, respectively). • Tissue dissociation solution (TDM): On the day of undertaking thyroid follicle isolation, mix Ca2+/Mg2+-free HBSS with collagenase type A (3.9 Wu¨nsch U/50 mL) and dispase (50 mg/ 50 mL) (see Note 1). Sterilize by filtration, and place in 37 °C water bath. Fifty milliliters (50 mL) is sufficient for completing isolation from one thyroid (see Note 2). • Collagenase type A (from Clostridium histolyticum). • Dispase (neutral protease from Bacillus polymyxa). • Dulbecco’s Modified Eagle Medium (DMEM): This base medium (Life Technologies #11965) contains 110.34 mM NaCl, 44.05 mM NaHCO3, 5.33 mM KCl, 0.81 mM MgSO4 (anhydrous), 1.80 mM CaCl2, 2.48 mM Fe(NO3)3.9H2O, 25 mM D-Glucose, and 0.04 mM phenol red. The formulation includes a mixture of vitamins and amino acids, including 4.0 mM L-glutamine (see Note 3). In a 5% CO2 atmosphere, pH is 7.4. • Fetal bovine serum (FBS): The preferred product is Life Technologies #26140079 (see Note 4).

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• Thyroid-stimulating hormone stock (TSH, 1 IU/mL, 1000×): Dissolve 10 IU in 10 mL HBSS with 0.1% bovine serum albumin to prepare TSH stock. Store frozen at -20 °C in conveniently scaled aliquots. • Growth medium (GM): This comprises DMEM supplemented with FBS (10%), thyroid-stimulating hormone (final concentration, 1 IU/L), and penicillin-streptomycin (50 U/mL and 50 μg/mL). Mix components, filter to sterilize, and store at 4 °C. • Antiseptic surgical scrub (such as Betadine): use if surgically excising thyroids from freshly euthanized animals [9]. 2.2 Plasticware and Dissection Tools

Obtain the following items as pre-packaged, sterile units: • Filtration units for sterilization of solutions (pore size, 0.22 μm): We typically use units suitable for filtering volumes ranging from 50 mL to 1 L. • Sterile serological pipettes. • Tissue culture dishes (100 mm diameter). • Sterile conical centrifuge tubes (15 and 50 mL). • Cell strainers with mesh pore size of ~100 μM. Units obtained from several sources yield comparable results. • Tissue culture-treated T25 flasks. • Permeable growth supports: Corning Costar Snapwell® inserts (12-mm-diameter membrane; 1.12 cm2 growth area) are the growth support of choice if the downstream use is continuous monitoring of epithelial transport via short-circuit current measurements (see Note 5). Snapwells are convenient for use in suitably modified Ussing chambers. The epithelial resistance procedure described uses a measuring chamber designed to accept Snapwell® inserts. • Scalpel blades. In addition, have available the following sterilized items: • Single-edged razor blades. • Blunt and fine forceps. • Dissection scissors. • Scalpel handle. • Weighing spatula. • Borosilicate Pasteur pipettes.

2.3 Required Equipment for Cell Culture

For the cell culture procedure, the following equipment is used: • Class II A2 biosafety cabinet fitted with a vacuum line. • 5% CO2, 37 °C humidified incubator.

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• Cell counter. • 37 °C water bath. • Automatic pipettor. • Dissection microscope and light source. • Inverted light microscope. • Cold blocks. • Tabletop centrifuge: A low-speed centrifuge with a rotor accommodating 50 mL conical tubes is adequate. 2.4 Solutions and Required Equipment for Measuring Epithelial Integrity

This protocol uses electrical resistance as a proxy for epithelial integrity. Measurements of transepithelial resistance use either a Hepes-buffered Ringer solution (HBRS) or a phosphate-buffered Ringer solution (PBRS). These single time-point measurements are performed at room temperature (see Note 6). • HBRS comprises 140 mM NaCl, 2.7 mM KCl, 1.8 mM CaCl2, 1.06 mM MgCl2, 12.4 mM Hepes (free acid), 5.1 mM glucose, pH 7.4 using NaOH. • PBRS contains 140 mM NaCl, 2.125 mM K2HPO4, 0.375 mM KH2PO4, 2 mM CaCl2, 1.2 mM MgSO4, 5 mM glucose. With this combination of di- and monobasic phosphate, pH should be close to 7.4. Adjust with NaOH if necessary. • Epithelial voltmeter (EVOM; World Precision Instruments): See Fig. 2 for an image of the front panel. • Electrode assembly of choice (EndOhm-24SNAP) and connectors. Figure 3c details the EndOhm-24SNAP chamber electrode arrangement. • Small (2.5 mm blade) flathead screwdriver. • Two blank Snapwell® inserts: Mark one to be used under non-sterile conditions and the other to be used under sterile conditions. • Blunt curved forceps for transferring Snapwells to EndOhm24SNAP. • Full-strength household bleach (5% sodium hypochlorite). • 70% ethanol/30% double-distilled water.

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Methods Adhere to aseptic technique whenever possible throughout the procedure. Undertake tissue mincing and follicle isolation, and culture in a Class II A2 Biological Safety Cabinet. Maintain cultures in a humidified 5% CO2 atmosphere, at 37 °C.

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For the novice, conceptualize the protocol by considering the four general processes, described by Wills [14]. These are tissue isolation, mechanical and enzymatic dissociation of the tissue, selection for the proliferative units desired, and seeding. 3.1 Transporting Freshly Excised Thyroid Glands

If thyroids are obtained locally, ensure that all protocols and procedures for handling and euthanasia adhere to approved institutional protocols for animal care and use. Transfer excised thyroids to prechilled sterile HBSS, then seal containers tightly, and place on ice for transport to the laboratory. Thyroids tolerate overnight storage at 4 °C before successful follicle harvest. We find that thyroids shipped on ice by overnight express can produce similar results. All volumes stated in subsequent sections of this protocol are scaled for processing of one thyroid. For each thyroid harvested, aseptically dispense 30 mL HBSS into a labeled 50 mL conical tube and keep chilled on ice (see Note 7).

3.2 Coarse Tissue Mincing

• Transfer the tube to the biosafety cabinet, and rinse the thyroid three times with sterile TWS. Aspirate TWS completely between rinses. • Place the thyroid into a fresh 100 mm culture plate. • Mince the thyroid into ~2 mm × 2 mm pieces using a singleedged razor blade (see Note 8). • Using a sterile weighing spatula, transfer the minced thyroid pieces into a 15 mL conical tube containing sterile TWS. Rinse by shaking, allowing the pieces to settle, and removing excess TWS. • Repeat the washes, shaking well in between, for a total of five washes (see Note 9).

3.3 Isolation of Follicle Fragments

• Remove the HBSS from the last wash and rinse three times with Ca2+-Mg2+-free HBSS. • After the last rinse, add 10 mL of warmed TDM. Incubate in a 37 °C water bath. Agitate intermittently (every 15 min) by briskly shaking for ~20 s. Allow digest to proceed 1 h. • At the 1-h time point, shake again before allowing the pieces to settle. Collect the supernatant (see Note 10) and transfer to a sterile 15 mL tube, marking this as fraction 1 (F1). Note the volume collected, and then neutralize the enzymes by adding FBS (0.5%; for a 10 mL collection, add 50 μL). Set the tube containing the collected F1 on ice. • Add 10 mL fresh TDM into the tube containing the residual loosely pelleted tissue fragments. Shake well and return the tube to the 37 °C water bath. Allow the digest to proceed above for an additional 30 min. Shake vigorously at 15 min and again at 30 min.

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• Collect the second fraction (F2), as described above (see Note 11). Repeat the procedure until no solid tissue remains. • Pool all collected fractions and centrifuge at 200 g × 5 min. Discard the supernatant and suspend the pellet in 10 mL DMEM. • Place the suspension on ice for 1 h to sediment the purified follicle fragments. • Prepare Snapwell® plates by pipetting pre-warmed GM into the lower, and then upper, chambers. Add 1.8 mL to each lower chamber and 0.5 mL to each upper chamber. Keep warm and equilibrate pH by transferring to a humidified, 37 °C, 5% CO2 incubator. • The follicles settle into a loose pellet in ~1 h. Carefully remove the supernatant. • Add 10 mL fresh DMEM and resuspend the purified follicles. • Place a 100 μm mesh cell strainer into a 50 mL conical tube. Slowly filter 5 mL of the suspension through this assembly. Rinse the strainer with 5 mL DMEM. The final volume will be ~10 mL. • Place a second strainer into another 50 mL conical tube, then filter the remaining 5 mL follicle suspension and rinse with 5 mL DMEM. You will now have two 50 mL tubes, containing ~10 mL of 0.5× follicle suspension (~5 mL initial suspension +5 mL DMEM) in each (see Note 12). Ensure both tubes contain equal volumes; if not, balance before the next step. • Spin the filtered suspension at 200 g × 5 min. Remove the supernatant, add fresh DMEM, and repeat the wash. • Resuspend the final pellets in 1–4 mL GM (see Note 13). Figure 1a is a light micrograph showing a preparation satisfactorily depleted of single cells and enriched in follicles. 3.4 Counting and Seeding Follicle Fragments

• Count the follicles and estimate the total yield. Remove the medium from the upper chambers before seeding >1 × 104 follicles (or fragments)/Snapwell® (see Note 14). For example, if plating on Snapwell® inserts (1.12 cm2 growth area), plate ~1.25 × 104 fragments/well. Follicles also can be used directly for immunofluorescence staining and confocal imaging (Fig. 1c). • Refresh GM 24 h after seeding. Remove media from lower chamber first before removing from upper chamber. When replenishing GM, reverse the order: upper chamber first, then lower chamber (see Note 15). Feed the cells every 2 days thereafter. Juvenile pig thyroid cultures studied between days 12 and 18 post-seeding consistently produce transepithelial resistances (Rte > 1000 Ohms.cm2) suitable for measurement of shortcircuit current in Ussing chambers. Preparations from adult

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Fig. 1 Light microscopic images shown in panels A and B document follicle release during isolation procedure. (a) shows open follicles released by enzymatic fractionation. Note partially open follicle at left, marked with the arrowhead; scale bar: 50 μm; original image acquired at 40× magnification. (b) provides a closer view of the open follicle shown within area of Panel A marked by square; image acquired at 100× magnification. Note the delineation of follicular edges by cuboidal epithelial cells. Visualization of tight junctions in panels C and D: (c) acutely isolated neonatal pig thyroid follicle, rabbit anti-ZO-1 primary antibody, donkey anti-rabbit AlexaFluor 594 secondary antibody. (d) Confluent pig thyroid epithelial monolayer, rabbit anti-ZO-1 antibody, donkey anti-rabbit Cy3-labeled secondary

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pig thyroids can yield higher Rte. Polarized monolayers grown on Snapwell® Clear supports lend themselves well to visualization by confocal microscopy, and ZO-1 staining confirms preservation of ability to form tight junctions (Fig. 1d). 3.5 A Simple Tool for Assessing Epithelial Integrity

A choice of methods exists for assessing epithelial integrity. Each investigator should determine which best suits their anticipated downstream application. These options, described by [14], include assessment of dye permeation by absorbance measurement, as well as measurement of epithelial resistance. Our lab favors the latter, as our cultures are destined for transport studies using open- and short-circuit current measurements. The following is the procedure our lab uses to assess epithelial resistance using an epithelial voltmeter (EVOM; World Precision Instruments, Sarasota, FL), fitted with an EndOhm-24SNAP electrode chamber (also from World Precision Instruments) (see Note 16). This protocol customizes for our applications the instructions furnished by the supplier, incorporating specific details, adjustments, and notes. Technical data are available by consulting the supplier’s device manuals. So performed, the resultant resistance measurements are particularly useful for evaluations of primary pig thyroid cultures obtained from neonatal to adult pigs. • Test the EVOM before use: Switch “Mode” to R (resistance) setting, power on, and then press “Test R.” The meter should read 1000 Ω or 1 kΩ, depending on the “Range” toggle setting (either 2000 Ω or 20 kΩ, respectively; see Note 17). • Position the upper electrode of the EndOhm-24SNAP chamber: Place a blank Snapwell® (“dummy”) into the EndOhm24SNAP. Loosen the nut on the cap, and then adjust the upper electrode height by rotating the cap so that there is 1–2 mm clearance between the electrode and the membrane. Secure the nut. There should be no need to change the position once set, assuming exclusive and continued use of Snapwell® supports. • Inspect the electrode surfaces. If they show shiny patches, they require re-chloriding. We use a disposable plastic Pasteur pipette to apply full-strength household bleach directly over the bottom electrode assembly (Fig. 3a; see Note 18). Invert the upper electrode assembly, carefully balance on the connector end, and apply a small volume on the electrode (Fig. 3b). Allow 15 min, and then rinse all surfaces thrice with distilled water. • Sterilize the equipment by wiping surfaces well with 70% ethanol/30% double-distilled water (see Note 19). • Transfer EVOM, EndOhm-24SNAP, and connectors to biosafety cabinet (i.e., if there is intention to perform repeated measurements over time to monitor development of resistance). This step is not strictly necessary if measurements are terminal.

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• Rinse the EndOhm-24SNAP well with sterile measuring solution. Aspirate to remove. • Test the electrodes: Fill the cup with ~3 mL of HBRS (or PBRS), and ensure there is fluid continuity between the top and bottom electrodes. • On the EVOM, set “Range” to 2000 Ω, and turn on the power. The meter should read 0. If it does not, use a small flathead screwdriver to adjust the “Zero Ω” screw. • Once the meter reads zero, press “Measure R” (see Note 20). This reflects the fluid resistance in the compartment. • Power off the EVOM. • Correct for blank membrane resistance: Aspirate to remove the solution. Insert a dummy Snapwell® into the chamber, and pipette 0.25 mL HBRS or PBRS into the top, then 2.5 mL to the bottom, chamber. Ensure there are no bubbles. • The expected background resistance will be low, so “Range” remains at the 2000 Ω setting (see Note 21). • Power on the EVOM and push the “Measure R” button. This is the background resistance of the blank membrane filter. In HBRS, it should be ~20 Ω. Record the measured value. • Switch off the EVOM before removing the blank insert. It is a good idea to have the electrodes in contact with fresh HBRS or PBRS. • The system now is ready to measure resistance of the epithelial cultures. • Prepare the culture inserts for measurement: Remove growth medium from both chambers. After washing both chambers twice with the measuring solution of choice (HBRS or PBRS) at room temperature, refresh the top chamber with 0.25 mL measuring solution. • With the EVOM switched off, replace the media in the EndOhm bottom chamber with 2.5 mL HBRS or PBRS. • Using a pair of blunt, curved forceps, transfer the culture insert (already containing 0.25 mL apical solution) directly to the EndOhm-24SNAP, ensuring there are no bubbles trapped underneath the insert as you do so. Place the upper electrode cap assembly over the Snapwell®, and check the top electrode makes contact with the apical solution. • Ensure that mode is set on R and the correct range is selected (see Note 21). • Turn on the EVOM, and push the “Measure R” button. • Record the displayed value.

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• Switch the unit off. Remove the Snapwell® and replace in its original plate well. Aspirate the measuring solution from the EndOhm lower chamber, and then replace with fresh solution. Repeat the procedure for all samples. • After the final measurement, perform another background resistance measurement on the blank Snapwell®. • If further growth is intended for measured cultures, aspirate off measuring medium, and replace with GM before returning to incubator. • Switch off the meter, detach the connectors, aspirate recording solution from EndOhm-24SNAP chamber, and then rinse electrode surfaces and chamber interior three times with doubledistilled water. Dry by blotting gently with a KimWipe, and then allow to air dry before storage. • Average the background resistance measurements taken before and after epithelial monolayer measurements. Make sure to subtract the average background resistance from values recorded for epithelial monolayers. From the corrected resistance values, calculate the unit area resistance (see Note 22).

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Notes 1. Note the lot number and activity of Collagenase A from the information supplied by the provider. Use this information to calculate the appropriate mass prior to preparing TDM. 2. Do not save TDM for later use. Discard any that remains at the end of the procedure. 3. DMEM (Life Technologies #11965) contains the following amino acids (in mM): 0.4 glycine, 0.4 L-arginine hydrochloride, 0.2 L-cystine.2HCl, 4 L-glutamine, 0.2 L-histidine hydrochloride.H2O, 0.8 L-isoleucine, 0.8 leucine, 0.7 L-lysine hydrochloride, 0.2 L-methionine, 0.4 L-phenylalanine, 0.4 Lserine, 0.8 L-threonine, 0.08 L-tryptophan, 0.4 L-tyrosine disodium salt dehydrate, 0.8 L-valine. Vitamins contained in this product are (in μM) 28.6 choline chloride, 8 D-calcium pantothenate, 9 folic acid, 32.8 niacinamide, 19.4 pyridoxine hydrochloride, 1.1 riboflavin, 11.9 thiamine hydrochloride, 40 i-inositol. A powdered form of this medium is also available (Life Technologies #12100). 4. FBS growth factor composition varies by lot. Fortunately, pig thyroid cultures do not appear to be sensitive to this, and we find that multiple lots of Life Technologies #26140079 yield comparable results. FBS from other suppliers likely will yield similar results.

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5. Two versions of Corning Costar Snapwell® inserts (0.4 μm pore size) are available. Cells attach well to both without the need for collagen pre-coating. If the downstream application is limited to transport experiments requiring rapid diffusion from the serosal chamber, the translucent polycarbonate (PC) membrane Snapwell® inserts (Corning 3407; 1 × 108 pore density) carry an advantage over Snapwell® Clear (Corning 3801), a similar product fitted with transparent polyethylene terephthalate (PET) membranes that have a lower pore density, 4 × 106. This difference does not appreciably affect monolayer growth properties and development of electrical resistance. However, the latter can be useful if transport measurements, followed by post-experimental staining and microscopy, are part of the experimental design. Snapwell® inserts comprise two parts: a scaffolding support and a shallow cup (see Fig. 3c for a schematic). The cup incorporates the growth membrane. It can be detached readily without edge damage to the monolayer and then directly positioned into Ussing chambers for transport measurements. Transwell Clear® (Corning 3460) is fitted with the same membrane as Snapwell® Clear. Although Transwells do not have the benefit of a two-piece, pull-apart support, overall they are more economical when the sole downstream purpose involves membrane excision, such as immunostaining and microscopic visualization. Resistance measurements of epithelial monolayers grown on Transwell Clear® supports require EndOhm-12, a measuring chamber having a geometry differing slightly from that of EndOhm-24Snap. 6. Both Hepes- and phosphate-buffered Ringer solutions are convenient for performing EVOM measurements, allowing them to be conducted in room air. GM, which utilizes the HCO3-/CO2 buffering system, requires maintenance of CO2 tension. The use of solutions containing alternative buffers renders this condition dispensible. Another advantage of using HBRS and PBRS in resistance measurements is their lack of FBS; this minimizes protein accumulation on electrodes. Because measurements are performed at room temperature, resistance values are expected to be slightly greater than those measured at 37 and 39 °C, temperatures used to perform short-circuit and open-circuit current measurements on pig thyroid epithelial cultures [11, 12]. HBRS and PBRS are not recommended for long-term maintenance in culture. If monolayers are to be returned to the incubator for growth, change media back to GM as soon as possible. 7. Use of individual tubes enables easy tracking of thyroids from different experimental groups.

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8. These dimensions serve primarily as guidelines. Finely mincing the tissue increases surface to volume ratio and promotes more efficient enzymatic dissociation. 9. Repeated washes effectively remove blood cells from the preparation. 10. The F1 contains primarily connective tissue remnants, single cells, a minority population of follicles, and thyroglobulin. These components contribute to the murky appearance of F1. Depending on the thoroughness of the initial cleaning (Subheading 3.2), excess fat may also visibly float at the surface. 11. Follicles and follicle fragments are most apparent in F2 and F3. One can monitor follicle release by pipetting a small volume (< 20 μL) onto a tissue culture dish and viewing on an inverted microscope at low power. 12. In some preparations, residual connective tissue fragments in the fractions may clog cell strainer filters, obstructing passage. Resist the temptation to add suspension to the strainer too quickly. It may be necessary to use additional strainers. Alternatively, we find it helpful to have sterile fine forceps and a small culture plate on hand. Carefully remove visible connective tissue fragments from the strainers with the forceps. Transfer these fragments to the culture plate for disposal at the end of the procedure. 13. To confirm depletion of single cells, dispense a small volume of the isolate onto a 35 mm tissue culture dish and inspect by light microscopy. If single cells are present, incorporate a 30-min- to 1 h-long pre-plate step. Single cells, and in particular fibroblasts, adhere to tissue culture-treated plastic within this period. Remove the follicle and fragment suspension for counting and seeding. 14. As an alternative to seeding directly on permeable supports, cultures may be grown in T25 flasks (~2.5 × 104 follicles per 25 cm2). At 80% confluence, passage the primary cultures to P1. If using Snapwells®, plate at ~1 × 105 cells/cm2. In our experience, P1 cells display electrical properties and pharmacological responsiveness comparable to primaries grown by seeding follicles and follicle fragments. 15. To minimize changes in hydrostatic pressure that may disrupt monolayer integrity, adhere to the stated order of media removal and replenishment. 16. The EVOM readout (resistance) reflects the voltage change measured upon application of a fixed square current pulse to the monolayer. The EndOhm-24SNAP is fitted with fixed current and voltage electrodes to measure the apical and basolateral potentials, respectively. Its fixed geometry eliminates

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Display panel: Shows value of parameter measured

Measure R: push to measure resistance

Zero Ω: correct background resistance Mode: switch between resistance and voltage measurements

Test R: Not for measurement; use only for instrument testing Range: Toggle to select range Zero V: not used for resistance measurements Power switch

Electrode Port: Connection for electrode assembly

Fig. 2 EVOM front panel; annotations in black font are described in protocol. Note that the “Test R” function (in blue) is used only for initial instrument testing

variability encountered with using “chopstick”-style paired electrodes. In the latter case, deviation from precise positioning results in inconsistent sensing of the current pulse when the “Measure R” function is chosen. For further discussion, see [14]. 17. Do not use the “Test R” function at any other time during resistance measurement; use it only during initial instrument testing. Refer to information on Fig. 2. 18. A properly chlorided electrode will not be shiny and will have a uniform, dull brownish cast. Apply bleach to re-chloride the electrode, as described in the procedures. Applying with a plastic Pasteur pipette gives adequate control without risk of electrode surface scratching (Fig. 3). 19. Sterilization and performing measurements with sterile technique are important if repeated measurements on the same monolayers are desired. Do not use organic solvents other than 70% ethanol/30% water, and make sure to rinse the EndOhm chamber well with water to remove any residual ethanol. 20. Do not confuse “Measure R” with the “Test R” function; see Note 17.

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B

Voltage electrode

Current electrode

C Upper electrode connector

Snapwell® insert • Support • Membrane “cup”

Lower electrode connector Fig. 3 (a) shows overhead view of bottom chamber and electrode pair at center. Note bleach applied over electrode surface for purpose of chloriding. The brown spot in the center is the voltage electrode, and the larger brown circle encircling it is the current electrode. (b) provides a side view of the upper chamber assembly, inverted so that its electrode assembly can be chlorided. The surface tension of the bleach (white arrow) ensures only the electrode region is exposed. Although of smaller dimension, the arrangement of the voltage and current electrodes mirrors that shown for the bottom chamber (see inset). (c) diagrams the measuring arrangement. Note the positioning of the Snapwell® in the chamber; the solution volume in the bottom chamber should contact the bottom of the Snapwell® membrane, but the level must not exceed the level with the well itself. The upper electrode level is preset so that it contacts the solution in the well without touching the monolayer itself. The upper and lower electrodes connect to the EVOM via the electrode port

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21. There are two resistance ranges available: 2000 Ω and 20 kΩ. In general, the first is adequate for neonatal cultures up to 21 days. This can be useful if the investigator wishes to perform a time course charting acquisition of functional polarity. Adult pig thyroid can achieve resistances in excess of 2000 Ω. If these are being measured more than 14 days after seeding, choose the 20 kΩ setting. 22. Readings reflect the area of Snapwell® membrane supports (1.12 cm2). To enable comparisons of data obtained from monolayers grown on different sizes of supports, the convention uses unit area resistance, expressed as Ω cm2. Obtain the unit area resistance by multiplying the background-corrected meter readings by 1.12 cm2. For example, a measurement of 1000 Ω taken from a monolayer grown on a Snapwell® has a unit area resistance of 1120 Ω cm2.

Acknowledgments I thank the Department of Anatomy and Physiology at Kansas State University College of Veterinary Medicine for supporting ongoing studies in my laboratory. References 1. Chow SY, Kunze D, Brown AM, Woodbury DM (1970) Chloride and potassium activities in luminal fluid of turtle thyroid follicles as determined by selective ion-exchanger microelectrodes. Proc Natl Acad Sci U S A 67(2): 998–1004. https://doi.org/10.1073/pnas. 67.2.998 2. Mauchamp J, Margotat A, Chambard M, Charrier B, Remy L, Michel-Bechet M (1979) Polarity of three-dimensional structures derived from isolated hog thyroid cells in primary culture. Cell Tissue Res 204(3):417–430. https://doi.org/10.1007/BF00233653 3. Steele RE, Preston AS, Johnson JP, Handler JS (1986) Porous-bottom dishes for culture of polarized cells. Am J Phys 251(1 Pt 1):C136– C139. https://doi.org/10.1152/ajpcell. 1986.251.1.C136 4. Chambard M, Verrier B, Gabrion J, Mauchamp J (1983) Polarization of thyroid cells in culture: evidence for the basolateral localization of the iodide “pump” and of the thyroid-stimulating hormone receptor-adenyl cyclase complex. J Cell Biol 96(4):1172–1177. https://doi.org/ 10.1083/jcb.96.4.1172 5. Nilsson M, Bjorkman U, Ekholm R, Ericson LE (1990) Iodide transport in primary

cultured thyroid follicle cells: evidence of a TSH-regulated channel mediating iodide efflux selectively across the apical domain of the plasma membrane. Eur J Cell Biol 52(2): 270–281 6. Armstrong J, Matainaho T, Cragoe EJ Jr, Huxham GJ, Bourke JR, Manley SW (1992) Bidirectional ion transport in thyroid: secretion of anions by monolayer cultures that absorb sodium. Am J Phys 262(1 Pt 1):E40–E45. https://doi.org/10.1152/ajpendo.1992.262. 1.E40 7. Armstrong JW, Cragoe EJ Jr, Bourke JR, Huxham GJ, Manley SW (1992) Chloride conductance of apical membrane in cultured porcine thyroid cells activated by cyclic AMP. Mol Cell Endocrinol 88(1–3):105–110. https://doi. org/10.1016/0303-7207(92)90014-w 8. Dumont JE, Lamy F, Roger P, Maenhaut C (1992) Physiological and pathological regulation of thyroid cell proliferation and differentiation by thyrotropin and other factors. Physiol Rev 72(3):667–697. https://doi.org/10. 1152/physrev.1992.72.3.667 9. Lillich JD, Fong, P. (2018) Isolation and culture of juvenile pig thyroid follicular epithelia. In: Baratta M (ed) Epithelial cell

Cultured Pig Thyroid Follicular Cells: Electrical Evaluation of Epithelial. . . culture: methods and protocols. methods in molecular biology, vol 1817. Methods in molecular biology. Humana Press, New York, p 9–18 10. Williams DW, Wynford-Thomas D (1997) Human thyroid epithelial cells. Methods Mol Biol 75:163–172. https://doi.org/10.1385/ 0-89603-441-0:163 11. Li H, Ganta S, Fong P (2010) Altered ion transport by thyroid epithelia from CFTR-/pigs suggests mechanisms for hypothyroidism in cystic fibrosis. Exp Physiol 95(12): 1132–1144. https://doi.org/10.1113/ expphysiol.2010.054700 12. Li Y, Ganta S, Fong P (2012) Endogenous surface expression of ΔF508-CFTR mediates

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cAMP-stimulated cl- current in CFTRΔF508/Δ pig thyroid epithelial cells. Exp Physiol 97(1):115–124. https://doi.org/10.1113/ expphysiol.2011.060756 13. Ericson LE, Nilsson M (2000) Deactivation of TSH receptor signaling in filter-cultured pig thyroid epithelial cells. Am J Physiol Endocrinol Metab 278(4):E611–E619. https://doi. org/10.1152/ajpendo.2000.278.4.E611 14. Wills NK (1996) Epithelial cell culture. In: Wills NK, Reuss L, Lewis SA (eds) Epithelial transport: a guide to methods and experimental analysis. Chapman and Hall, London, pp 236–255 F508

Chapter 3 Human Minor Salivary Glands: A Readily Available Source of Salivary Stem/Progenitor Cells for Regenerative Applications Caitlynn M. L. Barrows, Danielle Wu, Simon Young, and Mary C. Farach-Carson Abstract Resident stem/progenitor cells within the secretory salivary glands offer a potential therapeutic resource for use in the regeneration of salivary glands needed to restore saliva production in patients with chronic xerostomia, or dry mouth. Methods were developed previously to isolate human stem/progenitor cells (hS/PCs) from major salivary glands (parotid/submandibular). Abundant minor salivary glands located in readily accessible locations in the oral cavity and lip could provide an additional valuable therapeutic resource. An advantage of this cell resource is that these minor glands about the size of grape seeds can be harvested from healthy donors using minimally invasive surgical procedures. The disadvantage of using minor glands is that they contain many fewer cells than do major glands, and thus harvested cells need to be expanded in the lab to create a therapeutic resource. While earlier work has described isolation of proliferative cell populations from minor salivary glands that could be used in regenerative medicine, most of these expanded cells possess properties of mesenchymal cells rather than the epithelial population that secretes salivary products. Here, we describe in detail our recently established methods to isolate and expand hS/PCs isolated from human labial minor salivary glands. Expanded hS/PC populations are epithelial assessed by their expression of epithelial progenitor markers K5 and K14. Like expandable cell populations previously isolated from the major salivary glands, these cells also express nuclear p63, consistent with their ability to be expanded after explant culture. When hS/PCs with these properties are encapsulated into a customized 3D biomimetic hyaluronic acid-based hydrogel, they will assemble into microstructures that retain some progenitor markers while also beginning to differentiate. The increased expression of secreted mucin MUC-7 was used to demonstrate differentiation and secretory potential in assembled hS/PC microstructures. Compared to hS/PCs from major glands, those from minor salivary glands tend to be more heterogeneous in early passage; thus, use of K5/K14/p63 as an early quality assessment tool is highly recommended. Additionally, hS/PCs from minor glands are sensitive to stress and if mishandled will demonstrate a stress response that leads to their transitioning to a flat, squamous cell-like appearance that is of limited utility in regenerative medicine applications. We conclude that properly handled hS/PCs from minor salivary glands represent a powerful new source of therapeutic cells for applications including treating patients with chronic xerostomia. Key words Salivary gland, Minor salivary glands, Xerostomia, Cell isolation, Regenerative medicine, 3D hydrogel culture

Mario Baratta (ed.), Epithelial Cell Culture: Methods and Protocols, Methods in Molecular Biology, vol. 2749, https://doi.org/10.1007/978-1-0716-3609-1_3, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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Introduction We previously reported our methods for the isolation of primary human stem/progenitor cells (hS/PCs) from major salivary glands, parotid and submandibular, as a cell source for salivary tissue engineering and restoration of salivation for patients with chronic xerostomia associated with hyposalivation disorders [1, 2]. The major salivary glands reside deep in the maxillofacial region and contain abundant nerves and blood vessels, making them challenging to reach surgically for tissue harvest. Methods we used to date thus typically involved harvesting normal tissue regions from donors undergoing surgical resection for pathological conditions [1]. This offers a resource-dependent workflow to harvest normal hS/PCs for regenerative medicine applications. In contrast, minor salivary glands are numerous; humans have approximately 800–1000 distributed throughout the oral cavity, with many residing in easily accessible areas including the buccal mucosa, labial mucosa, lingual mucosa, soft/hard palate, and floor of mouth [3]. This feature, along with their small grapeseed-like structure within a well-contained membranous capsule, makes these miniglands a useful source for regenerative applications. Here we describe a method that we developed to isolate and expand hS/PCs harvested from minor labial salivary glands from consented, cancer-free donors who are being treated for acute injuries to the maxillofacial complex. Compared to donors who consent for normal region tissue harvest from major glands with pathological conditions, donors for minor salivary gland harvest tend to be younger and healthier, thus making these cells an excellent source for regenerative cell therapies. We describe the methods we used to demonstrate that the phenotype of hS/PCs derived from minor salivary glands is similar to those isolated previously from major glands including cytokeratin profiling and staining for the cell expansion marker p63 and is not primarily mesenchymal as has been reported for other cell preparations from minor glands [4]. We also report how we encapsulated minor gland-derived hS/PCs into hyaluronic acid (HA)-based hydrogels for threedimensional (3D) culture showing their potential for cell assembly into microstructures [1] and mucin 7 production, which is part of the normal salivary proteome [5, 6].

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Materials

2.1 Human Minor Salivary Gland Harvest and Handling

• Tissue handling and processing: small, sharp, stainless-steel scissors and forceps, P1000 pipette plus sterile tips. • General rinse solution: Ca2+- and Mg2+-free phosphate buffered saline (1×PBS).

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• Wash solution #1: ice-cold DMEM/F-12 medium, supplemented with 1% (v/v) penicillin-streptomycin, 1% (v/v) amphotericin B, 1% (v/v) Betadine solution (from 10% povidone iodine stock), and sterile filtered with a 0.22 μm polyethersulfone (PES) filter. • Wash solution #2: ice-cold DMEM/F-12 medium supplemented with 1% (v/v) penicillin-streptomycin and 1% (v/v) amphotericin B and sterile filtered with a 0.22 μm PES filter. • Cell culture disposables: 10 cm petri dishes, T-25 culture flasks, 1.5 mL Eppendorf tubes. • Cell passage: 0.125% (v/v) trypsin-EDTA, 1 mg/mL (w/v) soybean trypsin inhibitor. • Humanized salivary media (HSM): Williams’ Medium E base without L-glutamine (Sigma, W4128), 1 mg/mL (w/v) albumin from human serum (Sigma, A1887), 1% (v/v) penicillinstreptomycin, 1% (v/v) CTS™ GlutaMAX™ -I Supplement, 1% (v/v) insulin-transferrin-selenium (InVitria, 777ITS032), 0.1 μM (v/v) dexamethasone, 10 ng/mL (v/v) human epidermal growth factor (hEGF) (Gibco, PHG0311). 2.2 Hydrogel Preparation and Cell Encapsulation

• Peptide synthesis reagents: matrix metalloproteinase (MMP)cleavable peptide (“PQ”) (GenScript, KGGGPQGIWGQGK) and integrin binding peptide (RGD) (GenScript, GRGDS), acrylate-polyethylene glycol (PEG)-succinimidyl valerate (Laysan Bio, ACRL-PEG-SVA-3400, 3400 g/mol), 1 N sodium hydroxide (NaOH), Milli-Q® water. • N-(2-hydroxyethyl)piperazine-N′-(4-butanesulfonic acid) (HEPBS) buffer solution: [mM]: 20 HEPBS, 100 NaCl, 2 CaCl, 2 MgCl in Milli-Q® water, pH 8. • Peptide synthesis equipment and supplies: lyophilizer and FastFreeze® flasks and adapters, analytical balance, 50 mL amber conical tubes, 4 L glass beakers, pH probe, dialysis membranes for RGD (132590, 3.5 kDa MWCO, Spectrum™ Spectra/ Por™ #6) and “PQ” (132655, 6–8 kDa MWCO, Spectrum™ Spectra/Por™ #1), orbital shaker, sterile filters (0.22 μm PES), aluminum foil, argon gas, stir plates, and stir bars. • Polydimethylsiloxane (PDMS) molds and design: SYLGARD™ 184 silicone elastomer kit, Illustrator, CO2 laser cutter, microscope cover glass, autoclave. • HEPES buffer solution: 25 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) in 1×PBS (pH 7.8). • Hydrogel synthesis: 10 mg/mL Glycosil® Version 2.0 (Advanced Biomatrix, GS222F) in 1xPBS, 30 mg/mL Ac-PEG-PQ-PEG-Ac in HEPES buffer solution, 70 mg/mL Ac-PEG-RGD in HEPES buffer solution.

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• Cell encapsulation: 0.125% (v/v) trypsin-EDTA, 1 mg/mL (w/v) soybean trypsin inhibitor, Eppendorf tubes (1.5 mL), sterile pipettes and tips, humanized salivary medium (HSM) (see Subheading 2), PDMS molds, glass microscope slides, 26G needles, forceps, stainless-steel spatulas, #9 scalpel blades, and blade handle. All surgical tools are sterilized prior to use. 2.3

Cell Phenotyping

• Antibodies and cell dyes: Keratin 5 (K5, 905501, BioLegend,), Keratin 14 (K14, CBL197, Millipore), TP63 (4892S, Cell Signaling), Mucin-7 (ab55542, Abcam), goat anti-mouse Alexa Fluor™ 568 (A-11031, Invitrogen), goat anti-rabbit Alexa Fluor™ 488 (A-11034, Invitrogen), 4′,6-diamidino-2-phenylindole (DAPI) (D3571, Invitrogen). • Immunostaining: 4% (w/v) paraformaldehyde (PFA) in Hank’s Balanced Salt Solution (1×HBSS) with calcium/magnesium; permeabilization solution: 0.2% (v/v) Triton X-100 in 1×HBSS; blocking solution: 10% (v/v) goat serum in permeabilization solution; wash solution: 1×HBSS.

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Methods

3.1 Tissue Harvest and Preparation for Transport

1. All procedures were conducted under approved Institutional Review Board guidelines at UTHealth Houston. All patient donors provided signed informed consent for tissue collection prior to undergoing the procedures, and all collections were immediately deidentified except for sex and age. 2. Minor labial salivary glands are removed from the lips of patient donors undergoing transoral mandibular surgical procedures to repair acute injuries. Typically, each donor provides three to six minor salivary glands, obtained from the surgical incision site at the time of wound closure. Both male and female patient donors with ages ranging from 18 to 62 donated tissue for development of these methods. 3. Harvested tissues are placed into sterile containers with a small volume (3–10 mL) of sterile saline to ensure tissue samples remain hydrated during transport. Samples at the collection site are kept in a sealed, insulated container protected from light at 4 °C on wet ice until transfer to the processing location.

3.2 Preparation and Isolation of Cells from Fresh Minor Salivary Glands (Fig. 1)

1. Upon arrival at the laboratory, glands are removed from the transport container with sterile forceps and placed into a 1.5 mL Eppendorf tube and washed twice using a P1000 pipette with cold 1×PBS in a cell culture hood using aseptic technique. 2. 1×PBS is replaced with cold wash solution #1, and glands are washed twice more for a total of 3 min.

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Fig. 1 Overview of the tissue processing process used for minor salivary glands. (a) Minor salivary glands are removed from the transport container and placed into a sterile 1.5 mL Eppendorf tube for wash. (b) After the final wash, tissue with residual wash buffer #2 is placed into the lid of a 10 cm cell culture dish. (c) Minor glands are minced with fine stainless-steel scissors. (e) Appearance of glands after mincing. (f) Tissue pieces are moved to a T-25 flask with p1000 pipette. (f) Minced tissue in T-25 before placement into the cell culture incubator

3. Wash solution #1 is replaced with cold wash solution #2, and glands are washed twice more for a total of 3 additional minutes. 4. Minor glands are transferred using sterile forceps or a P1000 pipette onto the lid of a sterile 10 cm petri dish. 5. Using sharp, small fine scissors, the grapeseed-like minor glands are carefully minced until the pieces are small enough to fit through the orifice of a P1000 pipette. If they are not small enough to fit through, the tissue should be minced further. 6. Once finely minced, the tissue can be transferred to a T-25 tissue culture flask, and a maximum of 2.5 mL of HSM is added. Place the flask in a humidified tissue culture incubator at 37 °C and 5% CO2 (see Note 1). 7. Explants should be left to grow for 5 to 7 days largely undisturbed (see Note 2). 8. After 5–7 days, the flasks can be removed from the incubator and observed. At this time, the medium is removed and any floating explants are collected and centrifuged at 233 g for 2 min. Fresh HSM (5 mL) is added to the original explant

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flask to support the growth of the attached explant cultures. The centrifuged floated explants can be replated into fresh medium in another T-25 tissue culture flask (see Note 3). 9. After another 5–7 days, the medium should be removed from both T-25 culture flasks. Any remaining unattached tissue can be disposed of using appropriate biohazardous waste protocols. 10. The first explant flask typically is ready for the first passage at this time. Healthy margins of cells that have migrated from the attached tissue should be seen around approximately half of the attached tissues (see Note 4). 3.3 Culture, Passage, and Expansion of Primary Minor Salivary Gland Cells

1. For the first passage of cells from the explants, wash the T-25 explant flask with pre-warmed 1×PBS. 2. Add warm 0.125% trypsin (1 mL) and leave for 2–3 min, watching carefully through the microscope. Monitor the cultures, and do not leave cells in trypsin for too long. Radial cell outgrowth from the tissue explant is multilayered near the explant core and monolayered at the peripheral margins. Trypsinize the peripheral cell monolayers to avoid lifting of the explant (see Note 5). 3. Once the cells at the periphery of the explant begin to visibly loosen or lift off around the explant, add 1 mL of soybean trypsin inhibitor to the culture flask. 4. Use a 5 mL serological pipette to dislodge the cells from the flask by gently pipetting up and down. Transfer detached cells to a sterile 15 mL conical tube. 5. Add fresh HSM (5 mL) to the T-25 flask. The remaining cells around the explant should continue growing. Explant flasks can be maintained in culture as above and cells harvested for more than five passages if these procedures are carefully followed. This practice will increase the yield of cells per minor gland. 6. Centrifuge detached cells in the 15 mL tube at 233 g for 2 min. Do not over-centrifuge. Remove the supernatant and add pre-warmed salivary gland culture medium (5 mL) to the cell pellet. Gently pipette up and down to resuspend the cell pellet, and plate resuspended cells into a new T-25 flask. A smaller flask can be used if numbers appear low (see Note 6). 7. Allow the replated cells to grow in the incubator for 7–14 days. Once cells in the flask reach 60–70% confluence, they can be harvested again using the same trypsinization protocol described above. For cell expansion, these cells now can be plated into a larger flask, typically a T-75, and allowed to grow once more. Do not allow them to overgrow (see Note 7).

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8. If cells are growing slowly but appear healthy, as occurs for some specimens, continue feeding once a week leaving them largely undisturbed until they reach 60–70% confluency. 9. Cells in the T-75 at this confluency can be frozen in liquid nitrogen using standard cell culture freeze down procedures (see Note 8). 3.4 Preparation of Customized Hydrogels for hS/PC Culture

1. Prepare synthetic peptides as described previously [7, 8]. 2. Using amber tubes (50 mL), separately reconstitute PQ and RGD peptides and PEG-SVA in HEPBS buffer solution to obtain the following molar ratios: 2 PEG-SVA: 1 PQ; 1 PEG-SVA: 1.2 RGD. 3. Use 1 M NaOH dropwise to bring the pH of the peptide solutions to 8.0. 4. Add PEG-SVA dropwise to each peptide-containing tube. 5. Flush with argon to prevent oxidation and allow reaction to occur at 4 °C on an orbital shaker. Monitor peptide solutions hourly and adjust pH to 8.0 until pH remains stable (see Note 7). Allow to continue to react overnight on a slow shaker at 4 °C. 6. The next day, prepare the dialysis membranes, stir plates, stir bars, and 4 L glass beakers to receive each reacted solution. PQ reaction is added to the 6–8 kDa MWCO membrane, after thoroughly wetting and removing protective glycerol, and securely fastened with dialysis clips. RGD reaction is added to the pre-wetted 3.5 kDa MWCO membrane and securely fastened with dialysis clips. An air bubble should be trapped to encourage a vertical position of the reaction containing dialysis tubes. 7. Refresh the dialysis water with pure Milli-Q® water hourly for the first 2 h, and then leave to continue dialyzing overnight at room temperature with stirring. 8. The next day, complete three more exchanges, one every 2 h. 9. Sterile-filter the dialyzed peptides with 0.22 μm PES membrane into an amber tube and freeze at -80 °C overnight. Alternatively, flash freeze tube in liquid nitrogen with a tilted rotation to increase surface area for evaporation. 10. Lyophilize the frozen, dialyzed peptide solutions to remove all water and moisture; purge with argon to remove oxygen from tube, parafilm the lid, and store frozen at -80 °C until use.

3.5 Encapsulation of hS/PCs in 3D Hydrogels

1. Prepare casting molds on glass microscope slides as described previously [9]. 2. Cells were encapsulated in 3D similar to previously published methods [1, 7] with the customizations below.

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3. Add 1×PBS (1 mL) to Glycosil® (HA-SH) Version 2.0. Place in a heated bead bath at 50 °C and vortex briefly every 10–15 min over 1 h. 4. Reconstitute peptides in sterile HEPES buffer solution. We use 30 mg/mL lyophilized PQ reaction (Ac-PEG-PQ-PEG-Ac) and 70 mg/mL of lyophilized RGD reaction (Ac-PEG-RGD) for these experiments (see Notes 7, 8, and 9). 5. Calculate volume of hydrogel components needed for encapsulation. Follow volume ratios [4:1:1] of reconstituted Glycosil® (HA-SH), RGD, and PQ, respectively. 6. Calculate the number of cells needed to achieve a seeding density of three million cells/mL of hydrogel volume determined in Step 5. 7. Once hydrogel components are ready and calculations are complete, trypsinize hS/PCs and harvest as described above in Subheading 3.3, and count cells using a hemocytometer. 8. Spin cells down at 233 g for 2 min. Remove media including the residual media above the cell pellet with a P200 or P10. (a) In the same tube with cell pellet, add four-parts volume of HA-SH to the cell pellet, and gently pipette up and down to achieve even cell suspension. (b) Add one-part volume of RGD solution and gently pipette up and down until well mixed. (c) Add one-part volume of PQ and gently pipette up and down until well mixed. Let the hydrogel mixture react until right before it becomes too viscous to pipette (see Note 8). Gently pipette 50 μL of cell-laden hydrogel into each 6 mm circular molds. 9. Place the molds into the incubator at 37 °C with 5% CO2 for 1 h to allow them to finish gelation. 10. Once the hydrogels complete gelation, remove them from the incubator. Test the hydrogels by placing a small drop of medium onto the hydrogel. If the medium stays as a small droplet, the hydrogel is ready to be removed from the PDMS mold. If it sinks in the hydrogel, allow the hydrogel to set for another 15–20 min. 11. Place a small amount of media (20–50 μL) on top of each hydrogel. Gently score each hydrogel with a 26 G needle by tracing the perimeter of the mold with slight pressure to reach the glass microscope slide at the base. Do this for each hydrogel in the mold. 12. Using sterile forceps, lift the mold from a corner and gently pull toward the opposite end. The hydrogels should release cleanly from the mold, if thoroughly scored out, and stick to the underlying glass microscope slide.

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13. Add additional media to the hydrogels prior to keep hydrogels hydrated while transferring to culture plates. Gently release and lift the hydrogels with a thin flat tool to support the full width of the hydrogel, i.e., scalpel or spatula, to be transferred to culture well plates, typically 1 hydrogel to each well of a 24-well plate. Alternatively, softer hydrogels can be pushed directly from the microscope glass into the well plates. 14. Add 1 mL of medium to each culture well, and gently sink each hydrogel to the bottom of the well with a spatula. Ensuring all hydrogels are fully submerged prevents them from drying out. 15. Continue to feed cells through exchange of culture medium once each week for the duration of the work to be performed. Hydrogel cultures are stable in culture for several weeks to months. 16. Cells were phenotyped in 3D following similar methods as found in Subheading 3.6. 3.6 Phenotyping Cells Isolated from Minor Salivary Glands

1. Major salivary gland hS/Pcs were phenotyped recently by our laboratory and shown to be positive for hallmark biomarkers TP63 and cytokeratins 5 and 14 [2]. 2. Minor salivary gland-isolated hS/PCs at passage 3 from five different patient donors were used for this phenotypic study. 3. Cells were trypsinized from T-75 culture flasks (see Subheading 3.3) and plated into multiple wells in a 24-well cell culture plate (see Note 10). 4. Replated cells were grown to confluency and then rinsed with 1×HBSS with calcium and magnesium to support firm attachment. 5. Cells were fixed with 4% PFA in 1×HBSS with calcium and magnesium for 10 min and then washed through three exchanges each for 5 min using 1×HBSS. 6. Cells were permeabilized with 0.2% Triton X-100 in 1×HBSS through three exchanges each for 5 min. 7. After blocking with 10% goat serum in 1×HBSS for 1 h at room temperature, primary antibodies (at 1:100) were added in blocking solution overnight at 4 °C under gentle rotation. 8. To remove primary antibodies, cells were washed with HBSS for 5 min each through three complete exchanges. 9. Secondary antibodies (1:1000) and 1 μg/mL DAPI were added. To prevent photobleaching, cells were protected from light by wrapping in aluminum foil and left at room temperature under gentle rotation. At the end of the incubation, cells were washed with 1×HBSS through three exchanges for 5 min each.

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Fig. 2 Phenotyping of stem/progenitor cells from minor salivary glands. K5, K14, and p63 immunostaining of hS/PCs that were maintained properly after explant. Top row: cells from donor M62 at passage 2; bottom row: cells from donor M26 at passage 4. Left column: K14 (red) p63 (green) and DAPI (blue). Middle column: K5 (red) and DAPI (blue). Right column: representative secondary antibody control

10. Results of phenotyping are shown in Figs. 2 and 3. Minor salivary gland-derived hS/PCs, when carefully treated using the procedures we describe, expressed markers similar to those isolated from major salivary glands (submandibular, parotid) (Fig. 2). Stem/progenitor cells isolated from minor salivary glands express K5, K14, and p63 in virtually all cells for multiple passages. hS/PCs are rounded and uniform in structure. 11. We found that hS/PCs from minor glands are more prone to undergo unwanted differentiation without proper maintenance and remain more diverse over passages compared to the more homogeneous major gland-derived cells. hS/PCs that have been overgrown or over-trypsinized can form large plaque-like structures (Fig. 3, M62 P3 and F25) with lower levels of K14 and K5. Additionally, some cells undergo a squamous like transition with aberrant expression of K5 and/or K14, and the cells often are large and flat and resemble “fish scales” in appearance (Fig. 3, M18) similar to cultured lung epithelial cells [10].

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Fig. 3 Effects of stress on cells from minor salivary gland explants. K5, K14, and p63 immunostaining of cells that were maintained poorly after explant. Top row: cells from donor M62 at passage 3; middle row: cells from donor M18; last row: cells from donor F25. Left column: K14 (red), p63 (green), and DAPI (blue). Middle column: K5 (red) and DAPI (blue). Right column: representative secondary antibody control

12. For 3D cell encapsulation studies, the same methods are followed except the fixation time is increased to 30 min and all washes are increased to 15 min. Antibody concentrations, blocking concentration, and reagents like 1×HBSS all remained the same. Results for 3D phenotyping are shown in Fig. 4. hS/PCs in hydrogels form microstructures that lose expression of most p63, consistent with their lower proliferation rates, and have lower levels of K14 as seen previously using encapsulated cells from major glands. K5 levels remain high in cell microstructures, consistent with retention of an epithelial

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Fig. 4 Progenitor and differentiation markers for minor hS/PCs after encapsulation. In gels, cells begin assembly to form microstructures. Left column: donor M62; right column: donor M26. Top row: K14 (red), p63 (green), DAPI (blue); second row: K5 (green), DAPI (blue); third row: salivary secretory protein MUC7 (red), DAPI (blue); bottom row: representative secondary antibody control DAPI (blue) and differential interference contrast (DIC)

phenotype. Additionally, MUC7 is expressed in encapsulated cells and especially in microstructures in 3D, indicating the hS/PCs can differentiate and secrete salivary products. 13. The availability and phenotypic properties of hS/PCs from minor salivary glands offer an expanded tissue source with broad potential for regenerative therapies for hyposalivation disorders and associated xerostomia.

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Notes 1. It is important not to exceed 2.5 mL of medium as the minced tissue will float and not land efficiently nor adhere to the plates to allow cell explants to begin forming. 2. It is critically important not to touch the new explant cultures or move the flasks during this time period as any movement can disrupt the adherence of the minced tissue pieces or the newly emergent cells. 3. Many viable salivary hS/PCs will come from the second explant flask. Sometimes the second flask is more successful than the first. Throwing away these “floaters” greatly reduces the total number of hS/PCs that can be obtained from minor glands. 4. We have achieved better results by passaging cells off an explant at this time and moving them to a smaller plate such as one well of a six-well plate. Cells initially will grow slowly so patience will pay off. 5. Minor salivary cells tend to be sensitive to stress during early stages of culture. Over-trypsinization, plating cells too sparsely, or allowing the explant flask to overgrow all can result in the minor cells losing their stem/progenitor properties, undergoing senescence or differentiating into flat, “fried egg” looking cells that share visible features of squamous epithelial cells. 6. It is not recommended to freeze cells directly off an initial explant flask because there are too few cells during the expansion phase. We note that the first few harvests of hS/PCs often are heterogenous and do not always survive freeze down well. We recommend freezing cells from the T-75 flask stage at the earliest as the purity and viability of the hS/PCs increases between this passage and passage 3. 7. Peptides are light sensitive and hydrophilic. Be sure to process peptides in amber vials protected from light and flush with argon after checking the pH every time. 8. Peptides react with the hyaluronic acid through a Michael addition between the thiol groups on the hyaluronic acid and the acrylate group on the modified peptides. This reaction is pH sensitive and requires a slightly basic pH to reach completion. Gelation is quickened by an increase in pH. It is helpful to form a small test hydrogel first to assess the reaction speed for a given batch of reagents. Pipetting into the molds too early before the hydrogel has set can result in cells sinking to the bottom. Pipetting too late can result in sheared cells in torn or warped gels as gelation has progressed too far. HEPES can be added to the PBS buffer to help stabilize the pH.

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9. Any pegylated peptide with an adhesion motif can be used in place of the RGD peptide we use. However, complete elimination of the peptide can result in changes to the hydrogel chemistry. Different amounts of PQ peptides can be used to alter the cross-linking and properties of the hydrogel. Hydrogels with PQ only tend to shrink. Choice and concentration of peptides can be adjusted to alter hydrogel properties for other cells and cell types. 10. Salivary hS/PCs, especially those derived from minor salivary glands, do not always adhere well to uncoated glass cover slips; therefore, phenotyping was performed on culture plates to which they adhere well. Funding Acknowledgments CCTS TL1 (TR003169). OMS Foundation. NIH R01 DE032364. References 1. Wu D, Chapela P, Farach-Carson MC (2018) Reassembly of functional human stem/progenitor cells in 3D culture. In: Epithelial cell culture, p 19–32 2. Wu D, Chapela PJ, Barrows CML et al (2022) MUC1 and polarity markers INADL and SCRIB identify salivary ductal cells. J Dent Res 101:983–991 3. Kessler AT, Bhatt AA (2018) Review of the major and minor salivary glands, part 1: anatomy, infectious, and inflammatory processes. J Clin Imaging Sci 8:47 4. Andreadis D, Bakopoulou A, Leyhausen G et al (2014) Minor salivary glands of the lips: a novel, easily accessible source of potential stem/progenitor cells. Clin Oral Investig 18: 847–856 5. Siqueira WL, Salih E, Wan DL et al (2008) Proteome of human minor salivary gland secretion. J Dent Res 87:445–450 6. Huang C-M, Zhu W (2009) Profiling human saliva endogenous peptidome via a high

throughput MALDI-TOF-TOF mass spectrometry. Comb Chem High Throughput Screen 12:521–531 7. Sablatura LK, Tellman TV, Kim A et al (2022) Bone marrow endothelial cells increase prostate cancer cell apoptosis in 3D triculture model of reactive stroma. Biology 11:1271 8. Fong ELS, Wan X, Yang J et al (2016) A 3D in vitro model of patient-derived prostate cancer xenograft for controlled interrogation of in vivo tumor-stromal interactions. Biomaterials 77:164–172 9. Fong ELS, Martinez M, Yang J et al (2014) Hydrogel-based 3D model of patient-derived prostate xenograft tumors suitable for drug screening. Mol Pharm 11:2040–2050 10. Gianotti A, Delpiano L, Caci E (2018) In vitro methods for the development and analysis of human primary airway epithelia. Front Pharmacol 9:1176

Chapter 4 Salivary Organotypic Tissue Culture: An Ex-vivo 3D Model for Studying Radiation-Induced Injury of Human Salivary Glands Akshaya Upadhyay , Migmar Tsamchoe, and Simon D. Tran Abstract An organotypic tissue culture model can maintain the cellular and molecular interactions, as well as the extracellular components of a tissue ex vivo. Thus, this 3D model biologically mimics in vivo conditions better than commonly used 2D culture in vitro models. Here, we provide a detailed workflow for generating live 3D organotypic tissue slices from patient-derived freshly resected salivary glandular tissues. We also cover the processing of these tissues for various downstream applications like live-dead viability/cytotoxicity assay, FFPE sectioning and immunostaining, and RNA and protein extraction with a focus on the salivary gland radiation injury model. These procedures can be applied extensively to various solid organs and used for disease modeling for cancer research, radiation biology, and regenerative medicine. Key words Ex vivo models, Disease modeling, Organotypic slice culture, Radiation injury, Salivary gland regeneration, 3D tissue culture

1

Introduction Biological processes involve complex cellular and acellular interactions among various cell types through multiple pathways. Cellular and molecular interactions can have a significant effect during physiological as well as stress responses like radiation injury and cancer. Most biological studies utilize in vitro disease models due to their efficiency and ease. However, most in-vitro models cannot replicate the complex microenvironment of tissues and organs. In vivo animal models are the second-best choice before fully fledged clinical trials are conducted. However, these models are less humane, especially for painful injury models, and are timeconsuming [1]. Thus, developing ex vivo models which can biomimic in vivo conditions is becoming increasingly essential. Various organ systems have applied organotypic tissue culture models to study developmental, physiologic, pathologic, as well as therapeutic

Mario Baratta (ed.), Epithelial Cell Culture: Methods and Protocols, Methods in Molecular Biology, vol. 2749, https://doi.org/10.1007/978-1-0716-3609-1_4, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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pathways. Most commonly, this culture system is employed in neurological systems [2–4], while others include the pancreas [5] and liver [6]. The crucial role of microenvironment and precision medicine has come to light with these model systems. These models maintain structural integrity, phenotypic characteristics, and functional activity of the tissues. Preclinical salivary gland research is challenging due to the limited availability of salivary gland disorder models. Radiotherapy-induced side effects in head and neck cancer patients affect salivary glands most drastically due to their anatomical position within the radiation zone and high radiosensitivity. Salivary gland cells, particularly acinar cells, were considered postmitotic quiescent mature cells. Nevertheless, their proliferative and reparative role was recently identified in response to stress or injury [7, 8]. Still, it remains a challenge to understand the mechanism of radiation damage, which can involve multiple cell types and molecular mechanisms. A reliable biomimicry model of radiation injury is required to identify potential therapies to ameliorate this response. Furthermore, acinar cells are the most radiosensitive cells, but their culture in vitro is scarce [9]. A crucial water channel protein, AQP5 expression, and its apical localization are lost during 2D cell culture using traditional techniques. Our lab previously established a 3D organotypic tissue slice culture model cultured over 14 days [10]. Here we provide the detailed protocol established in our lab to successfully reciprocate clinical radiation injury using a human donor-derived salivary gland tissue culture model. GammaH2AX is a robust marker of DNA damage. It is expressed at double-stranded break sites after radiation, the most common pathway for cellular DNA damage. With this model, we were able to maintain apical expression of AQP5 (indicating the normal function of acinar cells) and replicate salivary gland-specific radiation injury (downregulation of acinar function through AQP5) and pathways of DNA damage (GammaH2AX expression recovery). This model can be used for salivary glands and other exocrine glands like the pancreas and various solid organs.

2

Materials

2.1 Tissue Preparation and Slicing (Table 1)

1. Scalpel blade. 2. Autoclaved forceps, scissors, and spatula. 3. Metallic molds. 4. Heat top with magnetic stirrer. 5. Dropper.

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Table 1 Key resources Reagent

Source

Reference

Mammary epithelial basal medium (EpiMax1)

Wisent, Inc.

002-010-CL

Live/dead assay kit

Thermo Fischer

L3224

Universal block

BioGenX

HK085-5K

mirVana PARIS kit

Thermo Fischer

AM1556

SOD colorimetric activity kit

Invitrogen

EIASODC

Equipment

Source

Reference

Vibratome

Leica Biosystems

1000plus

Spectrophotometer

Variable

Variable

X-ray biological irradiator

Rad Source Tech Inc.

RS 2000

Nanodrop

Thermo Fischer

ND 2000

CO2 incubator, biological safety cabinet (class 2)

Variable

Variable

6. 30-mm-diameter, 0.4-um-pore-size (PICM03050, Millipore).

membrane

insert

7. 6-well plates. 8. Wet ice. 9. Pipettes and tips. 10. Vibratome. 11. Phosphate Buffered Saline (PBS)/Hanks’s Balanced Salt Solution (HBSS) with 2% antibiotics. 12. Epithelial cell growth media with 10% Fetal Bovine Serum (FBS) and 2% antibiotics. 13. Agarose gel powder (low melting). 14. Instant glue. 15. Paint brush. 2.2

Live-Dead Assay

1. Pipettes and tips. 2. Eppendorf tubes. 3. Live-dead viability/cytotoxicity kit. 4. Aluminum wrap. 5. PBS. 6. Aqueous mounting media and slides.

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Table 2 Antibodies Antibody

Species

Localization

Dilution

Reference

E-cadherin

Mouse monoclonal

Cell membrane

1/1000

ab231303

AQP5

Rabbit polyclonal

Cell membrane

1/500

PA5-99403

Alpha-SMA

Mouse monoclonal

Cytoplasm

1/2000

ab7817

Vimentin

Rabbit monoclonal

Cytoplasm

1/500

EPR3776

Gamma H2AX

Rabbit monoclonal

Nucleus

1/500

ab81299

Primary

2.3 Immunostaining (FFPE Sections)

1. Slide holder and glass jars. 2. Pipettes and tips. 3. Paraformaldehyde (PFA) 4%. 4. Pap pen. 5. Aluminum foil. 6. Cover slips. 7. Water bath set to 95–100 °C. 8. Humidified chamber. 9. IF/confocal microscope. 10. CitriSolv solution. 11. Ethanol (EtOH) (100%, 90%, 80%). 12. Wash buffer (1× PBST). 13. Permeabilization solution (PBS/0.5% Triton-×100). 14. Universal block/BSA. 15. Antigen retrieval solution (10 mM sodium citrate, 0.05% Tween 20, pH 6.0). 16. Antibodies and DAPI (Table 2). 17. Mounting media.

2.4 RNA and Protein Extraction

1. RNase-free pipette tips and Eppendorf. 2. Pipettes. 3. Metal spatula/tweezer. 4. Mortar and pestle. 5. RNAse-free working hood. 6. mirVana PARIS Kit (AM1556). 7. 100% ethanol (ACS grade). 8. Nuclease-free water.

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1. Pipette and pipette tips. 2. Amicon Ultra 0.5 mL centrifuge filters (10 kDa cutoff). 3. Superoxide Dismutase (SOD) Colorimetric Activity kit.

2.6 Irradiator (Optional)

3

Methods

3.1 Tissue Preparation and Slicing

3.1.1 Connective Tissue Removal (1.5 h)

Overview Salivary glands are soft glandular tissues with multiple epithelial lobes intertwined in mesenchymal stroma. The lobes are interconnected through epithelial ducts. The whole gland and groups of lobes are covered by connective tissue capsules providing physical resilience to the tissues. For tissue culture, the pieces of glands are embedded in a gel with a comparable tensile strength to the gland to facilitate cutting freshly resected tissues while maintaining viability. Vibratome is an instrument that makes cutting thin sections of live tissues possible. The more rigid connective tissues must be removed as it can hinder the cutting process. The yield or the number of slices obtained will vary from patient to patient as there can be structural changes due to the age, condition, and gender of the patient.

1. The issues should be carried and processed over ice as soon as possible after the resection (see Notes 1–4). 2. Prepare 1XPBS buffer with 2% antibiotics and culture media (EpiMax with supplements, 10% FBS) (see Notes 5 and 6). 3. Prepare 3% agarose: For 100 mL, add 3 g low-melting agarose to 100 mL PBS at room temperature with constant mixing over a magnetic stirrer. Once most of the agarose dissolves, increase the temperature to 100 °C until the solution becomes transparent. Store sealed at room temperature or maintain at 37 °C immediately before embedding. 4. Wash glands with PBS (2%Anti) to clean blood and surgical debris in tissue culture plate using 10 mL pipettes, or vortex them in 50 mL tubes (see Notes 7 and 8). 5. Using forceps and a scalpel, gently remove the connective tissue (translucent slimy capsule over the gland and lobes) in the tissue culture plates over ice (see Notes 9 and 10) (Fig. 1). 6. The connective tissue is mostly removed once the gland looks dark pink and does not slip when picked up with forceps. 7. Cut the tissues into 3–6 mm pieces.

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Fig. 1 Schematic workflow of tissue preparation and culture of organotypic tissue slices. (a). The connective tissue is dissected and removed using forceps and scissors; tissues are cut into 3–6 mm pieces. Individual or multiple pieces are embedded in 3% agarose gel. (b). Agarose gel with the tissues is fixed on the Vibratome platform with ice-cold buffer; 150–200-micron-thick slices are cut and picked up with a paintbrush. For the final step, the slices are washed and cultured over a transwell membrane with culture media on the bottom, providing an air-liquid interface to support epithelial cells. (Created with BioRender.com) 3.1.2 Embedding (30 min)

1. Maintain the 3% low agarose gel at 37 °C temperature over a heated top (see Notes 12 and 13). 2. Prepare a cold flat surface for keeping the tissues after embedding them with ice or an ice pack. 3. Using a plastic dropper, put a thin layer of liquid agarose into the metal mold (the dropper tip can be cut to facilitate). 4. Carefully pick up the tissue pieces and place them into the mold. Multiple or single tissue pieces can be placed depending on the sample size and the mold.

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5. Once the placement is satisfactory, fill the mold with more liquid agarose to cover up the tissue completely. 6. Transfer the mold on top of the ice/ice pack with quick but stable movement. Agarose will undergo gelation quickly, preventing any movement of the tissues. 7. Wait for 10 min before taking out the embedded agarose gel using a spatula. 8. Place in transfer buffer (PBS/HBSS) in a tissue culture dish, and carry over ice to a Vibratome machine. 3.1.3 Precision Cut Tissue Slicing (4–6 h)

1. Prepare the Vibratome by placing a razor blade, adding ice in the chamber, sterilizing the platform using 70% ethanol, and waiting until it dries. 2. Place a tissue culture dish with transfer buffer (HBSS) over ice. 3. Set Vibratome 1000 plus to a speed of 0.075 mm/s, frequency of 75–85 Hz. This model already has the blade at an angle of 15°; adjust the angle accordingly if using another model. 4. Fix the tissue section on the platform using instant glue and let it dry (see Note 14). 5. Add ice-cold PBS with 2% antibiotics before slicing the embedded tissue. 6. The tissue section can be manually adjusted to a 150–200 mm thickness by rotating the knob clockwise to move the platform up or anticlockwise to go down accordingly. 7. Carefully move the platform upward till the blade almost touches the tissue. Discard the first slice, as its thickness is unknown. 8. Collect the slices as they separate from the rest of the embedded tissues upon cutting using a paintbrush. 9. Gently transfer them into the tissue culture dish containing the ice-cold transfer buffer. 10. Repeat steps 4–8 until all the sections are cut or sufficient slices are obtained (see Notes 15–17). 11. Wash the slices three times with PBS (2% antibiotics) (see Note 18). 12. Prepare 6-well plates by inserting trans-well membranes using forceps. 13. Add 1000 μL 10% FBS EpiMax media at the bottom of the trans-well membrane. 14. Pick up the slices from the tissue culture dish with forceps or spatula, and place them carefully over the membranes. 15. Remove excess PBS from the top of the membrane, if any.

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Fig. 2 (a) and (b). Live-dead assay: Live are stained green, dead cells are stained red, and the nuclei are blue. Intensity analysis was conducted using ImageJ software to assess the viability of the tissues over different days. (c). FFPE sections: Hematoxylin and eosin staining and immunofluorescence using E-cadherin (a marker for epithelial cells) show that the integrity in cellular connections of the tissues is maintained. (d). qPCR for salivary gland related genes: Salivary gland-specific genes were expressed throughout the 14-day culture [10], but the optimal expression was seen within 3 days of the culture. So, we recommend a 2–5-day protocol for experimental setups

16. Cover the plate and transfer it to a humidified cell culture incubator at 37 °C with 5% CO2. 17. Culture for 24 h to 14 days as required (see Fig. 2 legend). 18. For prolonged cultures, discard the old media from below the trans-well inserts using pipette, and add 1000 μL culture media (10% FBS EpiMax) every 2–3 days. 3.1.4 Radiation (Optional) (20–30 min)

1. Carry the 6-well tissue culture plates in a thermally insulated box to the irradiator (see Note 19). 2. For RS 2000 X-ray biological irradiator, calculate the radiation dose using the following equation: Minutes = target dose in Gy/dose rate for RS 2000 according to the traget level Secondes = decimal remainder from the minutes × 60 3. Place the tissue culture plate in the middle of the irradiator platform at appropriate level (level 3). 4. Close the radiator and start the cycle according to the time calculated. 5. Transfer the tissue culture plates back to the incubator as soon as possible.

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3.2 3.2.1

Assays Live-Dead Assay

47

Overview: As the name suggests, the live-dead assay is a two-color assay that distinguishes viable and nonviable cells. It is based on assessing the plasma membrane integrity (Ethidium Homodimer1) and esterase activity (Calcein). ED-1 penetrates the dead cells and stains the nucleus red, while Calcein gives a florescent green color in response to active esterase activity in live cells. With a third stain added for the total number of nuclei, we can analyze the exact number of cells in the tissue slice (Fig. 2a). Since this assay stains the cells in real time, it can be used as a direct measurement to assess the viability of the tissues rather than other metabolic tests, which can be inconsistent if the size or composition of the tissue slices varies. • Pre-stain wash: 15–20 min 1. Thaw Calcein, ED-1, and Hoechst at room temperature (see Note 20). 2. Prepare appropriate concentrations of the stains (Calcein = 2 mM, ED = 4 mM, Hoechst = 4 mM) in PBS. Vortex to mix. 3. Add the slices to an Eppendorf tube and wash three times with PBS, 5 min each (see Note 21). • Staining: 60 min 4. Add 200 μL of the prepared staining solution in each Eppendorf. 5. Cover with aluminum foil and incubate for 60 min at 37 °C in a CO2 incubator. 6. Vortex from time to time to ensure homogenous staining. • Post staining wash and mounting: 20–30 min 7. Wash three times with PBS, 5 min each. 8. Transfer the slices to the top of the glass slides. Dry the surroundings using Kim wipes. 9. Add mounting media and cover with a cover slip. 10. Image using a confocal microscope (100–150 μm depth) within 24 h of staining. Store overnight at 4 °C if required. 11. For image analysis, do imaging at 10× with 10 μm z-stacks and at 20× with 2–5 μm. Intensity analysis is conducted using ImageJ software.

3.2.2

Immunostaining

Overview: Tissue culture slices can be directly stained after overnight fixation or embedding in paraffin (FFPE) and cut further into 5 μm sections. We recommend FFPE section staining as multiple sections can be cut and visualized for different stains maintaining the cellular morphology and function (Fig. 2b). Direct staining can

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also be applied, but it gives higher background, long-term storage of samples is not possible, and it is more challenging to image due to the higher depth (100–150 μm). 1. Tissue culture slices can be fixed in 4% PFA overnight to a maximum of 1 week at 4 °C, followed by paraffin embedding and cutting into 5 mm sections (see Note 22). • Deparaffinization/rehydration: 30 min 2. Bake slides at 60 °C for 1 h. 3. Remove slides from the oven and let cool at room temperature, 5–10 min. 4. CitriSolve: 2 × 5 min. 5. EtOH 100%: 2 × 5 min. 6. EtOH 95%: 5 min. 7. EtOH 90%: 5 min. 8. EtOH 80%: 5 min. 9. dH2O: 2 × 5 min. • Antigen retrieval: 60–90 min 10. Keep in 95 °C antigen retrieval buffer. 11. Let it boil for 30 min. 12. Carefully remove the container from water bath and put at room temperature. Give 30 min to cool down. 13. Wash slides in dH2O: 2 × 5 min. • Blocking and permeabilization: 90 min (/overnight) 14. Prepare a humid chamber for slide incubations. 15. Remove the peroxidase block from 4 °C and keep it on ice. 16. Surround the tissue with a hydrophobic pap pen and return to the water. 17. Once all slides are circled with a pen, carefully shake off excess water and place it in a humid chamber. 18. Incubate slides with permeabilization solution (1% Triton): for cell surface marker, 5–10 min; cytoplasmic markers, 15 min; and nuclear markers, 20 min. 19. Wash slides with PBS: 3 × 5 min. 20. Incubate slides with peroxidase block (200 μL/slide) for 20 min at room temperature (not required for immunofluorescence staining). 21. Wash slides with PBST: 2 × 5 min. 22. In a humidified chamber, incubate the slides with blocking solution (universal block, 200 μL/slide) for 60 min at room temperature or overnight at 4 °C.

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• Primary antibody staining: overnight 23. Remove blocking solution and directly add primary antibodies with recommended/optimized dilutions (20–50 μL, depending on the size of the tissue). 24. In a humidified chamber, incubate for 1 h at room temperature or overnight at 4 °C. • Secondary antibody staining: 90–120 min 25. Remove primary antibody solution. 26. Wash with PBST: 3 × 5 min. 27. Add diluted secondary antibody (according to primary antibody). 28. Incubate in humidified chamber for 60–90 min at room temperature (in dark). 29. Wash slides with PBST: 3 × 5 min. 30. Add DAPI and incubate in humidified chamber for 5–10 min at room temperature (in dark). 31. Wash slides with PBS: 2 ×1 min. • Mounting: 10 min 32. Add one to two drops of DAKO aqueous mounting media at the edge of the slide. 33. Slowly angle the cover slip and let it cover the slide. Let mounting media spread and remove bubbles by carefully generating pressure points on cover slip. 34. Store at 4 °C overnight and visualize the following day. 35. Once visualized, seal with nail polish and store for short term at 4 °C or -20 °C for longer term. 3.2.3 RNA and Protein Extraction

Overview: Since tissues from human subjects are usually limited and the tissue slices are quite small, we employed low input RNA extraction kit (0.5 mg starting material), which could recover native proteins simultaneously. We employed RNA organic extraction followed by solid-phase extraction to enable the robustness of organic extractions while maintaining the efficiency and sensitivity of solid-phase extraction. The proteins and RNA yielded from this protocol can be used for various downstream applications. • Tissue disruption: 15–20 min 1. Measure or estimate the weight of frozen sample. Aliquot 100–625 μL (six to eight volumes per tissue mass) of ice-cold cell disruption buffer into an Eppendorf. Buffer volume can be adjusted according to the lysate required for protein (see Notes 23 and 24).

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2. Transfer the tissue slices (pellet) to the Eppendorf containing ice-cold cell disruption buffer. 3. Vortex until tissues visibly digest and the homogenate seems cloudy. 4. Separate the lysate equally for protein and RNA extraction. If the slices clog the pipette tip during the transfer, cut off the tip slightly. 5. Lysate for protein extraction can be incubated for 5 min more and then transferred to -80 fridge. 6. Transfer the lysate that will be used for RNA isolation to a tube containing an equal amount of 2× denaturing solution at room temp. • Organic extraction: 30 min 7. Immediately mix thoroughly using vortex. Incubate the mixture on ice for 5 min. 8. Add a volume of acid-phenol: chloroform equal to the total volume of the sample lysate plus the 2× denaturing solution (see Note 25). 9. Centrifuge for 5 min at maximum speed (≥10,000 × g) at room temp to separate the mixture into aqueous and organic phases. After centrifugation, the interphase should be compact; if it is not, repeat the centrifugation. 10. Carefully take out the aqueous (upper) phase without disturbing the lower phase or the interphase and transfer it to a fresh tube. • Final RNA isolation: 20–30 min 11. Note the volume recovered. Add 1.25 volumes of room temperature 100% ethanol to the aqueous phase. Mix thoroughly using vortex. 12. For each sample, place a filter cartridge into one of the collection tubes. Transfer the lysate/ethanol mixture onto the filter cartridge (maximum 700 μL at a time). 13. Centrifuge for 30 s at 10,000 g and discard the flow through. Repeat until all of the lysate/ethanol mixture has passed through. 14. Wash the column according to the manufacturer’s protocol. Elute RNA with 50 μL DEPC water. 15. Store the eluate (RNA) at -20 °C for short storage and -80 °C for longer.

Ex-vivo Organotypic Salivary Gland Tissue Culture 3.2.4

SOD Activity Assay

51

Overview: One of the first responses to radiation damage is the increased production of super oxide dismutase (SOD) to reduce free radicals generated inside the cells. SOD1 is the major intracellular SOD, SOD2 is mitochondrial, and SOD3 is the primary extracellular SOD. Depending on the target, we can use the culture media at the bottom of the trans-well membranes (SOD3) or the proteins extracted in the previous procedure (SOD1, SOD2, and SOD3). For culture media, the media concentration using ultracentrifugation through filters might be required (if the SOD activity is below the detection level). • Sample and solution preparation: 60 min 1. Prepare samples: For culture media, concentrate the samples using Amicon Ultra 0.5 mL centrifugal filters (10 kDa cutoff) (see Notes 26 and 27). For tissue derived proteins, ultracentrifugation is required only if the protein concentration is very low. 2. Dilute the samples 1:3 in assay buffer (triplicate); 10 μL is required for each well. 3. Prepare the SOD standard as per the manufacturer’s instructions. 4. Prepare 1× xanthine oxidase (XO) and substrate according to requirement of number of plates. • Plating: 60 min 5. To each well in the 96-well plate provided, add 10 μL of standard or diluted samples. 6. Add 50 μL of 1× substrate. 7. Read blank at 450 nm. 8. Add 25 μL of XO into each well. 9. Incubate for 20 min at room temperature. • Read out and analysis: 15 min 10. Read absorbance at 450 nm. 11. Using the standard curve, derive the concentration for each unknown sample after removing the background (blank) across all wells. 12. Multiply by appropriate factor to correct for dilution.

4

Notes 1. Ethical approval and clearance must be obtained before conducting studies involving human tissues. Proper consent is mandatory from the patients or the family of the diseased patients.

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2. Tissue collected from the hospitals should be immediately carried over ice and stored at 4 °C not more than 24 h postsurgery/postharvest. 3. All procedures should be performed in sterile conditions inside Biological Safety Cabinet (BSC) Class 2. 4. Always wear double gloves while handling human tissues; the top glove can be changed frequently. 5. Prepare the buffers and culture media before the tissue collection. EpiMax mammary growth media come with two supplements; additionally, 5–10% FBS is observed to have a better survival of the slices over multiple days of culture. 6. Agarose gel should be prepared a day before and maintained at 37 °C before embedding as higher temperatures can cause damage to the tissues, while at lower temperatures, gelation occurs, leading to the insufficient or improper embedding of the tissues. 7. Cauterized vessels can be cut using scissors to remove necrotic tissues and accelerate cleaning. Additionally, heparin can also be added to prevent blood coagulation (1 mL in 49 mL of PBS). 8. The weight and size of the glands can be measured at this point by transferring them into a tissue culture dish. 9. Removal of connective tissue and fibers is crucial as they can hinder the slicing and dislodge the tissue sections from agarose gel at the time of slicing. 10. Always work with tissues over ice, and do not let them dry. Add drops of PBS as required. A semidry state is best for identifying and removing connective tissues, as they can be quite sticky. 11. Embedded tissue size can vary based on the samples and age of the patient. For more fibrotic glands, cut smaller sections to prevent dislodgment. Glands from older females usually are more fibrotic, but the scientists can decide on the size with their experience. 12. Heat top can be transferred temporarily into the BSC after sterilization with 70% alcohol. 13. Metal molds can be placed over the heat top and the agarose gel container to keep the gel liquid when embedding. 14. Ensure the agarose gel is present below the tissue piece before fixing it over instant glue. Do not let the tissue come in direct contact with the tissue, as it is toxic. 15. Cut the excess agarose gel as close to the tissue as possible to reduce the amount of surrounding gel, as it can take up more space on the trans-well membrane and hinder downstream applications like RNA and protein extraction.

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16. Always wait for the platform to dry before applying glue for subsequent pieces. 17. On average, 20–30 slices can be obtained in 45–60 min. 18. Always discard PBS after washing using 1000 μL pipettes instead of suctioning since tissue slices can get lost due to the high suction pressure. Pipette tips can be cut from the top since the tissue slices can sometimes block the pipette tips. 19. The tissue can be radiated any time, preferably after 24 h of culture. Treatment can be given before or after the radiation into the culture media according to the experimental requirements. 20. The staining solutions should be freshly prepared for each staining, not more than 1 h before the procedure. 21. Use pipettes to remove the solutions while washing, as the slices can get lost in the suction tube due to higher pressure. 22. For preprocessing for paraffin embedding, the tissue slices must be enclosed carefully in a blotting paper before putting into the cassettes since, due to the small size of the tissues, they can get lost as they float out of the cassettes. 23. Once the tissue is removed from the -80 °C freezer during RNA isolation, it is crucial to process it immediately without partial thawing. Alternatively, else the ice crystals can rupture the cell releasing RNases. 24. Agarose gel should be reduced as much as possible because the polysaccharide chains can precipitate upon ethanol addition at the time of elution of RNA. It can reduce the final RNA yield. Adding sodium salts can somewhat reduce this effect (refer to plant-based tissue extraction protocols). 25. Be sure to withdraw the bottom phase containing acid-phenol: chloroform, not the aqueous buffer that lies on top of the mixture. Vortex for 30–60 s to mix. 26. Wash the Amicon filter with 50 mM ammonium bicarbonate before adding the samples. The filters can be reused as long as they are kept moist with buffer or water. 27. SOD activity can be normalized to pre-radiation conditions for each sample to eliminate the chances of error due to differences in the number of slices in each group. Alternatively, protein concentration can be measured after ultracentrifugation and samples diluted accordingly to standardize the protein levels across all samples.

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Acknowledgments We are thankful to all the donors of the discarded salivary tissues, without whom this work would not have been possible. We are grateful to the dedicated surgeons (Drs. H Abo Sharkh, M El-Hakim, J Gigliotti, and A Zeitouni) at McGill University, Montreal, who harvested the glands for research. We are thankful to Dr. T Stroh at the McGill Neurological Institute (MNI) for providing the Vibratome and confocal imaging at the MNI Microscopy Facility. We are grateful to Drs. J Peng and J Wu for providing access to the radiator at the University of Montreal. We are thankful to Drs. P. Metrakos and A Lazaris for RNA related training and lab space. Additionally, we are grateful to the McGill Centre for Bone and Periodontal Research, as well as the McGill Goodman Cancer Institute, Histology Core, for assisting us in the handling and processing of the tissue slices. All the work presented was supported by a research grant from the Canadian Institutes of Health Research. References 1. Su X, Liu Y, Bakkar M, ElKashty O, El-Hakim M, Seuntjens J, Tran SD (2020) Labial stem cell extract mitigates injury to irradiated salivary glands. J Dent Res 99(3): 2 9 3 – 3 0 1 . h t t p s : // d o i . o r g / 1 0 . 1 1 7 7 / 0022034519898138 2. Parker JJ, Lizarraga M, Waziri A, Foshay KM (2017) A human glioblastoma organotypic slice culture model for study of tumor cell migration and patient-specific effects of antiinvasive drugs. J Vis Exp 125. https://doi. org/10.3791/53557 3. Kondru N, Manne S, Kokemuller R, Greenlee J, Greenlee MHW, Nichols T, Kong Q, Anantharam V, Kanthasamy A, Halbur P, Kanthasamy AG (2020) An ex vivo brain slice culture model of chronic wasting disease: implications for disease pathogenesis and therapeutic development. Sci Rep 10(1): 7640. https://doi.org/10.1038/s41598020-64456-9 4. Falsig J, Sonati T, Herrmann US, Saban D, Li B, Arroyo K, Ballmer B, Liberski PP, Aguzzi A (2012) Prion pathogenesis is faithfully reproduced in cerebellar organotypic slice cultures. PLoS Pathog 8(11):e1002985. https://doi. org/10.1371/journal.ppat.1002985 5. Misra S, Moro CF, Del Chiaro M, Pouso S, Sebestye´n A, Lo¨hr M, Bjo¨rnstedt M, Verbeke CS (2019) Ex vivo organotypic culture system of precision-cut slices of human pancreatic

ductal adenocarcinoma. Science 9(1):2133. https://doi.org/10.1038/s41598-01938603-w 6. Kenerson HL, Sullivan KM, Labadie KP, Pillarisetty VG, Yeung RS (2021) Protocol for tissue slice cultures from human solid tumors to study therapeutic response. STAR Protoc 2(2):100574. https://doi.org/10.1016/j. xpro.2021.100574 7. Weng PL, Aure MH, Maruyama T, Ovitt CE (2018) Limited regeneration of adult salivary glands after severe injury involves cellular plasticity. Cell Rep 24(6):1464–1470.e1463. https://doi.org/10.1016/j.celrep.2018. 07.016 8. Aure MH, Konieczny SF, Ovitt CE (2015) Salivary gland homeostasis is maintained through acinar cell self-duplication. Dev Cell 33(2):231–237. https://doi.org/10.1016/j. devcel.2015.02.013 9. Su X, Pillai S, Liu Y, Tran SD (2022) Isolation, culture, and characterization of primary salivary gland cells. Curr Protoc 2(7):e479. https://doi.org/10.1002/cpz1.479 10. Su X, Fang D, Liu Y, Ramamoorthi M, Zeitouni A, Chen W, Tran SD (2016) Threedimensional organotypic culture of human salivary glands: the slice culture model. Oral Dis 22(7):639–648. https://doi.org/10.1111/ odi.12508

Chapter 5 Differentiation of Pig Gastric Primary Cells into Mucus Producing Epithelial Cells Macarena P. Quintana-Hayashi, Sinan Sharba, and Sara K. Linde´n Abstract There is a growing interest in the development of in vitro models that mimic the intrinsic characteristics of cells in vivo to replace and/or reduce the use of experimental animals. The stomach is lined with mucus secreting epithelial cells, creating a thick mucus layer that protects the underlying epithelial cells from acid, pathogens, and other harmful agents. Mucins are a main component of the mucus layer, and their secretion is an important protective feature of epithelial cells in vivo. Here, we present a method that differentiates pig gastric primary cells into mucin secreting epithelial cells by culturing the cells on polyester membranes under semi-wet interface for 14 days, using differentiation medium containing the N-[(3,5-difluorophenyl) acetyl]-L-alanyl-2-phenyl]glycine-1,1-dimethylethyl ester (DAPT) in the basolateral compartment for the first 7 days and subsequent 7-day culture in non-differentiation medium. The in vitro mucosal surfaces created by these cells are harvested 2 weeks post confluence, and two preservation methods are described to fix the monolayers for further analysis. Key words Pig, Primary cells, Spheroids, Gastric, Antrum, Mucus, Mucins, Epithelial cells

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Introduction To mimic the mucosal surfaces, the ability of the cells to form an adherent continuous polarized layer is a prerequisite to ensure bacteria interact with the apical surface of the cells and avoid introducing nonnatural targets by limiting access to the basolateral surface. Epithelial cells lining the gastrointestinal tract continuously secrete a mucous layer. Often, in vitro cell cultures are very different from in vivo mucosal surfaces as they do not polarize or form tight epithelia, lack important components of the glycocalyx, and lack the essential adherent mucus layer. Although some cell lines produce mucins, they do not form the adherent mucus layer present in vivo. We have previously described a method for an in vitro model derived from the HT29-MTX-E12

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human intestinal cancer epithelial cell line that reproduces key characteristics of mammalian intestinal mucosal surfaces in vivo, promoting disorganized/non-polarized/non-in vivo like cells to develop a mucus producing organized tight epithelium with an in vivo like cell morphology [1]. Pig intestinal organoids have been used to develop epithelial monolayers with tight junctions and Muc2 mucin expression [2]. Monolayers derived from human gastric glands also had tight junctions and expressed mainly MUC6 but a low amount of MUC5AC [3]. Most gastric cell lines do not polarize, form a tight epithelium, or secrete mucins when cultured under standard or semi-wet conditions with and without mechanical and chemical stimulation with a Notch γ-secretase inhibitor [4, 5], highlighting the need for robust in vitro experimental systems to analyze mucus production, secretion, regulation, and host-pathogen interactions in the stomach. Here, we propose a method where pig gastric primary cells are differentiated into a polarized mucus producing in vitro epithelium. Initially, pig gastric cells are isolated from pig antral gastric tissue and expanded in spheroid culture in vitro (Fig. 1a–c), using a method adapted from a previously published protocol for the isolation of mouse gastrointestinal stem cells [6]. To achieve an adequate cell seeding density, pig gastric primary cells are seeded on Matrigel-coated polyester membrane inserts that allow better visibility of the cells and monolayer formation compared to polycarbonate membranes. The cells are incubated in a 5% CO2 environment at 37 °C until confluent; after which, cells are differentiated into mucus producing cells by culture under a semi-wet interface with chemical stimulation. Pig gastric primary cells are chemically stimulated by culture in differentiating medium containing the N-[(3,5-difluorophenyl)acetyl]-L-alanyl-2-phenyl]glycine1,1-dimethylethyl ester (DAPT) in the basolateral compartment, resulting in the differentiation of epithelial cells into mucus producing cells (Fig. 1d, e). In this in vitro model, we analyzed mucin production using PAS/Alcian blue staining of neutral and acidic mucins (Fig. 1d) and metabolic labeling of newly synthetized mucins with N-azidoacetylgalactosamine (GalNAz) (Fig. 1e). Overall, the in vitro mucus producing pig gastric epithelium aims to be a tool that can be used to increase the knowledge on mucin regulation and host-pathogen interactions at mucosal surfaces, ultimately decreasing the use of experimental animals. Tissue can be harvested from pigs that are slaughtered for other purposes, and the large size of the organs makes the isolation of large number of cells straightforward.

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Fig. 1 Pig gastric spheroids and primary cells differentiated into mucus producing cells. Morphology of pig gastric spheroids embedded in Matrigel and cultured in 50% conditioned medium for 2 days (a) 4× and (b) 20× objective. Close-up image of a single spheroid embedded in Matrigel (c). PAS/Alcian blue staining of neutral (magenta) and acidic (blue) mucins in the pig gastric monolayer at 2 weeks post confluence; cells fixed with 4% PFA. Although the mucus layer is not present in the images due to loss of secreted material during the tissue processing, a shiny, slimy layer is visible on the apical surface during culture. M = polyester membrane (e). Mucus producing monolayer derived from pig gastric primary cells stained with GalNAz (red) and counterstained with DAPI (blue) at 2 weeks post confluence; cells fixed with methanolic Carnoy’s (d). Scale bar = 50 μm

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Materials 1. Collagenase solution: 2 mg/mL collagenase type I, 50 μg/mL gentamicin dissolved in washing medium (see number 3 below) and sterile filtered (0.22 μm). 2. Freezing medium: 80% (v/v) washing medium, 10% (v/v) fetal bovine serum (FBS), and 10% (v/v) dimethyl sulfoxide (DMSO). 3. Washing medium: DMEM/F12 with HEPES supplemented with 10% (v/v) FBS, 2% (v/v) penicillin-streptomycin (P/S), and 1% GlutaMAX (Gibco, New York, USA) [6]. Pre-warmed to 37 °C. 4. 50 μg/mL gentamicin 5. 50% conditioned medium: prepared in-house from L-WRN cells (mouse L-fibroblast cells) according to a previously published protocol [6]. Pre-warmed to 37 °C. 6. Differentiation medium: 50% conditioned medium containing 10 μM Rock inhibitor and supplemented with 10 μM of the γ-secretase inhibitor DAPT dissolved in dimethyl sulfoxide (DMSO). 7. Dulbecco’s phosphate-buffered saline (DPBS): w/o Ca++ and Mg++. 8. 0.5 M EDTA ultrapure, pH 8.0. 9. Matrigel Membrane Matrix (Corning, New York, USA). 10. 10% Matrigel solution: 10% (v/v) Matrigel Membrane Matrix in cold DPBS. 11. 12-well Nunc™ cell culture treated multi-well plate with the Nunclon™ Delta Surface treatment. 12. Transwell™ multi-well plate with polyester membrane inserts: 12 inserts in a 12-well plate with a 0.4 μm pore size polyester membrane (Corning, New York, USA). 13. TrypLE™ Express Enzyme (1×), no phenol red (Gibco, New York, USA). 14. Rock inhibitor (ROCKi): Y-27632 (Bio-Techne, Minneapolis, USA).

dihydrochloride

15. ALK receptor inhibitor: inhibits the transforming growth factor-β (TGF-β) type I receptor/ALK, SB 431542 (Bio-Techne, Minneapolis, USA). 16. Trypan blue solution: 0.4% phosphate-buffered saline. 17. 4% paraformaldehyde (PFA). 18. 70% ethanol (EtOH).

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19. Methanolic Carnoy’s fixative: 60% (v/v) dry methanol (max. 0.003% H2O), 30% (v/v) chloroform, and 10% (v/v) acetic acid. 20. Methanol dried: max. 0.003% H2O.

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Methods Pig gastric antrum cells are isolated and expanded and primary cells cultured onto permeable polyester membrane inserts under semiwet interface in combination with chemical stimulation, resulting in the formation of a monolayer with mucus producing epithelial-like gastric cells.

3.1 Pig Antrum Gastric Cell Isolation

1. Excise a 2 × 2 cm piece of pig stomach antrum with a scalpel. 2. Transport pig stomach biopsy in cold washing medium supplemented with 10 μM ROCKi. 3. Scrape mucus from antrum with a glass slide, and remove connective tissue and muscle layers with scissors on a small Petri dish. 4. Cut the tissue into 3–5 mm pieces. 5. Add 1 mL of collagenase solution to the tissue and pipette vigorously. 6. Incubate tissue in the Petri dish at 37 °C for 30 min, pipetting every 10 min to dissociate crypts until the majority of the crypts are isolated/separated from tissue pieces. 7. Pipette the cell suspension on top of a 70 μm pore size cell strainer placed on top of a 50 mL conical tube, followed by washing medium. 8. Centrifuge at 100 × g for 5 min at room temperature (RT). 9. Discard the supernatant and resuspend pellet in washing medium. 10. Centrifuge at 200 × g for 5 min at RT. 11. Discard the supernatant and resuspend the pellet in cold Matrigel while carefully pipetting to avoid the formation of bubbles (see Note 1). 12. While working on ice, seed cells on a 12-well cell culture plate by quickly adding a drop of cell suspension in Matrigel (using a 1 mL pipette tip) at the center of each well, and then spread the cells in Matrigel in a spiral motion using the same 1 mL pipette tip, avoiding the edges of the well (see Note 2). 13. Cover the 12-well plate with its lid and turn the plate upside down, and incubate for 10 min at 37 °C in a 5% CO2 and atmospheric O2 environment (see Note 3).

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14. Add 1 mL per well of 50% conditioned medium supplemented with 10 μM ROCKi and 1 μM ALK receptor inhibitor (see Note 4). 15. Incubate at 37 °C in a 5% CO2 and atmospheric O2 environment changing the 50% conditioned medium supplemented with inhibitors as needed (see Note 5). 3.2 Expansion of Spheroid Cultures

1. To passage the cells, discard the 50% conditioned medium and rinse the Matrigel embedded cells with DPBS containing 0.5 mM EDTA while scraping/dislodging the cells from the bottom of the well (see Note 6). 2. Centrifuge the pooled cell suspension at 200 × g for 5 min at RT. Discard the supernatant. 3. Add TrypLE Express enzyme and dissolve any cell clumps (see Note 7). 4. Incubate cells for 3 min in a water bath at 37 °C. 5. Inactivate the enzyme by adding washing medium to the dissociated cells, pipetting well enough to obtain a homogenous solution (see Note 8). 6. Proceed to steps described in Subheading 3.1 (Steps 8 to 15; see Note 9).

3.3 Semi-Wet Interface Culture of Pig Gastric Primary Cells

1. Working on ice, pre-coat each polyester membrane insert of a Transwell™ plate with 200 μL of 10% Matrigel solution. Incubate the plate at 37 °C for 20–30 min, and then discard the Matrigel solution from each insert (see Note 10). 2. Discard the 50% conditioned medium from each well of the 12-well cell culture plate containing cells, and proceed as described in Subheading 3.2 (Steps 1–5). 3. To remove any leftover debris from the spheroid culture, filter cells in washing medium using a 40 μm pore size cell strainer placed on top a 50 mL Falcon tube. Add more washing media if needed. 4. Centrifuge cells at 200 × g for 5 min at RT. 5. Discard supernatant and resuspend cells in washing medium supplemented with 50 μg/mL gentamicin. 6. Centrifuge at 500 × g for 5 min at RT and discard supernatant. 7. Resuspend cells in 50% conditioned medium containing 10 μM Rock inhibitor and 50 μg/mL gentamicin. 8. Determine the number of viable cells using the Trypan Blue exclusion technique. Dilute the cells accordingly in 50% conditioned medium containing 10 μM Rock inhibitor to obtain 18 × 104 cells per 200 μL medium.

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9. Seed 200 μL of cells suspended in 50% conditioned medium on the apical side of each Matrigel-coated Transwell™ polyester membrane insert. Add 800 μL of 50% conditioned medium containing 10 μM Rock inhibitor in the basolateral compartment. 10. Incubate plates at 37 °C in a 5% CO2 and atmospheric O2 environment for 6 days until confluent, changing the apical and basolateral medium every other day and checking the cells under a microscope. 11. A semi-wet interface with chemical stimulation is achieved by leaving 25 μL of medium in the apical compartment and replacing the basolateral medium with 800 μL of differentiation medium to differentiate the apical cells into mucus producing cells. 12. Incubate the plates at 37 °C in a 5% CO2 and atmospheric O2 environment for 7 days, changing the basolateral differentiation medium daily. 13. After 7 days, replace the basolateral medium with 800 μL of 50% conditioned medium containing 10 μM Rock inhibitor. The basolateral medium should be changed every 2 days in the beginning and daily toward the end, for a period of 7 days. Throughout this period, the membranes should continue to be incubated at 37 °C in a 5% CO2 and atmospheric O2 environment (see Note 11). 3.4 Fixation and Preservation of the Pig Gastric Monolayer 3.4.1 In-Well Fixation with 4% PFA

The cells can be fixed using two methods as per below.

1. Add 4% paraformaldehyde (PFA) from the wall of the polyester membrane insert, and leave for 1 h. 2. Replace PFA with 70% EtOH until further processing. 3. Cut out and “sandwich” the membranes working with two membranes at a time (both membranes should have undergone the same experimental treatment), cut out the membranes from their inserts using a surgical scalpel, and with the help of a small forceps, carefully place one membrane on top of the other membrane making sure that both apical surfaces of the membranes are facing each other. Place the sandwiched membranes between two foam biopsy pads inside an embedding cassette (see Note 12). 4. Immerse the membranes in 70% EtOH until processing. 5. In a tissue processor, infiltrate membranes in paraffin wax starting with changes of increasing EtOH (70%, 80%, 95%, and three changes of 100% EtOH), followed by three changes of Histo-Clear and three changes of paraffin wax (1 h each step).

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6. Dewax membranes at 65 °C for 5 min. Carefully remove biopsy pads, and embed membranes into paraffin blocks as we previously described [1]. 3.4.2 Methanolic Carnoy’s Fixation

1. In this method, the membranes need to be cut out from their inserts and “sandwiched” before fixation; therefore, proceed to Step 3 described in Subheading 3.4.1 (see Note 13). 2. Immerse the membranes inside the embedding cassettes in methanolic Carnoy’s fixative for 24 h. 3. Immerse the membranes in dried methanol for 30 min. 4. In a tissue processor, infiltrate membranes in paraffin wax starting with two changes of Histo-Clear (10 min each), followed by three changes of paraffin wax (15 min each). 5. Proceed to Step 6 described in Subheading 3.4.1.

4

Notes 1. The total volume of Matrigel is calculated based on 30 μL per well to be seeded. If necessary, cut off the tip of a 1 mL pipette tip to prevent clogging. 2. Matrigel and the 12-well plate should be kept cold by placing them on ice before seeding the cells. 3. Incubating the plate upside down prevents the cells from attaching to the bottom of the well. 4. Conditioned media should be added carefully on the edge of the well so to not disturb the cells in Matrigel. 5. Pig gastric spheroids can be frozen in freezing medium and placed at -70 °C for 1 week and then transferred to -150 °C or liquid nitrogen. 6. This step requires scraping the bottom of the wells using a 1 mL pipette tip. Additional DPBS containing 0.5 mM EDTA can be added to the wells at the end of this procedure ensuring detachment of all cells from the wells. 7. The total volume of enzyme to be added is based on 100 μL per well containing cells. 8. Volume of washing medium should double the volume of enzyme. After this step, the number of viable cells can be determined using the Trypan Blue exclusion technique if needed. Addition of gentamicin as a supplement to the washing medium and 50% conditioned medium can further aid in the prevention of bacterial contamination of the cultures if necessary.

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9. In our experience, cells grow better when split to a maximum ratio of 1:3 at a time. Splitting the cells twice will result in more than enough cells to seed 6 12-well membrane inserts. 10. Transparent polyester membranes, unlike polycarbonate membranes, facilitate the observation of cells and assessment of the monolayer under a microscope. 11. Several pig cytokines are commercially available and can be added to the basolateral medium to treat the monolayer. For bacterial infection studies, add the pathogen on the apical side of the membrane insert, and replace the basolateral medium with antibiotic-free 50% conditioned medium. 12. Single membranes can be “sandwiched” with filter paper cut out to match the size of the membrane. 13. Methanolic Carnoy’s is the preferred fixative to retain/preserve mucus during tissue processing; however, in-well fixation of the membranes cannot be performed as this fixative dissolves the plastic insert.

Acknowledgments This work was supported by the Swedish Research Council (201901152) and the Magn. Bergvall Foundation (2021-04307). References 1. Quintana-Hayashi MP, Linden SK (2018) Differentiation of gastrointestinal cell lines by culture in semi-wet interface. Methods Mol Biol 1817:41–46. https://doi.org/10.1007/978-14939-8600-2_5 2. van der Hee B, Loonen LMP, Taverne N, Taverne-Thiele JJ, Smidt H, Wells JM (2018) Optimized procedures for generating an enhanced, near physiological 2D culture system from porcine intestinal organoids. Stem Cell Res 28:165–171. https://doi.org/10.1016/j.scr. 2018.02.013 3. Boccellato F, Woelffling S, Imai-Matsushima A, Sanchez G, Goosmann C, Schmid M, Berger H, Morey P, Denecke C, Ordemann J, Meyer TF (2019) Polarised epithelial monolayers of the gastric mucosa reveal insights into mucosal homeostasis and defence against infection. Gut 68(3):400–413. https://doi.org/10.1136/ gutjnl-2017-314540

4. Linden SK, Driessen KM, McGuckin MA (2007) Improved in vitro model systems for gastrointestinal infection by choice of cell line, pH, microaerobic conditions, and optimization of culture conditions. Helicobacter 12(4): 3 4 1 – 3 5 3 . h t t p s : //d o i . o r g / 1 0 . 1 1 1 1 / j . 1523-5378.2007.00509.x 5. Navabi N, McGuckin MA, Linden SK (2013) Gastrointestinal cell lines form polarized epithelia with an adherent mucus layer when cultured in semi-wet interfaces with mechanical stimulation. PLoS One 8(7):e68761. https://doi.org/ 10.1371/journal.pone.0068761 6. Miyoshi H, Stappenbeck TS (2013) In vitro expansion and genetic modification of gastrointestinal stem cells in spheroid culture. Nat Protoc 8(12):2471–2482. https://doi.org/10. 1038/nprot.2013.153

Chapter 6 Isolation, Culture, and Microscopic Imaging of Guinea Pig Primary Gastric Tissue Cells Weronika Gonciarz and Magdalena Chmiela Abstract In this chapter, the procedure of isolation and propagation of guinea pig gastric tissue primary cells in cell culture in vitro is presented. Selected methods of microscopic imaging of cells are shown, including monitoring the ability of cells to migrate as a determinant of their activity. The primary cells that expanded in cell cultures in vitro have characteristics of natural cells and facilitate studying both the spontaneous and induced biological processes on the cellular level. Particularly, the primary cells derived from the guinea pig stomach were found to be a good model for studying the effects of bacteria-host interactions and the development of inflammatory responses driven by gastric pathogen Helicobacter pylori. Key words Primary cells, Cell culture, Gastric tissue

1

Introduction The ability to isolate and culture defined cell types in vitro provides a valuable tool for studying the complexity of biological processes at the cellular level when these phenomena are difficult to study in a living organism. Primary cell cultures may be established by enzymatic dissociation of cells from a given tissue and placing these cells into culture medium for further expansion or by allowing cells to migrate from tissue specimens (explants) that have been placed into culture medium [1–4]. Primary culture-derived cells have characteristics of naturally occurring cells. They may respond by activation to different stimuli and proliferate; however, the number of cell divisions in vitro is limited [5].

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Materials

2.1 Media and Reagents for Cell Cultures and Microscopic Imaging

1. Medium 1: DMEM (Dulbecco’s Modified Eagle Medium) supplemented with 100 U/mL penicillin, 100 U/mL streptomycin, and 0.025 μg/mL amphotericin B. 2. Medium 2: Ham’s F-12 medium and DMEM (1:1) supplemented with 10% heat inactivated FCS (fetal calf serum), 30 min, 56 °C in water bath, 100 U/mL penicillin, 100 U/mL streptomycin, 0.025 μg/mL amphotericin B, L-glutamine 2 mM/ mL, 1% HEPES (N-2-hydroxyethylpiperazine-N-2-ethane sulfonic acid), 0.01 μg/mL EGF (epidermal growth factor), 0.005% dexamethasone solution in RPMI (Roswell Park Memorial Institute)-1640 medium. 3. Solution 1: Hanks’ buffer (for cell culture) supplemented with penicillin (100 U/mL), streptomycin (100 μg/mL), and amphotericin B (0.025 μg/mL). 4. Solution 2: 2% BSA (bovine serum albumin) solution in Hanks’ buffer. 5. Solution 3: 5% BSA solution in DMEM medium (Medium 1) supplemented with penicillin (100 U/mL), streptomycin (100 μg/mL), and amphotericin B (0.025 μg/mL). 6. Solution 4: 0.25% trypsin solution in RPMI-1640 medium. 7. Solution 5: PBS (phosphate-buffered saline) pH 7.4, liquid, sterile filtered, suitable for cell culture. 8. Solution 6: 4% formaldehyde solution in PBS. 9. Solution 7: DAPI (4′,6-diamidino-2-phenylindole) solution in PBS, for fluorescent staining. 10. Solution 8: Phalloidin Texas Red solution in PBS, for fluorescent staining. 11. Six-well cell culture plates. 12. 96-well black cell culture plates with glass bottom. 13. Sterile volumetric pipettes and Pasteur pipettes. 14. Cell culture incubator with controlled temperature, humidity, and CO2 supply. 15. Inverted field light microscope. 16. Confocal microscope.

3

Methods

3.1 Isolation and Culture of Guinea Pig Primary Gastric Tissue Cells

1. Isolate the stomach from the euthanized animal (overdose of sodium barbiturate), according the Animal Research: Reporting of In Vivo Experiments (ARRIVE) guidelines and guidelines and regulations of EU directive (Directive 2010/63/EU

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of the European Parliament and of the Council of 22 September 2010 on the protection of animals used for scientific purposes) [6]. 2. Cut the organ along the greater curvature. 3. Wash the organ three times in Solution 1 (see Note 1). 4. Divide the stomach into smaller pieces and homogenize them with an electric homogenizer in 10 mL Solution 1. 5. Centrifuge at 250 × g at room temperature. 6. Remove the supernatant. 7. Add Solution 4 and mix thoroughly. 8. Incubate for 15 min at room temperature. 9. Centrifuge at 250 × g at room temperature. 10. Remove the supernatant. 11. Add Solution 2 and mix thoroughly. 12. Centrifuge at 250 × g at room temperature. 13. Remove the supernatant. 14. Add Solution 3 and mix thoroughly. ˝rker 15. Suspend the cells in Medium 2, count the cells in the Bu chamber, and adjust the cell suspension to the required density. Determine cell viability in trypan blue exclusion assay or the MTT (-(4,5-Dimethyl-2-thiazolyl)-2,5-diphenyl-2H-tetrazolium Bromide) reduction assay (see Note 2). 16. Add 1 mL of cell suspension at a density of 2 × 106 cells/mL to the wells of six-well plates. 17. Incubate for 24 h (5%CO2, 37 °C). 18. Wash out unbound cells three times with Solution 5 (add 1 mL Solution 5, and wash thoroughly and remove) (see Note 3). 19. Add Medium 1. 20. Incubate for 48 h (5%CO2, 37 °C). 21. Evaluate the cell morphology under the light microscope, at magnification ×10 (see Fig. 1). 22. Use the cells for the planned experiment. 3.2 Imaging of Guinea Pig Primary Gastric Tissue Cells in Confocal Microscope

1. Culture primary cells in Medium 1. 2. Bring the cell suspension to a density of 2 ×104 cell/mL in Medium 1 (see Note 2). 3. Add 100 mL of cell suspension in a well of a 96-well black plate with glass bottom. 4. Incubate overnight (5%CO2, 37 °C). 5. Monitor cells under the light inverted microscope. 6. Remove Medium1 from the wells.

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Fig. 1 Representative microscopic image of primary gastric tissue cells derived from the stomach of guinea pig in inverted field microscope, ×10 magnification

7. Wash the cell monolayer once with 100 μL of Solution 5 (see Note 3). 8. Fix the cells with 100 μL of Solution 6. 9. Incubate for 15 min (room temperature). 10. Remove Solution 6 from the wells. 11. Wash the cell monolayer three times with 100 μL of Solution 5 (see Note 3). 12. Add 100 μL of Solution 7 to the wells (see Note 6). 13. Incubate at room temperature in the dark (see Note 6). 14. Remove Solution 7 from the wells. 15. Wash the cell monolayer three times with 100 μL Solution 5 (see Note 3). 16. Add 100 μL of Solution 8 to all wells (see Note 6). 17. Incubate at room temperature in the dark (see Note 6). 18. Remove Solution 8 from the wells. 19. Wash the monolayer three times with 100 μL Solution 5 (see Note 3). 20. Add 100 μL of Solution 5 to the wells (see Note 6). 21. Image cells in a confocal microscope at wavelengths dedicated to a specific fluorescent dye: DAPI excitation 340 nm, emission 488 nm, Phalloidin Texas Red excitation 591 nm, emission 608 nm (see Notes 4–6) (Fig. 2). 3.3

Migration Assay

1. Culture primary cells in Medium 1. 2. Adjust cell suspension to a density of 2 × 106 cell/mL in Medium 1. 3. Add 1 mL of cell suspension to the wells of six-well culture plate. 4. Incubate overnight (5%CO2, 37 °C).

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Fig. 2 Representative microscopic image of primary gastric tissue cell derived from the stomach of guinea pig in a confocal microscope, stained with DAPI (nucleus-blue) and Phalloidin Texas Red (actin – red), ×64 magnification [6]

5. Monitor the cell confluence in the light inverted field microscope. 6. Remove Medium1 from the wells. 7. Wash the cell monolayer once with 100 μL of Solution 5 (see Note 3). 8. Make a scratch in the cell monolayer with a sterile pipette tip. 9. Wash the cell monolayer once with 100 μL of Solution 5 (see Note 3). 10. Add to the wells 1 mL of fresh Medium 1. 11. Make a photo of wounds at time 0. 12. Incubate the plates overnight (5%CO2, 37 °C). 13. Evaluate wound healing by migrating cells after 24, 48, and 78 h (Fig. 3).

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Conclusions Primary cells from gastric tissue of guinea pig obtained by the above procedure have been used in numerous studies in vitro to examine the adhesion of H. pylori to gastric tissue cells, to establish the role of these bacteria in gastric barrier disintegration due to induction of oxidative stress, followed by the increased number of cells undergoing apoptosis. This model facilitated the monitoring of secretion alarming cytokines, including interleukin (IL)-33, in response to H. pylori infection and to follow the cell pro-regenerative activity [6–10].

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Fig. 3 The images of primary gastric tissue cells derived from the stomach of guinea pig migrating in the “wound” area, in the cell monolayer (images from inverted field microscope, ×10 magnification). The arrows show the percentage of wound healed

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Notes 1. When washing the stomach, make sure you have removed all its content since food remnants may affect the successful cell isolation. 2. Remember that the number of cell cycles of primary cells is limited. Check the cell morphology and cell viability by trypan blue exclusion assay prior to each experiment. Cell viability during various experimental procedures might be determined on the basis of the cell metabolic activity in the reference MTT reduction assay (ISO: ISO norm 10993-5) [7]. 3. The primary cells are more sensitive than neoplastic cells to experimental conditions, so handle them carefully particularly when washing. This will prevent against cell monolayer detachment before further specific treatment of cells. 4. For cell imaging, use the fluorescent labels DAPI and Phalloidin Texas Red, in a concentration and time recommended by the manufacturer. 5. Imaging cells in a confocal microscope should preferably be developed immediately or no later than 7 days due to bleaching of the fluorescent marker. If you can’t image fluorescently stained cells immediately, keep them in the refrigerator (4 °C). 6. If you are not currently using cells, store them in -80 °C in sterile freezing tubes. Prepare stocks in standard freezing medium (mix 1 mL cell culture dimethyl sulfoxide 10% and 9 mL heat inactivated FCS). Grow cells from the stock for the next experiment. Total number of cell passages is 10.

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References 1. Ganjibakhsh M, Aminishakib P, Farzaneh P et al (2017) Establishment and characterization of primary cultures from Iranian oral squamous cell carcinoma patients by enzymatic method and explant culture. J Dent (Tehran, Iran) 14:191 2. Peehl DM (2005) Primary cell cultures as models of prostate cancer development. Endocr Relat Cancer 12:19–47 3. Farnie G, Clarke RB, Spence K et al (2007) Novel cell culture technique for primary ductal carcinoma in situ: role of Notch and epidermal growth factor receptor signaling pathways. J Natl Cancer Inst 99:616–627 4. Pu Y, Li Y, Han Y, Yuan C et al (2001) Rat keratinocyte primary cultures based on conductive polypyrrole primary cell culture technique. J Biomed Eng 18:416–418 5. Asad S, Kuroda M, Aoyagi Y et al (2011) Ceiling culture-derived proliferative adipocytes retain high adipogenic potential suitable for use as a vehicle for gene transduction therapy. Am J Phys Cell Phys 301:C181–C185 6. Gonciarz W, Walencka M, Moran AP et al (2019) Upregulation of MUC5AC production

and deposition of Lewis determinants by helicobacter pylori facilitate gastric tissue colonization and the maintenance of infection. J Biomed Sci 26:23 7. Gonciarz W, Pia˛tczak E, Płoszaj P et al (2022) Salvia cadmica extracts rich in polyphenols neutralize a deleterious effects of oxidative stress driven by helicobacter pylori lipopolysaccharide in cell cultures of gastric epithelial cells or fibroblasts. Ind Crop Prod 178:114633 8. Gonciarz W, Krupa A, Hinc K et al (2019) The effect of helicobacter pylori infection and different H. pylori components on the proliferation and apoptosis of gastric epithelial cells and fibroblasts. PLoSOne 7(1):e0220636 9. Gonciarz W, Krupa A, Chmiela M (2020) Proregenerative activity of IL-33 in gastric tissue cells undergoing helicobacter pylori-induced apoptosis. Int J Mol Sci 21:1801 10. Gonciarz W, Krupa A, Moran P et al (2021) Interference of LPS H. pylori with IL-33 driven regeneration of Caviae porcellus primary gastric epithelial cells and fibroblasts. Cell 10:1385

Chapter 7 Method for Two-Dimensional Epithelial Monolayer Formation Derived from Mouse Three-Dimensional Small Intestinal Organoids Yuta Takase and Toshio Takahashi Abstract The intestinal epithelium is composed of two distinct structures, namely, the villi and crypts. The base of the crypts contains intestinal stem cells (ISCs), which support the high regenerative capacity of the intestinal epithelium. With the establishment of the three-dimensional (3D) organoid culture method, the cellular and molecular mechanisms of differentiation, proliferation, and maintenance of ISCs have been widely analyzed. However, the sphere-like morphology of the 3D organoids prevents access to the apical side of the epithelium. To overcome this limitation, two-dimensional (2D) monolayer cultures derived from 3D organoids have been attempted; however, 2D culture methods for the mouse small intestine have not been well established. In this study, we developed a simple method that uses only commercially available materials, for the formation of 2D epithelial monolayers from mouse 3D small intestinal organoids. Using this method, confluent 2D epithelial monolayers were established within 4 days. This monolayer showed stable tight junction and included ISCs and differentiated intestinal cells. It also showed physiologically relevant transepithelial electrical resistance values. On the basis of these findings, this method opens a novel platform for analyzing the physiology of the intestinal epithelium, its interaction with microbes, and mechanisms of villus formation. Key words Mouse small intestine, 2D epithelial monolayer, 3D organoid, Sphere-like morphology, Tight junction, Intestinal stem cells, Cell differentiation, Transepithelial electrical resistance

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Introduction The surface of the small intestine is lined by an epithelial monolayer, which is organized into two distinct structures: villi and crypts [1, 2]. The villi contain a variety of differentiated cells that are responsible for the digestion and absorption of ingested food; they also serve as a barrier against various microbes [1, 2]. The base of the crypts contains proliferative leucine-rich repeat-containing G-protein-coupled receptor 5 (Lgr5)+ intestinal stem cells (ISCs), which mainly give rise to five types of differentiated intestinal cells (viz., absorptive epithelial cells, enteroendocrine cells, tuft

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cells, goblet cells, and Paneth cells) [3, 4]. In mice, intestinal cells are newly generated in the crypts and are lost within 5 days following apoptosis at the tips of the villi [1, 2, 4]. Thus, homeostasis of the small intestinal epithelium is supported by the differentiation, proliferation, and maintenance of ISCs. The establishment of the three-dimensional (3D) intestinal organoid culture method has accelerated the cellular and molecular analyses of the intestinal epithelium [4–6]. The intestinal organoids obtained by culturing in 3D environments that mimic the conditions of in vivo systems exhibit a spherical structure with crypt-like buddings. As 3D organoids do not contain neurons or immune cells, they are useful for the analysis of cell-cell interactions within the intestinal epithelium [6]. For example, our group has reported that nonneuronal acetylcholine (ACh) secreted from the intestinal epithelium regulates ISC proliferation and differentiation via muscarinic and nicotinic ACh receptors [7, 8]. However, the apical side of the 3D organoids faces the lumen, and no villus structures are present. Therefore, the 3D organoids are not suitable for studies on the physiology of the intestinal epithelium, interaction with microbes, and mechanism of villus formation. Various approaches have been adopted to address these issues. For example, methods of injecting substances into the lumen of a 3D organoid or turning the organoid inside out have been reported; however, these methods are difficult to perform and inefficient [9, 10]. Furthermore, there is a well-known analysis system that uses human colon carcinoma cell line (Caco-2)-derived monolayers [11, 12]. However, Caco-2-derived monolayers differ from the small intestinal epithelium in terms of gene expression and physiological characteristics [12, 13]. Recently, the formation of 2D epithelial monolayers from 3D intestinal organoids has been attempted as an experimental system that can solve such problems. For human small intestinal cell monolayers, several methods using 3D intestinal organoid- or induced pluripotent stem cell (iPSCs)derived intestinal epithelial cells have been reported [13–15]. In contrast, monolayer formation from mouse 3D small intestinal organoids is difficult because of rapid cell turnover. In a few cases, methods requiring primary tissue (myofibroblast or enteric nervous system)-derived culture medium or long culture periods have been reported [16, 17]. Herein, we report a method for the efficient formation of 2D epithelial monolayers from mouse 3D small intestinal organoids using only commercially available reagents and media (Fig. 1). In this method, single cells or small clusters derived from growing 3D organoids are cultured in the translucent inserts that are coated with a thin layer of extracellular matrix (ECM). Using this method, a confluent monolayer with stable intercellular junctions was formed within 4 days. This monolayer also contained ISCs and differentiated intestinal epithelial cells. Additionally, the

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Fig. 1 Schematic diagram of 2D epithelial monolayer formation from mouse 3D intestinal organoids

transepithelial electric resistance (TEER) values of this monolayer were within the expected physiological range (50–100 Ω cm2) for the mouse small intestine [18]. This method will serve as a novel platform for analyzing the physiology of the intestinal epithelium, its interaction with microbes, and mechanisms of villus formation.

2

Materials

2.1 Crypt Isolation and 3D Organoid Formation

1. C57/BL6: a standard wild-type strain of Mus musculus (see Note 1). 2. 24-well plates. 3. Matrigel (Corning). 4. 10,000 U/mL penicillin-streptomycin (pen-strep). 5. 10 mg/mL gentamicin sulfate solution (gentamicin). 6. Advanced Dulbecco’s Modified Eagle Medium/Ham’s F-12 (DMEM/F-12). 7. 200 mM L-glutamine (100× stock). 8. Phosphate-buffered saline (PBS, pH 7.4). 9. 0.5 M ethylenediaminetetraacetic acid (EDTA)/PBS (pH 8.0). 10. PBS + antibiotics (PBS-ABx): PBS containing 100 U/mL pen-strep and 50 μg/mL gentamicin.

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11. Complete DMEM: advanced DMEM/F-12 containing 100 U/mL pen-strep, 50 μg/mL gentamicin, and 1% (v/v) fetal bovine serum (FBS). 12. Sorbitol DMEM: 2% (w/v) sorbitol/complete DMEM. 13. 20 μg/mL mouse epidermal growth factor (EGF) (1000× stock; PeproTech). 14. 250 μg/mL recombinant mouse R-Spondin 1 (500× stock; R&D Systems). 15. 20 μg/mL recombinant mouse noggin (200× stock; R&D Systems). 16. 3D organoid culture medium: advanced DMEM/F12 containing antibiotics (50 U/mL pen-strep and 25 μg/mL gentamicin), 2 mM L-glutamine, and growth factors (20 ng/mL EGF, 100 ng/mL noggin, and 500 ng/mL R-Spondin1). 2.2 Maintenance of 3D Organoids

1. Cell Recovery Solution (Corning).

2.3 TwoDimensional Epithelial Monolayer Formation

1. MatriMix 511 (Nippi) (see Note 2).

2. 15 mL low retention tube.

2. Transparent cell culture inserts for 24-well plates (pore size, 0.4 μm; Greiner). 3. Translucent cell culture inserts for 24-well plates (pore size, 0.4 μm; Corning). 4. TrypLE Express Enzyme (Gibco). 5. Cell Counter CDA-1000 (Sysmex). 6. IntestiCult Organoid Growth Medium for Human (Veritas). 7. 0.8 M valproic acid (1000× stock). 8. 1 mM Y-27632 (ROCK inhibitor) (100× stock). 9. 2D monolayer culture medium: IntestiCult Organoid Growth Medium containing 50 U/mL pen-strep, 25 μg/mL gentamicin, 0.8 mM valproic acid, and 10 μM Y-27632 (see Note 3).

2.4 Fluorescent Immunohistochemistry

1. 4% (w/v) paraformaldehyde/PBS. 2. 0.2% (v/v) Triton X-100/PBS (PBST). 3. Blocking solution: 1% (w/v) BSA/PBST. 4. Primary antibodies. 5. Fluorescent secondary antibodies. 6. 1 mg/mL Hoechst 33342. 7. VECTASHIELD Vibrance Antifade Mounting Medium (Vector Laboratories). 8. FV3000 confocal microscopy (EVIDENT).

Method for Epithelial Monolayer Formation from 3D Organoids

2.5 Transepithelial Electric Resistance Measurement

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1. Millicell ERS-2 Voltohmmeter (Merck).

Methods

3.1 Crypt Isolation and 3D Organoid Formation

Crypt isolation and 3D organoid formation and maintenance were performed as described previously [19, 20]. Before crypt isolation and 3D organoid culture, frozen Matrigel was dissolved at 4 °C and placed on ice. 1. Isolate the small intestine, including the duodenum and proximal half of the jejunum. Remove the fat tissues from the small intestine. 2. Using a 5 mL syringe, flush the small intestine with cold PBS-ABx to clear the luminal content. 3. Cut the tissue open lengthwise, and then wash it with cold PBS-ABx. 4. Cut the tissue into 5 × 5 mm pieces, and then wash them with cold PBS-ABx. Collect the fragments in a 50 mL tube. 5. Wash the fragments again with 25 mL of cold PBS-ABx. 6. Remove PBS-ABx, add 25 mL of cold PBS-ABx containing 2 mM EDTA, and then incubate the samples for 30 min on ice. 7. During the incubation period, place a 24-well plate in a 37 °C incubator. 8. Remove EDTA solution, add 25 mL of fresh cold PBS-ABx, and then manually shake the tissue fragments up and down vigorously 30–40 times to disintegrate the crypt-villus complexes (see Note 4). 9. Pass the resulting suspension through a 70 μm cell strainer to remove residual villous material. 10. Centrifuge the isolated crypts at 390 × g for 3 min at 4 °C, and then discard the supernatant. 11. Resuspend the pellet in 20 mL of sorbitol DMEM with pipetting, and then divide the solution into two 10 mL aliquots in 15 mL tubes. 12. Centrifuge the isolated crypts at 80 × g for 3 min at 4 °C, and then discard the supernatant (see Note 5). 13. Add 10 mL of sorbitol DMEM to each tube. After suspension, centrifuge the isolated crypts at 80 × g for 3 min at 4 °C. 14. Repeat Step 13. 15. Discard the supernatant, add 10 mL of complete DMEM, and wait for 1 min (see Note 6).

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16. After 1 min, collect each 10 mL aliquot (total 20 mL) of supernatant and pass through a 70 μm cell strainer. 17. Count the purified crypts under an inverted microscope, and then collect sufficient crypt samples (see Note 7). Centrifuge the mixture at 290 × g for 3 min at 4 °C. 18. Suspend the pellets with 40 μL Matrigel per well, and then seed in the pre-warmed 24-well plate. 19. Incubate the 24-well plate for 15 min at 37 °C. 20. After the polymerization of Matrigel, add 500 μL of 3D organoid culture medium. Begin 3D organoid culture at 37 °C. The medium is changed every 2–3 days. 3.2 Maintenance of 3D Organoids

The growing 3D organoids were maintained once for 4–7 days. The passage ratio is one well to two to three wells. 1. Discard the old culture medium and wash the samples three times with 1 mL of cold PBS. 2. Add 1 mL of Cell Recovery Solution, and then mechanically peel off the Matrigel. Collect the samples in a 15 mL low retention tube (see Note 8). 3. Incubate the samples for 15 min on ice with shaking. During the incubation period, place a 24-well plate in a 37 °C incubator. 4. Centrifuge the samples at 200 × g for 1 min at 4 °C and carefully discard the supernatant. 5. Add 1 mL of cold PBS and mechanically dissociate the samples into single crypt domains using a 1 mL pipette. 6. Centrifuge the samples at 200 × g for 2 min at 4 °C, and carefully discard the supernatant. 7. Suspend the pellets with 40 μL Matrigel per well, and then seed the samples in the pre-warmed 24-well plate. 8. Begin 3D organoid culture (as described in Steps 19–20, Subheading 3.1).

3.3 TwoDimensional Epithelial Monolayer Formation

Two-dimensional (2D) epithelial monolayers are formed using growing 3D organoids (Fig. 2a). The 2D epithelial monolayers are stably formed until at least the tenth passage. Use 2 wells of 3D organoids to form 1 well of 2D epithelial monolayers (approximately 200,000–400,000 cells per well). As a translucent insert induces a more confluent 2D epithelial monolayer than a transparent one, use translucent inserts, except to examine the processes of 2D epithelial monolayer formation (Fig. 2b, c).

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Fig. 2 Processes of 2D epithelium monolayer formation derived from mouse 3D organoids (a) Representative image of 3D intestinal organoids at the sixth passage for generating 2D epithelial monolayers. Scale bar: 200 μm. (b) Representative images of the processes of 2D intestinal epithelial monolayer formation for 4 days. Scale bar: 100 μm. (c) Representative images of 2D monolayers on day 4. Upper panels show the images of the monolayer cultured in the transparent insert. Bottom panels show the images of the monolayer in the translucent one. Nuclei were observed using Hoechst 33342 staining (cyan). Scale bars: 500 μm

1. Coat 24-well culture inserts with 75 μL of MatriMix (511) solution, according to the manufacturer’s instructions (see Note 9). 2. Incubate the 24-well plate for 90 min at 37 °C. 3. Isolate 3D organoids from the Matrigel (as described in Steps 1–4, Subheading 3.2).

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4. Add 3 mL of cold Advanced DMEM/F12 and mechanically dissociate the samples into single crypt domains using a 1 mL pipette. 5. Centrifuge the samples at 200 × g for 2 min at 4 °C and carefully discard the supernatant. 6. Suspend the pellets in 2 mL of TrypLE Express Enzyme. 7. Incubate the samples at 37 °C for 3–5 min and mechanically dissociate them into single cells or small clusters by vigorous pipetting. 8. Add 2 mL of cold Advanced DMEM/F12 and mix the samples thoroughly by pipetting. Centrifuge the samples at 300 × g for 2 min at 4 °C and carefully discard the supernatant. 9. Resuspend the pellet of the dissociated cells in 1 mL of cold Advanced DMEM/F12, and then determine the cell concentration using cell counter CDA-1000. 10. Centrifuge the samples at 300 × g for 2 min at 4 °C, and then carefully discard the supernatant. 11. Resuspend the pellet in 100 μL/well of cold 2D monolayer culture medium. 12. After the polymerization of MatriMix (511) thin layer, remove the excess solution from the inserts. 13. Add 500 μL of 2D monolayer culture medium to the bottom wells, and then gently add 100 μL of the cell suspension to the upper inserts. Begin 2D epithelial monolayer culture at 37 °C. 14. Change the medium every 1–2 days (see Note 10). The 2D monolayers generally reach 100% confluency within 4 days. 3.4 Fluorescent Immunohistochemistry

Examine the epithelial characteristics (tight junctions, ZO-1; adherence junctions, E-cadherin; apical and lateral membranes, F-actin) and intestinal cell differentiation (stem cells, LGR5; Paneth cells, lysozyme; tuft cells, DCLK1) in 2D epithelial monolayers on day 4 with the markers mentioned (Figs. 3a, b, 4). Basically, the buffer/reagent changes are performed at volumes of 100 μL for the upper inserts and 500 μL for the bottom wells. 1. Discard the old culture medium and wash the samples three times with cold PBS. 2. Fix the samples in 4% paraformaldehyde/PBS for 60 min at 4 °C. 3. Wash the samples three times for 5 min each in PBS. 4. Incubate the samples in 0.2% PBST for 5 min. 5. Block the 2D monolayers with blocking solution for 30 min.

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Fig. 3 Epithelial characteristics of 2D intestinal epithelium monolayers. (a) Low-magnification confocal images of ZO-1 (magenta, tight junctions) of 2D epithelial monolayers on day 4. The nuclei were stained with Hoechst 33342 staining (cyan). Scale bar: 50 μm. (b) High-magnification confocal images of E-cadherin (green, adherence junctions), ZO-1 (magenta), and F-actin (white) of 2D epithelial monolayers on day 4. The nuclei were stained with Hoechst 33342 staining (cyan). Upper and bottom panels show the X-Y and X-Z sections, respectively. Scale bars: 10 μm. (c) Transepithelial electrical resistance (TEER) values of 2D epithelial monolayers during days 4–7 (n = 5 each)

6. Add 100 μL of primary antibodies diluted with blocking solution to the upper inserts, and replace the bottom well with flesh blocking solution. Incubate the samples overnight at 4 °C. 7. Wash the samples three times for 5 min each in PBS. 8. Add 100 μL of secondary antibodies and Hoechst 33342 diluted with blocking solution to the upper inserts, and add 500 μL of blocking solution to the bottom wells. Incubate the samples for 60 min at room temperature (RT, 20–26 °C).

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Fig. 4 Intestinal cell differentiation characteristics of 2D intestinal epithelium monolayers. High-magnification confocal images of intestinal cell differentiation markers (a: LGR5, ISCs; b: lysozyme, Paneth cells; c: DCLK1, tuft cells) in 2D epithelial monolayers on day 4. The nuclei (cyan) and F-actin (white) were stained with Hoechst 33342 and phalloidin, respectively. Scale bars: 20 μm

9. Wash the samples three times for 5 min each in PBS. 10. Cut the insert membranes, and then place them on a glass slide with the monolayer-side up. 11. Mount the membranes with VECTASHIELD Vibrance Antifade Mounting Medium. 12. After at least 1 h at RT, observe and take images of the stained monolayers using FV3000 confocal microscopy.

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Measure TEER using confluent 2D monolayers on days 4–7 (Fig. 3c). Use MatriMix (511)-coated insert (without any cells) as a blank. 1. Replace the culture medium with flesh 2D monolayer culture medium, and then incubate the samples at 37 °C for at least 60 min. 2. Allow the samples to reach RT (approximately 30 min). 3. Prepare Millicell ERS-2 Voltohmmeter, according to the manufacturer’s instructions. 4. Measure the resistance of the blank and sample inserts. Repeat the process three times independently. 5. Calculate the TEER values.

4

Notes 1. This study was approved by the Suntory Animal Ethics Committee (APRV000561), and all animals were maintained in accordance with the guidelines of the Committee for the Care and Use of Laboratory Animals. The mice used for small intestinal crypt isolation were 8–12 weeks old. 2. Although 2–5% Matrigel solutions can also induce 2D monolayers, we recommend Matimix (511) because of its obvious composition and fewer lot differences. 3. This medium induces a more confluent 2D epithelial monolayer than 3D organoid culture medium. 4. After suspension, aliquot 20 μL of the solution into a 3 cm Petri dish. Examine the separated crypts and villi in the aliquot using an inverted microscope (Nikon). The following steps should be performed quickly. 5. As the pellet can disintegrate easily, do not aspirate too much. Leave 2 mL of the supernatant in the tube. 6. This step is important because of the purified crypts obtained. 7. To estimate the number of crypts, aliquot three 20 μL portions of the solution into a 3 cm Petri dish. After counting the crypts, calculate the volume of the solution required for 3D organoid formation (100 crypts/well). 8. 3D organoids tend to adhere to tubes. To reduce the loss of samples, a low retention tube should be used. 9. MatriMix (511) solution is prepared using the protocol for thin-layer coating. 10. Gently perform this step because the mouse 2D monolayers can easily detach. Additionally, to reduce damage to the monolayers, the old medium in the upper inserts is not completely removed.

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Acknowledgments This work was supported by Grants-in-Aid for Scientific Research (C) (grant numbers JP19K16140 and JP22K06238) from the Japan Society for the Promotion of Science (JSPS). References 1. Clevers H (2013) The intestinal crypt, a prototype stem cell compartment. Cell 154(2): 274–284 2. Barker N (2014) Adult intestinal stem cells: critical drivers of epithelial homeostasis and regeneration. Nat Rev Mol Cell Biol 15(1): 19–33 3. Barker N, van Es JH, Kuipers J et al (2007) Identification of stem cells in small intestine and colon by marker gene Lgr5. Nature 449(7165):1003–1007 4. Beumer J, Clevers H (2021) Cell fate specification and differentiation in the adult mammalian intestine. Nat Rev Mol Cell Biol 22(1): 39–53 5. Sato T, Vries RG, Snippert HJ et al (2009) Single Lgr5 stem cells build crypt-villus structures in vitro without a mesenchymal niche. Nature 459(7244):262–265 6. Boonekamp KE, Dayton TL, Clevers H (2020) Intestinal organoids as tools for enriching and studying specific and rare cell types: advances and future directions. J Mol Cell Biol 12(8): 562–568 7. Takahashi T, Shiraishi A, Murata J (2018) The coordinated activities of nACHR and Wnt signaling regulate intestinal stem cell function in mice. Int J Mol Sci 19(3):E738 8. Takahashi T, Shiraishi A, Murata J et al (2021) Muscarinic receptor M3 contributes to intestinal stem cell maintenance via EphB/ephrin-B signaling. Life Sci Alliance 4(9):e202000962 9. Wilson SS, Tocchi A, Holly MK et al (2015) A small intestinal organoid model of non-invasive enteric pathogen-epithelial cell interactions. Mucosal Immunol 8(2):352–361 10. Co JY, Margalef-Catala` M, Li X et al (2019) Controlling epithelial polarity: a human Enteroid model for host-pathogen interactions. Cell Rep 26(9):2509–2520.e4 11. Sambuy Y, De Angelis I, Ranaldi G et al (2005) The Caco-2 cell line as a model of the intestinal

barrier: influence of cell and culture-related factors on Caco-2 cell functional characteristics. Cell Biol Toxicol 21(1):1–26 12. Sun H, Chow ECY, Liu S et al (2008) The Caco-2 cell monolayer: usefulness and limitations. Expert Opin Drug Metab Toxicol 4(4): 395–411 13. Negoro R, Takayama K, Kawai K et al (2018) Efficient generation of small intestinal epithelial-like cells from human iPSCs for drug absorption and metabolism studies. Stem Cell Reports 11(6):1539–1550 14. Sasaki N, Miyamoto K, Maslowski KM et al (2020) Development of a scalable coculture system for gut anaerobes and human colon epithelium. Gastroenterology 159(1): 388–390.e5 15. Yamashita T, Inui T, Yokota J et al (2021) Monolayer platform using human biopsyderived duodenal organoids for pharmaceutical research. Mol Ther Methods Clin Dev 22:263– 278 ˜ aga E, Tosi S et al (2019) Self16. Altay G, Larran organized intestinal epithelial monolayers in crypt and villus-like domains show effective barrier function. Sci Rep 9(1):10140 17. Puzan M, Hosic S, Ghio C et al (2018) Enteric nervous system regulation of intestinal stem cell differentiation and epithelial monolayer function. Sci Rep 8(1):6313 18. Srinivasan B, Kolli AR, Esch MB et al (2015) TEER measurement techniques for in vitro barrier model systems. J Lab Autom 20(2): 107–126 19. Takahashi T (2019) New trends and perspectives in the function of non-neuronal acetylcholine in crypt–villus organoids in mice. Methods Mol Biol 1576:145–155 20. Takase Y, Fujishima K, Takahashi T (2023) The 3D culturing of organoids from murine intestinal crypts and a single stem cell for organoid research. J Vis Exp 194:e65219

Chapter 8 Human Hepatic Spheroid Coculture Model for the Assessment of Drug-Induced Liver Injury Linhao Li and Hongbing Wang Abstract Accurate evaluation of potential drug risks such as drug-induced liver injury (DILI) continues to be a challenge faced by pharmaceutical industry and regulatory agencies. Preclinical testing has served as a foundation for the evaluation of the potential risks and effectiveness of investigational new drug (IND) products in humans. However, current two-dimensional (2D) in vitro human primary hepatocyte (HPH) culture systems cannot accurately depict and simulate the rich environment and complex processes observed in vivo, while animal studies present inherited species-specific differences and low throughput scales. Thus, there is a continued demand to establish new approaches that can better characterize DILI during drug discovery and development. Among others, the three-dimensional (3D) hepatic spheroid model comprising self-aggregated primary human hepatocytes cocultured with non-parenchymal cells (NPCs) appears to be a more accurate representation of the natural hepatic microenvironment with intercellular interactions between hepatocytes, stellate cells, Kupffer cells, liver sinusoidal endothelial cells (LSECs), and other cell types. This model holds the potential to improve the ability for long-term functional and toxicological studies. Here, we provide methodological details for this human hepatic spheroid coculture model system. Key words Human liver spheroids, 3D cell culture, Drug-induced liver injury (DILI)

1

Introduction Traditional two-dimensional (2D) human primary hepatocyte (HPH) culture has been routinely used to study liver biological functions, such as evaluation of drug absorption, distribution, metabolism, and excretion (ADME) properties and potential drug-induced liver injury (DILI) during drug development. However, this terminally differentiated 2D monolayer culture of HPHs is often associated with rapid loss of specific hepatic function and gene expression. Therefore, it is impractical to mimic the physiological environment of liver organ over the long term using the

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conventional 2D hepatocyte model. To improve the preclinical prediction of drug safety and efficacy, researchers continue to develop better model systems to stabilize liver functionality and to promote the use of improved cell- and tissue-based assays for more accurate representation of human hepatic susceptibility to drug response. Normal cell physiology and function relies predominantly on cell-cell and cell-extracellular matrix (ECM) interactions in the 3D tissue environment [1]. Over the past decades, researchers have developed various in vitro complex cell culture systems for more accurate representation of human hepatic microenvironment and better prediction of DILI, from simple 2D sandwich cultures to more complicated 3D hepatic spheroid coculture model [2, 3] and from single cell type static 3D models to multicell type coculture 3D models equipped with microfluidic flow control. Each provides different advantages and limitations. Compared to 2D cell culture models, the emerging 3D models exhibit improved resembling of the natural organ/tissue microenvironment, experienced by cells under physiological and/or pathophysiological conditions, and offer a greater potential in assessing drug disposition and pharmacokinetics (PKs) that influence drug safety and efficacy at an early stage of drug development [4–6]. In recent years, 3D hepatic spheroid coculture system has been extensively utilized in long-term functional and toxicological studies. For example, the potential of remdesivir (RDV)-induced hepatotoxicity was determined by using sandwich cultured HPHs and 3D human liver spheroids containing HPHs and NPCs. The 50% cytotoxic concentration (CC50) of RDV, ranging from 7.5 to 22.66 μM in sandwich cultured HPHs from three different liver donors, was shifted to 4.24–7.96 μM in the 3D spheroid coculture model [7]. Given that the 3D spheroid coculture maintains higher metabolic function of hepatocytes, the increased intracellular conversion of RDV to the toxic triphosphate metabolite (GS-443902) may contribute to the observed shift of RDV-induced hepatotoxicity. Here, we provide step-by-step instructions for how to generate this 3D human liver spheroid coculture model.

2 2.1

Materials Equipment

1. 51300 B2 biological safety cabinet (BSC) (Thermo Fisher Scientific, Asheville, NC) for all cell operation. 2. CO2 incubator MCO-17AIC (SANYO, Wood Dale, IL) for cell culture.

Human Hepatic Spheroid Coculture Model for the Assessment. . .

2.2 Materials, Reagents, and Solutions

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1. Costar CLS7007 96-well, Clear Round Bottom Ultra-Low Attachment Microplate (Corning, Corning, NY). 2. Seeding medium for 3D hepatic spheroids from freshly isolated hepatocytes by BioIVT: 1. INVITROGRO™ CP medium (BioIVT) 250 mL, and add 5.5 mL TORPEDO Antibiotic Mix (BioIVT). 2. William’s Medium E (Sigma-Aldrich) 500 mL, and add 5 mL ITS+ (BD Biosciences, Bedford, MA), 5 mL L-glutamine (Invitrogen), 5 mL penicillin/streptomycin (Invitrogen), 5 μL dexamethasone (10 mM) (Sigma-Aldrich), and 50 mL fetal bovine serum (FBS) (Invitrogen). 3. Seeding medium for 3D hepatic spheroids from cryoplateable spheroid qualified hepatocytes (BioIVT): INVITROGRO™ Spheroid Plating medium (BioIVT) 250 mL; add 1 mL Spheroid Media Supplement A (BioIVT) and 5.5 mL TORPEDO Antibiotic Mix (BioIVT). 4. INVITROGRO™ Spheroid Spin medium (BioIVT) 50 mL for recovery of cryoplateable spheroid qualified hepatocytes. 5. 3D hepatic spheroid maintenance medium: William’s Medium E 500 mL, and add 5 mL ITS+, 5 mL L-glutamine, 5 mL penicillin-streptomycin, and 5 μL dexamethasone (10 mM). 6. Percoll® (Sigma-Aldrich) 25 mL for post-thaw debris removal for cryopreserved non-parenchymal cells (NPCs).

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3.1 Recovery of Cryoplateable Spheroid Qualified Hepatocytes

1. Pre-warm INVITROGRO™ Spheroid Spin medium and INVITROGRO™ Spheroid Plating medium to 37 °C. 2. Transfer 25 mL of warm INVITROGRO Spheroid Spin medium to a sterile 50 mL conical tube. 3. Carefully remove the vial from the shipping container or liquid nitrogen storage. Immediately immerse the vial into a 37 °C water bath. This step can take 90–120 s. When the cells pull away from the vial wall, transfer the content of vial into the INVITROGRO Spheroid Spin medium in the 50 mL conical tube. The vial may be rinsed by transferring 1 mL of medium. 4. Resuspend the hepatocytes in the INVITROGRO™ Spheroid Spin medium by gently inverting the tube three times. 5. Spin the tube at 100 g for 10 min. 6. Aspirate supernatant and resuspend cells in 5 mL of INVITROGRO™ Spheroid Plating medium.

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3.2 Recovery and Post-thaw Debris Removal for Cryopreserved Nonparenchymal Cells (NPCs) (BioIVT)

1. In a clean BSC, the following was added to a sterile 50 mL conical tube: (a) 16.7 mL INVITROGRO™ CP medium (b) 3.3 mL isotonic Percoll (90% Percoll) 2. This makes a 15% Percoll solution in CP medium. 3. Allow 15% Percoll to equilibrate to room temperature in the BSC (should take around 30 min). 4. Allow INVITROGRO™ CP medium to equilibrate to 37 °C in a water bath. 5. Thaw NPC cell vial in a 37 °C water bath. 6. In the BSC, decant or pipette cells into the 15% Percoll solution. Rinse the empty cell vial two times to ensure maximal cell recovery. 7. Centrifuge cells at 500 × g for 10 min. 8. In the BSC, aspirate dead cells and debris from the top of the 15% Percoll; continue aspirating until only the cell pellet remains. 9. Resuspend cell pellet in 1 mL of species relevant CP. 10. Count cells and bring to the final cell concentration (based on application) in warm (37 °C) INVITROGRO™ CP medium.

3.3 3D Human Hepatic Spheroid Coculture Using Cryoplateable Spheroid Qualified Hepatocytes and Cryopreserved NPCs

1. After recovery, the cells were seeded at a ratio of 4:1 (HPHs/ NPCs) into ultralow attachment 96-well plates, and a total of 1875 viable cells (HPHs 1500; NPCs 375) were diluted in 100 μL INVITROGRO™ Spheroid Plating medium per well. 2. Spin plate at 250 g for 2 min. 3. Check the bottom of the plates for cell aggregation in each well. 4. Place the seeded plate in the incubator at 37 °C for 5 days (checking cell aggregation every other day). 5. On day 5 when the spheroids become sufficiently compact, carefully remove 50 μL of media from each well without disturbing the spheroid, and add 50 μL of 3D hepatic spheroid maintenance medium. 6. Change the medium every other day.

3.4 3D Human Hepatic Spheroid Coculture Using Freshly Isolated Hepatocytes and Cryopreserved NPCs

1. Spin the fresh hepatocyte transport tube at 100 g for 10 min. 2. Aspirate supernatant and resuspend hepatocytes in 40 mL of either INVITROGRO™ CP medium or complete William’s Medium E with 10% FBS. 3. Follow the steps in 3.3.

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Notes To analyze the integration of NPCs in the hepatic spheroid coculture system, mRNA and protein expression levels of Vimentin and CD68, the biomarker for hepatic stellate cells (HSCs) and Kupffer cells, respectively, were detected by RT-PCR and immunofluorescence staining with the following methods. 1. Total RNA isolated from around 40 spheroids using TRIzol reagent (ThermoFisher, Rockford, IL) was reverse transcribed to cDNA using a High Capacity cDNA Archive Kit (Applied Biosystems, Foster City, CA) following the manufacturer’s instructions. RT-PCR assay was performed to detect Vimentin and CD68 mRNA expression as shown in Fig. 1a on an ABI StepOnePlus Real-Time PCR System (Applied Biosystems) using SYBR Green PCR Mastermix (Qiagen). 2. Twenty spheroids were collected and fixed with 4% paraformaldehyde overnight at 4 °C, followed by cryoprotection in 30% sucrose overnight at 4 °C and embedded in Tissue-Tek OCT compound. Spheroid cryosections (eight microns) were stained for Vimentin and CD68 proteins (Fig. 1b) and viewed using confocal microscopy (Nikon) with a FITC and Texas Red filter at 10 × magnification.

B

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8 6 4 2 0 HPHs alone

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Fig. 1 Non-parenchymal HSCs and Kupffer cells could be successfully integrated into the hepatic spheroid coculture system. Expression of Vimentin (VIM) and CD68 mRNA and protein was measured using RT-PCR assay (a) and immunofluorescence staining (b) on day 17 of the spheroid culture. Results are expressed as mean ± SD (n = 3) (**p < 0.01)

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References 1. Lin RZ et al (2006) Dynamic analysis of hepatoma spheroid formation: roles of E-cadherin and beta1-integrin. Cell Tissue Res 324(3): 411–422 2. Bell CC et al (2016) Characterization of primary human hepatocyte spheroids as a model system for drug-induced liver injury, liver function and disease. Sci Rep 6:25187 3. Hurrell T et al (2020) Human liver spheroids as a model to study Aetiology and treatment of hepatic fibrosis. Cell 9(4):964 4. Breslin S, O’Driscoll L (2013) Threedimensional cell culture: the missing link in

drug discovery. Drug Discov Today 18(5–6): 240–249 5. Dame K, Ribeiro AJ (2021) Microengineered systems with iPSC-derived cardiac and hepatic cells to evaluate drug adverse effects. Exp Biol Med (Maywood) 246(3):317–331 6. Mittal R et al (2019) Organ-on-chip models: implications in drug discovery and clinical applications. J Cell Physiol 234(6):8352–8380 7. Liu K et al (2023) Dexamethasone mitigates remdesivir-induced liver toxicity in human primary hepatocytes and COVID-19 patients. Hepatol Commun 7(3):e0034

Chapter 9 In Vitro Porcine (Explant) Colon Culture Matheus de Oliveira Costa and Michael K. Dame Abstract Models have been extensively used to investigate disease pathogenesis. Animal models are costly and require extensive logistics for animal care, and samples are not always suitable for different analytical techniques or to answer the research question. In vitro cell culture models are generally focused on recreating a specific characteristic of an organ and are limited to a single cell population that does not display the characteristic tissue architecture of the source organ. In addition, such models do not account for the many interactions between pathogens and the diverse cell subsets that are normally present in a given organ. Conclusions based on conventional 2D cell culture methods are limited, requiring extrapolation from a reductionist model to understand in vivo events. In vitro organ culture (IVOC) offers a way to overcome some of these limitations. Explants conserve important in vivo characteristics, such as different cell types and complex tissue architecture. This in vitro (ex vivo) organ culture protocol of the swine large intestine aims at maintaining viable colonic mucosa for up to 5 days. The protocol described herein applies a combination of methods used for immortalized cell culture and stem cell stimulation to support the physiological cellular flow inherent of the intestinal mucosa. Required equipment includes a hyperoxic chamber and culture at the air-liquid interface. Key words Organ culture, Ex vivo, IVOC, Swine, Colon, Explant, In vitro, Colon model

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Introduction The intestinal mucosa is one of the main interaction interfaces between the host and the exterior world, including microorganisms and other potentially harmful agents. It is a complex assemblage of specialized tissues that interact in numerous ways. A sophisticated model that recapitulates the network of processes that develop during healthy and diseased states is essential to further develop the subjects of veterinary microbiology, physiology, internal medicine, and others. Access to the intestinal mucosa for in vivo sampling, however, is anatomically, technically, and ethically challenging. In addition, animal models of the swine gastrointestinal tract are expensive to maintain, require intensive logistics, and are often financially inaccessible. Thus, laboratory models of the

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intestinal mucosa have been developed in recent years [1– 3]. Two-dimensional cell culture models, particularly for immortalized cell lines, are common resources because they are easy to manipulate, and they efficiently maintain cell viability. However, this simplistic method has inherent limitations. The technique is generally focused on recreating a specific characteristic of the intestinal mucosa but harbors only one or a limited number of intestinal cell subsets, and does not account for the many interactions between the different tissue layers that form the intestines. In contrast, three-dimensional gut organoids include different cell types of the colonic epithelium, but do not include complex niche components, both cellular and structural, which regulate stem cell renewal and drive differentiation. In addition, the supporting layers of the muscularis mucosa and submucosa are absent. Due to their three-dimensional nature, organoids require that inoculation/ exposure to agents be performed on the basolateral side or after previous disruption of the organoid epithelial lining [4, 5]. In vitro (or ex vivo) organ culture (IVOC) offers the convenience of cell culture models, such that they are easily sampled and manipulated, while maintaining the complex architecture of the in vivo mammalian organ [6–8]. IVOC is suitable for studies involving infectious agents, pharmacological treatments, and nutritional and toxicological evaluations. The objective of this protocol is to consistently maintain, ex vivo, the porcine colonic mucosal/submucosal architecture and cellularity observed in vivo for a prolonged period (5 days). It relies on the combination of air-liquid interface culture and supplementation with growth factors that support Lgr5+ stem cell renewal [9]. As an alternative to the high oxygen environment used, a microfluidic device that constantly supplies explant with gassed culture media has been developed [10]. Tissue samples are collected immediately after euthanasia from healthy animals. The tissue is cleaned to remove intestinal contents, and the mucosa is manually separated from the serosa to improve its in vitro viability. Finally, explants are cultured on air-liquid interface using a specific combination of gas and liquid media. The protocol is divided into Subheading 3.1 media preparation and Subheading 3.2 organ processing and culturing.

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Materials

2.1 Tissue Handling and Dissection

1. 100 mL beaker (3×). 2. Toothless forceps (1×). 3. Toothed forceps (2×). 4. Mayo scissors (1×).

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5. Scalpels (blade no. 20, 2×). 6. Sterile large Petri dish, 60 × 15 mm. 7. Styrofoam box (approximately 30 × 30 cm, including lid with a shallow depression). 8. Ice supply. 9. Cell culture incubator (37 °C). 2.2 Culture Apparatus

1. Modular Incubator Chamber (MIC-101, Billups-Rothenberg, Inc.). 2. 99% O2, 1% CO2 gas mix (10 L). 3. Sterile six-well plate. 4. 100 μM cell strainer.

2.3 Organ Transport Media (500 mL)

1. Hank’s Balanced Salt Solution (HBSS, Gibco, Canada). 2. 100 mL sterile glass bottle. 3. 10 mL sterile syringe. 4. 0.22 μM syringe filter. 5. Calcium chloride (CaCl2), final concentration 1.5 mM.

2.4 Organ Culture Media (500 mL)

1. Keratinocyte culture media (KBM Bullet Kit, Lonza, Walkersville, MD, USA), 1 bottle (500 mL). 2. Calcium chloride (CaCl2); final concentration 1.5 mM. 3. Fungizone (InvivoGen InvivoGen, Burlington, ON, Canada), 10 μg/mL.

2.5 Stem Cell Conditioned Media

1. Advanced DMEM/F12 (500 mL). 2. Fetal bovine serum (127 mL). 3. Penicillin, 10,000 units/mL. 4. Streptomycin, 10 mg/mL. 5. G418 (Gibco, Burlington, ON, Canada), 0.5 mg/mL. 6. Hygromycin, 0.5 mg/mL. 7. Trypsin-EDTA (0.25% trypsin, 1 mM EDTA, Invitrogen, Burlington, ON, Canada). 8. L-WRN cells (ATCC CRL3276). 9. 150 cm2 sterile culture flask.

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Methods

3.1 Media Preparation

All the procedures below should be performed inside a biosafety cabinet, and although samples might not be sterile at all times, sterile technique should be performed at all times to minimize contamination.

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1. Conditioned media preparation. (a) Prepare the L-cell medium as in Miyoshi et al. [11]. (b) Add penicillin (100 units/mL), streptomycin (0.1 mg/ mL), FBS (20%, vol/vol), and L-glutamine (2 mM) to DMEM F12 (500 mL, L-cell medium). (c) Aliquot 25 mL of L-cell medium and prewarm it at 37 °C using a 50 mL centrifuge tube. (d) Warm a tube of L-WRN cells at room temperature (or at 37 °C). (d.1) Immediately transfer the cells to the prewarmed L-cell medium once thawed. (e) Transfer the seeded L-cell medium to at 150 cm2 culture flask. (e.1) Incubate the flask at 37 °C for 24 h. (f) Add G418 (500 μg/mL) and hygromycin (500 μg/mL) to a prewarmed 25 mL aliquot of L-cell medium. (f.1) Change the medium in the flask using this selective L-cell medium. (g) Grow cells until confluence (100%, 48 or 72 h). (h) Wash the cells twice with 10 mL of PBS (phosphatebuffered saline) and discard the wash. (i) Add 1 mL of trypsin-EDTA and incubate at 37 °C until cells are resuspended (5 min; tapping may be necessary to release the cells). (j) Recover the cells in 12 mL of L-cell medium. (k) Split it 1:10 into 150 cm2 flasks (adjustable as per research design). (l) Add 25 mL of L-cell medium without antibiotics (critical step) into one of the new 150 cm2 flasks seeded with the selected L-WRN cells. (m) Incubate flask for 48 h or until cells are over-confluent. (m.1) Recover the conditioned medium into a 50 mL centrifuge tube. (m.2) Add fresh medium to the cell culture flask and return it to the incubator. (n) Centrifuge the conditioned medium at 1000 × g for 10 min. (n.1) Use a syringe filter to sterilize the media. (n.2) Aliquot in 10 mL tubes and store at -80 °C. (o) Repeat Steps 1.12 and 1.13 daily for up to 12 days if desired.

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2. Transport media preparation (500 mL). (a) A CaCl2 stock solution (100×) can be prepared by mixing 1.665 g of CaCl2 into 100 mL of sterile molecular grade water. (b) Attach the 10 mL syringe to the 0.22 μM syringe filter. (b.1) Sterilize the content by passing the solution through the filter. (c) Using a sterile pipette, remove 5 mL from the HBSS bottle (500 mL), and add the same amount of CaCl2 solution to the bottle (final concentration of 1.5 mM Ca). (d) Keep at 4 °C until use. 3. Culture media preparation. (a) Thaw the KBM Gold Bullet kit components and one vial (1.5 mL) of Fungizone to room temperature. (b) Using a sterile pipette, remove and discard 17.5 mL of KBM media from the bottle (500 mL). (c) Using 1 mL sterile micropipettes, add the Gold Bullet kit components and 500 μL of Fungizone to the KBM bottle. (d) Add 12.5 mL of previously prepared L-WRN conditioned media to the bottle. (e) Add 5 mL of previously prepared (see Subheading 2) CaCl2 sterile stock solution (final concentration of 1.5 mM of Ca2+). (f) Keep at 4 °C until use. 3.2 Organ Processing and Culturing

1. Pre-euthanasia. 2. A Styrofoam box lid (or any other thermal insulation material) can be used to create a refrigerated surface for tissue manipulation. (a) Use crushed ice cubes to create a layer of approximately 5 cm of thickness on top the insulated surface (shown in Fig. 1). (b) Place a large Petri dish on the ice. (c) Add 5 mL of chilled transport media to the Petri dish lid, not to cover the entire surface but to provide moisture. 3. Place the transport media on ice, in the open Styrofoam box. 4. Transfer 3 mL of culture media into each well of a six-well plate. (a) Place the six-well plate in the incubator at 37 °C. 5. See Note 1.

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Fig. 1 Luminal exposure by opening the colon segment along the mesenteric border

6. Use the mayo scissors to collect a 10 cm segment of colon immediately after euthanasia. 7. Place the colon segment in the chilled transport media, in a Styrofoam box. 8. See Note 2. 9. Place the colon segment on the previously prepared large Petri dish. (a) Using toothed forceps, hold the segment on one of the edges, trying to minimize damage while grasping the organ. (b) Using mayo scissors, carefully open the colon segment along the mesenteric border. 10. Add ~10 mL of transport media to the three clean beakers, and without releasing the forceps from its previous position (Subheading 3.1), place the colon into the beaker, and shake it vigorously to free the mucosa from any fecal matter and mucus (Fig. 1). (a) Move on to the next beaker once the media becomes saturated. (a.1) If necessary, discard the used media and replace it with a clean aliquot. (a.2) Repeat Step 4.1 until the transport media do not become cloudy after submersion of the segment.

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Fig. 2 Separation of the mucosa and serosa using the two-forceps technique. The serosa-mucosa interface becomes more apparent as it is separated (A)

(a.3) Discard the large Petri dish lid and place the dish on the Styrofoam box lid ice. (a.4) Add 5 mL of chilled transport media to the Petri dish. (a.5) Place the opened colon segment on the Petri dish, mucosa side down (serosa side up). (a.6) Using two toothed forceps, carefully separate the mucosa from the serosa. (a.7) Starting a corner, make sure you can clearly see the serosa/mucosal delineation (Fig. 2a). (a.8) See Note 3. (b) Discard the serosa appropriately and turn the segment mucosa side up. (b.1) Using two scalpels moving simultaneously on opposite directions, remove the edges of the mucosa where it was previously held by the forceps (more likely to have been damaged). (c) Using the same technique, further divide the mucosa in as many explants as required. (d) Explants should be no bigger than 2 × 2 cm.

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11. Retrieve the prewarmed six-well plates and place one cell strainer into each well. (a) See Note 4. 12. Using a toothless forceps, place explants into a six-well plate, one explant per well (Fig. 4). (a) If necessary, adjust the liquid media volume so that it touches the bottom aspect of the cell strainer, but does not invade its inner aspect. 13. Close plates and move them to the modular incubator. 14. Tightly seal the modular incubator and plug in the hyperoxic hose while leaving the output plug free. (a) Flush the incubator continuously for 3 min at 10 liters/ min. (b) Seal the output plug tightly and monitor for inflation of the incubator (3–5 s). (c) Seal the input plug once it inflates. 15. Move the modular incubator, now properly gassed, to the incubator. 16. Culture explants at 37 °C (Fig. 3). 3.3 Daily Culture and Gas Media Change

1. Using the water bath, warm the culture media to 37 °C for 15 min. 2. Retrieve new six-well plates and place them inside the incubator used for organ culture for 15 min to prewarm. 3. After this period, bring both culture media and six-well plates to the biosafety cabinet, and transfer 3 mL of prewarmed culture media to each well.

Fig. 3 Air-liquid interface explant setup. The organ is gently deposited on top of the cell restrainer. The liquid media should never flood the inner chamber of the cell strainer where the tissue is deposited (A)

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Fig. 4 Final organ culture setup on a six-well plate

(a) Cover the plates and keep them in the biosafety cabinet. 4. Retrieve the six-well culture plates that require media replacement. (a) Open the modular incubator carefully to prevent any splashing. 5. In the biosafety cabinet, transfer the cell strainers from the previously used six-well plates to a freshly prepared six-well plate (Fig. 4). 6. Repeat Steps 12 to 15 from Subheading 3.2 to reestablish the hyperoxic atmosphere. 7. During the culture period (this protocol has been tested up to 7 days; Fig. 5), monitor explants for the following signs of contamination: 8. Bacterial colony-like growth on the tissue. 9. Culture medium turns cloudy. 10. Mold-like growth on the cell strainer. (a) If any of the signs above are observed, we suggest that individual explants should be removed and the remaining healthy explants be placed on a new, sterile six-well plate. 11. Explant sampling should be performed according to the researcher’s choice of analytical technique (e.g., samples for histology should be immersed in 10% buffered formalin).

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Fig. 5 H&E stained sections of a colon explant from a 5-week-old pig cultured for 7 days at higher magnification/cross section (a) and lower magnification/longitudinal section (b). Necrotic and sloughed off epithelial cells are seen accumulated in the crypts and on the luminal aspect of the explants

4 Notes 1. At any time while handling the colon segment, use the forceps gently and try to always target the same location for holding it. 2. It is critical that the transportation time between the euthanasia site and the processing laboratory is kept as short as possible (5–15 min). Viability of explants is greatly diminished if the total processing time (euthanasia to incubator) is longer than 2 h. 3. Do not attempt to remove the segment’s entire serosa in one movement. It is safer, and less damaging to the tissue, to slowly remove by holding the serosa on its closest attachment point to the mucosa, and then pull. 4. Depending on the manufacturer, you may have to cut part of the plastic strainer border to fit the culture well properly. This should be done observing proper sterile technique. References 1. McOrist S, Jasni S, Mackie RA, Berschneider HM, Rowland AC, Lawson GH (1995) Entry of the bacterium ileal symbiont intracellularis into cultured enterocytes and its subsequent release. Res Vet Sci 59(3):255–260 2. Naresh R, Song Y, Hampson DJ (2009) The intestinal spirochete Brachyspira pilosicoli attaches to cultured Caco-2 cells and induces pathological changes. PLoS One 4(12):e8352.

https://doi.org/10.1371/journal.pone. 0008352 3. Skjolaas KA, Burkey TE, Dritz SS, Minton JE (2007) Effects of salmonella enterica serovar typhimurium, or serovar choleraesuis, lactobacillus reuteri and bacillus licheniformis on chemokine and cytokine expression in the swine jejunal epithelial cell line, IPEC-J2. Vet Immunol Immunopathol 115(3–4):299–308.

Porcine Colon Culture https://doi.org/10.1016/j.vetimm.2006. 10.012 4. Zhang YG, Wu S, Xia Y, Sun J (2014) Salmonella-infected crypt-derived intestinal organoid culture system for host-bacterial interactions. Physiol Rep 2(9). https://doi. org/10.14814/phy2.12147 5. Wilson SS, Tocchi A, Holly MK, Parks WC, Smith JG (2015) A small intestinal organoid model of non-invasive enteric pathogenepithelial cell interactions. Mucosal Immunol 8(2):352–361. https://doi.org/10.1038/mi. 2014.72 6. Autrup H, Stoner GD, Jackson F, Harris CC, Shamsuddin AK, Barrett LA et al (1978) Explant culture of rat colon: a model system for studying metabolism of chemical carcinogens. In Vitro 14(10):868–877 7. Browning TH, Trier JS (1969) Organ culture of mucosal biopsies of human small intestine. J Clin Invest 48(8):1423–1432. https://doi. org/10.1172/JCI106108 8. Dame MK, Veerapaneni I, Bhagavathula N, Naik M, Varani J (2011) Human colon tissue

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in organ culture: calcium and multi-mineralinduced mucosal differentiation. In Vitro Cell Dev Biol Anim 47(1):32–38. https://doi.org/ 10.1007/s11626-010-9358-3 9. Sato T, Vries RG, Snippert HJ, van de Wetering M, Barker N, Stange DE et al (2009) Single Lgr5 stem cells build crypt-villus structures in vitro without a mesenchymal niche. Nature 459(7244):262–265. https:// doi.org/10.1038/nature07935 JM, 10. Eslami Amirabadi H, Donkers Wierenga E, Ingenhut B, Pieters L, Stevens L et al (2022) Intestinal explant barrier chip: long-term intestinal absorption screening in a novel microphysiological system using tissue explants. Lab Chip 22(2):326–342. https:// doi.org/10.1039/d1lc00669j 11. Miyoshi H, Stappenbeck TS (2013) In vitro expansion and genetic modification of gastrointestinal stem cells in spheroid culture. Nat Protoc 8(12):2471–2482. https://doi.org/ 10.1038/nprot.2013.153

Chapter 10 Bioencapsulation of Oocytes and Granulosa Cells Massimo Faustini, Stella Agradi, Daniele Vigo, Maria L. Torre, and Giulio Curone Abstract A protocol for the encapsulation in sodium alginate of granulosa cells in primary culture and coculture of oocyte-cumulus complexes is reported. Sodium alginate forms strong gels when jellified with barium ions, allowing the self-organization of cells into a 3D structure. This method of encapsulation is simple and cheap, allowing the culture of cells in a three-dimensional fashion. Key words Encapsulation, Granulosa cells, Alginate, 3D culture

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Introduction During recent decades, several attempts have been made to allow oocyte in vitro maturation (IVM) and/or fertilization (IVF) culture systems with the reaching of successful yield. Traditional culture methods are based on protein and growth factor-enriched media with the purpose of replacing the biological environment for the isolated oocytes, but more recently, alternative oocyte maturation systems have been designed, based on two functional keypoints 1. Oocytes are surrounded by a cellular environment (e.g., cell cumulus), with a dedicated extracellular matrix. 2. In vivo cells present a three-dimensional (3D) organization. A culture cell technique that considers these principles as its keystone more closely approaches the biological maturation system and forms one of the most recent advances in tissue engineering. These features have been well reviewed by Gilchrist et al. [1] and more recently demonstrated by Hussein et al. [2]; oocyte-cumulus cellular contacts and paracrine factors are the basis for optimal oocyte development and competence. Following these principles, a number of

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three-dimensional granulosa/oocyte coculture systems were tested and, in some cases, patented; in this field, encapsulation seems to be a valid technology for obtaining a 3D environment suitable for coculture. Pangas et al. [3] developed an alginate bead 3D culture system designed for cumulus-oocyte complexes (COCs); such technology yielded good results in terms of structural development and meiosis resumption for murine ova. Further successes were achieved in rat oocytes after modifying the cell adhesion properties of the alginate with the use of the tripeptide RGD (Arg-Gly-Asp), an ubiquitary adhesion sequence for several extracellular matrix proteins [4]. This latter system aims to mimic the extracellular matrix functions more closely. In the last years, several advanced culture methods have been designed: In the recent times, the microfluidic systems gained a great attention in 3D cell culture. Starting from polydimethylsiloxane-based systems (PDMS), evolving to thermoplastics or glass-based devices, in order to reduce the times of preparation, and increasing the lab-to-lab reproducibility, poor features when PDMS is adopted [5–7]. Moreover, microfluidic techniques can be coupled with 3D printing and 3D extrusion in order to tailor the process of culture architecturing [8, 9]. Nevertheless, microfluidic systems can show some aspects in order to adapt the macrosystem cell culture to microfluidics; this transition phase is not immediate, since a number of cell culture protocols have been optimized on macroscopic cell culture. In this chapter, a simple, practical methodology for granulosa cell and cumulus-oocyte complex (COC) encapsulation and culture is illustrated (Figs. 1 and 2), adopting the technique employed for boar semen encapsulation [10]. The method implies a macroscopic cell culture with a limited use of materials and disposables, allowing the 3D self-organization of cells, in particular COCs and granulosa complexes.

2

Materials NaCl 0.9% w/v. Polypropylene 5 mL disposable syringe. Capillary tube. Falcon 50 mL tubes. Tissue culture medium 199 (TCM199). 10% fetal calf serum (FCS). Penicillin-streptomycin.

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Fig. 1 Schematic figure of the process for the production of encapsulated granulosa cells-oocyte: the granulosa/barium chloride suspension is dropped into a continuously stirred sodium alginate solution. The resulting suspension is set aside in continuous stirring for 1 h, and then injected with the oocyte. The capsule is then immersed into the well to culture

TCM199, containing Earle salts, l-glutamine, and NaHCO3. Barium chloride (anhydrous) in saturated aqueous solution. Medium viscosity (3500 cP) sodium alginate. Protamine sulfate. 3,17-Androstenedione. All solutions must be freshly prepared. Store culture media at 4 °C; otherwise, respect the label indications.

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Methods All procedures are carried out at room temperature if not otherwise stated. The following protocol has been adopted for the maintaining of granulosa cells collected from sows but can be extended to other species; the results are encouraging in order to design new simple three-dimensional culture systems for granulosa cells, providing a suitable milieu for oocyte maturation with a minimal use of hormones.

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Fig. 2 Photographs of three phases of encapsulated granulosa cells-oocyte co-culture 3.1 Granulosa Encapsulation

Ovaries at different stages of development are collected from sows aged 6–11 months in a slaughterhouse; ovaries are then washed at 30 °C with NaCl 0.9% w/v, and follicles with a diameter of 2–6 mm are identified on the surface of the ovaries; follicular fluids containing GC are aspirated using a polypropylene disposable syringe and mixed in a Falcon tube in order to obtain a pool. The cellular suspension is then centrifuged (600 × g, 10 min) and washed twice with 10 mL of tissue culture medium 199 (TCM199) + 10% fetal calf serum (FCS) + 1% penicillinstreptomycin. The cell concentration in the resulting granulosa cell pellet is generally determined using a Makler counter. Cell count in the pellets is generally in the range between 1 × 105 and 5 × 105 cells/mL. The centrifuged cell suspension is dispersed in a xanthan gum solution (Satiaxane®, SKW Biosystems, France) 0.5% in TCM199, containing Earle salts, l-glutamine, and sodium bicarbonate (Sigma-Aldrich, Milan, Italy), to provide a suspension xanthan gum ratio of 1:3. A saturated solution of BaCl2 is then added to the suspension to reach a concentration of 20 mM (see Note 1). The suspension is extruded at 25 °C through a 25-gauge needle by a peristaltic pump

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or by syringe and dropped into a medium viscosity (3500 cP) sodium alginate solution (Sigma-Aldrich, Milan, Italy) 0.5% in culture medium stirred at 30 rpm for 60 min (see Note 2). The resultant capsules are collected; washed at least twice with TCMI99 containing Earle salts, l-glutamine, and sodium bicarbonate; and suspended in the same medium. Aiming to increase the capsule mechanical resistance, capsules are treated with a solution of protamine sulfate 1% in TCM199 (containing Earle salts, l-glutamine, and sodium bicarbonate) for 30 min at 25 °C. All employed solutions must be sterilized by filtration. Each single capsule is put into a well for cell culture and suspended in 600 μL of culture medium (TCMI99 + 10% FCS + 1% penicillinstreptomycin +100 mg 1 3,17-androstenedione). All culture wells are maintained in an incubator at 38.5 °C, 5% CO2, and 90% humidity. Oocytes can be easily recovered from ovaries collected at the abattoir, and their maturation process can be carried out through the coculture with granulosa cells. Thus, the obtained cumulus-oocyte complexes can be directly injected into the 3D granulosa capsules via a capillary tube connected to a syringe. 3.2 Oocyte Encapsulation

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Oocytes can be easily recovered from ovaries collected at the abattoir, and their maturation process can be carried out through the coculture with granulosa cells. Thus, the obtained cumulus-oocyte complexes can be directly injected into the 3D granulosa capsules via a capillary tube connected to a syringe.

Notes 1. In a sealable flask, add anhydrous (or dihydrate, BaCl2·2H2O) to distilled water until a sediment is formed. Set aside for 1 day at room temperature. The precipitate should be maintained during all the use of the solution. 2. Dissolution time of alginate can vary: Alginate should be dissolved into water a few hours before using it, continuously stirring and avoiding the warming of the forming solution.

References 1. Gilchrist RB, Ritter LJ, Armstrong DT (2004) Oocyte-somatic cell interactions during follicle development in mammals. Anim Reprod Sci 82–83:443–446 2. Hussein TS, Thompson JG, Gilchrist RB (2006) Oocyte-secreted factors enhance oocyte developmental competence. Dev Biol 296:514–521

3. Pangas SA, Saudye H, Shea LD et al (2003) Novel approach for the three-dimensional culture of granulosa cell-oocyte complexes. Tissue Eng 9(5):1013–1021 4. Kreeger PK, Woodruff TK, Shea LD (2003) Murine granulosa cell morphology and function are regulated by a synthetic Arg-GlyAsp matrix. Mol Cell Endocrinol 205:1–10

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5. Torino S, Corrado Iodice M, Coppola G (2018) PDMS-based microfluidic devices for cell culture. Inventions 3:1–14 6. Heo YS, Cabrera LM, Song JW, Futai N, Tung YC, Smith GD, Takayama S (2007) Characterization and resolution of evaporation-mediated osmolality shifts that constrain microfluidic cell culture in poly(dimethylsiloxane) devices. Anal Chem 79:1126–1134 7. Regehr KJ, Domenech M, Koepsel KT, Carver KC, Ellison-Zelski SJ, Murphy WL, Schuler LA, Alarid ET, Beebe DJ (2009) Biological implications of polydimethylsiloxane-based microfluidic cell culture. Lab Chip 9:2132– 2139

8. Waheed S, Cabot JM, Macdonald NP, Lewis T, Guijt RM, Paull B, Breadmore MC (2016) 3D printed microfluidic devices: enablers and barriers. Lab Chip 16:1993–2013 9. Gross BC, Erkal JL, Lockwood SY, Chen C, Spence DM (2014) Evaluation of 3D printing and its potential impact on biotechnology and the chemical sciences. Anal Chem 86:3240– 3253 10. Vigo D, Villani S, Faustini M et al (2005) A follicle-like model by granulosa cell encapsulation in a barium alginate/protamine membrane. Tissue Eng 11:709–714

Chapter 11 Culturing and Differentiation of Patient-Derived Ectocervical Epithelial Stem Cells Using Air-Liquid Interphase and Matrigel Scaffold Rajendra Kumar Gurumurthy, Naveen Kumar, and Cindrilla Chumduri Abstract The ectocervix acts as a multilayered defense barrier, protecting the female reproductive system from external pathogens and supporting fertility and pregnancy. To understand the complex cellular and molecular mechanisms of cervical biology and disease, reliable in vitro models are vital. We present an efficient method to isolate and cultivate epithelial stem cells from ectocervical tissue biopsies. This method combines enzymatic digestion, mechanical dissociation, and selective culturing to obtain pure ectocervical epithelial cells for further investigation. The protocol accommodates both 2D stem cell monolayer and advanced 3D culture systems, such as air-liquid interface and Matrigel scaffolds, using a defined media cocktail, making it highly versatile. The primary ectocervical epithelial cells retain their native characteristics, enabling the exploration of ectocervical epithelial tissue behavior and pathology. This chapter provides step-by-step guidelines for setting up 2D and 3D cultures, facilitating adoption across different laboratories, and advancing cervical biology and disease research. Key words Cervical tissue biopsies, Epithelial cells, Ectocervix, Cell isolation, Primary cell culture, 2D culture, 3D culture, Air-liquid interface (ALI), Organoids

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Introduction The emergence of 3D epithelial organoids derived from adult stem cells has revolutionized regenerative medicine, offering remarkable tools to study infections, disease progression, and drug efficacy [1– 5]. In particular, these organoids have proven highly promising for investigating various aspects of the female reproductive tract, with the ectocervix playing a crucial role as both a barrier and facilitator of fertilization [2]. Recent groundbreaking developments have led to the creation of long-term, expandable organoid models explicitly tailored for the ectocervix, providing invaluable insights into cervical homeostasis and metaplasia dynamics [6].

Mario Baratta (ed.), Epithelial Cell Culture: Methods and Protocols, Methods in Molecular Biology, vol. 2749, https://doi.org/10.1007/978-1-0716-3609-1_11, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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Different techniques, such as Matrigel [6] and Transwells [7], have been employed to establish these ectocervical organoids, effectively mimicking the intricate architecture of the ectocervical tissue found in living organisms. Both mouse- and human-derived ectocervical organoids closely resemble the in vivo tissue structure, exhibiting multilayered KRT5+ epithelia containing basal proliferative TP63+ cells and more differentiated cells toward the inner lumen [8]. The differentiation process in these organoids is primarily regulated by Notch signaling, and inhibiting this pathway results in a failure of stratification, leading to a TP63+ monolayer. Matrigel-based organoids position the stem cell compartment on the outer surface with the lumen toward the center, while Transwell-based air-liquid interface (ALI) cultures attach the stem cells to the Transwell membrane, placing the lumen on the outer surface. These methods also enable the development of complex cocultures involving fibroblasts and immune cells, enhancing the versatility of organoid models. Success in cultivating these organoids relies on precise customization of nutrients and growth factors in the culture medium to accurately mimic the stem cell microenvironment. Significantly, these organoids have been successfully utilized to model infections caused by human papillomavirus (HPV), herpes simplex virus (HSV), and Chlamydia trachomatis, leading to groundbreaking insights into molecular mechanisms and potential therapeutic targets [3, 9]. Coinfections, such as chlamydia and HPV, can impact the host epithelium and immune response, potentially influencing disease progression and treatment outcomes. The findings discussed here underscore the tremendous potential of organoids as powerful tools for investigating cervical cancer, infections, and personalized medicine approaches tailored to individual genetic profiles. The use of 3D epithelial organoids in research and medical applications promises to advance our understanding of these conditions and provide tailored treatments for patients based on their unique genetic makeup.

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Materials Maintain a sterile environment when preparing all solutions and reagents, ensuring they are of cell culture grade and stored as per the manufacturer’s instructions. Dispose of waste materials following appropriate regulatory guidelines.

2.1 Tissue Samples from the Human Ectocervix

Human biopsies for scientific experiments require institutional human ethics permission and informed patient consent. For the primary epithelial cell cultures, it is essential to obtain human tissue samples from the ectocervix regions within 2–3 h of surgical removal. It is of utmost importance to perform all procedures for

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primary epithelial cell isolation and culture in containment level 2 facilities to mitigate any risk of infection from viruses or other pathogens originating from human tissue samples and body fluids. Following the isolation of primary epithelial cells, tissue residues and cell suspension should be decontaminated with Korsolex prior to autoclaving. Any disposable materials used during tissue preparation for epithelial cell culture must be sterilized by autoclaving. 2.2 2.2.1

Cell Lines Culture Reagents

3 T3-J2 fibroblast cell line is essential for successful coculture of ectocervical epithelial cells in a 2D environment. 1. Use Dulbecco’s Phosphate Buffered Saline-1X (DPBS-1X) for washing tissue samples. 2. For culturing the 3 T3-J2 cell line, prepare complete Dulbecco’s Modified Eagle Medium (DMEM) by adding a final concentration of 10% (vol/vol) heat-inactivated FCS, 2 mM Lglutamine, 1 mM sodium pyruvate, and 1% (vol/vol) penicillin-streptomycin solution. Store at 4 °C for up to 2 weeks (see Note 1). 3. Mitomycin C solution (0.4 mg/mL): Dissolve 2.0 mg of mitomycin C in 5 mL of sterile water, and then filter-sterilize. Store 1 mL aliquots at 4 °C in the dark for up to 3 weeks (see Note 2). 4. For human cervical epithelial isolation, prepare a 0.5 mg/mL collagenase type 2 solution by dissolving 100 mg of collagenase type 2 in 200 mL of Hank’s balanced salt solution. Filtersterilize, make 5 mL aliquots in 15 mL tubes, and store at 20 °C for up to 2 months. 5. To prepare human ectocervical organoid media and steps involved in the harvesting and passaging of organoids, use Advanced DMEM/F-12 with HEPES and Glutamax (ADF+ +). Add a final concentration of 10 mM HEPES and 1% (vol/vol) Glutamax to Advanced DMEM/F-12 media (Gibco™), and store at 4 °C for up to 1 month. 6. For human ectocervical epithelial 2D culture, prepare a fresh working solution of collagen type 1 (4 mg/mL) in warm DPBS by diluting it at a 1:100 ratio. 7. Prepare a collagen type 1 coated plate or flask, prewarm the collagen type 1 working solution in a 37 °C water bath, and add 1 mL per well of a 6-well plate, 2 mL per T25 flask or 4 mL per T75 flask. Incubate the culture dish or flask at 37 °C for 1–3 h or overnight at 4 °C. Remove the collagen solution completely using a 1 mL pipette, air-dry the coated surface for 10 min, and use for cell culture (see Note 3). 8. Prepare WNT-3a or Rspondin-1 conditioned media: Prepare the Wnt-3a and Rspondin-1 conditioned media as previously described [8], or procure recombinant proteins from a certified supplier (see Note 4).

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9. To prepare the human ectocervical 3D medium for air-liquid interface (ALI) and organoid culturing, add a final concentration of 12 mM HEPES, 1% (vol/vol) GlutaMax, 1% (vol/vol) B27, 1% (vol/vol) N2, 0.5 μg/mL hydrocortisone, 10 ng/mL human EGF, 100 ng/mL human noggin, 100 ng/mL human FGF-10, 1.25 mM N-acetyl-L-cysteine, 2 μM TGF-β R kinase inhibitor IV, 10 μM ROCK inhibitor (Y-27632), 10 mM nicotinamide, and 10 μM FSK to ADF, and then filter sterilize. Store the media at 4 °C for up to 2 weeks (see Note 5). 10. For ectocervical organoid cultivation, use Matrigel. Thaw the required volume on ice for 2 hours and always keep it on ice (see Note 6). 11. Use the collagen solution from bovine skin, Sigma Aldrich, cat. No. C4243-20 mL. 12. For the 10× reconstitution buffer, initially prepare 0.062 N NaOH by adding 1.86 mL of 10 N NaOH to 300 mL of sterile distilled water. Mix 2.2 g of sodium bicarbonate and 4.77 g of HEPES with 75 mL of 0.062 N NaOH, bring the final volume to 100 mL with 0.062 N NaOH, filter-sterilize, and store at -20 °C. 13. For 10× DMEM medium, dissolve 6.69 g of DMEM powder (Gibco™) in 50 mL of sterile distilled water and store at -20 °C.

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Methods

3.1 Isolation of Human Ectocervical Epithelial Cells from Tissue Biopsies

1. Follow the protocol steps as illustrated in Fig. 1. Transfer the tissue sample into a sterile 10 cm Petri dish containing sterile DPBS. To clean the tissue, gently agitate it with sterile forceps within the DPBS solution to remove mucus or blood. Subsequently, discard the used DPBS and replace it with fresh DPBS. Repeat this step until the tissue is thoroughly free from mucus and blood. 2. Mince the tissue into smaller pieces with thicknesses less than 0.5 mm3 using sterile scissors and forceps without DPBS (see Note 7). 3. Place the minced tissue into a 15 mL Falcon tube containing 5 mL of prewarmed collagenase II solution (refer to Subheading 2.2.1). Then, incubate it horizontally within the orbital shaker for 2.5 h at 37 °C and 180 rpm (see Note 8). 4. Centrifuge the contents at 1000 g for 5 min at 4 °C to pellet down cells. Discard the supernatant and resuspend the resultant cell pellet in 5 mL of warm TrypLE solution using a 5 mL pipette. Incubate it horizontally within the orbital shaker for 15 min at 37 °C and 180 rpm (see Note 8).

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Fig. 1 Schematic representation of isolation of ectocervical cells, cultivation of 2D stem cell, and 3D Transwell- and Matrigel-based ectocervical epithelial tissue. (a). Schematic representation of the human female reproductive tract. (b–g). Single cells isolated from ectocervical tissue (b–c) cultured in 2D to enrich stem cells (d) and long-term propagate by coculturing on fibroblast (e). (e–g). Phase-contrast images of pure human ectocervical epithelial stem cells (e) and propagated on 3 T3-J2 fibroblast cells (g). (h–i). Establishment of Transwell-based air-liquid interphase cultures from 2D stem cells (h) and organoid cultures by embedding into Matrigel (i). (j–l). Representative confocal images of human ectocervical epithelial tissue biopsy (j) air-liquid interphase cultures (k) and organoids (l) stained for epithelial marker E-cadherin and nuclei (Hoechst)

5. Following incubation, dissociate the cell aggregates by pipetting up and down five times with a 5 mL pipette. Next, add 5 mL of ADF++ (refer to Subheading 2.2.1), and continue to pipette up and down five times using a 5 mL pipette.

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6. Let the tube sit for 2 min to allow tissue debris to settle by gravity. Transfer the upper supernatant containing the dissociated cells through a 40 μm cell strainer into a new 50 mL Falcon tube. 7. Resuspend the settled tissue debris in 5 mL of ADF++, and then repeat the previous step twice. 8. Use a Neubauer chamber to count the cell numbers in the 50 mL Falcon tube (see Note 9). 9. Pellet down the desired number of cells for 2D (up to 0.1 million cells/T25 flask) or 3D (10,000 cells/50 μL Matrigel/ well of 24-well plate) culturing via centrifugation at 1000 g for 5 min at 4 °C. Discard the supernatant and proceed to next steps. 3.2 Culturing and Long-Term Expansion of Human Ectocervical Stem Cell in 2D 3.2.1 2D Culturing of Ectocervical Stem Cells

1. Begin by isolating the ectocervical epithelial cells. Follow the protocol steps outlined in Subheading 3.1, steps 1 through 9. 2. Resuspend the resultant cell pellet in 4–5 mL of human ectocervical 3D media (refer to Subheading 2.2.1). Transfer the resuspended cells into a collagen type 1 coated plate/flask, allocating 2 mL for one well of a six-well plate or 5 mL for one T25 flask. Refer to Subheading 2.2.1 for the preparation protocol of the collagen type 1 coated plate/flask. 3. Place the plate/flask in an incubator set at 37 °C with 5% CO2 incubator for 2–3 weeks. During this period, replace the medium with fresh human ectocervical 3D medium every 3–4 days. 4. Once the cells reach 80% confluency, passage them. For longterm stem cell maintenance, coculture the cells with the 3 T3J2 fibroblast cell line using the following method.

3.2.2 Fibroblast Cell Preparation for Epithelial Coculture and Long-Term Epithelial Stem Cell Maintenance

Note: When splitting the primary 2D ectocervical epithelial cells, preparing the 3 T3-J2 cell line in advance is crucial. Prepare 3 T3-J2 cell line culture 1–2 weeks before splitting the ectocervical 2D epithelial cells from 2D passage 0–2D passage 1. 1. Inoculate a T25 flask with 1 × 105 3 T3-J2 cells in 5 mL of complete DMEM medium (refer to Subheading 2.2.1), and place it in a humidified incubator with 5% CO2 at 37 °C. Once the cells achieve 90% confluence, decant the medium and rinse the cell monolayer two to three times with warm 1× DPBS. 2. Apply 1 mL of warm trypsin-EDTA solution to the cell monolayer to ensure an even distribution across the entire surface of the T25 flask. Transfer the flask to a 5% CO2 incubator at 37 °C for 3–4 min or until the cells begin to dissociate from the flask.

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3. Neutralize the trypsin solution by adding 10 mL of complete DMEM, and then centrifuge at 300 g for 5 min at 4 °C. After discarding the supernatant, replate the cell pellet in a T25 flask at the necessary splitting ratio (see Note 10). 4. When the 3 T3-J2 cells reach 70–80% confluence, proceed with gamma irradiation to induce cell cycle arrest, applying a total dose of 30 Gy. Thereafter, store the flask in a 5% CO2 incubator at 37 °C for use within 1 day (see Note 11). 3.2.3 Coculturing of Human Ectocervical Stem Cell with Fibroblast Cells in 2D

1. Upon achieving 80–90% confluence of the ectocervical epithelial cells, remove the medium and rinse twice with 5 mL of warm DPBS (see Note 12). 2. Apply 4 mL of prewarmed TryplE solution to the epithelial monolayers, followed by incubation at 37 °C in a 5% CO2 incubator for 8–15 min (see Note 13). 3. Transfer the cell suspension into a 15 mL Falcon tube. Utilize a heat-narrowed glass Pasteur pipette or a 26 G needle to segregate the cells by pipetting the suspension up and down six times. 4. Add 5 mL of ADF++ medium and centrifuge at 1000 g for 5 min at 4 °C. Discard the supernatant and resuspend the cell pellet in 4 mL of human ectocervical organoid medium (refer to Subheading 2.2.1). 5. Transfer the resuspended cells to the T25 flask containing irradiated or mitomycin C treated 3 T3-J2 feeder cells (refer to Subheading 3.2.2). Incubate in a 37 °C incubator with 5% CO2 until the epithelial stem cells reach 70–80% confluence. Refresh the medium every 3–4 days (see Note 14). 6. Once the desired epithelial cell confluence is reached, remove the medium and wash twice with warm DPBS. To selectively detach 3 T3-J2 cells, add 2 mL of prewarmed TrypLE for 1–2 min at 37 °C in a CO2 incubator. Verify the detachment of all feeder cells under a microscope. 7. Maintain the flask in an upright orientation and add 5 mL of warm DPBS to the bottom of the flask. Tilt the flask horizontally and gently cleanse the monolayer by swaying it from side to side. Finally, remove and discard the supernatant. Avoid applying DPBS solution directly onto the monolayer’s surface to prevent epithelial cell detachment from the flask. 8. Repeat steps 2–3 from Subheading 3.2.3, and centrifuge at 1000 g for 5 min at 4 °C. Discard the supernatant and utilize the cell pellet for expansion via coculture with a splitting ratio of 1:4 to 1:5 or for 3D ALI, organoid culture, or cryopreservation.

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3.3 Human Ectocervical 3D Epithelial Tissue Culture from 2D Stem Cells Using Air-Liquid Interface Approach 3.3.1 Human Ectocervical Air-Liquid Interface Cultures Using Transwells

1. Prepare the collagen mixture. Take the required number of 3 T3-J2 feeder cells (2.5 × 105 per ml of collagen mixture is necessary) (refer to steps Subheading 3.2.2). Centrifuge the cells at 300 g for 5 min at 4 °C to pellet them, and then discard the supernatant and place the pellet on ice. Important: To avoid collagen solution solidification, perform the following steps on the ice. 2. Resuspend the feeder cells in 1/10 volume of prechilled 10x reconstitution buffer, followed by 1/10 volume of prechilled 10× DMEM, 8/10 volume with prechilled collagen from bovine skin, and 2.4 μL of sterile 10 N NaOH per mL of collagen mixture (see Note 15). 3. Position the cell culture inserts (hydrophilic PTFE, 0.4 μm, Merck KGaA) into a 6-well or 24-well plate. Introduce the collagen-fibroblast mixture into the insert, using 2.5 mL per well in a 6-well plate and 350 μL per well in a 24-well plate. 4. Incubate the plates in a 5% CO2 incubator at 37 °C for 3 h to solidify the mixture. 5. Once solidified, add 2.5 mL per well of the human ectocervical medium in the 6-well plate (1 mL in the insert, 1.5 mL outside the insert) or 0.5 mL per well in the 24-well plate (100 μL in the insert, 400 μL outside the insert). 6. Resuspend the ectocervical epithelial stem cells (refer to 3.2.3) in the human ectocervical medium at 1 × 106 per mL concentration. Add 1 mL per well for a 6-well plate or 0.25 mL per well for a 24-well plate of the resuspended cells on top of the insert. Incubate in a 5% CO2 incubator at 37 °C. 7. On the third day, remove all media from the insert’s top and bottom. Add fresh human ectocervical media only to the bottom of the insert: 1.5 mL per well in a 6-well plate or 400 μL per well in a 24-well plate. 8. Every 2 days, replace all media from both sides of the insert, adding fresh media only to the bottom of the insert until the ALI culture is ready for harvesting on day 17.

3.3.2 Harvesting, Fixation, Embedding, and Preparation of ALI Culture Sections for Staining

1. Remove the medium from both inside and outside the insert, and then add 3.7% paraformaldehyde (PFA) into these regions. Allow it to incubate overnight at room temperature (RT). On the following day, discard the PFA and rinse the Transwell insert twice with PBS. Each rinse should last 5 min and be carried out at RT. Store the inserts at 4 °C until ready for embedding. 2. For embedding, excise the Transwell membrane with ALI culture from the plastic casing using a scalpel, and then halve it using a pair of scissors. To the Biopsy Cryomold, add 500 μL of

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preheated (5 min at 95 °C) Histogel (Richard-Allan ScientificTM HistoGel™ Specimen Processing Gel). Place the halved ALI membrane in an upright position in the Histogel, holding it with forceps until the Histogel solidifies on ice. 3. Transfer the solidified Histogel to a plastic embedding cassette, seal the lid, and label the cassette using a pencil. Place this cassette inside the processing chamber of the Leica TP1020 tissue processor. Proceed with the paraffinization procedure overnight, immersing the tissue in a series of reagents for 1 h each with agitation. This series includes 50% ethanol, 70% ethanol, 90% ethanol, 100% ethanol × 2, 100% isopropanol × 2, 100% xylene, and molten paraffin (see Note 16). 4. Embed the dehydrated and paraffinized Histogel block with the ALI membrane using a paraffin embedding machine. After cooling for at least 1 hour, extract the paraffin-embedded Histogel with ALI membrane with the cassette. These paraffinized blocks can be stored for years at 25 °C (see Note 17). 5. Make 3–5-μm-thick sections using a paraffin rotation microtome on coated slides. These sections are then ready for further staining processes. 3.4 Human Ectocervical 3D Organoids from 2D Stem Cells Using Matrigel Scaffold

1. Thaw the Matrigel on ice before the start of the experiment, and preheat 24 well plates at 37 °C for a minimum of 1 h (see Note 18).

3.4.1 Generation of Human Ectocervical 3D Organoids from 2D Stem Cells

3. In an Eppendorf tube, prepare a cell suspension equal to the desired number of wells for 3D organoid culture (10,000 cells per well/50 μL of Matrigel) (see Note 19).

2. Resuspend the cell pellet from Subheading 3.2.3. and perform cell count.

4. Centrifuge at 1000 g for 5 min at 4 °C to pellet the cells, and then discard the supernatant. Allow the cell pellet to chill on ice for at least 5 min (see Note 20). 5. Mix the cell pellet with 50 μL of Matrigel per well, and dispense a 50 μL droplet onto the center of the prewarmed 24-well plate. Incubate the plate in 5% CO2 at 37 °C for a minimum of 15 min to enable Matrigel polymerization (see Note 21). 6. Following Matrigel polymerization, supplement with 500 μL of prewarmed human ectocervical medium, and continue to incubate in a 5% CO2 at 37 °C incubator. Refresh the medium every 3–4 days. Perform cryopreservation on day 3 of organoid culture. After approximately 14 days, once the organoids have reached a size of roughly 150 μm, proceed to split the organoids as outlined below.

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3.4.2 Splitting and Passaging Ectocervical Organoids

1. Add 500 μL of ice-cold ADF++ to each Matrigel well. Mix well using a 1 mL pipette tip before transferring the suspension into a Falcon tube that contains 4 mL of cold ADF++. Centrifuge the Falcon tube at 300 g for 5 min at 4 °C. Discard the supernatant using a 5 mL pipette and remove the Matrigel layer using a 1 mL pipette tip without disturbing the organoid pellet (see Note 22). 2. Following this, add 1 mL of warm TrypLE to the resultant pellet, and resuspend the pellet by aspirating 200 μL three to four times. Incubate in a shaker set at 37 °C for 30 min at 100 rpm. 3. Next, fragment the organoids using a Pasteur pipette or a 26 G needle, and then add 3 mL of ADF++. Count the cells and then aliquot 10,000 cells for seeding each well. 4. Centrifuge the mixture at 1000 g for 5 min at 4 °C. Discard the supernatant while retaining 500 μL, and then resuspend the pellet before transferring the mixture to an Eppendorf tube. 5. Repeat the centrifugation and discard the supernatant. Resuspend the pellet in Matrigel, utilizing 50 μL per well, and then place the droplet on a 24-well plate. Place the plate in an incubator for 15 min at 37 °C and 5% CO2. Following this, add 500 μL of the warm ectocervical organoid medium. Continue incubating for 10–14 days, refreshing the medium every 3–4 days.

3.4.3 Harvesting and Fixation of Organoids

1. Medium removal and initial wash: Remove the medium from the 24-well plate. Add ice-cold 1× PBS with 0.1% BSA and gently mix up and down three to four times using a 1 mL pipette tip. Transfer the solution into glass tubes. 2. Settling of organoids: Add 4 mL of ice-cold 1× PBS with 0.1% BSA to the glass tube. Allow the organoids to settle down by gravity while keeping them on ice for 15 min. 3. Supernatant removal and repeated washing: Carefully remove the supernatant using a 5 mL pipette, leaving behind 1 mL of solution. Add 5 mL of ice-cold 1× PBS with 0.1% BSA to the glass tube. Allow the organoids to settle down while keeping them on ice for 15 min. 4. Repeat this process three times to remove all the Matrigel from the organoids. Perform a final wash with 5 mL of ice-cold 1× PBS without 0.1% BSA. These organoids can be PFA fixed or used for other studies. 5. PFA fixation: Add 5 mL of prewarmed 4% PFA and incubate for 1 hour at 25 °C. Exchange the PFA with 1× PBS and proceed with the dehydration and paraffinization process as described previously [8].

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Notes 1. The penicillin-streptomycin solution is harmful; avoid skin contact. 2. Do not use the precipitated solution as it induces cell toxicity. Avoid skin contact and inhalation. 3. Prepare collagen-coated plates/flasks beforehand. Alternatively, dry the collagen-coated plates/flasks for 1 h a day in advance, and store them at 4 °C. 4. Self-produced Wnt-3a and Rspondin-1 conditioned media should be tested for activity and microbial contaminants. 5. Adherence to the following guidelines is vital to ensure optimal organoid growth. Reconstitute the growth factors at the recommended concentrations. Ensure their usage before they reach their expiry date. Meticulously confirm the inclusion of each growth factor while preparing the 3D medium. Failure to include any essential growth component or using expired ones could result in diminished growth in 3D, or even its complete absence. 6. Thaw Matrigel stock overnight on ice at 4 °C. Mix well and make 1 mL aliquots in prechilled 1.5 mL Eppendorf tubes on ice. Store at -20 °C until the expiry date specified by the manufacturer. Always keep thawed Matrigel on ice. 7. Strive to mince as finely as possible, aiming for a size less than 0.5 mm3. This is to prevent inadequate digestion in subsequent steps and ensure a higher recovery of epithelial cells. 8. Ensure the Falcon tube cap is securely sealed with parafilm to prevent any solution leakage during incubation. 9. It is crucial to accurately calculate and seed an appropriate quantity of cells per Matrigel. Overseeding epithelial cells can lead to the formation of overcrowded organoids and escalate the acidity of the medium, indicated by a color change to yellow. This altered condition can impose stress on the epithelial cells, potentially leading to the abrupt death of organoids due to an insufficient supply of growth components from the medium. 10. For coculture purposes, divide the 3 T3-J2 cells at a ratio of 1: 2. For passaging, the ratio can be up to 1:10. 11. Ensure adherence to operating guidelines provided by an authorized operator for the irradiation machine. Calculate the duration necessary to reach a 30 Gy dose based on the irradiation machine’s current dose rate (Gy/min). Alternatively, use 0.1 mL of mitomycin C solution (refer to Subheading 2.2.1) in 5 mL of complete DMEM medium for 3 h to

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arrest the cells. Discard the medium, wash the flask four times with warm DPBS, and use the cells in coculture. Arrested cells can also be stored in 8 mL of complete DMEM for 2 days in a 37 °C incubator with 5% CO2. 12. Be cautious to prevent the cells from reaching full confluence, as it may lead to cell stress. 13. Ensure the detachment of all cells by observing under the microscope. 14. For coculturing, approximately 0.1 million primary cells could be seeded in a single T25 flask, typically reaching confluency in about 6–8 days. If the epithelial cells take longer than 8 days to become confluent, it’s advisable to replenish the 3D ectocervical medium with the 3 T3-J2 cell line (refer to Subheading 3.2.2 but ensure to use TryPLE for detachment and subsequently wash with ADF++ before use). Alternatively, to counteract the slow growth of primary epithelial cells, the initial cell seeding density could be increased. This approach ensures the desired confluence is reached in 6–8 days. 15. Ensure to mix the contents by inverting the tube to avoid bubble formation. 16. This procedure can also be performed manually using the same series of reagents under a fume hood. 17. Position the flat side of the ALI membrane facing upward in the Histogel to facilitate sectioning through the filter’s midsection. Ensure the entire Histogel is on the same plane before adding the paraffin. 18. Ensure to completely thaw the Matrigel stock overnight at 4 °C or, at a minimum, 2 h prior to usage in ice. Thorough mixing of the Matrigel is crucial; without this step, the Matrigel may collapse after the addition of media, resulting in the growth of epithelial cells as monolayers. 19. If the volume of the cell suspension exceeds 500 μL, initially use a 15 mL Falcon tube for centrifugation, and then transfer the cell pellet into the Eppendorf tube with less than 500 μL of ADF++ medium. 20. It’s crucial to meticulously remove the majority of the medium, leaving behind only 10–30 μL. Exercise care to avoid any additional medium accumulating on the tube walls during the prechilling process on ice. This step helps to prevent unnecessary dilution and the subsequent collapse of Matrigel when new media are added. 21. It is crucial to avoid air bubble formation to prevent uneven organoid distribution and potential interference with imaging. Furthermore, incubating for less than 15 min could lead to Matrigel droplet dissolution upon medium addition.

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22. To prevent the loss of organoids during Matrigel removal, avoid using a vacuum aspirator. Instead, a 1 mL pipette tip is recommended for cautious extraction of Matrigel postcentrifugation. Moreover, using low-protein-binding tubes and low-retention pipette tips can help reduce potential organoid loss or their adhesion during the transfer process.

Acknowledgments Cindrilla Chumduri received financial support from Deutsche Forschungs-gemeinschaft (DFG CH2527/2-1, GRK2157, and CRC DECIDE). References 1. Fatehullah A, Tan SH, Barker N (2016) Organoids as an in vitro model of human development and disease. Nat Cell Biol 18(3):246–254. https://doi.org/10.1038/ncb3312 2. Chumduri C, Turco MY (2021) Organoids of the female reproductive tract. J Mol Med (Berl) 99(4):531–553. https://doi.org/10.1007/ s00109-020-02028-0 3. Koster S, Gurumurthy RK, Kumar N, Prakash PG, Dhanraj J, Bayer S, Berger H, Kurian SM, Drabkina M, Mollenkopf HJ, Goosmann C, Brinkmann V, Nagel Z, Mangler M, Meyer TF, Chumduri C (2022) Modelling chlamydia and HPV co-infection in patient-derived ectocervix organoids reveals distinct cellular reprogramming. Nat Commun 13(1):1030. https://doi. org/10.1038/s41467-022-28569-1 4. Kessler M, Hoffmann K, Fritsche K, Brinkmann V, Mollenkopf HJ, Thieck O, Teixeira da Costa AR, Braicu EI, Sehouli J, Mangler M, Berger H, Meyer TF (2019) Chronic chlamydia infection in human organoids increases stemness and promotes age-dependent CpG methylation. Nat Commun 10(1):1194. https://doi.org/10.1038/ s41467-019-09144-7 5. Hoffmann K, Berger H, Kulbe H, Thillainadarasan S, Mollenkopf HJ, Zemojtel T, Taube E, Darb-Esfahani S, Mangler M, Sehouli J, Chekerov R, Braicu EI, Meyer TF, Kessler M (2020) Stable expansion of high-grade serous ovarian cancer organoids requires a low-Wnt environment. EMBO J 39(6):e104013. https://doi.org/10.15252/ embj.2019104013

6. Chumduri C, Gurumurthy RK, Berger H, Dietrich O, Kumar N, Koster S, Brinkmann V, Hoffmann K, Drabkina M, Arampatzi P, Son D, Klemm U, Mollenkopf HJ, Herbst H, Mangler M, Vogel J, Saliba AE, Meyer TF (2021) Opposing Wnt signals regulate cervical squamocolumnar homeostasis and emergence of metaplasia. Nat Cell Biol 23(2):184–197. h ttps://d oi.org/10.1038/s415 56-02000619-0 7. Zadora PK, Chumduri C, Imami K, Berger H, Mi Y, Selbach M, Meyer TF, Gurumurthy RK (2019) Integrated phosphoproteome and transcriptome analysis reveals chlamydia-induced epithelial-to-mesenchymal transition in host cells. Cell Rep 26(5):1286–1302. e1288. https://doi.org/10.1016/j.celrep.2019. 01.006 8. Gurumurthy RK, Koster S, Kumar N, Meyer TF, Chumduri C (2022) Patient-derived and mouse endo-ectocervical organoid generation, genetic manipulation and applications to model infection. Nat Protoc 17(7):1658–1690. https:// doi.org/10.1038/s41596-022-00695-6 9. Lohmussaar K, Oka R, Espejo Valle-Inclan J, Smits MHH, Wardak H, Korving J, Begthel H, Proost N, van de Ven M, Kranenburg OW, Jonges TGN, Zweemer RP, Veersema S, van Boxtel R, Clevers H (2021) Patient-derived organoids model cervical tissue dynamics and viral oncogenesis in cervical cancer. Cell Stem Cell 28(8):1380–1396. e1386. https://doi. org/10.1016/j.stem.2021.03.012

Chapter 12 Ovine Trophoblast Cells: Cell Isolation and Culturing from the Placenta at the Early Stage of Pregnancy Paola Toschi, Irene Viola, Isabella Manenti, Silvia Miretti, Elisabetta Macchi, Eugenio Martignani, Paolo Accornero, and Mario Baratta Abstract Embryo development is dependent upon the exchange of oxygen and nutrients through the placenta, mainly composed of peculiar epithelioid cells, known as trophoblast cells. Normal trophoblast functionality plays a key role during the whole pregnancy, especially in the first stage of placentation. This chapter explains the techniques to obtain sheep primary trophoblast cells from the early placenta. Overall, procedures for cell isolation, culture, characterization, and cryopreservation are described. Key words Early placenta, Primary trophoblast cells, Sheep, Cell isolation, Morphology

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Introduction Cell culture is an invaluable tool for research in numerous fields. It facilitates the analysis of biological properties and processes that are not readily accessible at the level of lived organisms [1]. This is particularly true for research in the field of placentation as the organ will be easily accessible only at term or in case of pathological miscarriage making it challenging to study early placenta development [2]. Alternatively, the biology of placental cell development can be investigated using cell lines. Several trophoblast cells from human and rodent species have been widely used [3, 4], whereas a few cell lines from other mammals have been currently established [5, 6]. Since the sheep is a well-recognized model for studying placenta physiology [7], the present chapter focuses on the establishment of an in vitro model from the early ovine placenta (16–23 days of pregnancy). In this view, we have developed a primary cell system presenting typical features of sheep trophoblast, including invasiveness and progesterone secretion [8, 9].

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2.1 Placenta Collection

The following materials are necessary to collect placenta in the early stage of placentation in sheep (it means from 16 to 40 days of pregnancy) at the slaughterhouse. All procedures are carried out under an airflow workbench, except the collection of tissue. 1. Sterile gloves. 2. Polystyrene box: In case of hot temperatures and long distances from the laboratory, it is recommended to use an ice pack. 3. Sterile sampling bag. 4. Sterilized saline solution (0.9% NaCl), or Ringer’s lactate, or sterile PBS pH 7.2. 5. Penicillin and streptomycin solution 100× (100 UI/L penicillin +100 μg/L streptomycin). 6. 70% ethanol spray. 7. 500 mL sterile backers. 8. Sterile surgical instruments: forceps, scissors, and surgical blades. 9. 10 and 15 cm sterile cell culture dishes.

2.2 Placenta Processing and Trophoblast Cell Isolation

All the reagents and materials used under the airflow workbench must be sterile and previously sterilized, respectively. 1. 1000 and 100 μL pipettes. 2. Pipet-Aid. 3. 1000 and 100 μL sterile tips. 4. 50 mL sterile tubes. 5. 25 and 5 mL sterile serological pipettes. 6. 10 cm plastic cell culture dishes. 7. 20 mL sterile syringe. 8. 100 μm mesh-size cell strainer. 9. Sterile syringe filter with 0.22 μm pore size. 10. Centrifuge adapted from 50 mL tubes. 11. Thermostatic bath. 12. Thermostatic shaker. 13. Incubator supplied with 5% CO2 and 95% air. 14. Cell counter (i.e., Bu¨rker counting chamber).

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15. Sterile phosphate-buffered saline (PBS): 140 mM NaCl, 27 mM KCl, 7.2 mM Na2HPO4, 14.7 mM KH2PO4. Adjust to pH 7.2 and autoclave for 20 min at 121 °C. 16. Sterile ultrapure water (high-quality grade water 18 Megohm prepared using ion-exchange and reverse-osmosis apparatus is recommended). 17. Minimal Essential Medium (MEM). 18. 20 mg/mL bovine serum albumin (BSA): Dissolve and filter 1 g BSA in 50 mL PBS. 19. 5 mg/mL trypsin-EDTA. 20. 10 mg/mL collagenase IV: Dissolve the collagenase powder in MEM mL and filter-sterilize it with a 0.22 μm pore size filter unit. 21. Growth medium solution: Dulbecco’s Modified Eagle’s/F-12 Medium (DMEM/F-12), 1 UI/L penicillin and 1 μg/L streptomycin, 10% heat-inactivated fetal bovine serum (FBS), 2 mM L-glutamine, 1 mM sodium pyruvate, 0.1 mM nonessential amino acids (NEA). 2.3 Trophoblast Cell Culture

1. 1000 and 100 μL pipettes. 2. 1000 and 100 μL sterile tips 3. Pipet-Aid or vacuum pump. 4. 15 mL sterile tubes. 5. 1.5 mL sterile cryogenic vials. 6. Sterile cell culture plastic dishes. 7. Centrifuge adapted from 15 mL tubes. 8. Thermostatic bath. 9. Incubator supplied with 5% CO2 and 95% air. 10. Cell counter (i.e., Bu¨rker counting chamber). 11. PBS (Subheading 2.2, item 15). 12. Dimethyl sulfoxide (DMSO). 13. Heat-inactivated fetal bovine serum (FBS). 14. 5 mg/mL trypsin-EDTA. 15. Growth medium solution (Subheading 2.2, item 21). 16. Cryopreservation chamber. 17. Liquid nitrogen storage container. 18. Inverted phase contrast microscope. 19. Laminar flow tissue culture hood.

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1. Prepare the 1% transport solution by adding an antibiotic solution to the chosen sterile solution (0.9% NaCl, or PBS). The best option is to add 5 mL of 100 UI/L penicillin +100 μg/L streptomycin (1×) to 500 mL of sterile solution. 2. At the slaughterhouse, take with gloved hands the whole uterus (from ovaries to vagina) directly from the carcass. During this step, avoid the uterus keeping contact with other parts of the body, in particular the digestive tract, adipose tissue, and skin. When removing the uterus, beware of the bladder and the rectum because urine and feces can contaminate the sample (Fig. 1a) (see Note 1). 3. Put the uterus in a sterile sample box by adding an amount of transport solution. The used volume could be different depending on the size of the uterus; generally, the transport solution can be added to fully cover the sample. This step is necessary to keep the sample moisturized in a sterile environment until the processing step in the laboratory. If you collect more than one uterus, it is strongly recommended not to preserve them together to avoid any cross contamination. Once arrived in the laboratory, open the sterile sample box under the laminar flow tissue culture hood, and transfer the uterus in a 500 mL backer with a new transport solution to remove excess blood. Repeat these steps two times. 4. Then place the uterus in a glass Petri dish and clean it quickly with 70% ethanol for a few seconds. This passage helps to reduce the outside-inside contamination during the following step (Fig. 1b).

Fig. 1 Placenta collection. The uterus is placed into a sterile dish to check the presence of corpus luteus on the ovary (a). Then, the uterine horns are carefully opened with a full-thickness incision on the cervix; the whole tissue complex (arrowhead) including the placenta, extraembryonic annexes, and embryo immediately flows out like a transparent liquid-filled membrane (b). Finally, the tissue complex is gently moved into a new sterile dish to clean it from the blood and examine the structure (c)

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5. Select the uterine horn ipsilateral to the corpus luteum. The pregnant-uterine horn always appears more expanded “sphereshaped.” 6. Using sterile surgical instruments to separate the uterus from the placenta. Open the selected horn following the suggestions: Make with a surgical blade a full-thickness orthogonal incision 1–1.5 cm below the expanded region on the uterine horn basis; exert gentle pressure from the upper part of the horn to make easier the placenta release from the horn. During the early stage of placentation, the placenta tissue is not strongly adherent to the endometrium, so this procedure allows for exteriorization gently keeping the whole tissue complex undamaged (placenta, extraembryonic annexes, and embryo) (Fig. 1b). 7. Move the tissue complex into a new plastic cell culture dish, so use sterile scissors and forceps to excise only the placenta, separating it from embryo annexes (amnios and yolk sac) without cutting the embryo (Fig. 1c) (see Note 2). 8. Repeat an additional washing of the placenta in the transport solution. The placenta must be preserved in the washing solution until the following step. 3.2 Placenta Processing and Trophoblast Cell Isolation

The placenta processing includes double digestion methods, that is, mechanical and enzymatic, respectively. 1. Before starting the placenta processing, prepare the digestive solution in a 50 mL sterile tube, and add the following: (a) 5 mL of 5 mg/mL trypsin-EDTA (final concentration of 0.5 mg/mL) (b) 2.5 mL of 20 mg/mL BSA (final concentration of 1 mg/ mL) (c) 2.5 mL of 10 mg/mL collagenase IV (final concentration of 0.5 mg/mL) (d) 40 mL MEM (see Note 3) 2. Mechanical digestion: Place the placenta in a 10 cm cell culture, and quickly add a small amount of MEM to avoid excessive tissue drying. Using sterile scissors and a surgical blade, finely mince the placenta into pieces small enough to be drawn with a 10 mL disposable pipet. 3. Enzymatic digestion: Transfer the obtained viscous and bloody mush to a 50 mL sterile tube by adding a 25 mL digestion solution. Incubate the tube for 90 min at 37 °C in a shaking incubator adjusted to 100 moves/min. Vortex the tube every 30 min for 30 s at medium speed. After digestion to better dissolve any remaining cellular aggregates, draw the solution using a 1000 μL cutting tip and pipetting many times (see Note 4).

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4. Filter the cell suspension through a 100 μm mesh-size cell strainer in a clean 50 mL sterile tube, and stop the enzymatic digestion by adding a 10% FBS of the total filtered volume. Centrifuge the filtrate at 300 g per 10 min. 5. Discard the supernatant and add onto the pellet 5 mL of sterile UP water for 5–10 s to wash and remove blood cell contamination as the blood cells undergo hemolysis in the water. Immediately after, add an equal amount of 2× PBS (5 mL) to equilibrate the solution osmolarity, and centrifuge the tube again at 300 g for 10 min (see Note 5). 6. During the centrifugation, prepare the growth medium solution as described in Subheading 2.2, item 21. Filter into a new 50 mL sterile tube the growth medium solution using a 20 mL sterile syringe with a 0.22 μm pore size filter, and put it at 37 °C thermostatic bath (see Note 6). 7. Discard the supernatant and mix the cell pellet with 8 mL of the growth medium solution, and place it in a 10 cm cell culture dish. Gently swirl the culture dish to evenly distribute the cell suspension, and then incubate at 37 °C with 5% CO2. 3.3 Trophoblast Cell Culture and Maintenance

1. Daily check cell suspension under the inverted microscope to follow cell development and control any microorganism contamination. If the culture medium appears clean and no bacteria are found, discard the growth medium every 48 h, wash cells with 2–3 mL PBS (×2) at room temperature, and then replace the medium previously heated at 37 °C in a thermostatic bath. Normally, cells reach an 80–90% confluence until 4 days of culturing (see Note 7). 2. To expand cells, discard the medium, wash 2–3 mL PBS (×2), and digest the intercellular cell junctions by adding 500–600 μL of trypsin-EDTA. Gently move the plate to cover the whole surface, and then put it for 3–5 min in the incubator at 37 °C with 5% CO2. Once all cells are detached, stop the digestion by adding at least 1000–1200 μL of growth medium. Transfer the suspension to a 15 mL sterile tube and centrifuge at 1200 rpm for 10 min. 3. Discard the supernatant and resuspend the cell pellet with 1 mL of growth medium. Use a cell counter for establishing the total number of cells (see Note 8). Once the total number of cells has been established, place at least 1 × 106 cells in a 10 cm plastic cell culture dish with an 8 mL growth medium. Gently swirl the culture dish to evenly distribute the cell suspension, and then incubate at 37 °C with 5% CO2. Follow the cell culturing as explained in Subheading 3.1, step 1 (see Note 9).

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4. To maintain cells for further experiments, trophoblast cells may be routinely frozen, stored frozen in liquid nitrogen, and then retrieved for the establishment of new cultures. (a) Freezing and storage: Repeat Subheading 3.3, step 2. Then, discard the supernatant and equilibrate by adding 900 μL FBS + 100 μL DMSO (total volume of 1 mL) to each 1 × 106 cell. Then transfer 1 mL of the cell suspension into 1.5 mL sterile cryovials, and position them within a cooler cryopreservation chamber precooled to 4 °C. Recover the cryopreservation chamber at -80 °C for 24 h at least (vials may be conserved until 1 month at 80 °C), and then move the frozen vials to the liquid nitrogen storage container, where they may be stored indefinitely. (b) Upon retrieval, firstly frozen aliquots should be rapidly thawed at 37 °C. To remove the freezing and storage medium, transfer the cell suspension to a 15 mL sterile tube, and centrifuge at 1200 rpm for 5 min. Discard the supernatant, resuspend the cell pellet in a suitable volume of the trophoblast growth medium, and continue from Subheading 3.2, step 7. 3.4 Trophoblast Cell Morphology

During the cell culture (Subheading 3.3), trophoblast cell growth and development must be followed and daily checked by microscope to ensure the trophoblast cell population shows typical properties (Fig. 2). Differentiated trophoblast cells are easy to recognize and distinguish from undifferentiated trophoblast stem cells. In particular, the trophoblast cell population presents different morphological characteristics in terms of cell size, shape, and number of nuclei. In this view, the following pictures are reported from ovine trophoblast cells from day 1 to day 7 of culture to better describe which kind of trophoblast cells may be identified in an in vitro ovine trophoblast cell model during cell culture time.

3.5 Trophoblast Cell Characterization

As outlined above, sheep primary trophoblast cells show typical morphological properties directly detected by microscope during cell culture. Besides cell morphology, trophoblast cell differentiation may be assessed by monitoring changes in gene expression of specific markers, hormone production, and typical trophoblast cell activity such as migration and invasiveness. Firstly, sheep trophoblast cells express different structural and functional placental markers. Immunocytochemistry and PCR analyses may be applied to detect cytokeratin-7 and interferon-tau (Fig. 3). Steroid and peptide hormones accumulate in the culture medium accompanying the differentiation of trophoblast cells. Progesterone is the major

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Fig. 2 Ovine trophoblast cell development (day 1–7). On day 1, trophoblast cells appeared as mononuclear cells (MNC, arrowhead) with epithelial cell-like proprieties. MNC showed a round or polygonal shape with a diameter of 20–50 μm (a). At 48 h of culture, MNCs started proliferating and forming a 5–10 smaller aggregated cellular cluster (arrowhead). Moreover, 10–20% of trophoblast cells become 10× larger than other MNCs (b). The MNC-circled clusters were distinguishable as a “single island” (arrowhead) on the whole plate surface after 5 days of culture. MNCs of each cluster were characterized by an evident nucleus and a thin ring of cytoplasm and showed a typical cobble-stone organization. Binucleate trophoblast cells (BNCs, arrow) with a diameter of 300–500 μm and an enlarged nucleus and prominent nucleoli were observed in the surrounding area of each MNC cluster (c–d). After 1 week of culture, the number of MNC clusters (arrowhead) increased as well as the number of differentiated BNCs (arrow) until almost covered the whole plate surface (e–f). The pictures shown in A, B, C, and E were captured at 5×; in D and F at 10× magnification. Scale bar 100 μm

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Fig. 3 Ovine trophoblastic marker detection. The panel displays the expression of ovine trophoblast marker in 21-day-old primary trophoblast cells compared to sheep fibroblast cells, using it as a negative control. The immunochemistry shows cytokeratin-7 (CK7) positively as a specific trophoblastic marker in ovine primary trophoblast cells, and vimentin (VIM) as a marker of fibroblast cells. The pictures were acquired at 20× magnification using a confocal fluorescent microscope. Scale bar 100 μm

steroid product and may be measured with a commercially available ELISA kit. Release of the prolactin family’s polypeptide hormones (i.e., placental lactogen) is monitored by Western blotting. Moreover, the migrative and invasive phenotype of trophoblast can be assessed by determining the directional movement of cells through a Transwell chamber system.

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Notes 1. The carcass is usually hung from the limbs, so the risk of contamination from the gut, bladder, and skin is lower. It is possible to take the uterus immediately after the abdominal incision and digestive tract removal (Fig. 4). 2. At this stage of placentation, embryo identification is easy with its 0.5–0.8 cm “bean-shape” pinkish aspect. You can’t see the cotyledons, but the placenta appears as a transparent-white tissue in the external part of the tissue complex. 3. Additionally, the digestion solution can include dispase II (final concentration of 5 mg/mL) and DNase (final concentration of 0.2 mg/mL). 4. To not lose time, remember to set up the correct temperature of the shaking incubator before starting the Subheading 3.2, step 2. 5. Alternatively, add 1 mL of red blood cell lysis buffer, incubate for 2–3 min at 37 °C, and then centrifuge. In that way, you don’t use UP water and 2× PBS, so this step carries on with more confidence.

Fig. 4 Sheep placenta is collected from the carcass at the slaughterhouse. Once the abdomen is open, the uterus (arrowhead) appears between the bladder (arrow upper) and gastrointestinal organs (arrow below)

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6. To not lose time, remember to set up the selected temperature of the thermostatic bath before starting the growth medium solution. Then, the residual growth medium solution may be stored at 4 °C for a few days. 7. Considering that trophoblast cell isolation begins from the whole placenta tissue, any mixed population may occur. After 4 days of culture, fibroblast cells may be present in the culture. In this case, to select only trophoblast cell types, it is recommendable to expand cells at least more than one time or until fibroblasts disappear. 8. Every cell counter manufacturer provides a diagram and counting instructions that should be consulted prior to carrying out cell counts for the first time. 9. Primary trophoblast cells may be used until passages 8–9.

Acknowledgments This work is supported by TOSP RILO 20 01 and TOSP GFI 22 01 F. References 1. Pan C, Kumar C, Bohl S, Klingmueller U, Mann M (2009) Comparative proteomic phenotyping of cell lines and primary cells to assess preservation of cell type-specific functions. Mol Cell Proteomics 8:443–450. https://doi.org/10.1074/ mcp.M800258-MCP200 2. Li L, Schust DJ (2015) Isolation, purification and in vitro differentiation of cytotrophoblast cells from human term placenta. Reprod Biol Endocrinol 13:71. https://doi.org/10.1186/ s12958-015-0070-8 3. Sahgal N, Canham LN, Konno T, Wolfe MW, Soares MJ (2005) Modulation of trophoblast stem cell and giant cell phenotypes: analyses using the Rcho-1 cell model. Differentiation 73:452–462. https://doi.org/10.1111/j. 1432-0436.2005.00044.x 4. Msheik H, Azar J, El Sabeh M, Abou-Kheir W, Daoud G (2020) HTR-8/SVneo: a model for epithelial to mesenchymal transition in the human placenta. Placenta 90:90–97. https:// doi.org/10.1016/j.placenta.2019 5. Kim JY, Burghardt RC, Wu G, Johnson GA, Spencer TE, Bazer FW (2011) Select nutrients in the ovine uterine lumen. VII. Effects of arginine, leucine, glutamine, and glucose on

trophectoderm cell signaling, proliferation, and migration. Biol Reprod 84:62–69. https://doi. org/10.1095/biolreprod.110.085738 6. Nakano H, Shimada A, Imai K, Takezawa T, Takahashi T, Hashizume K (2002) Bovine trophoblastic cell differentiation on collagen substrata: formation of binucleate cells expressing placental lactogen. Cell Tissue Res 307:225– 235. https://doi.org/10.1007/s00441-0010491-x 7. Barry JS, Anthony RV (2008) The pregnant sheep as a model for human pregnancy. Theriogenology 69:55–67. https://doi.org/10.1016/ j.theriogenology.2007.09.021 8. Seo H, Bazer FW, Burghardt RC, Johnson GA (2019) Immunohistochemical examination of trophoblast syncytialization during early placentation in sheep. Int J Mol Sci 20:4530. https:// doi.org/10.3390/ijms20184530 9. Viola I, Toschi P, Manenti I, Accornero P, Baratta M (2023) Modulatory role of mTOR in trophoblast adaptive response in the early stage of placentation in sheep. Reproduction 165: 313–324. https://doi.org/10.1530/REP22-0356

Chapter 13 Amniotic Membrane and Amniotic Epithelial Cell Culture Angelo Canciello, Adrian Cervero`-Varona, Maura Turriani, Valentina Russo, and Barbara Barboni Abstract Amniotic membrane (AM) is considered an important medical device for applications in regenerative medicine. The therapeutic properties of AM are due to its resistant extracellular matrix and to the large number of bioactive molecules released by its cells. To this regard, ovine amniotic epithelial cells (AECs) are a subset of placental stem cells with great regenerative and immunomodulatory properties. Indeed, either oAEC or AM have been object of intense study for regenerative medicine, thanks to several advantages in developing preclinical studies on a high value translational animal model, such as sheep. For this reason, a critical standardization of cultural practices is fundamental in order to maintain, on one hand, AM integrity and structure and, on the other hand, oAEC native properties, thus improving their in vivo therapeutic potential and clinical outcomes. In addition, freshly isolated AECs or AM can be exploited to produce enriched immunomodulatory secretomes that had been used with success into cell-free regenerative medicine procedures. To this aim, here is described an improved oAEC cultural protocol able to preserve their native epithelial phenotype also after the in vitro amplification and an innovative AM in vitro cultural protocol design to prolong the integrity and the biological properties of this tissue in order to collect stable conditioned media enriched with immunomodulatory factors. Key words Amniotic membrane, Amniotic epithelial cells, Epithelial-mesenchymal transition, Progesterone, Cell culture, Stem cells, In vitro amplification

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Introduction Placenta represents an important source of stem cells, easy to obtain without any ethical concerns [1–3]. Ovine amniotic epithelial cells (oAEC) are a subset of fetal stem cells located into the inner layer of the amniotic membrane (AM) and are among the most studied placental stem cells because of their peculiar properties [1, 4, 5]. In this regard, oAEC possess an epithelial phenotype and show a typical cobblestone-like morphology. Moreover, oAEC display embryonic markers such as SSEA-1, SSEA-3, SSEA-4, TRA-1-60, and TRA-1-81 and several surface adhesion molecules (CD29,

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CD49f, CD58, and CD166) and express pluripotent genes (Oct4, Sox2, Nanog, and Tert), while they result in negative for CD117 and for any hemopoietic markers (e.g., CD14, CD31, and CD45) [1, 5, 6]. Several evidences have demonstrated the ability of both human and animal derived AEC to differentiate into derivatives of all three germ layers (endoderm, mesoderm, and ectoderm), thus supporting their high plasticity [1, 3, 6, 7]. Moreover, since placenta is the natural site where fetusmaternal immune tolerance is played, oAEC display low immunogenicity. Indeed, oAEC show low expression of MHC I antigens and of no detectable levels of MHC II [1, 5, 6]. Furthermore, the long persistence of oAEC in host tissues is also helped by the extensive immunomodulatory and anti-inflammatory secretory activities of these cells. In fact, it was largely demonstrated the ability of oAEC to suppress in vitro the proliferation of stimulated peripheral blood mononuclear cell (PBMC), to reduce in vivo the infiltration of inflammatory cells as well as to stimulate the activation of M2 macrophage subpopulation [7–11]. Analogously, freshly isolated AM have been demonstrated to have an immune modulatory activity mainly mediated from the release of PGE2. Of note, differently from other source of mesenchymal stem cells, human AM are able to express an intrinsic immunomodulatory action as verified on T cell proliferative test [12] independently of any preliminary inflammatory activation. For these reasons, freshly isolated AECs or AM can be exploited to produce enriched immunomodulatory secretomes that had been used with success into cellfree regenerative medicine applications [13–15]. Increasing evidences revealed that both human and ovine AEC phenotype and biological activities can be strongly affected by cultural protocol and in vitro amplification [4, 16, 17]. Indeed, oAEC during in vitro expansion undergo epithelial-mesenchymal transition (EMT), a trans-differentiation process whereby they spontaneously lose the epithelial phenotype by progressively acquiring the mesenchymal one [4]. Beside phenotypical shift, oAEC also experienced a dramatic reduction of their immunomodulatory properties, upon EMT [4]. Conversely, the in vitro supplementation of progesterone (P4) – the main pregnancy hormone that has been previously involved in the regulation of EMT [18– 21] – is able to prevent the spontaneous EMT in oAEC and to preserve the native biological properties [4]. To this regard, recently it has been proposed a novel cultural approach able to maintain in vitro the morphological and biological properties of amniotic membrane (AM) that involved the use of P4. Indeed, the supplementation of P4 to the AM culture has been proven preserving tissue morphology and architecture even after 3 days of culture [22].

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Materials Use only sterile materials. Prepare all reagents at room temperature and store them at 4 °C (unless indicated otherwise). Use all reagents at 38 °C. Diligently follow all waste disposal regulations when disposing waste materials.

2.1 Cell Isolation, Growth, and Cryopreservation

1. Clean with soap and tap water the scalpels, the forceps, and the surgical and watchmaker tweezers. Rinse with distilled and double-distilled water and then sterilize in oven. Store in a sterile container. Before the use, sterilize the metallic instruments and with a glass bead sterilizer set at 200 °C for 15 s (see Note 1). 2. Antibiotic buffer for membrane isolation (use outside the laminar flow hood): 1% of 10.000 UI/mL penicillin-streptomycin in sodium chloride solution suitable for cell culture (see Note 2). Store at room temperature. 3. Antibiotic buffer for cell isolation (use inside the laminar flow hood): 1% of 10.000 UI/mL penicillin-streptomycin in phosphate buffered saline (PBS) without calcium, without magnesium, suitable for cell culture (see Note 3). Store at room temperature. 4. P4 stock solution: Weigh 100 mg of P4 and transfer in a sterile 15 mL tube (see Note 4). Add absolute ethanol to a volume of 10 mL and gently mix the solution. Work under fume hood. The concentration of P4 stock solution is 31.8 mM. Store a 4 ° C. 5. P4 working solution: Dilute 5 mL of P4 stock solution in 5 mL of absolute ethanol and gently mix the solution. Work under fume hood. The concentration of P4 working solution is 15.9 mM. Store a 4 ° C. 6. Growth medium: Alpha-Minimum Essential Eagle Medium supplemented with 20% fetal bovine serum (FBS), 1% Ultraglutamine, 10.000 UI/mL penicillin-streptomycin, and 2.5 μg/mL amphotericin (see Note 5). Add P4 at final concentration of 25 μM by discarding the equivalent volume of growth medium to exactly maintain the correct P4 concentration. Gently mix the solution (see Note 6). Filter the medium in a 0.22 μm filter, and equilibrate in an incubator at 38.5 °C, 30 min before the use. 7. Freezing medium solution: 10% (v/v) of dimethyl sulfoxide (DMSO) in FBS. Filter the solution with a 0.22 μm filter.

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2.2 AM Tissue Culture and Viability Assay

1. Ultralow attachment or nonadherent culture plates (see Note 7). 2. Growth medium: Alpha-Minimum Essential Eagle Medium supplemented with 20% FBS, 1% Ultraglutamine, 10.000 UI/mL penicillin-streptomycin, and 2.5 μg/mL amphotericin. Add P4 at final concentration of 25 μM by discarding the equivalent volume of growth medium to exactly maintain the correct P4 concentration and gently mix the solution. Filter the medium in a 0.22 μm filter and equilibrate in an incubator at 38.5 °C, 30 min before the use. 3. Calcein AM: Prepare 4 mM Calcein AM stock solution in DMSO. Use Calcein AM at final concentration of 4 μM by directly diluting 1:1000 the stock solution in growth medium. 4. Propidium iodide: Prepare 2.4 mM propidium iodide stock solution in H2O. Use propidium iodide at final concentration of 2.4 μM by directly diluting 1:1000 the stock solution in growth medium. 5. 4′,6-Diamidino-2-phenylindole (DAPI): Prepare 5 mg/mL DAPI stock solution in H2O. Use DAPI at final concentration of 5 μg/μL by directly diluting 1:2000 the stock solution in growth medium.

2.3 oAEC Flow Cytometry Characterization

1. Unconjugated primary antibodies marked with FITC by using Zenon Antibody Labelling Kit (Gibco, Invitrogen, Carlsbad, CA, USA), following the manufacturer’s instructions. 2. Washing buffer: 0.1% (v/v) of sodium azide and 0.5% (v/v) of bovine serum albumin (BSA) in PBS. 3. Fixation buffer: 0.5% (v/v) of paraformaldehyde in PBS.

2.4 Immunofluorescence

1. Fixation solution: 4% (v/v) paraformaldehyde in PBS. 2. Permeabilization solution: 0.2% (v/v) Triton ×-100 in PBS. 3. Blocking solution: 5% (w/v) BSA in PBS. 4. Antibody dilution buffer: 1% (w/v) BSA in PBS. 5. Primary antibodies: anti-cytokeratin-8, anti-α-SMA. 6. Secondary antibodies: Cy3 and Alexa Fluor 488 conjugated antibodies.

3

Methods Carry out all procedures at room temperature unless otherwise specified.

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oAEC Isolation

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1. Carefully prepare the uterus and the incision site with denatured alcohol (see Note 8). 2. Open the uterus wall with the aid of surgical forceps. Afterward, gently separate placenta from the uterus by manually detaching the cotyledons from caruncles. 3. Once the placenta is isolated from the rest of the uterus, roughly peel off with surgical and watchmaker tweezers the chorioallantois from the amnion. 4. Cut amnion pieces with the aid of surgical tweezers and forceps, and put them into antibiotic buffer for membrane isolation. 5. Move the amnion pieces under laminar flow hood and put them into antibiotic buffer for cell isolation. 6. Working into a 10 cm petri dish filled with antibiotic buffer for cell isolation, divide the amnion in smaller pieces of about 3–5 cm of length by using sterile watchmaker tweezer and scalpel. 7. Manipulate under a stereomicroscope in order to finely remove the residual parts of chorioallantois from the amnion with the aid of fine watchmaker forceps (see Note 9). 8. Dissect amnion with sterile watchmaker tweezer and scalpel to get smaller tissue pieces. 9. Rinse three times for 15 min the amnion pieces with antibiotic buffer for cell isolation (see Note 10). 10. Incubate amnion pieces into trypsinization flasks (see Note 11). Add 0.25% trypsin-EDTA solution and a magnetic stir bar. Place the trypsinization flasks on a magnetic stirrer in 38 ° C water bath for 30 minutes, with consistent agitation (see Note 12). 11. Add FBS 10% (v/v) to cell suspension in order to inactivate trypsin. Collect cell suspension, filter through a 40 μm cell filter, and pour into a 50 mL tube. Centrifuge the cell suspension at 2500 rpm for 10 min. 12. Discard the supernatant and resuspend the pellet in pre-warmed and equilibrated growth medium. 13. Dilute 1:2 cell suspension in Trypan Blue into a 0.5 micro tube and gently pipette the suspension. Count the cell suspension by using a Bu¨rker counting chambers (see Note 13). 14. Seed the cells at the final concentration of 3 × 103 cells/cm2 in growth medium supplemented with 25 μM of P4, in order to preserve the epithelial phenotype (Fig. 2) (see Note 14). 15. Carefully mix the dish by gentle agitation to obtain an equal seeding of the cells. 16. Incubate the culture dish in incubator at 38.5 °C in 5% CO2.

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3.2 AM Tissue Culture

1. The amnion pieces collected in step 9 of the previous section (oAEC isolation) represent the starting point of this protocol. 2. Cut amnion pieces into smaller pieces of approximately 2.5 × 2.5 cm (see Note 15). 3. Move the amnion pieces into ultralow attachment or nonadherent plates in growth medium supplemented with 25 μM of P4. 4. Incubate AM pieces at 38.5 °C in 5% CO2 up to 3 days. 5. Refresh daily the growth medium.

3.3 AM Viability Assay

1. Make sure working all the time in the dark. 2. Add Calcein AM at final concentration of 4 μM (1:1000) directly to the culture medium for 45 min (see Note 16). 3. After 40 min, add DAPI at final concentration of 5 μg/μL (1: 2000) directly to the culture medium (see Note 17). It takes 5 min to counterstain nuclei. 4. Finally, after 3 min, add propidium iodide at final concentration of 2.4 μM (1:1000) to the culture medium (see Note 18). It takes 2 min to counterstain nuclei. 5. With the aid of forceps, wash quickly each amnion piece in PBS solution to eliminate the excess of dye and culture medium. 6. Afterward, gently place each amnion pieces onto a coverslip and spread the edges with the aid of a forceps. 7. Analyze cell samples with the aid of a confocal microscope (Fig. 1).

Fig. 1 Viability of in vitro cultured AM. CTR and P4-treated AM were in vitro cultured up to 3 days with or without progesterone and then were subjected to staining by using Calcein AM (green), DAPI (blue), and propidium iodide (red) to assess cell viability and mortality. Representative 40× magnification is shown for CTR and P4 day 1 acquisition; 60× magnification for days 2 and 3. Scale bar: 50 μm. (Micrographs modified from Canciello et al. 2020)

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Table 1 List of antibodies used in flow cytometry analysis

3.4 oAEC Flow Cytometry Characterization

Antigen

Dilution

Host

Company details

CD14

1:10

Mouse

LifeSpan bioscience

CD29

1:100

Rabbit

VMRD

CD31

1:10

Mouse

abD Serotec

CD45

1:10

Mouse

abD Serotec

CD49f

1:100

Rat

Beckman coulter

CD58

1:10

Mouse

LifeSpan bioscience

CD117

1:100

Rabbit

Abcam

CD166

1:100

Mouse

Ancell

MHC I

1:50

Mouse

Novus biologicals

MHC II

1:50

Rabbit

Abcam

1. Stain 5 × 105 cells/sample by incubating them with 100 μL of 20 mM ethylenediaminetetraacetic acid (EDTA) at 37 °C for 10 min. 2. Wash in 3 mL of washing buffer and centrifuge at 4 °C, 400 × g for 8 min. 3. Resuspend cell pellet in 100 μL washing buffer containing the appropriate dilution of surface antibody (see Table 1), and incubate for 30 min at 4 °C in the dark. 4. Wash with 3 mL of washing buffer, and centrifuge at 4 °C, 400 × g for 8 min. 5. Suspend cell samples with 1 mL of fixation buffer, and incubate for 5 minutes at room temperature (RT). 6. Wash cell samples by centrifuging at 4 °C, 400 × g, for 8 min, and store at 4 °C in the dark until the acquisition. 7. Analyze cell samples on a FACS flow cytometer.

3.5 Culture, Passing, and Cryopreservation of oAEC

1. Change the growth medium each 2 days (see Note 19). 2. Split the cells when they reach about 80% confluency (see Note 20). 3. Discard the growth medium and rinse three times with PBS without calcium, without magnesium (see Note 21). 4. Discard the PBS and then add 0.25% trypsin-EDTA by ensuring that trypsin covers the entire surface of the culture dish. Place the cells at 38.5 °C in 5% CO2 for 8–10 min (see Note 22).

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5. Inactivate the trypsin with growth medium, and pipette vigorously the cell suspension to detach cells (see Note 23). Collect cells in a sterile tube, and centrifuge the suspension at 2500 rpm for 10 min. 6. Discard the supernatant and resuspend the pellet in pre-warmed and equilibrated growth medium. 7. Perform the cell count and seed the cells at the same density (3 × 103 cells/cm2) in growth medium supplemented with 25 μM of P4 for the next cultural passage. 8. Incubate the culture dish at 38.5 °C in 5% CO2. 9. Centrifuge the rest of oAEC not seeded at 2500 rpm for 10 min. Discard the supernatant and resuspend the pellet in freezing medium. Carefully mix the cell suspension. 10. Distribute 1 mL of cell suspension for each cryovials. Store the cells at -80 °C for 1 day and then move them in liquid nitrogen. 3.6 Assessing Epithelial Phenotype by Immunofluorescence

1. Plate oAEC in 35 mm petri dish with a sterile glass coverslip on the bottom (see Note 24). Culture the cells until reaching 50–60% confluency (see Note 25). 2. Rinse three times with PBS with calcium, with magnesium. 3. Incubate oAEC in fixation solution for 10 min at RT (see Note 26). Rinse three times with PBS. 4. Add permeabilization solution for 10 min at RT, with gentle agitation. 5. Block nonspecific sites with blocking solution for 1 h at RT, with gentle agitation. 6. Incubate with anti-cytokeratin-8 (1:200) or anti-α-SMA (1: 200) primary antibodies diluted in antibody dilution buffer, overnight, at 4 °C in gentle agitation (see Note 27). 7. Rinse three times with PBS at RT, with gentle agitation. 8. Incubate with Cy3 (anti-α-SMA) or Alexa Fluor 488 (anticytokeratin-8) conjugated secondary antibodies diluted 1:200 in antibody dilution buffer for 40 min at RT in the dark, with gentle agitation. 9. Stain nuclei with DAPI used at the final dilution of 1:5000 in PBS for 5 min in the dark, with gentle agitation. 10. Mount coverslips and analyze cell samples with the aid of a fluorescent microscope equipped with a CCD camera, configured for fluorescence microscopy and interfaced to a computer workstation, provided with an interactive and automatic image analyzer. Digital images are acquired using standard filters set up for Cy3, Alexa Fluor 488, or DAPI (Fig. 3).

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11. Determine the percentage of cytokeratin-8 and α-SMA positive cells by counting at least 100 cells for each sample (see Note 28).

4

Notes 1. Do not exceed 15 s of sterilization to avoid the metal from reaching too high temperature. High temperature could damage the tissue and negatively influence the yield of cell isolation because it increases the percentage of cell death. Moreover, metal objects exposed to high temperature became incandescent and could cause damage to the operators. 2. Prepare the antibiotic buffer for membrane isolation under a laminar flow hood. Use only sterile materials. Sodium chloride solution should be stored at room temperature to avoid precipitation of the salt. 3. Prepare the antibiotic buffer for cell isolation under a laminar flow hood. Use only sterile materials. PBS without calcium, without magnesium, should be stored at room temperature to avoid precipitation of the salts. PBS without calcium, without magnesium, guarantees good results in terms of oAEC detached from amniotic membrane, as adhesion proteins require divalent cations for proper function. 4. P4 is carcinogen and toxic for the reproduction (H351, H360 risk category). Use individual protection devices (DPI) such as safety glass clear lens, molded disposable mask with valve, gloves, and lab coat. Wear the mask when weighing the P4. To avoid exposing P4 to coworkers, weigh P4 directly in a 15 mL tube (by storing the container weight as tare weight), and close the tube when transporting it to the fume hood. Add 10 mL of absolute ethanol and filter the solution before storage. The used 15 mL tube should be disposed as hazardous waste. 5. Warm all reagents to 38 °C in the water bath in order to reach ovine cell physiological temperature. 6. Absolute ethanol is generally used as solvent for dissolving several drugs. However, it is possible that high concentrations of absolute ethanol can negatively influence cell culture. In agreement with other investigators, the final concentration of absolute ethanol in cell culture medium should be about 0.1%. The high dilution of P4 working solution selected (1:625) allows to obtain exiguous volume of absolute ethanol into growth medium. In these experimental conditions, the final concentration of absolute ethanol in the medium will be about 0.16%.

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7. AM tissue culture should be performed in ultralow attachment or nonadherent plates in order to avoid that the detached cells from the amniotic tissue will attach the plastic surface. 8. The ovine uteri are collected at slaughterhouse. For this reason, a proper sterilization of the incision site and the surgical instruments is strongly recommended because it significantly reduces the amount of contamination. Since the uteri are collected from animals intended for food, all the animals are preventively subjected to a prophylaxis against the principal sexually transmitted infections. 9. Ruminants are characterized by the presence of a chorioallantois, which is established by fusion between the allantoic wall and chorion. Chorioallantois is often recognizable by the presence of blood vessels. On the other hand, in ruminants, epithelial layer of the amnion is often recognizable by the presence of amniotic plaques that are stratified with glycogen-rich epithelial elevations from the inside inner epithelium of the amnion [23]. 10. These amnion pieces obtained in this step can be used for AM tissue culture. Alternatively, follow the protocol to isolate oAEC from amnion pieces. 11. Do not fill the trypsinization flask with a high number of AM pieces in order to allow a proper cell separation. Put about 10–15 pieces of AM for each 125 mL trypsinization flask. 12. Better results are obtained by using 15–20 mL of 0.25% trypsin-EDTA solution. Moreover, select a proper speed of magnetic stirrer (200–230 rpm) to allow a consistent agitation of magnetic stir. 13. Since oAEC tend to form clusters in suspension, vigorously pipette the cell suspension before performing the dilution with Trypan Blue solution. Count the cell suspension at least three times to be sure of the result of the counting. Indeed, oAEC are quite sensible to cell density and improper cell counting or seeding can negatively influence their growth. 14. This P4 concentration allows the preservation of the native epithelial phenotype of oAEC. Indeed, when cultured in absence of P4 (standard cultural conditions), oAEC experienced epithelial-mesenchymal transition (EMT) in culture. In particular, it is previously demonstrated that oAEC undergo EMT after three cultural passages (see Ref. Canciello et al. [4]). Conversely, when the culture medium is supplemented with 25 μM of P4, oAEC preserved their native epithelial phenotype in a long-term culture (Fig. 2). 15. Working in a 10 cm petri dish, cut amnion with the aid of a 2.5 × 2.5 cm square benchmark designed on the bottom of the petri dishes. This size is important because it avoids the

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Fig. 2 Morphology of oAEC at the end of the first passage. In absence of P4 (without progesterone), oAEC undergo EMT by losing their native epithelial morphology and by acquiring the mesenchymal one. On the contrary, when oAEC are cultured in growth medium which is supplemented with P4 (with 25 μM progesterone), they preserve the native epithelial phenotype. Scale bar: 50 μm

Fig. 3 Immunostaining for epithelial (cytokeratin-8) and mesenchymal (α-SMA) markers. At the end of the first passage, oAEC cultured in growth medium supplemented with progesterone preserve a high expression of cytokeratin-8 and a low appearance of α-SMA. Scale bar: 25 μm

excessive folding of the amnion pieces during the culture. Indeed, the accidental folding of amnion pieces could cause the death of the cells at the edges. Alternatively, amnion pieces bigger than 2.5 × 2.5 cm could easily fold on themselves, thus reducing the oxygen exchanges of the tissue. If accidental folding occurs, gently rotate culture dish to facilitate amnion pieces spreading (Fig. 3).

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16. Calcein AM, propidium iodide, and DAPI could be used as vital fluorescent dyes by adding them directly into the culture medium. Calcein AM is a cell-permeant dye that is generally used to determine cell viability. In live cells, the nonfluorescent Calcein AM is converted to a green-fluorescent Calcein, after acetoxymethyl ester hydrolysis by intracellular esterases. 17. DAPI is a fluorescent stain that binds strongly to A-T rich regions in DNA. As DAPI can pass through an intact cell membrane, it can be used to stain both live and fixed cells, though it passes through the membrane less efficiently in live cells. Alternatively, Hoechst 33342 dye can be used which is a popular cell-permeant nuclear counterstain that emits blue fluorescence when bound to dsDNA. 18. Propidium iodide is red-fluorescent nuclear dye which binds to DNA by intercalating between bases. Since propidium iodide is not permeant to live cells, it is also commonly used to counterstain dead cells. 19. At the first passage, the percentage of oAEC that adheres at the culture dish is lower than the number of seeded cells. For this reason, it is suggested to change the grow medium after 3 days at beginning of the cell culture in order to allow the cells to properly adhere. 20. Generally, oAEC take about 10–12 days to reach 80% confluency when freshly isolated. After the first cultural passage, oAEC take about 4–5 days to reach 80% confluency. It is not recommended to exceed the 80% confluency to avoid the morphological changes due to epithelial-mesenchymal transition. 21. The wash steps remove any traces of serum, divalent cations that would inhibit the dissociation action of trypsin. 22. For the epithelial nature of oAEC, they are usually difficult to detach from the culture dish. For this reason, it is recommended to use 0.25% instead of 0.05% of trypsin-EDTA. Moreover, in order to properly detach cells and increase the yield of trypsinization, incubate the culture dish for 8–10 min and check them microscopically every 3–4 min. It is advisable to gently tap the dish in order to facilitate cell detachment. 23. Add the equivalent volume of growth medium to the cell suspension to inactivate the trypsin action. Since growth medium contains 20% FBS, using an equal volume of it with respect to trypsin allows to obtain a final dilution of 10% FBS. As epithelial cells, oAEC tend to form clusters when in suspension. These clusters are generally hard to dissociate and can negatively influence cell count because they can be formed by a high number of cells. For this reason, vigorously pipette cell suspension at least 25–30 times to facilitate cluster dissociation.

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24. Put glass coverslips (22 × 22 mm) into a beaker and sterilize in oven at 180 °C for 2 h. Afterward, immerse glass coverslips in absolute ethanol under laminar flow hood and let them become air-dried. Put the coverslip into a 35 mm petri dish with the aid of sterile watchmaker tweezer. 25. A confluency greater than 60% could negatively influence cell fixation and interfere with the interpretation of the data due to the overlapping of fluorescent signals. 26. Be careful in handling paraformaldehyde because it is flammable and may cause skin irritation, allergic skin reaction, eye damage, and respiratory irritation. Use personal protective equipment and avoid contact with the skin and eye. In order to avoid the formation of dust and aerosol, work under a fume hood. 27. Carefully aspirate the excess of PBS from the edges of the petri dish leaving a thin layer of PBS only on the coverslip. It could be helpful to use a PAP pen (ab2601) which allows to create hydrophobic barrier when a circle is drawn around a specimen on a slide. It is recommended to carefully add a drop of approximately 50–60 μL of diluted primary antibodies. Cytokeratin-8 is an intermediate filament protein characteristic of the epithelial cells and is considered one of the first markers that disappears in the context of EMT [24]. On the contrary, α-SMA is an actin isoform expressed by vascular smooth muscle, myoepithelial, and mesenchymal cells and is well defined as a marker of an advanced stage of EMT [25]. 28. Snap micrographs with low magnification in order to count at least 100 cells in each field. To determine the total number of cells per field, count all the nuclei counterstained with DAPI. In order to assess the percentage of epithelial or mesenchymal cells, count the cytokeratin-8 or the α-SMA positive cells, and divide the obtained number by the total number of nuclei and then multiply the result by 100.

Acknowledgments This project has received funding from the European Union’s Horizon 2020 research and innovation program under the Marie Skłodowska-Curie grant agreement no. 955685 (www.helsinki.fi/ p4fit).

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References 1. Mattioli M, Gloria A, Turriani M et al (2012) Stemness characteristics and osteogenic potential of sheep amniotic epithelial cells. Cell Biol Int 36:7–19. https://doi.org/10.1042/ CBI20100720 2. Murphy S, Atala A (2013) Amniotic fluid and placental membranes: unexpected sources of highly multipotent cells. Semin Reprod Med 31:062–068. https://doi.org/10.1055/s0032-1331799 3. Koike C, Zhou K, Takeda Y et al (2014) Characterization of amniotic stem cells. Cell Reprogram 16:298–305. https://doi.org/10.1089/ cell.2013.0090 4. Canciello A, Russo V, Berardinelli P et al (2017) Progesterone prevents epithelialmesenchymal transition of ovine amniotic epithelial cells and enhances their immunomodulatory properties. Sci Rep 7:3761. https://doi. org/10.1038/s41598-017-03908-1 5. Barboni B, Russo V, Berardinelli P et al (2016) Applications of placenta-derived cells in veterinary medicine. In: Parolini O (ed) Placenta tree life. CRC Press, pp 217–284 6. Barboni B, Curini V, Russo V et al (2012) Indirect co-culture with tendons or tenocytes can program amniotic epithelial cells towards stepwise Tenogenic differentiation. PLoS One 7:e30974. https://doi.org/10.1371/journal. pone.0030974 7. Barboni B, Mangano C, Valbonetti L et al (2013) Synthetic bone substitute engineered with amniotic epithelial cells enhances bone regeneration after maxillary sinus augmentation. PLoS One 8:e63256. https://doi.org/ 10.1371/journal.pone.0063256 8. Barboni B, Russo V, Curini V et al (2014) Gestational stage affects amniotic epithelial cells phenotype, methylation status, immunomodulatory and stemness properties. Stem Cell Rev Rep 10:725–741. https://doi.org/10. 1007/s12015-014-9519-y 9. Muttini A, Russo V, Rossi E et al (2015) Pilot experimental study on amniotic epithelial mesenchymal cell transplantation in natural occurring tendinopathy in horses. Ultrasonographic and histological comparison. Muscles Ligaments Tendons J 5:5–11 10. Mauro A, Russo V, Di Marcantonio L et al (2016) M1 and M2 macrophage recruitment during tendon regeneration induced by amniotic epithelial cell allotransplantation in ovine. Res Vet Sci 105:92–102. https://doi.org/10. 1016/j.rvsc.2016.01.014

11. Shandley L, Alcorn D, Wintour EM (1997) Ovine amniotic and allantoic epithelia across gestation. Anat Rec 248:542–553 12. Rossi D, Pianta S, Magatti M et al (2012) Characterization of the conditioned medium from amniotic membrane cells: prostaglandins as key effectors of its immunomodulatory activity. PLoS One 7:e46956. https://doi.org/10. 1371/journal.pone.0046956 13. Ragni E, Papait A, Perucca Orfei C et al (2021) Amniotic membrane-mesenchymal stromal cells secreted factors and extracellular vesiclemiRNAs: anti-inflammatory and regenerative features for musculoskeletal tissues. Stem Cells Transl Med 10:1044–1062. https://doi. org/10.1002/sctm.20-0390 14. Yang M, Wang L, Chen Z et al (2022) Topical administration of the secretome derived from human amniotic epithelial cells ameliorates psoriasis-like skin lesions in mice. Stem Cell Res Ther 13:393. https://doi.org/10.1186/ s13287-022-03091-9 15. Jeng BH, Hamrah P, Kirshner ZZ et al (2022) Exploratory phase II multicenter, open-label, clinical trial of ST266, a novel secretome for treatment of persistent corneal epithelial defects. Transl Vis Sci Technol 11:8. https:// doi.org/10.1167/tvst.11.1.8 16. Alcaraz A, Mrowiec A, Insausti CL et al (2013) Autocrine TGF-β induces epithelial to mesenchymal transition in human amniotic epithelial cells. Cell Transplant 22:1351–1367. https:// doi.org/10.3727/096368912X657387 17. Caruso M, Evangelista M, Parolini O (2012) Human term placental cells: phenotype, properties and new avenues in regenerative medicine. Int J Mol Cell Med 1:64–74 18. Jeon S-Y, Hwang K-A, Choi K-C (2016) Effect of steroid hormones, estrogen and progesterone, on epithelial mesenchymal transition in ovarian cancer development. J Steroid Biochem Mol Biol 158:1–8. https://doi.org/10. 1016/j.jsbmb.2016.02.005 19. van der Horst PH, Wang Y, Vandenput I et al (2012) Progesterone inhibits epithelial-tomesenchymal transition in endometrial cancer. PLoS One 7:e30840. https://doi.org/10. 1371/journal.pone.0030840 20. Sumida T, Itahana Y, Hamakawa H, Desprez P-Y (2004) Reduction of human metastatic breast cancer cell aggressiveness on introduction of either form a or B of the progesterone receptor and then treatment with progestins. Cancer Res 64:7886–7892. https://doi.org/ 10.1158/0008-5472.CAN-04-1155

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Chapter 14 Evaluation of the Epithelial Barrier Integrity in Primary Cultures of Pig Mammary Epithelial Cells Chiara Bernardini, Debora La Mantia, and Monica Forni Abstract A major feature of epithelial and endothelial cells is the creation of biological barriers able to protect the body against stressors that could compromise homeostasis. The ability to characterize biological barriers in vitro is an important study tool especially used for the intestinal barrier, the blood-brain barrier, and the lung barrier. The strength and integrity of biological barriers may be assessed by the measurement of the transepithelial/transendothelial electrical resistance (TEER) that reflects the ionic conductance of the paracellular pathway. The TEER measurement is a quantitative, non-invasive, highly useful, and representative method that must be strictly standardized. Here we describe a quantitative protocol to assess the mammary epithelial barrier integrity by combining the TEER measurement with a test for studying the passage of the sodium fluorescein, that is, a hydrophilic paracellular marker. Being the swine species an excellent translational model, primary cultures of mammary epithelial cells, isolated from hybrid pig tissue collected at slaughterhouse, are used. Key words Mammary epithelial cells, Transepithelial/transendothelial electrical resistance, Sodium fluorescein, Apical compartment, Basolateral compartment, Pig model, Barrier study

1

Introduction The organs and tissues are protected from physical, chemical, and biological damage by the presence of the biological barriers that maintain body homeostasis. The biological barriers also represent important interfaces between organs and the outside environment, such as air or body fluids (e.g., blood and saliva) [1, 2]. The ability to create biological barriers is a prerogative of epithelial and endothelial cells that grow forming a continuous sheet of cells linked together by different junctional complexes [3–6]. The strength and integrity of biological barriers can be determined in vitro by the measurement of the transepithelial/transendothelial electrical resistance (TEER) that reflects the ionic conductance of the paracellular pathway in the epithelial/endothelial monolayer [7]. The TEER measurement is a quantitative technique that is quick, non-invasive,

Mario Baratta (ed.), Epithelial Cell Culture: Methods and Protocols, Methods in Molecular Biology, vol. 2749, https://doi.org/10.1007/978-1-0716-3609-1_14, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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and not related to specific molecule expression and cell physiological status. For these reasons, TEER measurement is widely accepted to be a robust marker for testing the cellular barrier compactness, before studies of chemicals and drug transport across biological barriers. The classical method to measure TEER consists of cells cultured as a monolayer on an insert filter (transwell) generally made of polycarbonate (PC), polyester (PE), or polyethylene terephthalate (PET). The cells that grow on the transwell create an apical (or upper) and basolateral (or lower) compartment, mimicking the natural polarization of epithelial cells. Many different instruments, known as “epithelial voltammeters,” are commercially available to the TEER measurement and consist of two electrodes that are placed one in the upper and the other one in the lower compartment, so theoretically applying direct current (DC), the ohmic resistance can be determined. Anyway, DC can damage cells and the electrodes; therefore, to overcome this problem, the instruments now available apply an alternating current (AC) voltage signal with a square waveform [7, 8]. The most studied in vitro biological barriers that have been characterized by employing the TEER measurement include the intestinal barrier [9, 10], the blood-brain barrier [11, 12], and the pulmonary barrier [13]. Overall, these studies indicated that, although the TEER measurement seems apparently a very simple method, the variables that can affect the results are numerous and must be carefully considered to avoid generating a useless and unpredictable model [7]. The choice of cell model primary cell line vs. immortalized cell line, the choice of optimal cell culture condition on transwell, and the standardization of the procedure of TEER measurement highly influence the development of a predictive model. Therefore, to the best evaluation of the barrier integrity, it’s recommended to associate the TEER measurement to a protocol for the paracellular permeability evaluation [14]. Among in vitro barrier models, the less investigated and still not fully characterized barrier there is the blood-milk barrier, and the results obtained in human model are very different whether primary cell line of mammary epithelial cells or immortalized cell line was used [15, 16]. In vitro animal models to study blood-milk barrier have been proposed [17]. Among the animal models, the most studied is the bovine species, where there is also evident economic interest in implementing research on the blood-milk barrier. However, this animal model has little translational value because of the physiological and metabolic differences mainly due to their polygastric nature. After a thorough review of the literature and physiological characteristics, the pig was indeed chosen by the IMI-European ConcePTION project as the animal model to study the transport of medicines across the mammary epithelial barrier [18, 19]. In this context, we recently published a new efficient method to isolate,

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expand, and culture the mammary epithelial cells from hybrid pig tissues (pMECs) [20]. The present protocol describes the method for the evaluation of the mammary epithelial barrier integrity using pMECs by TEER measurement. To optimize the barrier, studying the TEER measurement protocol is associated with a test for studying the passage of the sodium fluorescein, that is, a hydrophilic paracellular marker.

2

Materials All cell media and solutions must be sterile; therefore, prepare them under a laminar flow cabinet or sterilize them with 0.2 μm filtration. Prepare and store all reagents following the specific instructions; diligently follow all waste disposal regulations concerning biological and chemical waste.

2.1

Cell Culture

1. Laminar flow hood (LAF). 2. Water bath. 3. Optic inverted microscope. 4. CO2 incubator. 5. Manual cell counter chamber or automatic cell counter. 6. T-75 flasks and 24-well plate for primary cell culture (see Note 1). 7. 0.4 μm transparent PET membrane for 24-well plate (see Note 2). 8. Dulbecco’s Phosphate Buffered Saline (DPBS) with MgCl2 and CaCl2. Store at 4 °C. 9. 50 mg/mL insulin stock solution: Add 1 mL of DPBS sterile to 50 mg of lyophilized insulin recombinant human (for cell culture). Store in aliquots at -20 °C to avoid freeze/thaw cycle. 10. 500 μg/mL hydrocortisone stock solution: Add 1 mL absolute ethanol to dissolve 1 mg of lyophilized powder, and then dilute the solution by adding 1 mL of sterile DPBS. Store in aliquots at -20 °C to avoid freeze/thaw cycle. 11. 0.1 mg/mL human epidermal growth factor (hEGF) stock solution: Add 1 mL of sterile DPBS to dissolve 100 μg of lyophilized hEGF, human recombinant (for cell culture). Store in aliquots at -20 °C to avoid freeze/thaw cycle. 12. 100× antibiotic-antimycotic solution: 10,000 units of penicillin, 10 mg streptomycin, and 25 μg amphotericin B per mL, sterile-filtered for cell culture.

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13. Complete cell medium (500 mL): Mix 222,2125 mL Dulbecco’s Modified Eagle Medium with 222,2125 mL of Ham’s Nutrient Mixture F12 medium (DMEM/F12 medium). Add 50 mL of fetal bovine serum heat inactivated (FBS), 50 μL insulin (stock 50 mg/mL), 500 μL hydrocortisone (stock 500 μg/mL), 25 μL human epidermal growth factor (hEGF) (stock 0.1 mg/mL), and 5 mL antibiotic-antimycotic solution (stock 100×). The final composition is 1:1 DMEM-F12 medium +10% FBS + 5 μg/mL insulin +0.5 μg/mL hydrocortisone +5 ng/mL hEGF +1% anti-anti solution. Do not freeze; store in aliquots at +4 °C for 8 weeks. 2.2 TEER and Permeability Flux

1. Millicell ERS-2: epithelial volt-ohm meter or analog instrument. 2. Forceps, scissors, and scalpel must be previously sterilized. 3. Millicell ERS-2: epithelial volt-ohm meter and electrode tips. 4. Fluorescent multiplate reader (excitation wavelength of 485/490 nm and emission wavelength of 524/535 nm). 5. Transport buffer (250 mL): Dissolve 0.60 g HEPES and 0.88 g glucose in 250 mL Hanks’ Balanced Salt Solution (HBSS). Mix and adjust the pH with NaOH at 7.4. Sterilize the solution by 0.2 μm filtration. Store the solution in aliquots at 4 °C for 1 week. 6. 1 mg/mL (2.66 mM) sodium fluorescein solution: Dissolve 50 mg of sodium fluorescein powder in 50 mL of transport buffer. Sterilize the solution by 0.2 μm filtration. Store in aliquots at -20 °C for 1 week.

2.3

Immunostaining

1. 4% paraformaldehyde solution: Work under laboratory fume hood. Add 4 g of paraformaldehyde, reagent grade crystalline, to 50 mL of DPBS. Add 1 mL of 1 M NaOH and stir gently on a heating block at ~60 °C until the paraformaldehyde is dissolved. Add 10 mL of DPBS and allow the mixture to cool to room temperature. Adjust the pH to 7.4 with 1 M HCl (~1 mL), and then adjust the final volume to 100 mL with DPBS. Filter the solution through a 0.20 μm membrane filter to remove any particulate matter. Make the paraformaldehyde solution fresh prior to use. 2. Permeabilization solution 0.5%Triton-X100 (50 mL): Mix 250 μL Triton X-100 with 49.75 mL of DPBS until completely dissolved. Prepare only before the use. 3. Blocking solution 0.5%Triton-X 100, 10% FBS (50 mL): Mix 250 μL Triton X-100 with 5 mL FBS and 44.75 mL. Prepare only before the use. 4. Epifluorescence microscope.

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Methods The present protocol has been developed on primary cultures of mammary epithelial cells (pMECs) isolated from slaughtered hybrid commercial pig [20] (see Note 3). All cell manipulation procedures are conducted under a laminar flow cabinet respecting the good laboratory practice to work safely.

3.1

Cell Culture

1. Thaw the cells (1 × 106) and culture them for one passage in T-75 flask with complete cell medium at 38.5 °C, 5% CO2 (see Notes 4–6). 2. Trypsinize cells from maximally 70% confluent: Pre-warm the complete cell medium, DPBS, 0.25% trypsin-EDTA, and FBS in water bath. Aspirate and eliminate the exhaust culture medium, wash with 10 mL DPBS, rock gently, and aspirate off. Add 7 mL trypsin into a T-75 flask. Transfer in incubator for 7 min; after that, observe the cells under the optic inverted microscope, and if necessary, gently beat the flask. Inhibit the trypsin by adding the same volume of FBS (7 mL) (see Note 7). Transfer the cell suspension in a 15 mL conical tube. Wash the flask with DPBS (10 mL) to collect any remaining cell clusters, and add them to the cell suspension (final volume 24 mL). Centrifuge at 500 × g for 10 min at room temperature, soft stop. Discharge the supernatant by inversion, and resuspend the cell pellet with 3 mL of complete cell medium, and proceed with counting using manual counter chamber or automatic cell counter. 3. Seed the cells (3.3 × 105 cells/cm2) (see Notes 8–11) in the apical compartment on the 0.4 μm transparent PET membrane resuspended in 0.35 mL of complete medium (Fig. 1), and prepare at the same time the blank insert (no cells). Add 0.9 mL of complete medium in the basolateral chamber. 4. Incubate the cells at 38.5 °C, 5% CO2.

3.2 TEER Measurement

1. The day after seeding on transwell, start to measure the TEER. 2. Pre-warm transport buffer and DPBS. 3. Before starting the TEER measurement, sterilize the electrodes by soaking the electrode tips in 70% ethanol for 15 min. Allow to air dry for 15 s, wash the electrodes in sterile distilled water, and dry before the use. 4. Leave the plate 15 min under the laminar flow hood at room temperature before taking the measurements (see Note 12). 5. Measure, firstly, the resistance of the blank insert and then the resistance of inserts with cells (see Note 13).

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Fig. 1 Representative picture of cell seeding on transwell insert; the apical and the basolateral compartments are distinct. This figure was created using BioRender (https://www.biorender.com/) Table 1 Representative table format showing the predisposition of data necessary to calculate TEER Cells pMECs

Passage

Density (n cells/cm2)

Medium

Day

Insert

Rblank

Rcells

8

1 × 10

Transport buffer

1

A1

/

700

5

6. Lift the PET membrane insert helping with sterile forceps, discard at first the medium in basolateral compartment, and then aspirate it from the apical compartment (see Note 14). 7. Add the required volume of transport buffer (see Note 15): First, add 0.5 mL transport buffer in the apical compartment, and then add 1.2 mL transport buffer in the basolateral compartment (see Note 16). 8. Place the shorter electrode into the apical compartment and the other one in the basolateral compartment. The shorter electrode tip must not be in contact with the cell layer, and the longer tip should just touch the bottom of the outer well. Keep the electrode steady and at 90° angle with respect to the cell’s monolayer. Measure the resistance (R) (ohm) two times at opposite sides of the apical chamber, and use the mean value to calculate the TEER by Eq. 1 (see Notes 17 and 18). 9. Predispose a table format (as shown in Table 1) and the scheme of the plate (as in Fig. 2) that contain data necessary to calculate TEER. 10. Calculate the TEER value using Eq. 1:

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Fig. 2 Scheme of the plate to identify each insert

TEER Ω * cm2 = ½R cells ðΩÞ - R blank ðΩÞ] × Marea cm2

ð1Þ

where Rcells is the mean resistance measured in cell inserts, Rblank is the mean resistance measured in the blank inserts, and Marea is the area of the membrane. 11. Wash once the electrodes with DPBS between different well measurements. 12. At the end of experiment, wash and sterilize the electrodes as above reported. 13. Repeat the measurements every day to follow the TEER kinetic profile (Fig. 2). 14. At the end of the TEER measurements, the permeability flux assay begins. 3.3 Permeability Flux Assay

1. Pre-warm the sodium fluorescein solution (see Note 19) and transport buffer. 2. Lift the PET membrane insert helping with sterile forceps, discard at first the transport buffer in basolateral compartment, and then aspirate it from the apical compartment (see Note 20). 3. Add 0.5 mL sodium fluorescein solution in the apical compartment, and add 1.2 mL transport buffer in the basolateral compartment. Incubate for 1 h, by placing the plate in incubator at 38.5 °C and 5% CO2. 4. Lift the PET membrane insert helping with sterile forceps, and aspirate the sodium fluorescein solution from the apical compartment. Aspirate the transport buffer coming from the basolateral compartment, and collect this sample in a microtube (see Note 21). 5. Wash gently: Add 0.5 mL transport buffer in the apical compartment, and then add 1.2 mL transport buffer in the basolateral compartment.

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Table 2 Calibration curve points for the sodium fluorescein study Number

Concentration (μM)

1

13.300

2

6.650

3

3.325

4

1.663

5

0.831

6

0.416

7

0.208

6. Aspirate the transport buffer from the apical and from the basolateral compartment. Add 0.35 mL of complete medium in the apical compartment and 0.9 mL in the basolateral; leave the cells in incubation at 38.5 °C, 5% CO2, to continue kinetic analysis. 7. Start the analysis of the sodium fluorescein unknown samples (see Note 22). 8. Prepare the calibration curve of the sodium fluorescein at known concentrations to determine the concentration of the unknown samples. 9. Starting from the stock solution (2.66 mM), the sodium fluorescein must be diluted 1:100 to obtain the 26.6 μM solution. 10. Make a series of seven dilutions (see the table below) starting from the 26.6 μM (10 μg/mL) solution adding 500 μL of 26.6 μM solution to 500 μL of transport buffer in labeled microtubes (1–7) that match with the concentrations in Table 2. 11. Prepare a blank solution with only transport buffer. 12. Transfer 100 μL of each point of the calibration curve (from point 3 to 7 and blank) in a black 96-well plate, in duplicate, and use the mean value to calculate the sodium fluorescein concentration. 13. Cover the plate with aluminum, shielding it from light until analysis. 14. The samples must be read via fluorescence intensity, using a microplate reader, at the excitation wavelength of 485/490 nm and emission wavelength of 524/535 nm. 15. Predispose a table format (as shown as an example in Table 3) that contains data necessary to calculate the sodium fluorescein transport percentage (Table 3) (see Note 23).

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Table 3 Example showing the predisposition of data collected, necessary to calculate sodium fluorescein transport percentage

Cells

Density (n cells/ Passage cm2) Medium

pMECs 8

1 × 105

Transport buffer

C0 Day Insert (μM)

Csample (μM)

Vb (mL)

Va (mL)

1

5090

1.2

0.5

A1

2600

Fig. 3 Representative graph showing the TEER (blue) and sodium fluorescein transport (purple) profiles calculated in function of time (daily) obtained using primary cell cultures of pMECs

16. Calculate the percentage of sodium fluorescein transport following Eq. 2: Transport ð%Þ =

Csample Vb × 100 × C0 Va

ð2Þ

where Csample is the concentration of sodium fluorescein in the sample, C0 is the concentration of sodium fluorescein in the donor compartment, Vb is the volume of the basal compartment, and Va is the volume of the apical compartment (see Note 24). 17. Analyze data by reporting in a same graph TEER (Ω*cm2) values and sodium fluorescein transport (%) in function of time (Fig. 3).

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3.4 Immunostaining of Key Markers of Tight Junctions

When TEER measure is higher than blank (see Note 17) and sodium fluorescein transport reaches values 0.98, and then calculate the Csample (μM) by reverse formula. Consider whether the sample has been diluted too. Fluorescence intensity ðyÞ = a * Csample ðμMÞ þ b

ð3Þ

24. The monolayer of epithelial cells is compact when the transport percentage of sodium fluorescein is lower than 0.3%. 25. Take care not to damage the cell monolayer during the operation of detaching the membrane from the transwell. 26. Add a drop of mounting medium with DAPI on the imaging slide before placing the membrane. 27. Do not move the membrane on the glass before adding the coverslip.

Acknowledgments This work was financed by the University of Bologna RFO 2022 grant to Chiara Bernardini. References 1. Assimakopoulos SF, Triantos C, Maroulis I, Gogos C (2018) The role of the gut barrier function in health and disease. Gastroenterology Res 11:261 2. Choudhury H, Gorain B, Kesharwani P (2021) Chapter 4: physiology of the biological barriers. In: Kesharwani P, Taurin S, Greish K (eds) Theory and applications of nonparenteral nanomedicines. Academic Press, pp 79–95 3. Anderson JM (2001) Molecular structure of tight junctions and their role in epithelial transport. News Physiol Sci 16:126–130 4. Salama NN, Eddington ND, Fasano A (2006) Tight junction modulation and its relationship

to drug delivery. Adv Drug Deliv Rev 58:15– 28 5. Anderson JM, Van Itallie CM (2009) Physiology and function of the tight junction. Cold Spring Harb Perspect Biol 1:a002584 ´ lvarez-Ferna´ndez I, Blanco6. Garcı´a-Lino AM, A ´ lvarez AI (2019) Paniagua E, Merino G, A Transporters in the mammary glandcontribution to presence of nutrients and drugs into milk. Nutrients 11:2372 7. Srinivasan B, Kolli AR, Esch MB, Abaci HE, Shuler ML, Hickman JJ (2015) TEER measurement techniques for in vitro barrier model systems. J Lab Autom 20:107–126

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8. Benson K, Cramer S, Galla H-J (2013) Impedance-based cell monitoring: barrier properties and beyond. Fluids Barriers CNS 10:5 9. Lea T (2015) Caco-2 cell line. In: Verhoeckx K, Cotter P, Lo´pez-Expo´sito I, Kleiveland C, Lea T, Mackie A, Requena T, Swiatecka D, Wichers H (eds) The impact of food bioactives on health: in vitro and ex vivo models. Springer, Cham 10. Kannapin F, Schmitz T, Hansmann J, Schlegel N, Meir M (2022) Correction to: measurements of transepithelial electrical resistance (TEER) are affected by junctional length in immature epithelial monolayers. Histochem Cell Biol 157:119–119 11. Reichel A, Begley DJ, Abbott NJ (2003) An overview of in vitro techniques for blood-brain barrier studies. Methods Mol Med 89:307– 324 12. Vigh JP et al (2021) Transendothelial electrical resistance measurement across the blood-brain barrier: a critical review of methods. Micromachines (Basel) 12:685 13. Lehmann AD, Daum N, Bur M, Lehr C-M, Gehr P, Rothen-Rutishauser BM (2011) An in vitro triple cell co-culture model with primary cells mimicking the human alveolar epithelial barrier. Eur J Pharm Biopharm 77:398– 406 14. Hubatsch I, Ragnarsson EGE, Artursson P (2007) Determination of drug permeability and prediction of drug absorption in Caco-2monolayers. Nat Protoc 2:2111–2119 15. Kimura S, Morimoto K, Okamoto H, Ueda H, Kobayashi D, Kobayashi J, Morimoto Y (2006) Development of a human mammary epithelial cell culture model for evaluation of drug transfer into milk. Arch Pharm Res 29:424–429 16. Zhang T, Applebee Z, Zou P, Wang Z, Diaz ES, Li Y (2022) An in vitro human mammary

epithelial cell permeability assay to assess drug secretion into breast milk. Int J Pharma: X 4: 100122 17. Nauwelaerts N et al (2021) A comprehensive review on non-clinical methods to study transfer of medication into breast milk – a contribution from the ConcePTION project. Biomed Pharmacother 136:111038 18. Ventrella D, Forni M, Bacci ML, Annaert P (2019) Non-clinical models to determine drug passage into human breast milk. CPD 25:534–548 19. Ventrella D et al (2021) Animal models for in vivo lactation studies: anatomy, physiology and Milk compositions in the most used non-clinical species: a contribution from the ConcePTION project. Animals 11:714 20. Bernardini C et al (2021) Development of a pig mammary epithelial cell culture model as a non-clinical tool for studying epithelial barrier – a contribution from the IMI-ConcePTION project. Animals 11:2012 21. Kumura H, Tanaka A, Abo Y, Yui S, Shimazaki K, Kobayashi E, Sayama K (2001) Primary culture of porcine mammary epithelial cells as a model system for evaluation of milk protein expression. Biosci Biotechnol Biochem 65:2098–2101 22. Zheng Y-M, He X-Y (2010) Characteristics and EGFP expression of porcine mammary gland epithelial cells. Res Vet Sci 89:383–390 23. Sun YL, Lin CS, Chou YC (2006) Establishment and characterization of a spontaneously immortalized porcine mammary epithelial cell line. Cell Biol Int 30:970–976 24. Dahanayaka S, Rezaei R, Porter WW, Johnson GA, Burghardt RC, Bazer FW, Hou YQ, Wu ZL, Wu G (2015) Technical note: isolation and characterization of porcine mammary epithelial cells1,2. J Anim Sci 93:5186–5193

Chapter 15 Bovine Skeletal Muscle Satellite Cells: Isolation, Growth, and Differentiation Silvia Miretti, Isabella Manenti, Paola Toschi, Elisabetta Macchi, Eugenio Martignani, Paolo Accornero, and Mario Baratta Abstract Skeletal muscle in cattle occupies a large part of the animal’s body mass and develops into an important source of nutrients for human nutrition. Recently, the attention on bovine myogenic cells is increased to develop strategies of cultured in vitro meat as an alternative food source, more sustainable, ethical, and healthy than traditional meat production. At present, investigating the proliferation and differentiation of bovine skeletal muscle myogenic cells in vitro maintains its importance in the study of the mechanisms underlying the physiological and pathological events affecting the skeletal muscle, but it is of particular interest in animal husbandry and the food industry fields. In cell-based biological research, cell lines are one of the favored experimental tools because a population of cells could proliferate indefinitely in vitro under different stimuli, but they are limited to addressing the relevant biological properties of a cell population. On the other hand, primary cells from normal animal tissues undergo a limited number of divisions in vitro before they enter senescence but preserve their original characteristics and functions, and researchers can acquire the opportunity to study the individual donors and not just cells. In this chapter, we provide a basic protocol to isolate satellite cells from the skeletal muscle of cattle to obtain a good number of myogenic cells that can grow in in vitro conditions and undergo multiple rounds of cell division (myoblasts) before entering differentiation (myotubes). Furthermore, the robust expansion of these cells leads to the possibility to investigate physiological events or disorders related to the skeletal muscle tissue. Key words Skeletal muscle, Cattle, Satellite cells, Myoblasts, Growth, Differentiation

1

Introduction The muscle is primarily composed of a fixed number of multinucleated myofibers defined during the late gestation period [1]. This number remains constant during the postnatal life of animals, but a population of quiescent and self-renewal muscle cells, named satellite cells, could be activated and participate in muscle repair, muscle hyperplasia, and hypertrophy processes [2, 3].

Mario Baratta (ed.), Epithelial Cell Culture: Methods and Protocols, Methods in Molecular Biology, vol. 2749, https://doi.org/10.1007/978-1-0716-3609-1_15, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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Fig. 1 Cell culture of isolated bovine skeletal muscle primary myoblasts on a gelatine-coating plate during growth and differentiation. (a-d) The cultures on days 3 and 5 in growth medium (GM) and on days 8 and 10 in differentiation medium (DM), respectively. The days are calculated starting from the day of isolation and plating (day 0). (a) Round cells observed during the early culture days are proliferating myoblasts. (b) At 5 days of plating in GM, the cells start to acquire a spindle morphology, and when reaching 70–80% of confluence, the GM could be changed with the DM. (c) On day 8, some myotube is present, and on day 10 (d) the fusion of myotubes in syncytial structures is observed. Images in phase contrast were taken with a 10× magnification

Because of the difficulties to study muscle cell proliferation and differentiation in vivo, efficient isolation of satellite cells and their maintenance as myoblasts in vitro are fundamental to creating a reliable model to investigate molecular changes along growth and differentiation processes useful to explore physiological events such as hypertrophy or mechanisms underlying muscle disorders such as atrophy [4, 5]. This chapter provides a protocol to obtain a bulk population of satellite cells (Fig. 1) with a purity between 85 and 95% based on immunofluorescent staining for paired box transcription factor 7 (PAX7) and myogenic factor 5 (MyF5). Founded on the human and rodent antigens reported in the literature, the characterization through fluorescence-activated cell sorting (FACS) and magnetic cell isolation technology was initially attempted, but no antibodies

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with high performances in cattle were identified to this scope (e.g., mouse-antihuman CD56 not able to discriminate myoblasts to fibroblasts) [6, 7]. The satellite cells are very sensitive to stimuli, and the methodology employed to isolate and culture these cells can induce the beginning of the differentiation process with fusion of myoblasts into multinucleated myotubes. We have optimized our protocol with a lower number of steps and a well-defined timing to not incur unintentional activation of myoblastic cells. 1.1 Breed, Age of Animals, and Selection of Muscle

The setup of this protocol was born of the need to define better the genetic signature of the Piedmontese cattle breed by investigating molecules, such as microRNAs, that could be involved in the peculiar hypertrophic phenotype that characterizes these animals [8, 9]. To obtain satellite cells, the skeletal muscles of young adult Piedmontese beef cattle (15–17 months old) were selected [9], but the Friesian-Holstein breed has also been used in our studies following the same procedures [10]. The isolation was performed starting from three different muscles: sternocleidomastoid muscle, longissimus thoracis, and diaphragm. The easier muscle to collect is the sternocleidomastoid. This is due to its anatomical position and slaughter procedures. Sampling from the longissimus thoracis muscle requires obtaining the sample in a slaughterhouse with a cutting plant where boning and cutting of meat is performed. Isolation from the diaphragm muscle, initially selected for the low economic value of this anatomical section once it became meat, was abandoned because much more incurrent of the contamination of primary cultures was probably due to muscle anatomical position and related frequent laceration of the gastrointestinal viscera and its thickness. Figure 1 shows representative images of cattle myogenic cells isolated from the longissimus dorsi of a Piedmontese young adult bovine.

1.2 Myogenic Cell Characterization

The process of growth and development of skeletal muscle is complex. It requires proliferation, differentiation, and fusion of myoblasts into muscle fibers and involves a large number of changes in gene expression. Pax7 has been identified as a gene responsible for the condition of progenitor cells to the satellite cell lineage [11]. It is expressed in cultured satellite cells, and the temporal expression of the Pax7 mRNA in proliferating myoblasts has been defined, and a rapid downregulation of Pax7 transcripts upon myogenic differentiation has been described [11]. Pax7 is not expressed at detectable levels in a variety of non-muscle cell lines, and in general, quiescent satellite cells do not have any detectable levels of myogenic regulatory factors (MRFs) [12].

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Fig. 2 Immunostaining myogenic cells and differentiated myoblasts characterization. (a-c) Representative 20× phase-contrast and dark-field images of (2A) MyF5, (2B) Myogenin (F5D DSHB), and (2C) Myosin Heavy Chain (MF20 DSHB) immunostained cells after 3 days in DM. (1A-1B-1C) Cell nuclei of, respectively, phase-contrast and dark-field images are stained blue by 4′,6-diamidino-2-phenylindole (DAPI). Scale bar = 250 μm

Satellite cells have to exit their normal quiescent state to start proliferating. Their activation induces several rounds of proliferation, and in this stage, they are often referred to as myogenic precursor cells or adult myoblasts. The majority of the satellite cells differentiate and fuse to form myotubes. At the molecular level, activation of myoblasts is characterized by the rapid upregulation of MRFs and microRNAs (miRNAs). After the myoblasts’ proliferation phase, expression of Myogenin and MRF4 (MRF members) is upregulated in cells beginning their terminal differentiation program. The differentiation phase could be considered complete with the activation of muscle-specific proteins, such as MyHC (Fig. 2), and the myoblasts fuse with each other to form syncytial muscle myotubes (Fig. 2).

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miRNAs are small noncoding RNA molecules, posttranscriptionally downregulating target genes, in general ubiquitously expressed in all types of cells, but some are defined as tissue-specific when the expression is 20-fold higher than the mean expression in other tissues. MicroRNAs that are exclusively or preferentially expressed in striated muscle are called myomiRs [13]. MyomiRs play key roles in myogenesis, and miR-1 and miR-133 and miR-206 are critical regulators of muscle cell proliferation and differentiation [14, 15]. To characterize bovine myoblasts, we used immunofluorescence to define the amount of progenitor satellite cells and their transition in proliferation and differentiation (Fig. 2). Quantitative real-time PCR (qRT-PCR) was used to measure MRF and final differentiation genes (MyoG, MRF4, MEF2C, MyH1, and Myomaker) and myomiRs (miR-1, miR-133, and miR-206) expression with high throughput, accuracy, sensitivity, and reproducibility along cultured time [10].

2

Materials Store all reagents at room temperature, unless indicated otherwise. Warm at room temperature all medium before adding them to the cells. 1. PBS (phosphate-buffered saline) buffer: 800 mL of distilled water, 8 g of 0.137 M sodium chloride (NaCl), 0.2 g of 0.0027 M potassium chloride (KCl), 1.44 g of 0.01 M sodium phosphate dibasic (Na2HPO4), and 0.245 g of 0.0018 M potassium phosphate monobasic (KH2PO4). Adjust solution pH ≈ 7.4 with hydrochloric acid (HCl) and/or sodium hydroxide (NaOH). Add distilled water until the volume is 1 L and autoclave. 2. Pronase: 10 mL of sterile PBS and 100 mg of pronase dehydrated. This procedure must be done in ice. Filter with a 0.22 μm filter and then make aliquots of 500 μL in 1.5 mL Eppendorf. Pronase aliquots must be stored at -20 °C. 3. DMEM (Dulbecco’s Modified Eagle Medium; 1 g/L D-glucose, L-glutamine; 25 mM HEPES, pyruvate): supplemented with 5 mL of 10,000 U/mL penicillin, 10,000 μg/mL streptomycin, 5 mL of 100× 200 mM L-glutamine. When the term DMEM is used from here, it refers to DMEM with antibiotics and glutamine. 4. FBS (fetal bovine serum): original bottles stored at -20 °C. Once thawed and aliquoted (50 mL), store at -20 °C. 5. HS (horse serum): original bottles stored at -20 °C. Once thawed and aliquoted (10 mL), store at -20 °C.

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6. GM (growth medium): DMEM 10% HS, 20% FBS, 100× PSA. In a 50 mL falcon, mix 5 mL of HS (horse serum), 10 mL of FBS (fetal bovine serum), and 35 mL of DMEM. Store at +4 °C. 7. DM (differentiation medium): DMEM 2% HS. In a 50 mL falcon, mix 1 mL of HS (horse serum) with 49 mL of DMEM. Store at +4 °C. 8. Gelatine 2%: 2 g of gelatine powder in 100 mL of ultrapure water and then autoclave. Gelatine must be aliquoted and stored at +4 °C for many months. 9. Tris buffered saline pH 7.6: 800 mL of distilled water, 15.76 g of 0.1 M TRIS hydrochloride (Tris/HCl), and 8 g of sodium chloride (NaCl). Adjust solution pH ≈ 7.6 with hydrochloric acid (HCl) and/or sodium hydroxide (NaOH). Add distilled water until the volume is 1 L and store at +4 °C. 10. Antibody dilution buffer: 100 mL of Tris-buffered saline, 1 g of bovine serum albumin (BSA), and 100 mg of sodium azide (NaN3). Adjust pH ≈ 7.4 with hydrochloric acid (HCl) and/or sodium hydroxide (NaOH) and store at +4 °C.

3

Methods

3.1 Muscle Collection

The muscle sample is obtained at the slaughterhouse, immediately at the end of the slaughter chain. 1. Collect muscle samples with a sterile knife from the selected anatomical muscle through an extended incision. At this level, it is impossible to avoid the tissues that are not muscles; fat, vessels, and aponeurosis are present. The good amount of the sample to collect is approximately 500 g of weight. 2. Lay muscle sample in a sterile glass beaker (2 L of volume capacity) and cover it with ethanol 70%. Take the sample into the cell culture lab within 1 h.

3.2 Coating Cell Culture Dishes

All the next steps are under the biological hood, in a sterile condition. 1. Place an aliquot of gelatine stored at +4 °C in a water bath at 37 °C until liquefied. 2. Add sufficient mL of gelatine to cover the bottom of the dish, and swirl gently to avoid some areas being uncoated. 3. Leave the dish closed under the hood for at least 30 min. 4. Recover the gelatine and leave plates opened under the hood to dry for at least 30 minutes. Gelatine can be reused ten times without a decrease in cell adhesion and growth.

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All the next steps are under the biological hood, in a sterile condition. 1. Prewarm 50 mL of DMEM to 37 °C. 2. Transfer the 500gr muscle piece to a second sterile beaker (2 L). 3. Wash the muscle in PBS three times. 4. Transfer the muscle to a Petri dish of 15 cm in diameter. 5. Remove a layer of muscle 1 cm deep, and start to collect only muscle fragments from the core of the muscle piece avoiding blood vessels, fat, and connective tissue. 6. Transfer the dissected fragments to a new 15 cm Petri dish to remove the remaining connective tissue with sterile straight and curved fine-point forceps (see Note 1). 7. Finely chop the muscle and collect the debris of fibers in a 50 mL conical tube with 9.5 mL DMEM NO FBS, 100x PSA, and 500 μL pronase. 8. Shake in a water bath at 37 °C for 1 h at 290 rpm, pipetting every 20 minutes with a 25 mL pipette at least 20 times. The muscle sample mixture appears as a fine slurry. 9. Add 10 mL of DMEM 10% HS. 10. Centrifuge at 400 g for 5 min at room temperature. 11. With a 5 mL pipette, gently remove the supernatant without touching the pellet. 12. Resuspend the pellet in 10 mL of DMEM 10% HS, pipette 20 times, and let it settle for 10 min. 13. Collect the supernatant and transfer it to a new 50 mL conical tube. 14. Resuspend the pellet in 10 mL of new DMEM 10% HS, pipette 20 times, and let it settle for 10 min. 15. Collect the supernatant and combine it with the previous collection. 16. Resuspend the pellet in 10 mL of new DMEM 10% HS, pipette 20 times, and let it settle for 10 min. 17. Collect the supernatant and combine it with the previous two collections, and let it settle for 10 min. 18. Filter the supernatant with 40 μm cell strainer (more filters will be needed) in a new 50 mL conical tube. 19. Centrifuge at 1000 g for 10 min. 20. Gently remove the supernatant (see Note 2). 21. Resuspend the pellet in GM and then plate it on dishes with gelatine (see Note 3).

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22. In a 60 mm dish, add 3 mL of GM and culture the cells undisturbed in the incubator for 3 days. 23. Rinse the cultures five times with 5 mL of PBS before adding 3 mL of fresh GM. This helps to remove debris that in the primary cultures can be easily mistaken for contamination. This first time rinse the cultures very gently to avoid cell detachment. 24. Repeat the washing before replacement of the culture medium with fresh GM every day until the cells reach 70–80% of confluence. 25. After washing with PBS, add 3 mL of differentiation medium (DM) to cultures to start differentiating cells. 26. Rinse the cultures with 3 mL of fresh DM every day for 3 days. 3.4 Cell Culture Fixation and Immunostaining

1. Remove the culture medium, and add 5 mL of acetone/methanol solution 1:1 to completely cover the cells (see Note 4). 2. Remove the solution after 15–30 s, and quickly and gently wash twice the cells with Tris-buffered saline pH 7.6 at room temperature. 3. Discard and replace Tris-buffered saline. The cells can be kept in buffer at +4 °C for 1–2 weeks before staining them. 4. Remove the Tris buffer and quickly make a circle with PAP pen around the cells of interest to delimit the space. 5. Preblock the cells with diluted serum (10% in Tris-buffered saline) from the same secondary antibody species for at least 30 min. 6. Quickly wash with Tris-buffered saline. 7. Incubate with the minimum amount to cover the cells of the primary antibody diluted in antibody dilution buffer for 60 min at room temperature. 8. Wash the cells with Tris-buffered saline for 5 min. Repeat three times. 9. Incubate with the minimum amount to cover the cells of secondary antibody diluted in antibody dilution buffer for 60 min at room temperature (in the dark for immunofluorescence). 10. Wash the cells with Tris-buffered saline for 5 min. Repeat three times. 11. If the secondary antibodies are conjugated with an enzyme, add a suitable amount of a specific substrate, and incubate for 5–10 min checking frequently the staining density. 12. Counterstain the nuclei with hematoxylin (for immunohistochemical staining) for 1–2 min or with DAPI (for immunofluorescence) diluted in Tris-buffered saline for 10–15 min.

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13. Wash the cells with Tris-buffered saline. 14. Store the cells in Tris-buffered saline at +4 °C. The staining will last for several days.

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Notes Myogenic Cell Isolation 1. Surgical tools for harvesting pieces of muscle: straight and curve forceps with point-fine tips, tissue forceps, and Adson dissecting forceps serrated without teeth. For better results during cell isolation in terms of the survival rate of cells, we suggest working in pairs: two operators under a double sterile hood from Steps 5 to 7 (Subheading 3.3). One operator is dedicated to selecting pieces of muscle from the muscle core, and the second one processes the selected pieces removing the ulterior connective tissue that could be still present and fine-chopping the sample. In this way, excessive dryness of the fibers is avoided. 2. If cells are not plated but frozen whole, an aliquot of the supernatant (Subheading 3.3; Step 20) could be plated in a sterile no gelatine-coating 60 mm dish and stored in the incubator to check possible contamination. 3. At the end of the isolation procedure (Subheading 3.3; Step 21), usually, the pellet is used for one part to satellite growth plate it in a dish with GM, and the other four parts are resuspended in 4 mL of freezing medium (90% FBS and 10% DMSO (dimethyl sulfoxide)) to store them at 80°. Cell Culture Fixation 4. Acetone/methanol solution could affect the morphology of the cells or induce their detachment. Proceed very gently, working on the wall of the plate to add the solution (Subheading 3.4; Step 1). Check the cells after the removal of the solution.

References 1. Russell RG, Oteruelo FT (1981) An ultrastructural study of the differentiation of skeletal muscle in the bovine fetus. Anat Embryol (Berl) 162:403–417. https://doi.org/10. 1007/BF00301866 2. Hill M, Wernig A, Goldspink G (2003) Muscle satellite (stem) cell activation during local tissue injury and repair. J Anat 203:89–99.

https://doi.org/10.1046/J.1469-7580. 2003.00195.X 3. Gonzalez ML, Busse NI, Waits CM, Johnson SE (2020) Satellite cells and their regulation in livestock. J Anim Sci 98. https://doi.org/10. 1093/JAS/SKAA081 4. Grabowska I, Szeliga A, Moraczewski J et al (2011) Comparison of satellite cell-derived

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myoblasts and C2C12 differentiation in twoand three-dimensional cultures: changes in adhesion protein expression. Cell Biol Int 35: 1 2 5 – 1 3 3 . h t t p s : // d o i . o r g / 1 0 . 1 0 4 2 / CBI20090335 5. Baquero-Perez B, Kuchipudi SV, Nelli RK, Chang KC (2012) A simplified but robust method for the isolation of avian and mammalian muscle satellite cells. BMC Cell Biol 13:16. https://doi.org/10.1186/1471-2121-13-16 6. Agley CC, Rowlerson AM, Velloso CP et al (2015) Isolation and quantitative immunocytochemical characterization of primary myogenic cells and fibroblasts from human skeletal muscle. J Vis Exp. https://doi.org/10.3791/ 52049 7. Gromova A, Tierney M, Sacco A (2015) FACS-based satellite cell isolation from mouse hind limb muscles. Bio-Protoc 5. https://doi. org/10.21769/bioprotoc.1558 8. Miretti S, Martignani E, Accornero P, Baratta M (2013) Functional effect of mir-27b on myostatin expression: a relationship in piedmontese cattle with double-muscled phenotype. BMC Genomics 14. https://doi.org/ 10.1186/1471-2164-14-194 9. Tewari RS, Ala U, Accornero P et al (2021) Circulating skeletal muscle related microRNAs profile in Piedmontese cattle during different age. Sci Rep 11. https://doi.org/10.1038/ s41598-021-95137-w

10. Miretti S, Volpe MG, Martignani E et al (2017) Temporal correlation between differentiation factor expression and microRNAs in Holstein bovine skeletal muscle. Animal 11. https://doi. org/10.1017/S1751731116001488 11. Seale P, Sabourin LA, Girgis-Gabardo A et al (2000) Pax7 is required for the specification of myogenic satellite cells. Cell 102:777–786. https://doi.org/10.1016/S0092-8674(00) 00066-0 12. Charge´ SBP, Rudnicki MA (2004) Cellular and molecular regulation of muscle regeneration. Physiol Rev 84:209–238. https://doi.org/10. 1152/physrev.00019.2003 13. McCarthy JJ (2008) MicroRNA-206: the skeletal muscle-specific myomiR. Biochim Biophys Acta – Gene Regul Mech 1779:682–691. https://doi.org/10.1016/j.bbagrm.2008. 03.001 14. Chen JF, Mandel EM, Thomson JM et al (2006) The role of microRNA-1 and microRNA-133 in skeletal muscle proliferation and differentiation. Nat Genet 38:228–233. https://doi.org/10.1038/ng1725 15. Diniz GP, Wang DZ (2016) Regulation of skeletal muscle by micro RNAs. Compr Physiol 6:1279–1294. https://doi.org/10.1002/ cphy.c150041

Chapter 16 Equine Induced Pluripotent Stem Cell Culture Julia Falk and F. Xavier Donadeu Abstract Groundbreaking work by Takahashi and Yamanaka in 2006 demonstrated that non-embryonic cells can be reprogrammed into pluripotent stem cells (PSCs) by forcing the expression of a defined set of transcription factors in culture, thus overcoming ethical concerns linked to embryonic stem cells. Induced PSCs have since revolutionized biomedical research, holding tremendous potential also in other areas such as livestock production and wildlife conservation. iPSCs exhibit broad accessibility, having been derived from a multitude of cell types and species. Apart from humans, iPSCs hold particular medical promise in the horse. The potential of iPSCs has been shown in a variety of biomedical contexts in the horse. However, progress in generating therapeutically useful equine iPSCs has lagged behind that reported in humans, with the generation of footprint-free iPSCs using non-integrative reprogramming approaches having proven particularly challenging. A greater understanding of the underlying molecular pathways and essential factors required for the generation and maintenance of equine iPSCs and their differentiation into relevant lineages will be critical for realizing their significant potential in veterinary regenerative medicine. This article outlines up-to-date protocols for the successful culture of equine iPSC, including colony selection, expansion, and adaptation to feeder-free conditions. Key words Equine, iPSCs, Cell reprogramming, Feeder-free culture

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Introduction Induced pluripotent stem cells (iPSCs) have significantly transformed biomedicine, owing to their immense potential in disease modeling, drug screening, and cell-based therapies, in addition to highly exciting nonmedical applications such as wildlife conservation [1, 2]. Back in 2006, groundbreaking work by Takahashi and Yamanaka demonstrated that by introducing a combination of four transcription factors (OCT4, SOX2, c-MYC, and Klf4, referred to as OSKM), non-embryonic cells could be reprogrammed to a developmental state akin to embryonic stem cells (ESCs). As such, iPSCs are devoid of the ethical concerns associated with the use of embryos [3]. Remarkably, iPSCs have been derived from most cell types, including, for example, urine-derived cells [4], and

Mario Baratta (ed.), Epithelial Cell Culture: Methods and Protocols, Methods in Molecular Biology, vol. 2749, https://doi.org/10.1007/978-1-0716-3609-1_16, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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from a multitude of species, making them easily accessible for research purposes [1]. Moreover, through their large developmental plasticity and ability to differentiate into a variety of cell lineages, iPSCs provide powerful tools for a wide array of applications in biomedical research and therapy [1]. In addition to human medicine, iPSC technology holds significant promise in veterinary species, specifically in horses, in which stem cell therapies are already well established and could considerably benefit, in terms of improved efficacy, from the introduction of iPSCs [5]. Non-integrative or excisable reprogramming methods including Sendai virus, episomal vectors, RNA-based systems, or PiggyBac expression systems provide a feasible route to the therapeutic use of iPSCs and are now routinely employed to reprogram human cells [6–8]. However, the use of these approaches with horse cells has met with little success [9, 10], leading to the predominant use of integrating viral systems to generate equine iPSC lines [5]. Moreover, advancements in iPSC technology have led to the elimination of xeno- and animal-dependent cultures in favor of feeder-free conditions. This involves substituting feeders with a suitable coating matrix and using serum replacement or smallmolecule cocktails in the place of fetal bovine serum (FBS) [11]. Thus, numerous clinical-grade products are available to support growth of iPSCs from humans and rodents, but the culture of iPSCs from domestic species still relies heavily on the use of feeders [5, 12]. Yet, transitioning to feeder-free iPSCs is imperative for any future therapeutic applications for horses. Considering how slow the field of equine iPSCs has advanced compared to human and mouse, gaining a better understanding of the molecular pathways and critical factors involved in the generation and maintenance of equine iPSCs will be crucial. The protocols in this article have been successfully used over the years in our laboratory to grow equine iPSCs of different origins and generated using different approaches. Hopefully their use will aid future advancements in realizing the significant potential of these cells in veterinary regenerative medicine.

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Materials

2.1 Cell Culture Equipment

1. 70% ethanol for surface sterilization. 2. Class 2 biological safety hoods. 3. Water bath set at 37 °C. 4. Inverted microscope. 5. Neubauer hemocytometer (or other cell counting device). 6. Incubator, suitable for cell culture (set to 37 °C, 21% O2, 5% CO2, 95% relative humidity).

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7. Centrifuges (suitable for small tubes and 15–50 mL tubes). 8. Sterile cell culture plastic pipettes and pipette tips. 9. Sterile centrifuge tubes in various sizes. 10. Different size multi well tissue culture plates and flasks. 11. GammaCell 1000 irradiator. 12. Syringe with 18G × 1 1/2″ (1.2 × 40 mm) needle. 2.2 Cell Culture Reagents

1. Phosphate-buffered saline (PBS). 2. Mouse embryonic fibroblast (MEF) media: DMEM, high glucose (41965039, Thermo Fisher), 10%FBS, 0.1 mM MEM nonessential amino acids (NEAA), 2 mM L-glutamine, and penicillin-streptomycin (stable for up to 4 weeks at 4 °C). 3. Equine iPSC media: DMEM, high glucose (41965039, Thermo Fisher) containing 20% FBS or KnockOut™ Serum Replacement (10828028, Thermo Fisher), 1× GlutaMAX™ (35,050,061, Thermo Fisher) 0.1 mM β-mercaptoethanol, 0.1 mM MEM nonessential amino acids, and 1% penicillinstreptomycin supplemented with 10 ng/mL human bFGF and 1000 U/mL human LIF (should be prepared before use and stable for up to 7 days at 4 °C). 4. Equine embryonic fibroblast (EEF) media used to prepare conditioned media (CM): KnockOut™ DMEM (10,829,018, Thermo Fisher) containing 20% KnockOut™ Serum Replacement (10828028, Thermo Fisher), 1x GlutaMAX™ (35,050,061, Thermo Fisher), 0.1 mM β-mercaptoethanol, 0.1 mM MEM nonessential amino acids, and 1% penicillinstreptomycin (stable for up to 4 weeks at 4 °C). 5. Trypsin-EDTA (0.05%), phenol red (25300054, Thermo Fisher). 6. StemPro™ Accutase™ Cell Dissociation Reagent (A1110501, Thermo Fisher). 7. Corning® Matrigel® Matrix: Before use, thaw stock solution at 4 °C to avoid gel formation and dilute as appropriate in cold DMEM. Then coat tissue culture plates and flasks overnight at 4 °C or at 37 °C for 1 h, and use immediately or store sealed at 4 °C for up to a week. 8. Gelatin: Before use, dilute 2% gelatin solution (G1393, SigmaAldrich) in PBS to 0.1% working solution. Cover tissue culture plastic surfaces with working solution and incubate for 15 min at 37 °C, use immediately, or store sealed at 4 °C for up to a week. 9. Freezing media: 10% (v/v) DMSO (cell culture grade), 90% (v/v) FBS.

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Cell Stocks

1. Equine iPSCs. 2. MEFs. 3. EEFs.

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Methods

3.1 Preparation of MEF Feeder Cells

This protocol describes the preparation of irradiated MEF feeder cells for culture of equine iPSCs (see Note 1). 1. Use low passage MEFs grown in suitable culture vessel with MEF media. 2. When ready for irradiation, detach cells by incubating for 2–8 min with trypsin at 37 °C, then spin at 300 × g for 5 min, and resuspend in fresh pre-warmed media. Count cells in hemacytometer and replate in suitable vessel at 5.7 × 103 cells/cm2 to expand further, or proceed to irradiation as described below. 3. For irradiation, add desired cell number to 50 mL falcon in culture media, and expose to 12.5 Gray in gamma irradiator. 4. Plate cells immediately (see Step 7) or freeze down in freezing media for later use (see Note 2). 5. One day before plating iPSC culture, coat wells in a culture vessel of desired size with 0.1% gelatin solution, and incubate for 15 min at 37 °C. Use immediately or store sealed at 4 °C for up to 7 days. 6. Quickly thaw stock of irradiated MEFs by placing in 37 °C water bath, and then dilute in MEF culture media. Centrifuge cells 300 × g for 5 min at RT. Discard supernatant and resuspend in fresh MEF media. 7. Seed MEFs in gelatin coated surfaces (2 × 106 cells per six-well plate; see Note 3), and culture overnight to allow cell attachment before use for iPSC culture.

3.2 Picking, Expansion, and Culture of Equine iPSC Colonies on Feeder Cells

Equine iPSCs can be generated using different reprogramming approaches, and relevant protocols have been published [9, 13, 14]. The present is a protocol for establishing and maintaining iPSC lines in feeder culture once bona fide colonies have been generated using one of the approaches above and describes the selection of individual iPSC colonies and their subsequent expansion in culture. 1. Individual colonies should be selected for clonal expansion based on assessment by daily microscope inspection. Colonies selected for picking should show clear growth and appear

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Fig. 1 Representative bright field image of bona fide equine iPSCs grown on MEF feeders and showing compact colonies with defined edges. Colonies indicated by arrows displayed sustained growth over 2–5 days and were ready for picking

compact with well-defined edges. Colonies observed to increase in size for 2–4 days and devoid of a dark area in the center should be picked (Fig. 1). 2. One day before picking, prepare MEF feeders in 96- or alternatively, 24-well plates (see Subheading 3.1). 3. It is easiest to pick the colonies under a microscope or stereomicroscope, which is integrated into a laminar flow hood. Alternatively, a microscope can be brought into the hood while leaving the glass shield fully open and taking care to work as much as possible under aseptic conditions (e.g., avoid external airflow from people walking past, and proceed as quick as possible once culture dish is open). 4. Position the colony directly under the microscope, and use a 20uL pipette to carefully pick the colony by gentle aspiration. Then transfer onto well containing feeders in equine iPSC media, and identify the colony under microscope to confirm the transfer has been successful. If picking proves difficult, try using a syringe to cut the colony out before aspirating. 5. Repeat as above for other colonies as required, and place each in a separate well. 6. Culture picked colonies overnight and then confirm each has attached to feeder layer. Colonies should start proliferating 2–4 days after picking. If colony shows no sign of growth after 7 days of picking, they can be discarded. Once it has been growing for 3–5 days, and as long as the center is not

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turning dark or there are no signs of differentiation, a colony should be split for expansion into a larger vessel (24- or 12-well size) with fresh MEF feeders. Splitting can be done by mechanical disruption (see Subheading 3.2, step 7) or using Accutase (see Subheading 3.2, step 8). 7. To disrupt a colony mechanically, use a microscope set up as described in Subheading 3.2, step 3. Put the colony in focus under the microscope and use a syringe to carefully cut it into four to eight pieces. Carefully aspirate the pieces with a 20 or 100 uL pipette, and transfer into a well with fresh feeders. If multiple colonies in a well are disrupted, then media in the well should be aspirated in its entirety and spun for 4 min 200 × g, before carefully resuspending in fresh culture media. 8. To dissociate colonies with Accutase, carefully aspirate the media and wash with PBS. Add enough Accutase to cover the entire surface of the well. Place in incubator for 2–8 min and observe cells under microscope. When the outside of the colony starts to lift (cells will seem brighter) and colony becomes less compact, add equine iPSC media to stop reaction. Use 1 mL pipette to carefully scrape and disrupt iPSCs, before aspirating. Do not pipette cells up and down more than twice, to avoid disruption into single cells. Spin down cells for 4 min 200 × g, and then carefully resuspend in fresh culture media and plate on new feeders. 9. After splitting cells into larger vessel as described above, check daily to ensure colonies keep their compact morphology with clear edges. If cells become confluent, use Accutase as above to split into larger vessel (six-well plate). 10. iPSC cells can be maintained from then on in six-well plates using equine iPSC media (or other suitable media that has been previously optimized). Typically cells need to be split every 3–5 days at a ratio of 1:4–1:10 depending on the culture (see Note 4). 3.3 Growth of Equine iPSCs in Feeder-Free Culture

Feeder-free culture greatly facilitates work with iPSCs, and it is a requirement for many downstream applications, whether involving use as in vitro models or in prospective therapeutic applications. As reprogramming of equine cells is simpler using feeder layers, feederfree cultures usually result from adaptation of iPSC lines generated on feeders [13]. However, adaptation to feeder-free conditions can be challenging. Choosing the right coating substrate and culture media is crucial, and different combinations may need to be tested in each particular instance. The following protocol uses Matrigel and EEF CM. It was originally used to adapt iPSCs generated through reprogramming of equine keratinocyte with retroviral vectors and has later been successfully used with other equine iPSC lines in our laboratory [13].

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Fig. 2 Bright field images showing (a) equine iPSC colonies (indicated by arrows) which failed to adapt to feeder-free conditions as shown by distinct morphological changes and halted growth and (b) iPSC culture which adapted to feeder-free conditions successfully, as evidenced by highly proliferative, compact colonies with well-defined edges

1. Harvest CM from confluent layers of mitotically inactivated EEFs (see Note 5) in T175 flasks at 37 °C and 5% CO2. Do this by collecting and replacing EEF media daily from each flask for a total of 5 days. Stock store collected media at -20C until use. Discard cells after the 5 days. Before use, supplement the conditioned media with 10 ng/mL human bFGF and 1000 U/mL human LIF (keep at 4 °C for up to 7 days). 2. Start transition to feeder-free culture by gradually reducing feeder density during iPSC passaging. When preparing feeders before passaging iPSCs, plate 75% of the usual feeder number (e.g., if normally using 2 × 106 per six-well plate, then reduce to 1.5 × 106 per six-well plate). Split iPSC culture as usual using prepared feeders. 3. Monitor morphology of passaged iPSCs and use same feeder density for at least one more passage. After transition to reduced feeders, some iPSC colonies may undergo morphology changes indicative of loss of pluripotency (Fig. 2). In that case, try to remove abnormal colonies mechanically, and keep the rest of the culture growing. If on the contrary only a few colonies maintain typical morphology, then try to pick those and plate them in smaller vessel (see Subheading 3.2, step 1), and discard the rest. Once colonies are growing well, move on to next step. 4. As next step, reduce feeder density to 50% of the original following the same principle as in Subheading 3.4, step 2. In addition, when plating cells on reduced feeder density, replace half of equine iPSC media with CM collected in Subheading 3.4, step 1. Follow same procedures as in Subheading 3.4, step 3.

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5. Once colonies are growing well in 50% feeder density, then prepare a Matrigel coated culture dish and seed feeders at 25% of the original density. Passage iPSCs onto reduced feeders and culture in 100% CM. 6. Follow same procedures as in Subheading 3.4, step 3 (see Note 6). 7. Once colonies are growing well on a minimal amount of feeders, then passage onto Matrigel only and 100% CM. 8. Follow same procedures as in Subheading 3.4, step 3. 9. Continue to expand cells in feeder-free conditions as in Subheading 3.4, step 7, and confirm cells maintain normal karyotype once adapted.

4

Notes 1. In addition to MEFs, equine embryonic fibroblasts or SNLs can be used successfully as feeder layers to generate and maintain equine iPSCs. These cells can be inactivated either by the use of γ-irradiation or mitomycin C treatment [5, 15]. 2. It is useful to freeze irradiated MEFs at a “working concentration,” e.g., in vials each containing the required amount for seeding a six-well plate (2 × 106 MEFs). 3. Feeder cell density may need to be optimized based on the performance of iPSC cultures. 4. Once colonies have been expanded into multiple wells, further optimization of dissociation method and/or culture media can be performed if needed, by determining in the first instance growth rate and morphology of colonies under different conditions. Different media may be suitable for culture of equine iPSCs, including chemically defined media [5, 10]. 5. Our laboratory has always used fibroblasts derived from a single equine fetus [13]; therefore, the effects of fetal age (and other potential variables) on the suitability of EEFs for iPSC culture have not been determined. 6. If at this point colonies stop growing or loose typical iPSC morphology, then changing culture conditions should be attempted. Different coating matrix and culture media combinations can be tried to establish feeder-free culture. Matrigel can be replaced with vitronectin, Laminin-521 [16, 17] or other commercially available matrices, in combination with media such as Essential 8 (Gibco) or mTeSR plus (Stemcell Technologies) [18, 19].

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Acknowledgments JF holds a PhD studentship funded through an industrial contract with IC BIOSOLUTIONS. The Roslin Institutes receives funding from the Biotechnology and Biological Sciences Research Council through an institute strategic program grant. References 1. Shi Y, Inoue H, Wu JC, Yamanaka S (2017) Induced pluripotent stem cell technology: a decade of progress. Nat Rev Drug Discov 16(2). https://doi.org/10.1038/nrd. 2016.245 2. Katayama M, Fukuda T, Kaneko T et al (2022) Induced pluripotent stem cells of endangered avian species. Commun Biol 5(1). https://doi. org/10.1038/s42003-022-03964-y 3. Takahashi K, Yamanaka S (2006) Induction of pluripotent stem cells from mouse embryonic and adult fibroblast cultures by defined factors. Cell 126(4). https://doi.org/10.1016/j.cell. 2006.07.024 4. Steinle H, Weber M, Behring A et al (2019) Reprogramming of urine-derived renal epithelial cells into iPSCs using srRNA and consecutive differentiation into beating cardiomyocytes. Mol Ther Nucleic Acids:17. https://doi.org/10.1016/j.omtn.2019. 07.016 5. Donadeu FX, Esteves CL (2015) Prospects and challenges of induced pluripotent stem cells in equine health. Front Vet Sci 2:59. https://doi. org/10.3389/fvets.2015.00059 6. Ban H, Nishishita N, Fusaki N et al (2011) Efficient generation of transgene-free human induced pluripotent stem cells (iPSCs) by temperature-sensitive Sendai virus vectors. Proc Natl Acad Sci U S A 108(34). https:// doi.org/10.1073/pnas.1103509108 7. Okita K, Matsumura Y, Sato Y et al (2011) A more efficient method to generate integrationfree human iPS cells. Nat Methods 8(5). https://doi.org/10.1038/nmeth.1591 8. Anokye-Danso F, Trivedi CM, Juhr D et al (2011) Highly efficient miRNA-mediated reprogramming of mouse and human somatic cells to pluripotency. Cell Stem Cell 8(4). https://doi.org/10.1016/j.stem.2011. 03.001 9. Nagy K, Sung HK, Zhang P et al (2011) Induced pluripotent stem cell lines derived from equine fibroblasts. Stem Cell Rev Rep

7(3). https://doi.org/10.1007/s12015-0119239-5 10. Yu L, Wei Y, Sun HX et al (2021) Derivation of intermediate pluripotent stem cells amenable to primordial germ cell specification. Cell Stem Cell 28(3). https://doi.org/10.1016/j. stem.2020.11.003 11. Chen G, Gulbranson DR, Hou Z et al (2011) Chemically defined conditions for human iPSC derivation and culture. Nat Methods 8(5). https://doi.org/10.1038/nmeth.1593 12. Wiley LA, Burnight ER, Deluca AP et al (2016) CGMP production of patient-specific iPSCs and photoreceptor precursor cells to treat retinal degenerative blindness. Sci Rep:6. https://doi.org/10.1038/srep30742 13. Sharma R, Livesey MR, Wyllie DJA et al (2014) Generation of functional neurons from feederfree, keratinocyte-derived equine induced pluripotent stem cells. Stem Cells Dev 23(13). https://doi.org/10.1089/scd.2013.0565 14. Quattrocelli M, Giacomazzi G, Broeckx SY et al (2016) Equine-induced pluripotent stem cells retain lineage commitment toward myogenic and chondrogenic fates. Stem Cell Rep 6(1). https://doi.org/10.1016/j.stemcr. 2015.12.005 15. Conner DA (2001) Mouse embryo fibroblast (MEF) feeder cell preparation. Curr Protoc Mol Biol 51:23. https://doi.org/10.1002/ 0471142727.mb2302s51 16. Rodin S, Antonsson L, Hovatta O, Tryggvason K (2014) Monolayer culturing and cloning of human pluripotent stem cells on laminin-521based matrices under xeno-free and chemically defined conditions. Nat Protoc 9(10). https:// doi.org/10.1038/nprot.2014.159 17. Braam SR, Zeinstra L, Litjens S et al (2008) Recombinant Vitronectin is a functionally defined substrate that supports human embryonic stem cell self-renewal via αVβ5 integrin. Stem Cells 26(9). https://doi.org/10.1634/ stemcells.2008-0291

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18. Chen G, Gulbranson DR, Hou Z et al (2011) Chemically defined conditions for human iPSC derivation and culture. Nat Methods 8(5): 424–429 ˜ uelas R, Sanjurjo-Rodrı´guez C, 19. Castro-Vin ˜ eiro-Ramil M et al (2021) Tips and tricks Pin

for successfully culturing and adapting human induced pluripotent stem cells. Mol Ther Methods Clin Dev:23. https://doi.org/10. 1016/j.omtm.2021.10.013

INDEX A Air-liquid interface (ALI) ........................ 44, 92, 98, 110, 112, 115–117, 120 Alginate........................................................ 104, 105, 107 Amniotic epithelial cells (ALCs) .................................. 135 Amniotic membrane (AM) ........................ 135, 136, 138, 140, 143, 144, 146 Antrum ......................................................................59–60 Apical .........................................7, 16, 19, 40, 55, 57, 61, 63, 74, 79, 152, 156, 158, 160 Apical compartment............ 61, 155, 156, 158–160, 162

Encapsulation ..............................................27–28, 31–33, 35, 36, 104, 106–107 Epithelial cells............................................. 1–6, 8, 14, 34, 37, 41, 44, 46, 55–63, 73, 74, 100, 110–115, 119, 120, 146, 147, 152, 163 Epithelial-mesenchymal transition ............. 136, 144, 146 Equine................................................................... 176–181 Explant............................. 29, 30, 34, 35, 37, 65, 91–100 Ex-vivo ................................................................ 39–53, 92 Ex-vivo models ..........................................................39–53

F

B

Feeder-free culture ............................................... 179, 181

Barrier study ......................................................... 161, 162 Basolateral.......................................... 7, 8, 19, 55, 61, 63, 92, 152, 155, 158, 160 Basolateral compartment ...... 56, 61, 156, 158, 160, 162

G

C Cattle ............................................................................. 167 Cell culture .....................................1–6, 9–11, 27–29, 31, 32, 40, 46, 76, 92–94, 104, 107, 175–181 Cell differentiation ...........................................79, 82, 129 Cell isolation............................................. 59–60, 70, 111, 123–133, 137, 139, 143, 166, 171–173 Cell reprogramming.................................... 175, 178, 179 Cervical tissue biopsies ........................................ 112–114 Collagenase................................................. 3, 4, 9, 16, 58, 59, 111, 112, 125, 127 Colon ............................................................7, 74, 91–100 Colon model .............................................................91–99 Current .....................................8, 10, 13, 15, 18–21, 152

D Differentiation........................................ 1, 34, 36, 55–63, 74, 92, 109–121, 129, 165–173, 180 Disease modelling ........................................................... 39

E Early placenta ....................................................... 123–133 Ectocervix ............................................................. 109, 110

Gastric ................................................................. 55–63, 69 Gastric tissue....................................................... 56, 65–71 Granulosa cells...................................................... 103–107 Growth ................................................... 2, 4, 5, 8–10, 13, 16–18, 27, 30, 41, 52, 58, 66, 76, 92, 99, 103, 110, 119, 120, 125, 128, 129, 133, 137–146, 153, 154, 165–173, 176, 178, 179, 181

I Induced pluripotent stem cells (iPSCs) ................................................74, 175–182 Intestinal stem cells (ISCs) ................................ 73, 74, 82 In vitro ..................... 39, 40, 55, 56, 65, 69, 86, 92, 103, 123, 129, 136, 140, 151, 152, 161, 166, 179 In vitro amplification .................................................... 136 In vitro organ culture (IVOC) ....................................... 92

M Mammary epithelial cells (MECs)....................... 151–163 Minor salivary glands ................................................26–38 Morphology.................................47, 56, 57, 67, 70, 129, 135, 136, 145, 161, 162, 166, 173, 180, 181 Mouse small intestine ...............................................73–83 Mucins ................................................................ 26, 55–57 Mucus ........................................................ 55–63, 96, 112 Myoblasts.............................................................. 166–169

Mario Baratta (ed.), Epithelial Cell Culture: Methods and Protocols, Methods in Molecular Biology, vol. 2749, https://doi.org/10.1007/978-1-0716-3609-1, © The Editor(s) (if applicable) and The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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186 Index O

Organ culture .............................................. 92, 93, 96, 99 Organoids ................................................... 56, 73–83, 92, 109–113, 115, 117–121 Organotypic slice culture..........................................39–53

P Pig ............................ 7–22, 55–63, 65–70, 100, 151–163 Pig model ............................................................. 151–163 Primary cell..................................................55–63, 67–70, 120, 123, 152, 161, 162 Primary cell culture ...............................65, 153, 159, 161 Primary trophoblast cells ............................ 129, 131, 133 Progesterone ..............................123, 129, 136, 140, 145

R Radiation injury.........................................................39, 40 Regenerative medicine ..........................26, 109, 136, 176 Resistance .........................................8, 10, 11, 13, 15–20, 22, 75, 77, 83, 107, 151, 152, 155–157

S Salivary gland............................26, 28, 29, 32, 34, 39–53 Salivary gland regeneration ......................................26–38 Satellite cells ......................................................... 165–173 Serum-free medium ...................................................... 1–6 Sheep...........................................123, 124, 129, 131, 132 Skeletal muscle ..................................................... 165–173 Sodium fluorescein..................................... 153, 154, 156, 158–160, 162, 163

Spheroids ........................................ 56, 57, 60, 62, 85–89 Stem cells .................................................... 56, 79, 92, 93, 109–121, 135, 136, 175–181 Swine................................................................................ 91

T Three-dimensional (3D) culture ........26, 76–78, 83, 104 3D hydrogel culture........................................... 26, 31–33 3D organoid .................... 73–83, 92, 109, 110, 117–118 3D tissue culture ..................................... 39–53, 112–114 Thymic epithelial cells (TECs) ..................................... 1–6 Thyrocyte....................................................................... 7, 8 Tight junction ..................... 14, 15, 56, 79, 81, 160, 161 Transepithelial electrical resistance (TEER) .......... 75, 81, 83, 151–157, 159, 160, 162 Transepithelial/transendothelial electrical resistance (TEER) ........................................... 75, 77, 81, 83, 151–156, 159, 160, 162 Transport .................................................. 7, 8, 10, 12, 15, 18, 28, 29, 59, 88, 93–97, 126, 127, 152, 154–156, 158–160, 162, 163 Two dimensional (2D) culture........ 40, 76, 80, 111, 113 2D epithelial monolayer ...........................................73–83

V Voltage ......................................................... 8, 19, 21, 152

X Xerostomia.................................................................26, 36