Methods and Protocols

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Methods in Molecular Biology 1799

R. Lee Reinhardt Editor

Type 2 Immunity Methods and Protocols

Methods

in

M o l e c u l a r B i o lo g y

Series Editor John M. Walker School of Life and Medical Sciences University of Hertfordshire Hatfield, Hertfordshire, AL10 9AB, UK

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

Type 2 Immunity Methods and Protocols

Edited by

R. Lee Reinhardt Department of Biomedical Research, National Jewish Health , Denver, CO, USA; Department of Immunology and Microbiology, University of Colorado School of Medicine, Aurora, CO, USA

Editor R. Lee Reinhardt Department of Biomedical Research National Jewish Health Denver, CO, USA Department of Immunology and Microbiology University of Colorado School of Medicine Aurora, CO, USA

ISSN 1064-3745     ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-4939-7895-3    ISBN 978-1-4939-7896-0 (eBook) https://doi.org/10.1007/978-1-4939-7896-0 Library of Congress Control Number: 2018945080 © Springer Science+Business Media, LLC, part of Springer Nature 2018 This work is subject to copyright. All rights are reserved by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, express or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. Printed on acid-free paper This Humana Press imprint is published by the registered company Springer Science+Business Media, LLC part of Springer Nature. The registered company address is: 233 Spring Street, New York, NY 10013, U.S.A.

Preface Type-2 immunity is an evolutionary conserved response in mammals to repeated infection and ­ colonization by large, extracellular worms called helminths. Type-2 immunity is ­advantageous to the host as it limits both worm burden and the tissue pathology associated with chronic inflammation. Type-2 responses are common in many rural communities where frequent and repeated helminth exposure occurs. It is estimated that 1 in 4 people worldwide are infected with some form of soil-transmitted helminth. As such, helminthic infections contribute significantly to the global burden of disease, exceeding less neglected infectious conditions like malaria and tuberculosis in certain developing countries. In more industrialized societies, helminth exposure is rare, but type-2 immunity remains a common fixture. This is because type-2 immunity takes the form of inappropriate responses to innocuous antigens (termed allergens). These responses are often referred to as allergic or type-2 inflammation. Allergic inflammation can occur in many tissues and is associated with a spectrum of diseases including atopy, allergy, rhinitis, and allergic asthma. The incidence of allergic disease has increased over the past 60 years, and this trend is expected to continue, with 100 million new cases expected over the next decade in the United States alone. Thus, like helminth infections in rural communities, allergic disease represents a significant and increasing global health concern. Given that the type-2 immune response is remarkably similar between helminths and allergens, studying helminth colonization is likely to help elucidate key mechanisms driving allergic inflammation. Indeed, a good understanding of the disease hallmarks associated with type-2 immunity (mucus production, smooth muscle contractility, eosinophilia, and IgE) has led to the development of therapies that can ameliorate allergic symptoms and disease progression. However, these therapeutic approaches often fail to address the underlying basis of disease. Thus, a ­better understanding of the basic mechanisms initiating the development of type-2 i­nflammation remains an important area of research. A more basic understanding of type-2 immunity is likely to increase therapeutic options to those already afflicted, and will reveal novel ways to limit the onset or reduce susceptibility to allergic disorders. The development of novel tools, models, and experimental approaches to explore type-2 immunity in mice and humans lay at the heart of uncovering clinically relevant approaches to combating allergic disease and the pathology associated with high-burden helminth infections. In the last decade alone, basic research has revealed many new paradigms in type-2 ­immunity. To name just a few, these range from the discovery of innate lymphoid cells and their role in barrier homeostasis, tuft cell differentiation, and crosstalk with neurons at m ­ ucosal barriers; the role of the microbiome and helminths in influencing allergic disease ­susceptibility; the use of next-generation sequencing platforms, single-cell approaches, and systems biology to identify allergic endotypes; the increasing specialization of various CD4+ T cell subsets in settings of type-2 immunity; the use of mouse genetics and lineage tracing to reveal the ­developmental origins and effector functions of various myeloid and lymphoid populations; and our increasing understanding of the function and regulation of type-2 cytokines and the cells that produce them has spawned renewed interest in the d ­ evelopment of biologics to treat

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a spectrum of allergic diseases. These are just a few of the recent advances in type-2 immunity that are likely to have significant influence in the clinic in the coming years. The following chapters are provided by leading scientists in the field of type-2 i­ mmunity. In sum, the chapters reveal many of the key methods and tools used to identify the ­paradigms discussed in the preceding paragraphs. The initial chapters explore various models of type-2 immunity allowing researchers the opportunity to investigate various aspects of type-2 ­biology in different disease settings. Next, the ensuing chapters focus on the cellular ­protocols designed to identify, characterize, and assess the function of various adaptive and innate immune cells critical to the development of type-2 immunity. These approaches provide investigators the ability to isolate and evaluate specific cellular subsets at the genetic, epigenetic, and molecular level. The final chapters are devised to assess type-2 inflammation and its relationship to organismal and metabolic systems (e.g., microbiome) and explore the use of primary human cells and culture systems to assess relevance in humans. In all, the book is designed to provide a broad network of methods that can be used to develop a hypothesis and investigate its potential from bench to bedside. Denver, CO, USA

R. Lee Reinhardt

Contents Preface�����������������������������������������������������������������������������������������������������������������������   v Contributors��������������������������������������������������������������������������������������������������������������   xi 1 A Fungal Protease Model to Interrogate Allergic Lung Immunity����������������������    1 J. Morgan Knight, Evan Li, Hui-Ying Tung, Cameron Landers, Jake Wheeler, Farrah Kheradmand, and David B. Corry 2 Use of the Litomosoides sigmodontis Infection Model of Filariasis to Study Type 2 Immunity���������������������������������������������������������������  11 A. Fulton, S. A. Babayan, and M. D. Taylor 3 Production of Hymenolepis diminuta in the Laboratory: An Old Research Tool with New Clinical Applications����������������������������������������  27 Min Zhang, Amanda J. Mathew, and William Parker 4 A Mouse Model of Peanut Allergy Induced by Sensitization Through the Gastrointestinal Tract��������������������������������������������������������������������  39 Kelly Orgel and Michael Kulis 5 Induction and Characterization of the Allergic Eye Disease Mouse Model����������  49 Nancy J. Reyes, Rose Mathew, and Daniel R. Saban 6 Isolation and Purification of Epithelial and Endothelial Cells from Mouse Lung�������������������������������������������������������������������������������������  59 Hideki Nakano, Keiko Nakano, and Donald N. Cook 7 In Vitro and In Vivo IgE-/Antigen-Mediated Mast Cell Activation��������������������  71 Hae Woong Choi and Soman N. Abraham 8 The Use of Human and Mouse Mast Cell and Basophil Cultures to Assess Type 2 Inflammation���������������������������������������������������������������������������  81 Heather L. Caslin, Marcela T. Taruselli, Anuya Paranjape, Kasalina Kiwanuka, Tamara Haque, Alena P. Chumanevich, Carole A. Oskeritzian, and John J. Ryan 9 Isolation and Identification of Group 2 Innate Lymphoid Cells in Settings of Type 2 Inflammation������������������������������������������������������������  93 Jesse Charles Nussbaum and Jorge Felipe Ortiz-Carpena 10 Determination of the Fate and Function of Innate Lymphoid Cells Following Adoptive Transfer of Innate Lymphoid Cell Precursors�������������� 109 Timothy E. O’Sullivan and Joseph C. Sun 11 Characterization of Thymic Development of Natural Killer T Cell Subsets by Multiparameter Flow Cytometry��������������������������������������������� 121 Kathryn D. Tuttle and Laurent Gapin 12 Characterization of Mouse γδ T Cell Subsets in the Setting of Type-2 Immunity������������������������������������������������������������������������������������������� 135 Wanjiang Zeng, Rebecca L. O’Brien, Willi K. Born, and Yafei Huang

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13 The Identification of Allergen-Derived T Cell Epitopes�������������������������������������� 153 Véronique Schulten and Alessandro Sette 14 Generation of Allergen-Specific Tetramers for a Murine Model of Airway Inflammation�������������������������������������������������������������������������� 165 James J. Moon and Marion Pepper 15 The Generation and Use of Allergen-Specific TCR Transgenic Animals�������������� 183 Manon Vanheerswynghels, Wendy Toussaint, Martijn Schuijs, Leen Vanhoutte, Nigel Killeen, Hamida Hammad, and Bart N. Lambrecht 16 Using Cytokine Reporter Mice to Visualize Type-2 Immunity In Vivo��������������� 211 Mark Dell’Aringa and R. Lee Reinhardt 17 Live Imaging of IL-4-Expressing T Follicular Helper Cells in Explanted Lymph Nodes������������������������������������������������������������������������������� 225 Mark Dell’Aringa, R. Lee Reinhardt, Rachel S. Friedman, and Jordan Jacobelli 18 Imaging Precision-Cut Lung Slices to Visualize Leukocyte Localization and Trafficking������������������������������������������������������������������������������� 237 Miranda R. Lyons-Cohen, Hideki Nakano, Seddon Y. Thomas, and Donald N. Cook 19 Study of IgE-Producing B Cells Using the Verigem Fluorescent Reporter Mouse������������������������������������������������������������������������������������������������� 247 Zhiyong Yang, James B. Jung, and Christopher D. C. Allen 20 Chromatin Preparation from Murine Eosinophils for Genome-Wide Analyses�������������������������������������������������������������������������������� 265 Carine Bouffi, Artem Barski, and Patricia C. Fulkerson 21 A Sensitive and Integrated Approach to Profile Messenger RNA from Samples with Low Cell Numbers������������������������������������������������������ 275 Sandy Lisette Rosales, Shu Liang, Isaac Engel, Benjamin Joachim Schmiedel, Mitchell Kronenberg, Pandurangan Vijayanand, and Grégory Seumois 22 An Integrated and Semiautomated Microscaled Approach to Profile Cis-Regulatory Elements by Histone Modification ChIP-Seq for Large-Scale Epigenetic Studies���������������������������������������������������������������������� 303 Diana Youhanna Jankeel, Justin Cayford, Benjamin Joachim Schmiedel, Pandurangan Vijayanand, and Grégory Seumois 23 Library Preparation for ATAC-Sequencing of Mouse CD4+ T Cells Isolated from the Lung and Lymph Nodes After Helminth Infection����������������������������������������������������������������������������������� 327 Laura D. Harmacek, Preeyam Patel, Rachel Woolaver, R. Lee Reinhardt, and Brian P. O’Connor 24 Identification of Functionally Relevant microRNAs in the Regulation of Allergic Inflammation��������������������������������������������������������� 341 Marlys S. Fassett, Heather H. Pua, Laura J. Simpson, David F. Steiner, and K. Mark Ansel 25 The Use of Biodegradable Nanoparticles for Tolerogenic Therapy of Allergic Inflammation����������������������������������������������������������������������� 353 Charles B. Smarr and Stephen D. Miller

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26 Assessing the Mouse Intestinal Microbiota in Settings of Type-2 Immune Responses���������������������������������������������������������������������������� 359 Mei San Tang, Rowann Bowcutt, and P’ng Loke 27 The Use of CRISPR-Cas9 Technology to Reveal Important Aspects of Human Airway Biology��������������������������������������������������������������������� 371 Azzeddine Dakhama and Hong Wei Chu 28 A Consistent Method to Identify and Isolate Mononuclear Phagocytes from Human Lung and Lymph Nodes��������������������������������������������� 381 Sophie L. Gibbings and Claudia V. Jakubzick 29 Organoid Cultures for Assessing Intestinal Epithelial Differentiation and Function in Response to Type-2 Inflammation�������������������������������������������� 397 Bailey Zwarycz, Adam D. Gracz, and Scott T. Magness 30 Utilization of Air–Liquid Interface Cultures as an In Vitro Model to Assess Primary Airway Epithelial Cell Responses to the Type 2 Cytokine Interleukin-13��������������������������������������������������������������� 419 Jamie L. Everman, Cydney Rios, and Max A. Seibold Index������������������������������������������������������������������������������������������������������������������������   433

Contributors Soman N. Abraham  •  Department of Pathology, Duke University Medical Center, Durham, NC, USA; Department of Immunology, Duke University Medical Center, Durham, NC, USA; Molecular Genetics and Microbiology, Duke University Medical Center, Durham, NC, USA; Program in Emerging Infectious Diseases Duke-National University of Singapore, Singapore, Singapore Christopher D. C. Allen  •  Cardiovascular Research Institute, University of California, San Francisco, CA, USA; Sandler Asthma Basic Research Center, University of California, San Francisco, CA, USA; Department of Anatomy, University of California, San Francisco, CA, USA S. A. Babayan  •  Institute of Biodiversity, Animal Health and Comparative Medicine, College of Medical, Veterinary and Life Sciences, University of Glasgow, Glasgow, UK Artem Barski  •  Division of Allergy and Immunology, Department of Pediatrics Cincinnati Children’s Hospital Medical Center, University of Cincinnati College of Medicine, Cincinnati, OH, USA; Division of Human Genetics, Department of Pediatrics, , Cincinnati Children’s Hospital Medical Center, University of Cincinnati College of Medicine, Cincinnati, OH, USA Willi K. Born  •  Department of Biomedical Research, National Jewish Health, Denver, CO, USA; Department of Immunology and Microbiology, University of Colorado Health Sciences Center, Aurora, CO, USA Carine Bouffi  •  Division of Allergy and Immunology, Department of Pediatrics, Cincinnati Children’s Hospital Medical Center, University of Cincinnati College of Medicine, Cincinnati, OH, USA Rowann Bowcutt  •  Department of Microbiology, New York University School of Medicine, New York, NY, USA; UCB Celltech, Slough, UK Heather L. Caslin  •  Department of Biology, Virginia Commonwealth University, Richmond, VA, USA Justin Cayford  •  La Jolla Institute for Allergy and Immunology, La Jolla, CA, USA Hae Woong Choi  •  Department of Pathology, Duke University Medical Center, Durham, NC, USA Hong Wei Chu  •  Department of Medicine, National Jewish Health, Denver, CO, USA Alena P. Chumanevich  •  Department of Pathology, Microbiology and Immunology, University of South Carolina School of Medicine, Columbia, SC, USA Donald N. Cook  •  Immunity, Inflammation and Disease Laboratory, National Institute of Environmental Health Sciences, National Institutes of Health, Research Triangle Park, NC, USA David B. Corry  •  Department of Medicine, Baylor College of Medicine, Houston, TX, USA; Department of Pathology & Immunology, Baylor College of Medicine, Houston, TX, USA; Department of Biology of Inflammation Center, Baylor College of Medicine, Houston, TX, USA; Michael E. DeBakey VA Center for Translational Research on Inflammatory Diseases, Houston, TX, USA

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Azzeddine Dakhama  •  Department of Medicine, National Jewish Health, Denver, CO, USA Mark Dell’Aringa  •  Department of Immunology, Duke University Medical Center, Durham, NC, USA; Department of Biomedical Research, National Jewish Health, Denver, CO, USA Isaac Engel  •  La Jolla Institute for Allergy and Immunology, La Jolla, CA, USA Jamie L. Everman  •  Center for Genes, Environment, and Health, National Jewish Health, Denver, CO, USA Marlys S. Fassett  •  Department of Microbiology & Immunology, Sandler Asthma Basic Research Center, University of California, San Francisco, CA, USA; Department of Dermatology, University of California, San Francisco, CA, USA Rachel S. Friedman  •  Department of Biomedical Research, National Jewish Health, Denver, CO, USA; Department of Immunology and Microbiology, University of Colorado School of Medicine, Aurora, CO, USA Patricia C. Fulkerson  •  Division of Allergy and Immunology, Cincinnati Children’s Hospital Medical Center, Cincinnati, OH, USA; Department of Pediatrics, University of Cincinnati College of Medicine, Cincinnati, OH, USA A. Fulton  •  Institute of Immunology and Infection Research, School of Biological Sciences, University of Edinburgh, Edinburgh, UK; Centre for Immunology, Infection, and Evolution, University of Edinburgh, Edinburgh, UK Laurent Gapin  •  Department of Immunology and Microbiology, University of Colorado Anschutz Medical Campus, Aurora, CO, USA Sophie L. Gibbings  •  Department of Pediatrics, National Jewish Health, Denver, CO, USA Adam D. Gracz  •  Department of Genetics, University of North Carolina at Chapel Hill, Chapel Hill, NC, USA Hamida Hammad  •  VIB-UGent Center for Inflammation Research, Ghent, Belgium; Department of Respiratory Medicine, Ghent University, Ghent, Belgium Tamara Haque  •  Department of Biology, Virginia Commonwealth University, Richmond, VA, USA Laura D. Harmacek  •  Center for Genes, Environment, and Health, and the Department of Pediatrics, National Jewish Health, Denver, CO, USA; Department of Immunology and Microbiology, University of Colorado School of Medicine, Aurora, CO, USA Yafei Huang  •  Key Laboratory for Molecular Diagnosis of Hubei Province, The Central Hospital of Wuhan, Tongji Medical College, Huazhong University of Science and Technology, Wuhan, People’s Republic of China; Joint Laboratory for Stem Cell Engineering and Technology Transfer, School of Basic Medicine, Tongji Medical College, Huazhong University of Science and Technology, Wuhan, People’s Republic of China Jordan Jacobelli  •  Department of Biomedical Research, National Jewish Health, Denver, CO, USA; Department of Immunology and Microbiology, University of Colorado School of Medicine, Aurora, CO, USA Claudia V. Jakubzick  •  Department of Pediatrics, National Jewish Health, Denver, CO, USA; Department of Microbiology and Immunology, University of Colorado, Denver, CO, USA James B. Jung  •  Cardiovascular Research Institute, University of California, San Francisco, CA, USA; Sandler Asthma Basic Research Center, University of California, San Francisco, CA, USA

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Farrah Kheradmand  •  Department of Medicine, Baylor College of Medicine, Houston, TX, USA; Department of Pathology & Immunology, Baylor College of Medicine, Houston, TX, USA; Department of Biology of Inflammation Center, Baylor College of Medicine, Houston, TX, USA; Michael E. DeBakey VA Center for Translational Research on Inflammatory Diseases, Houston, TX, USA Nigel Killeen  •  VIB-UGent Center for Inflammation Research, Ghent, Belgium Kasalina Kiwanuka  •  Department of Biology, Virginia Commonwealth University, Richmond, VA, USA Mitchell Kronenberg  •  La Jolla Institute for Allergy and Immunology, La Jolla, CA, USA; Division of Biologic Sciences, University of California San Diego, La Jolla, CA, USA Michael Kulis  •  Division of Allergy, Immunology, and Rheumatology, Department of Pediatrics, University of North Carolina at Chapel Hill, Chapel Hill, NC, USA Bart N. Lambrecht  •  VIB-UGent Center for Inflammation Research, Ghent, Belgium; Department of Respiratory Medicine, Ghent University, Ghent, Belgium; Department of Pulmonary Medicine, ErasmusMC, Rotterdam, The Netherlands Cameron Landers  •  Department of Medicine, Baylor College of Medicine, Houston, TX, USA R. Lee Reinhardt  •  Department of Biomedical Research, National Jewish Health, Denver, CO, USA; Department of Immunology and Microbiology, University of Colorado School of Medicine, Aurora, CO, USA Shu Liang  •  La Jolla Institute for Allergy and Immunology, La Jolla, CA, USA Evan Li  •  Department of Medicine, Baylor College of Medicine, Houston, TX, USA P’ng Loke  •  Department of Microbiology, New York University School of Medicine, New York, NY, USA Miranda R. Lyons-Cohen  •  Immunity, Inflammation and Disease Laboratory, National Institute of Environmental Health Sciences, National Institutes of Health, Research Triangle Park, NC, USA Scott T. Magness  •  Department of Cell Biology and Physiology, University of North Carolina at Chapel Hill, Chapel Hill, NC, USA; Joint Department of Biomedical Engineering, University of North Carolina at Chapel Hill/North Carolina State University, Chapel Hill, NC, USA; Department of Medicine, University of North Carolina at Chapel Hill, Chapel Hill, NC, USA K. Mark Ansel  •  Department of Microbiology and Immunology, Sandler Asthma Basic Research Center, University of California, San Francisco, San Francisco, CA, USA Amanda J. Mathew  •  Department of Surgery, Duke University Medical Center, Durham, NC, USA Rose Mathew  •  Department of Ophthalmology, Duke University School of Medicine, Duke Eye Center, Durham, NC, USA Stephen D. Miller  •  Department of Microbiology-Immunology and Immunobiology Center, Feinberg School of Medicine, Northwestern University, Chicago, IL, USA James J. Moon  •  Center for Immunology and Inflammatory Diseases and Division of Pulmonary and Critical Care Medicine, Massachusetts General Hospital, Boston, MA, USA; Harvard Medical School, Boston, MA, USA J. Morgan Knight  •  Department of Medicine, Baylor College of Medicine, Houston, TX, USA

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Hideki Nakano  •  Immunity, Inflammation and Disease Laboratory, National Institute of Environmental Health Sciences, National Institutes of Health, Research Triangle Park, NC, USA Keiko Nakano  •  Immunity, Inflammation and Disease Laboratory, National Institute of Environmental Health Sciences, National Institutes of Health, Research Triangle Park, NC, USA Jesse Charles Nussbaum  •  Department of Internal Medicine, Infectious Diseases, University of California San Francisco, San Francisco, CA, USA Rebecca L. O’Brien  •  Department of Biomedical Research, National Jewish Health, Denver, CO, USA; Department of Immunology and Microbiology, University of Colorado Health Sciences Center, Aurora, CO, USA Brian P. O’Connor  •  Center for Genes, Environment, and Health, and the Department of Pediatrics, National Jewish Health, Denver, CO, USA; Department of Immunology and Microbiology, University of Colorado School of Medicine, Aurora, CO, USA Timothy E. O’Sullivan  •  Department of Microbiology, Immunology, and Molecular Genetics, David Geffen School of Medicine at UCLA, Los Angeles, CA, USA Kelly Orgel  •  Division of Allergy, Immunology, and Rheumatology, Department of Pediatrics, University of North Carolina at Chapel Hill, Chapel Hill, NC, USA Jorge Felipe Ortiz-Carpena  •  Department of Internal Medicine, Infectious Diseases, University of California San Francisco, San Francisco, CA, USA Carole A. Oskeritzian  •  Department of Pathology, Microbiology and Immunology, University of South Carolina School of Medicine, Columbia, SC, USA Anuya Paranjape  •  Department of Biology, Virginia Commonwealth University, Richmond, VA, USA William Parker  •  Department of Surgery, Duke University Medical Center, Durham, NC, USA Preeyam Patel  •  Department of Biomedical Research, National Jewish Health, Denver, CO, USA Marion Pepper  •  Department of Immunology, University of Washington School of Medicine, Seattle, WA, USA Heather H. Pua  •  Department of Microbiology & Immunology, Sandler Asthma Basic Research Center, University of California, San Francisco, CA, USA; Department of Pathology, University of California, San Francisco, CA, USA Nancy J. Reyes  •  Department of Ophthalmology, Duke University School of Medicine, Duke Eye Center, Durham, NC, USA Cydney Rios  •  Center for Genes, Environment, and Health, National Jewish Health, Denver, CO, USA Sandy Lisette Rosales  •  La Jolla Institute for Allergy and Immunology, La Jolla, CA, USA John J. Ryan  •  Department of Biology, Virginia Commonwealth University, Richmond, VA, USA Daniel R. Saban  •  Department of Ophthalmology, Duke University School of Medicine, Duke Eye Center, Durham, NC, USA; Department of Immunology, Duke University School of Medicine, Durham, NC, USA Benjamin Joachim Schmiedel  •  La Jolla Institute for Allergy and Immunology, La Jolla, CA, USA

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Martijn Schuijs  •  VIB-UGent Center for Inflammation Research, Ghent, Belgium; Department of Respiratory Medicine, Ghent University, Ghent, Belgium Véronique Schulten  •  Division of Vaccine Discovery, La Jolla Institute for Allergy and Immunology, La Jolla, CA, USA Max A. Seibold  •  Center for Genes, Environment, and Health, National Jewish Health, Denver, CO, USA; Department of Pediatrics, National Jewish Health, Denver, CO, USA; Division of Pulmonary Sciences and Critical Care Medicine, University of Colorado School of Medicine, Aurora, CO, USA Alessandro Sette  •  Division of Vaccine Discovery, La Jolla Institute for Allergy and Immunology, La Jolla, CA, USA Grégory Seumois  •  La Jolla Institute for Allergy and Immunology, La Jolla, CA, USA Laura J. Simpson  •  Department of Microbiology and Immunology, Sandler Asthma Basic Research Center, University of California, San Francisco, CA, USA Charles B. Smarr  •  Department of Microbiology-Immunology and Immunobiology Center, Feinberg School of Medicine, Northwestern University, Chicago, IL, USA; Translational Research Program, Benaroya Research Institute at Virginia Mason, Seattle, WA, USA David F. Steiner  •  Department of Microbiology and Immunology, Sandler Asthma Basic Research Center, University of California, San Francisco, CA, USA Joseph C. Sun  •  Immunology Program, Memorial Sloan Kettering Cancer Center, New York, NY, USA; Department of Immunology and Microbial Pathogenesis, Weill Cornell Medical College, New York, NY, USA Mei San Tang  •  Department of Microbiology, New York University School of Medicine, New York, NY, USA Marcela T. Taruselli  •  Department of Biology, Virginia Commonwealth University, Richmond, VA, USA M. D. Taylor  •  Institute of Immunology and Infection Research, School of Biological Sciences, University of Edinburgh, Edinburgh, UK; Centre for Immunology, Infection, and Evolution, University of Edinburgh, Edinburgh, UK; Ashworth Laboratories, Edinburgh, UK Seddon Y. Thomas  •  Immunity, Inflammation and Disease Laboratory, National Institute of Environmental Health Sciences, National Institutes of Health, Research Triangle Park, NC, USA Wendy Toussaint  •  VIB-UGent Center for Inflammation Research, Ghent, Belgium; Department of Respiratory Medicine, Ghent University, Ghent, Belgium Hui-Ying Tung  •  Department of Pathology & Immunology, Baylor College of Medicine, Houston, TX, USA Kathryn D. Tuttle  •  Department of Immunology and Microbiology, University of Colorado Anschutz Medical Campus, Aurora, CO, USA Manon Vanheerswynghels  •  VIB-UGent Center for Inflammation Research, Ghent, Belgium; Department of Respiratory Medicine, Ghent University, Ghent, Belgium Leen Vanhoutte  •  VIB-UGent Center for Inflammation Research, Ghent, Belgium; Transgenic Core Facility, VIB Center for Inflammation Research, Ghent, Belgium Pandurangan Vijayanand  •  La Jolla Institute for Allergy and Immunology, La Jolla, CA, USA; Faculty of Medicine, Clinical and Experimental Sciences, National Institute for Health Research, Southampton Respiratory Biomedical Research Unit, University of Southampton, Southampton, UK

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Contributors

Jake Wheeler  •  Department of Medicine, Baylor College of Medicine, Houston, TX, USA Rachel Woolaver  •  Department of Immunology and Microbiology, University of Colorado School of Medicine, Aurora, CO, USA Zhiyong Yang  •  Cardiovascular Research Institute, University of California, San Francisco, CA, USA; Sandler Asthma Basic Research Center, University of California, San Francisco, CA, USA Diana Youhanna Jankeel  •  La Jolla Institute for Allergy and Immunology, La Jolla, CA, USA Wanjiang Zeng  •  Department of Obstetrics and Gynecology, Tongji Hospital, Tongji Medical College, Huazhong University of Science and Technology, Wuhan, People’s Republic of China Min Zhang  •  Department of Surgery, Duke University Medical Center, Durham, NC, USA Bailey Zwarycz  •  Department of Cell Biology and Physiology, University of North Carolina at Chapel Hill, Chapel Hill, NC, USA

Chapter 1 A Fungal Protease Model to Interrogate Allergic Lung Immunity J. Morgan Knight, Evan Li, Hui-Ying Tung, Cameron Landers, Jake Wheeler, Farrah Kheradmand, and David B. Corry Abstract Allergic airway diseases (asthma and chronic rhinosinusitis) are among the most common of all human diseases in heavily industrialized societies. Animal models of asthma have provided remarkable insight into allergic disease pathogenesis and will continue to drive the discovery of new therapeutic insights. We provide in this chapter a detailed protocol for inducing allergic immunity in the lungs of mice using a purified fungal protease and include related protocols for assessing immune endpoints. Key words Allergic immunity, Asthma, Airway hyperresponsiveness, TH2 cell, Protease, Fungi

1  Introduction Allergic diseases of the respiratory tract, comprised largely of asthma (lower airway) and chronic rhinosinusitis (upper airway), are among the most common of all human diseases in heavily industrialized societies. The incidence and prevalence of these conditions continue to rise in the United States at the same time that therapy remains inadequate [1]. Although fundamental insight into the pathogenesis of allergic lung diseases has been made and has resulted in the development of novel therapeutic approaches [2], much additional study through experimental models will be required before salutary changes in allergic disease epidemiology ensue. All allergic disorders share a distinctive pattern of inflammation dominated by characteristic immune cells (e.g., T helper cells type 2 (TH2), eosinophils, innate lymphoid cells type 2 (ILC2)), cytokines (e.g., interleukin 4 (IL-4), IL-5, IL-13), chemokines (CCL7, CCL17, CCL26), and antibody responses (e.g., immunoglobulin E (IgE)) that often mediate common physiological alterations

R. Lee Reinhardt (ed.), Type 2 Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1799, https://doi.org/10.1007/978-1-4939-7896-0_1, © Springer Science+Business Media, LLC, part of Springer Nature 2018

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(e.g., airway and gut hypermotility or hyperresponsiveness and mucus secretion) [3]. The fundamental importance of protease activity for allergic inflammation was a transformative discovery in the pathogenesis of allergic lung disease [4]. Regardless of whether the allergen contains proteases or not, endogenous proteases such as thrombin are the likely danger signals that mediate allergic inflammation [5]. Nonetheless, data increasingly suggest that exogenous proteases, and particularly protease-secreting pathogens such as fungi, are of fundamental importance to human allergic airway diseases [6–8]. We provide in this chapter a detailed protocol for inducing allergic immunity in the lungs of mice using a purified fungal protease. In addition to describing the method for preparing and challenging mice with fungal protease, we include additional protocols for measuring lung mechanics (airway responsiveness), collecting airway immune cells via bronchoalveolar lavage, and processing blood and lung for immune studies.

2  Materials 2.1  Protease/ Allergen Model

1. Age-matched (ideally 4–12 weeks old), sex-matched mice of suitable strain for proposed study: C57BL/6, Balb/c, FVB/N, knockout strain wild type, etc. (see Note 1). 2. Phosphate-buffered saline (PBS). 3. Protease solution: 1.8 mg/mL protease from Aspergillus oryzae (Sigma: P6110), stored in 55 μL aliquots at −80 °C. Aliquots are resuspended to 550 μL with PBS for treatment of mice with 50 μL of protease (see Note 2). 4. Basic tabletop anesthesia machine and chamber 5. ISOTHESIA – isoflurane inhalant 6. Oxygen supply

2.2  Plethysmography

1. Assess airway responsiveness with custom built or commercially available plethysmography equipment (Buxco or FlexiVent). 2. Small animal ventilator. 3. PROTECTIV® Safety IV angiocatheters, 20-gauge × 1 1/4″, FEP polymer.

radiopaque,

4. 26-gauge needle for establishing tail IV or suitable equivalent as defined by lab equipment. 5. NERL™ Blood Bank Saline, buffered to a pH of 7.0–7.2 (Thermo Fisher Scientific). 6. Acetylcholine stock solution: 1.6 mg/mL acetylcholine (ACh) in PBS and stored in 10 mL aliquots at −80 °C.

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7. Sterile 1 mL syringes. 8. Amidate (etomidate). 2.3  Tissue Collection and Processing

1. 1 mL syringes. 2. 1.5 mL microcentrifuge tubes. 3. Neutral buffered saline. 4. Hemacytometer or cell counting equipment. 5. Cytospin. 6. Superfrost Plus Microscope Slides. 7. Formalin fixative solution: 10% formalin. 8. Hematoxylin and eosin (H&E) stain. 9. Complete media (DMEM or RPMI): DMEM or RPMI containing 10% FBS, penicillin (100 U/mL), and streptomycin (100 μg/mL). 10. 6-well tissue culture plates. 11. Falcon cell strainer, 40 micron. 12. 50 mL conical tubes. 13. 3 mL syringes. 14. RNAlater. 15. TRIzol. 16. Surgical thread. 17. Necropsy equipment.

3  Method 3.1  Murine Challenge with Protease

1. Adequate numbers of mice should be acquired to allow for 5–6 mice per treatment group (see Note 1). Naïve vehicle control (PBS) and positive control groups (protease challenged) should be assessed in every experiment, along with the desired test groups (drug treatment, adoptive transfer, knockout mice, etc.) to ensure changes in airway responsiveness and inflammation. 2. Defrost and bring protease solution to a final volume of 550  μL, a standard volume for treatment of ten mice (five/ group) and one extra dose for error (see Note 2). A tube of equivalent volume of PBS/diluent/vehicle should be prepared for treatment of naïve control groups (see Note 3). 3. Mice should be anesthetized in accordance to approved animal protocol, to sufficient depth for reliable intranasal (i.n.) treatment (tip: titrate anesthesia plane to one breath every 1–2 s (see Note 4)). Once sufficient sedation is achieved, apply 50 μL dose of protease/vehicle to the nares using a handheld pipette

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Fig. 1 Protease model. (a) Timeline for protease/allergen challenge in which mice are intranasally (i.n.) challenged every other day for seven challenges. (b) Exemplary graph of respiratory system resistance measurements (RRS) plotted against increasing concentrations of acetylcholine chloride (ACh) injected intravenously to C57BL/6 mice challenged with Aspergillus oryzae protease (PAO) or PBS. (c) Bronchoalveolar lavage (BAL) fluid total and differential inflammatory cell counts from the same mice undergoing airway mechanic testing (b) including total cells and total macrophages (Macro), eosinophils (Eos), neutrophils (Neutro), and lymphocytes (Lymph). N = 5 mice/group. *P 1.0 × 105 conidia/treatment prepared from Aspergillus spp. and others) are sufficient to induce the hallmarks of allergic disease assessed in this model. 4. Adequate sedation is critical to ensure that the mouse reliably inhales with sufficient force to deliver protease/buffer to the airway and minimizes swallowing. Too deep a sedation (>4 s between breaths) will result in death due to asphyxiation. Additionally, it is important to place a small drop of protease/ PBS solution on the nares to assess if solution is absorbed by sinuses or swallowed. If the drop is passively absorbed by the sinuses or swallowed, allow one breath to pass and test again. If the drop remains on the nares and is not slowly absorbed between breaths, then quickly apply the bulk of the dose to be delivered, and allow the mouse to inhale over 1–2 s. Immediately following inhalation, apply the remaining dose, and wait for complete inhalation of the dose to be delivered before returning the mouse to the cage. 5. Once thawed, four serial dilutions of 3.2 mL acetylcholine into 6.8 mL of buffered saline result in five tubes of increasing doses of acetylcholine (first dose, 0.01; second, 0.03; third, 0.32; fourth, 1.0; fifth, 3.2 μg/g). 6. Lung, spleen, blood, and bronchial airway lavage samples can be collected. Lung tissue can be partitioned into sections for creation of a single cell suspension for ELISPOT, ELISA, and/ or flow cytometric analysis or process for protein or mRNA. Whole lung inflation or partial lung inflation (i.e., cannulation of the airway prior to hydrostatic inflation with fixative) may be performed to assess goblet cell metaplasia. If mucus plugging is a significant observation, it is recommended that additional mice be included. Ideally, mice dedicated for histologic assessment should not undergo airway mechanic

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testing, due to the increased fluid movement/mucus flushing induced by high doses of acetylcholine. 7. An initial tidal volume of 0.15 cc is typically sufficient for most mice. It is best to continuously calibrate tidal volume at the beginning for each mouse, ensuring that observed airway resistance values are not artificially enhanced or reduced due to tidal volume error. 8. A factor of 24 times body weight is an overdose and ensures that the mouse is completely sedated for analysis and harvesting. A factor of 16 times body weight can be used and is sufficient to achieve sedation but allows the mouse to recover from analysis for experiments where survival is desired. For survival studies, mice should be allowed to recover in a cage supplied with oxygen and warmed by a heating pad. Additionally, mice should remain intubated until their respiration rate has increased and regained a strong twitch response. It is critical that the catheter is removed once the mouse has significantly recovered from sedation in order to prevent choking and death. 9. Higher doses may require additional time between injections to allow for sufficient recovery and return to baseline conditions. Repeating doses is acceptable, but void volume must be considered. Priming the IV with one lower dose before repeating the desired dose will ensure the proper dose delivered. 10. Total number of cells can vary significantly across treatment and control groups. As long as a clear monolayer of cells is placed on the slide in sufficient numbers to allow the differential count of at least a 100–200 cells, the number of cells spun down is not critically restricted to 2 × 105. Additionally, it is important to maintain fresh stocks of H&E stains to ensure proper staining of eosin for eosinophil identification. 11. The lobe selected for RNA or protein analysis should be consistent throughout the experiment. Ideally, a lobe is dedicated to RNA or protein. RNAlater allows sample collection and storage for future RNA isolation by TRIzol. If protein stability is an issue, it is strongly suggested that the lobe be partitioned and snapfrozen in liquid nitrogen or immediate homogenization and cell lysis in the presence of protease and phosphatase inhibitors. References 1. Akinbami LJ, Moorman JE, Bailey K, Zahran HD, King M, Johnson CA, Liu X (2012) Trends in asthma prevalence, health care use, and mortality in the United States, 2001–2010. NCHS Data Brief 94:1–8 2. Pawankar R, Hayashi M, Yamanishi S, Igarashi T (2015) The paradigm of cytokine networks in allergic airway inflammation. Curr Opin Allergy Clin Immunol 15(1):41–48

3. Lambrecht BN, Hammad H (2015) The immunology of asthma. Nat Immunol 16(1):45–56 4. Kheradmand F, Kiss A, Xu J, Lee SH, Kolattukudy PE, Corry DB (2002) A protease-­ activated pathway underlying Th cell type 2 activation and allergic lung disease. J Immunol 169(10):5904–5911

A Fungal Protease Model to Interrogate Allergic Lung Immunity 5. Millien VO, Lu W, Shaw J, Yuan X, Mak G, Roberts L, Song LZ, Knight JM, Creighton CJ, Luong A, Kheradmand F, Corry DB (2013) Cleavage of fibrinogen by proteinases elicits allergic responses through Toll-like receptor 4. Science 341(6147):792–796 6. Mak G, Porter PC, Bandi V, Kheradmand F, Corry DB (2013) Tracheobronchial mycosis in a retrospective case-series study of five status asthmaticus patients. Clinical Immunol 146(2):77–83

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7. Millien VO, Lu W, Mak G, Yuan X, Knight JM, Porter P, Kheradmand F, Corry DB (2014) Airway Fibrinogenolysis and the initiation of allergic inflammation. AnnalsATS 11(Suppl 5):S277–S283 8. Porter PC, Lim DJ, Maskatia ZK, Mak G, Tsai CL, Citardi MJ, Fakhri S, Shaw JL, Fothergil A, Kheradmand F, Corry DB, Luong A (2014) Airway surface mycosis in chronic TH2associated airway disease. J Allergy Clin Immunol 134(2):325–331

Chapter 2 Use of the Litomosoides sigmodontis Infection Model of Filariasis to Study Type 2 Immunity A. Fulton, S. A. Babayan, and M. D. Taylor Abstract Helminth parasites infect over 2 billion people worldwide resulting in huge global health and economic burden. Helminths typically stimulate Type 2 immune responses and excel at manipulating or suppressing host-immune responses resulting in chronic infections that can last for years to decades. Alongside the importance for the development of helminth treatments and vaccines, studying helminth immunity has unraveled many fundamental aspects of Type 2 immunity and immune regulation with implications for the treatment of autoimmunity and Type 2-mediated diseases, such as allergies. Here we describe the maintenance and use of Litomosoides sigmodontis, a murine model for studying host-parasite interactions, Type 2 immunity, and vaccines to tissue-dwelling filarial nematodes, which in humans cause lymphatic filariasis (e.g., Brugia malayi) and onchocerciasis (Onchocerca volvulus). Key words Filariasis, Helminth, Parasite, Infection model, Type 2 immunity

1  Introduction Infection of resistant (e.g., C57BL/6) and susceptible (e.g., BALB/c) strains of inbred mice with the filarial nematode Litomosoides sigmodontis provides a useful laboratory model for studying lymphatic filariasis and onchocerciasis [1]. As with the majority of helminth parasites, L. sigmodontis predominantly stimulates a Type 2 immune response, and resistance is dependent upon IL-4 and IL-5 [2, 3]. The model was developed in the laboratory of Dr Odile Bain [4, 5] and is the only filarial parasite that can undergo its full life cycle within inbred laboratory mouse strains. Additionally, L. sigmodontis lives within the cavity between the pleural membranes covering the lungs and ribcage, making it very easy to recover parasites and immune cells from the infection site. This has made it an invaluable model for studying immunity to human filarial nematodes, such as Brugia spp., Wuchereria bancrofti, Loa loa, and Onchocerca volvulus, which are host specific and do not fully develop in mice. The L. sigmodontis model has been R. Lee Reinhardt (ed.), Type 2 Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1799, https://doi.org/10.1007/978-1-4939-7896-0_2, © Springer Science+Business Media, LLC, part of Springer Nature 2018

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successfully used to study antifilarial vaccines, drug development, infection dynamics, the hygiene hypothesis, and infection-related and fundamental aspects of Type 2 immunity, immune suppression, and immune regulation. The vector for L. sigmodontis is the mite, Ornithonyssus bacoti, which transmits the infective third-stage larvae (L3s) to the host during a blood meal. L3s migrate from the skin via the lymphatics arriving in the pleural cavity from day 3 postinfection (pi). Approximately half to two-thirds of larvae die before reaching the pleural cavity, the survivors remaining in the intrapleural space between the pleural membranes for the remainder of the infection. The L3 larvae molt to their fourth larval stage (L4) between 8 and 12 days postinfection (pi), before going through a final molt to the adult stage from days 25 to 30 pi [1]. The timings of the molts can vary between mouse strains. In susceptible strains of mice, male and female parasites develop fully, and the females start releasing the transmission stage microfilaria (Mf) from days 50 to 55 pi onward, which migrates to the blood and circulates in the bloodstream. A fully developed (patent) infection is defined by having detectable levels of Mf in the bloodstream, at which point the host can transmit infection, and is typically assayed at day 60 pi. Although L. sigmodontis establishes patent infections in mice, infection is cleared after day 80–100 pi. Long-term chronic infections can be maintained in both gerbils (Meriones unguiculatus) and its natural host the cotton rat (Sigmodon hispidus) and as a result are used as reservoirs to maintain the life cycle. Here we describe a protocol for maintaining the L. sigmodontis life cycle and performing experimental infections based on the original protocol developed in the laboratory of Dr Odile Bain [4]. In this protocol infections are performed by subcutaneous (s.c.) injection of L3 larvae to allow precise control over the infection dose. However, this has the disadvantage that it bypasses the natural transmission process via mite feeding. Natural infections can be performed by exposing animals to infected mites [6], although it is more difficult to control the infection dose. More recently, a third approach has been described in which gerbils are naturally infected by exposure to L. sigmodontis-infected mites, and the L3 parasites are purified from the pleural cavity days 5–6 pi and subsequently used to infect mice [7]. This results in the recovery of large numbers of L3 with higher infectivity. However, it also changes the course of infection with mice developing a patent infection earlier.

2  Materials 2.1  Mite Colony Maintenance

1. 500 mL plastic Erlenmeyer flasks.

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2. 20-micron gauze. 3. Fine grade animal bedding. 4. Elastic bands. 5. Climate chamber or room capable of maintaining 27 °C and 70% humidity. 6. Mouse cages and water bottles. 7. Mouse diet. 8. 70% ethanol. 9. Juvenile and/or adult mice. 2.2  L. sigmodontis Infections

1. Climate chamber or room capable of maintaining 27 °C and 70% humidity. 2. 500 mL Erlenmeyer flasks. 3. Fine grade animal bedding. 4. 20-micron gauze. 5. Elastic bands. 6. Distilled water. 7. Small paintbrush. 8. Trimmed paintbrush. 9. Metal sieve. 10. 500–1000 mL beakers. 11. Water moat (e.g., large tray or shallow sink). 12. Small trays or beakers for use as platforms. 13. Gerbil cages with bedding, diet, and water bottles. 14. Sealable plastic box with ventilated lid large enough to contain a gerbil cage. 15. Autoclaved glass tubes (approx. 1.5 cm in diameter, 6 cm tall). 16. Filter paper rectangles (1 cm × 2 cm). 17. Nonabsorbent cotton wool. 18. Tissue paper. 19. Cling film. 20. 250 mL glass beaker. 21. Dupont no. 5 forceps. 22. RPMI-1640 media with or without 10% sera (see Note 15). 23. Glass petri dish (5 cm). 24. 500 mL beaker of hot water with added detergent. 25. Dissection microscope with dark field (magnification × 6.3–60). 26. 15 cm glass Pasteur pipette and bulb.

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27. Syracuse watch glasses. 28. 1 mL disposable syringes. 29. 0.6 × 25 mm (23 gauge (G) × 1 in.) needles. 30. Insect forceps. 31. Bunsen burner. 32. L. sigmodontis-infected gerbils or cotton rats. 2.3  Quantitating L. sigmodontis Infection Levels and Recovery of Immune Cells

1. Razor or sharp scalpel. 2. 0.5 mm × 16 mm (25G × 5/8 in.) needle. 3. 1.5 mL microcentrifuge tubes. 4. FACS lysing buffer (BD). 5. Hemocytometer. 6. Dissection kit. 7. PBS and/or cell culture media (RPMI-1640, 5% FBS, 100 U/ mL penicillin, 0.1 mg/mL streptomycin, 2 mM l-glutamine). 8. 70% ethanol (20 dpi). 10. 60-micron cell strainers. 11. Dissection microscope with dark field (magnification × 6.3–60). 12. Glass tissue grinder.

3  Methods 3.1  Maintenance of Uninfected O. bacoti Mite Colonies

An O. bacoti colony must be maintained as a source of uninfected mites for the experimental infections in Subheading 3.2. O. bacoti colonies are kept at 27 °C and 70% humidity in a temperature-­ controlled room or climate chamber (see Note 1). Water moats are used to contain mites, and all mite work is carried out on raised platforms in a tray of water or dedicated sink. For decontamination, mite-exposed equipment and surfaces are washed with hot water and 70% ETOH (see Note 2). Bedding containing mites, or excess mites, are kept at −20 °C overnight to kill mites prior to disposal. O. bacoti are hematophagous and can be maintained by blood feeding on juvenile (Subheading 3.1.1) or adult (Subheading 3.1.2) mice. Appropriate personal protective equipment including gloves and dedicated laboratory coats should be worn at all time to avoid contact with mites and when animal handling (see Note 3).

3.1.1  Maintaining Mites in Flasks and Feeding with Juvenile Mice

Maintenance of mite colonies in flasks requires strict management to prevent overcrowding. This ensures effective feeding and minimizes adverse effects to juvenile mice (see Note 4). One colony

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flask started with 200 mites will provide approximately 900 mites after 3 weeks. 1. Start a new colony flask by adding approximately 50 g clean animal bedding to a 500 mL plastic Erlenmeyer flask. Add approximately 200 large female mites to the flask (collected in Subheading 3.2.1). Seal the flask with 20-micron mesh secured with two elastic bands. Keep flasks at 27 °C and 70% humidity (see Note 5). 2. Feed the mites twice weekly for 2 weeks by introducing the appropriate number of juvenile mice to the flask for up to 18 h. After 18 h remove the juveniles, assess their condition, and cull (see Note 6). 3. In the third week, mites are harvested from the flask (Subheading 3.2.1) for L. sigmodontis infections or to start new colony flasks. Freeze and then dispose of any excess mites (see Note 7). 3.1.2  Maintaining Mite Colony in Animal Cages and Feeding with Adult Mice

1. Mites are kept in mouse cages containing animal bedding. The cages are positioned on raised platforms surrounded by a water moat to contain the mites (Fig. 1). 2. To feed the mites, introduce one adult mouse to each cage once a week for 48–72 h (see Note 8). Remove and cull the mouse after feeding. 3. Bedding should be refreshed every 3–4 weeks by replacing half with clean bedding. Freeze discarded mite-contaminated bedding prior to disposal (see Note 9).

Fig. 1 Large shallow sinks and platforms used to create water moats around animal cages containing O. bacoti

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Fig. 2 Working area showing moat and raised platforms for collecting uninfected mites from colony flasks 3.2  L. sigmodontis Infections

3.2.1  Collecting Uninfected Mites

To produce infective L. sigmodontis stage 3 larvae (L3s), mites are collected from the uninfected colony (Subheading 3.2.1) and infected with L. sigmodontis by allowing them blood feed on L. sigmodontis-infected gerbils (Subheading 3.2.2). Mf take 12 days to develop into the infective L3 stage once ingested by the mite, and infective L3 larvae are dissected out of the mite 12–14 days post-ingestion for infections (Subheading 3.2.3). Using one gerbil to infect 450 mites should yield 600–1000 L. sigmodontis L3s. This is based on the recovery of 250–350 mites following exposure to infected gerbils. Mites will contain 2–3 L3s each with mite survival at approximately 90%. 1. Place 5–10 g of animal bedding into the required number of 500 mL plastic Erlenmeyer flasks. For collection of mites from colony flasks, use steps 2–4, and for collection of mites from animal cages, go to step 5. 2. Set up a working area with a container for collecting mites and waste container surrounded by a water moat (Fig. 2). 3. Sieve the contents of a colony flask (from Subheading 3.1.1) into the collecting container, tapping the sieve on the side of the collecting container so that the bedding is retained in the sieve and mites fall through. Discard the bedding into the waste beaker. 4. Tilt and tap the collecting container so that the mites climb up one side. Using a paintbrush dampened in distilled water, pick 450 large female mites (Fig. 3) from the collecting container into the flask (see Note 10).

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Fig. 3 Large female mites and smaller protonymphs

5. Pick mites directly from animal cages using a paintbrush dampened in distilled water during the 48–72 h that adult mice are present (see Notes 10 and 11). Place 450 large female mites (Fig. 3) into each flask. Go to step 6. 6. Seal the flask with gauze and elastic bands pulling the gauze tight (see Note 5). Roll the animal bedding around the sides of the flask to remove any water droplets. 7. Keep the collected mites at 27 °C and 70% humidity for 6–9 days without feeding before infecting them with L. sigmodontis (Subheading 3.2.2). The starvation period ensures mites feed well when exposed to L. sigmodontis-infected gerbils resulting in higher infection levels. 3.2.2  Infection of Mites with L. sigmodontis

1. Select a gerbil that has been infected with L. sigmodontis for at least 90 days and has detectable microfilaria within the blood (Subheading 3.3.2), preferably greater than 800 Mf/ mL. Gerbil weights are taken before and after the procedure to monitor for adverse health effects (see Note 12). 2. To prevent mite escape, place the gerbil and its cage inside a sealable ventilated box. Add a small amount of bedding from the gerbil cage, and provide food and water (see Note 13). 3. Pour 450 uninfected mites and their bedding (collected in Subheading 3.2.1) into the gerbil cage and seal the plastic box. Leave the mites to feed on the infected gerbil for 12–18 h.

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Fig. 4 (a) O. bacoti infected with L. sigmodontis are collected into glass tubes. (b) The tubes are placed into beaker with damp tissue paper to humidify the local environment

4. Remove the gerbil from the cage, tapping the box on the sides and top prior to opening to dislodge any mites. Tap the lid of the animal cage to dislodge any mites back into the cage. Lift the gerbil cage out of the box, and tap to move the bedding to one side. Sieve the mites from the bedding into a collecting box (as in step 3 of Subheading 3.2.1) for picking. 5. Collect the infected mites from the ventilated box, and those that were sieved from the bedding, into glass tubes containing a small square of folded filter paper. Use a fine dry paintbrush to collect 50 large well-fed female mites into each tube. Use your gloved thumb to seal the tube to prevent mites escaping while collecting. To seal the tubes, form a plug using nonabsorbent cotton wool. The plug should be dense and fit tightly in the tube (Fig. 4a) (see Note 14). 6. Secure the tubes together with an elastic band. Dampen two sheets of tissue paper, and place in bottom of plastic beaker to maintain a local humidified environment. Place tubes in the beaker on top of tissue paper, and secure cling film over the top with an elastic band (Fig. 4b). Ensure tubes remain upright. Keep at 27 °C and 70% humidity for between 12 and 15 days to allow L. sigmodontis parasites to develop into the infective L3. 7. Decontaminate mite-exposed equipment, bedding, and surfaces. Equipment and surfaces are washed with hot water and ethanol. Place contaminated bedding at −20 °C overnight prior to disposal. 3.2.3  Infection of Animals with L. sigmodontis

1. Set up a working area surrounded by a water moat (Fig. 5a). Working area should include a large beaker full of hot water for decontamination of mite-contaminated items, a glass petri

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Fig. 5 (a) Setup of water moat and platforms for collection of mite dissection. (b) Drawn-out glass Pasteur pipettes. (c) Syracuse watch glasses

dish containing approximately 5 mL of RPMI media with or without 5–10% sera, and an upside-down 100 mL beaker to act as a small surface on which to collect mites (see Note 15). 2. Take a tube containing L. sigmodontis-infected mites (Subheading 3.2.2), tap sharply to knock mites down to the bottom, and remove the cotton wool. Place the tube upside down on the upside-down beaker, and using Dumont no. 5 forceps, carefully collect any mites remaining on the cotton wool, and tap mites into the petri dish containing RPMI without putting forceps into media. Discard cotton wool into the beaker of hot water (see Note 16). 3. Gently remove the filter paper from the tubes with forceps, pick the mites off with a fine paintbrush (moistened in RPMI), and place into the RPMI-filled petri dish. Place the filter paper into beaker of hot water together with cotton wool to kill any uncollected mites. 4. Keeping the tube upside down on the glass beaker, dislodge the remaining mites by tapping the tube against the beaker. Collect the live large adult mites and transfer to the RPMI-­ filled petri dish. 5. Repeat for remaining tubes collecting all mites into the same petri dish.

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6. Using a dissection microscope, crush the mites gently with fine forceps (Dumont no. 5), checking the abdomen has burst sufficiently to allow L3s to emerge (see Note 17). 7. Use a Bunsen burner to melt and draw out a glass Pasteur pipette to create a very fine end. Place the neck of the pipette into the Bunsen flame with the tip protruding from the flame and your finger just below the tip. When the tip falls onto your finger, pull the tip to lengthen and reduce the diameter, and simultaneously remove from the flame. Break off the end of the pipette to leave a hole just large enough to collect L3 (Fig. 5b). 8. Infective doses for each animal are counted out and collected individually. Typically, mice are infected with 20–40 L. sigmodontis L3 for experimental purposes, and gerbils are infected with 100 L. sigmodontis L3 for life cycle maintenance. Use the Pasteur pipette to collect the required number of L3 for each dose, and transfer them to a Syracuse watch glass (Fig. 5c) (see Note 18). 9. Let the L3 form a cluster in the bottom of the Syracuse watch glass. The L3s can be recounted using a dissection microscope at this point to double check the accuracy of the dose. Take up the L3 in a 1 mL syringe with 0.6 × 25 mm needle, aiming to collect the dose in 100–200 μL of RPMI media. To prevent the L3 clumping and sticking in the needle, finish by sucking up approximately 100 μL of air. 10. Infect gerbils intraperitoneally for life cycle maintenance and mice subcutaneously for experimental infections using a 0.6 × 25 mm needle (see Note 19). 3.3  Quantitating L. sigmodontis Infection Levels and Recovery of Immune Cells

3.3.1  Recovering Adult Parasites and Immune Cells from the Pleural Cavity of Mice

L. sigmodontis parasites migrate from the skin to the pleural cavity over the first 3–6 days of infection and can be isolated from the pleural cavity by pleural lavage from day 4 pi onward, along with immune cells from the infection site (Subheading 3.3.1). Adult female L. sigmodontis parasites start releasing Mf from day 50 pi, which can be measured by blood sampling and pleural lavage (Subheading 3.3.2). Skin-draining LN can be used to sample the immune response for the first 7–12 day pi. Pooled parathymic and posterior mediastinal lymph nodes can be used to sample the infection site (see Note 20). 1. Once the animal has been culled, open the peritoneal cavity taking care not to damage the diaphragm, or cut into the pleural cavity. Pull organs down away from the diaphragm. Cut off the xiphoid process from the sternum, which should cause a hole to open in the diaphragm next to the sternum. Lavage the peritoneal cavity 1 mL at a time with 10 mL PBS for the col-

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lection of parasites. Use culture media if both parasites and cells are to be collected (see Note 21). 2. Separate cells and parasites. Prior to day 10 pi, pour lavage fluid into a petri dish under a dissection microscope, and suck up parasites using a P20 pipette. From day 40 pi, parasites can be collected using insect forceps. Between days 10 and 40 of infection, parasites and cells can be separated by passing the lavage fluid through a 60-micron cell strainer. The parasites will remain in the cell strainer, while the cells will pass through. However, at later stages of infection, there is a risk of damaging parasite morphology using a cell strainer. Collect parasites into PBS and retain lavage fluid containing immune cells. 3. Count the number of parasites under a dissection microscope (see Note 22). 4. Fixation of parasites (if required). Prior to day 20 pi, fix parasites in 4% formalin. After day 20 pi, fix parasites using warm 70% ethanol (see Note 23). 3.3.2  Quantifying Levels of Mf in the Blood and Pleural Cavity

1. Aliquot 95 or 500 μL BD FACS lysing buffer (prepared as per manufacturer’s instructions) into a 1.5 mL Eppendorf tube to assess microfilaria in gerbils or mice, respectively. One tube per animal (see Note 24). 2. To quantify microfilaria in the blood, insert a needle into the tail vein, and collect 5 μL blood from gerbils or 30 μL blood from mice into the Eppendorf tube containing FACS lysing buffer. Shave the tail of gerbils prior to collection with a razor or scalpel. To quantify the Mf in the pleural cavity of mice, resuspend pleural lavage fluid in 5 mL, mix well, and sample 20 μL into an Eppendorf tube with FACS lysing buffer. 3. Use a hemocytometer and microscope to count microfilaria in gerbil samples. To count Mf in mouse samples, spin down the sample (17,000 g, 5 min), and remove as much liquid as possible with a pipette. Resuspend in the remaining liquid and smear onto a slide. Count all the Mf in the sample under an inverted microscope.

3.3.3  Preparation of L. sigmodontis Antigens

Soluble L. sigmodontis antigens can be prepared for in vitro immune cell restimulations and measuring L. sigmodontis-specific antibodies via ELISA. 1. On ice, crush L. sigmodontis parasites in sterile PBS in a glass tissue grinder. Remove the insoluble fraction by centrifuging at 17000 g for 5 min at 4 °C and collecting soluble fraction. Repeat 2–3 times to remove all insoluble material.

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2. Check the protein concentration using standard methods (e.g., Bradford assay), and adjust to the required concentration in sterile PBS. Aliquot and store at −70 °C. 3. Coat ELISA plates with L. sigmodontis antigen at 5 μg/mL for antigen-specific antibody ELISAs, and use sterile-filtered L. sigmodontis antigen at 10 μg/mL for in vitro immune restimulations.

4  Notes 1. Although O. bacoti requires high temperature and humidity (approx. 27 °C, 70% humidity), it does not have to be as precise as obtained by a climate chamber or specialized room. Colonies can be maintained without climate chambers or specialized facilities so long as the temperature and humidity levels can be raised. 2. Detergent can leave a residue that impairs mite survival. It is better to use non-detergent-based methods for cleaning equipment involved with mite maintenance. 3. O. bacoti will bite humans, although it is not able to survive feeding on humans. Precautions should be taken to prevent contact with mites and mite escape. O. bacoti tend to climb and will travel up equipment being held. Work quickly, frequently wash off tools (e.g., forceps) with water or place under water, and visually inspect gloves at frequent intervals to check for mites. Wear light-colored gloves and protective personal equipment to allow mites to be seen easily. Double-sided sticky tape can be placed around the cuff of gloves to form a mite barrier. 4. Overcrowding of mites in colony flasks is a major problem. It results in juvenile mice receiving a large number of bites that can cause unnecessary morbidity and may lead to mortality. It also results in mites failing to feed adequately, impairing their development and their survival at later stages in the protocol. The clinical condition of juvenile mice should be assessed immediately on their removal from the flasks, including mobility, anemia, and skin integrity. If any of these are poor, then steps should be taken to reduce the number of mites. Increasing the number of juveniles will not solve the overcrowding problem. The number of mites can also be directly assessed when they are sieved out of the bedding (Subheading 3.2.1). Subheading 3.1.1 should allow feeding of mites with negligible adverse effects for the juvenile mice. 5. Animal bedding should be light colored so that mites stand out against it and is best to be a uniform size for ease of sieving

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and collecting mites. We use a fine grade bedding (Gold Chip, LBS Biotechnology). For redundancy use two elastic to prevent the flask unsealing if one breaks. The gauze must be pulled tightly over the top of the flasks; otherwise mites, especially smaller protonymphs, will gather within creases in the gauze on the outside rim making escape more likely. Replace the gauze after each opening, and wash gauzes thoroughly in hot water. 6. Mite feeding can result in trauma to the juveniles, and the severity of trauma relates to the size of the juvenile, strain, and number of mites in the flask. Juvenile mice should be P8–P10 and over 6 g in weight. CD1 juveniles are a recommended strain. Mite feeding efficiency can be reduced if older juveniles are used, while younger or smaller juveniles are more susceptible to adverse health effects. In a well-controlled colony, the required juvenile numbers are predictable, and mite feeding can be performed with minimal adverse effects for the juvenile mice. Typically, two juveniles per flask are used for newly split flasks (feeds 1 and 2 in week 1), increasing to four juveniles per flask as mite numbers increase (feeds 3 and 4 in week 2). If using a less robust strain, such as C57BL/6, then juvenile numbers should be increased, e.g., to three and five, respectively. If mites are not required for L. sigmodontis infections, then the four feeds can be given once a week over 4 weeks. 7. Do not keep a colony flask longer than the four feeds as the mites will over expand. Start new flasks to maintain the colony. 8. Exposure to mites can result in stress in adult mice, and adding more than one mouse per cage can result in fighting and resource guarding. The condition of adult mice should be monitored for the period that they are exposed to mites. 9. As the mites live and lay eggs in the animal bedding, it is important not to disturb or change the bedding too frequently. As adult mice are only kept in the cage for 48–72 h, the bedding can be left for 3–4 weeks before partially renewing. The water bottles and cage lids are a favorite egg-laying site and so should only be changed when required. If multiple cages are used, then bridges can be made between cages to allow the migration and mixing of the mite colony. The moats should be cleared and cleaned regularly. 10. It is easier to pick up mites with a damp, rather than dry, paintbrush. However, if the brush is too wet, then this can result in mites trapped in water droplets, and water in the flasks can promote the growth of mold. Use the largest (female) mites for L. sigmodontis infections and starting new colony flasks. Transferring too many small protonymphs (Fig. 3) can result

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in overcrowding of colony flasks and unnecessarily increase the adverse effects suffered by gerbils. 11. The fed female mites seek egg-laying sites around the top of the cage, under the water bottle, and behind cage card, which are a good location to collect mites. 12. Although more L3s can be produced by using gerbils with higher numbers of Mf/mL, any gerbil with detectable Mf in the blood can be used. A gerbil can be reused to infect different batches of mites, although the number of L3s recovered per mite will decrease with each exposure. Typically, one gerbil can be used to infect four batches of mites before a significant decline in L3 recovery. Gerbils experience a transient weight loss following exposure to mites, and their condition should be monitored to verify that they have fully recovered before reexposure to mites. Resting gerbils for 2 weeks between exposures is recommended although 7 days can be sufficient. A system of rotating gerbils can be used to provide a constant supply of infectious L3. 13. The box vents need to let in air while preventing mite escape. If taking an infected gerbil from a cage containing multiple gerbils, it is important to co-transfer some cage bedding so that the gerbil recognizes their cage mates on reintroduction. Only put a small amount of food directly into the cage, sufficient for 18 h, as the presence of excess food pellets will make it more difficult to collect mites. 14. The paintbrush must be dry as any moisture in the tubes will cause mold growth killing the mites. Use a fine brush and splay the hairs to assist with picking. Avoid introducing bedding and animal dander to the tubes. 15. Once dissected out of the mite, L3 larvae only remain infective for a few hours, and infections should be performed as soon as possible after dissections. L3 collected in RPMI will remain infective for 3–5 h. Adding 10% fetal bovine serum or horse serum to the RPMI increases the length of time they remain infective. If planning to culture immune cells in vitro, then different types of sera should be used for inoculations and in vitro assays; otherwise a recall response may be observed against serum antigens. We prepare L3s in horse serum and perform in vitro restimulations with fetal bovine serum or mouse serum. 16. It is best to use a paintbrush to collect mites as mites will stick to forceps when placed in media and try to climb up the ­forceps. However, forceps are used for picking mites off the cotton wool. Trimming the paintbrush to a fine point aids the collection of mites.

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17. Once the mites are dissected, leave for around 30 min to allow all the L3s to emerge from mites. They will move to the edge of the dish making them easier to collect. 18. Collecting the L3s from around the edges and transferring to a clean petri dish can help separate L3s from mite debris and make collection easier. Approximately two-thirds of injected larvae are killed within 4 days of inoculation. Thus, infecting susceptible BALB/c mice with 20 or 40 L3s will result in the establishment of an average of 7 and 13 L3s, respectively. We recommend injecting 40 L3s for experiments assessing changes in parasite recovery (e.g., vaccination). Injecting 20 L3s is sufficient for experiments which only require the recruitment of immune cells. Using less than 20 L3s increases the risk of single sex infections, which will not result in microfilaria. Infecting with higher L3 numbers increases the magnitude of the immune response but can result in impaired parasite development [5, 8]. 19. S.c. infections of L. sigmodontis can be performed anywhere on the mouse. We recommend the nape of the neck as it allows a larger volume of liquid to be injected and is drained by the brachial LN, which are relatively large and easy to collect. 20. The precise timings of the L. sigmodontis life-cycle shows natural variation over time, and not all mice develop a patent infection or at the same time. In susceptible BALB/c mice, between 50% and 100% of mice can be expected to develop patent infections with female mice being more susceptible than male mice [9]. Although Mf can be detected from day 50 pi, this is not always the case. Typically, day 60 pi is considered the time point by which Mf should be robustly detectable if an infection has fully developed. If blood Mf have not been detected by day 70 pi, it is unlikely that a patent infection will develop. 21. It is recommended to exsanguinate mice prior to lavage to reduce blood leakage into the pleural cavity. Not doing this can result in coagulation and loss of immune cells. The presence of blood will also obscure the LN within the pleural cavity. It is best not to cut a hole directly in the diaphragm as it will enlarge during lavage resulting in the liquid escaping. Cut off as much as the xiphoid process as possible. If a hole does not immediately appear, then it can be opened by brushing up with tweezers just under the cut area. L. sigmodontis parasites are fragile and will be damaged if repeatedly aspirated up and down using a Pasteur pipette. Care must be taken if morphological analysis is required. 22. L. sigmodontis parasites tend to curl up into ball of complex knots. Placing on ice will slow down their movement while alive allowing them to be teased apart with paintbrushes.

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23. The use of formalin to fix large L. sigmodontis parasites results in contraction of the parasite making morphological analysis more difficult. Warm ethanol fixes the parasite while better retaining morphology. Ethanol can also be used to fix the earlier parasite stages. 24. Red blood cells must be lysed before fixation; otherwise they will obscure the Mf resulting in inaccurate counts. BD FACS lysing buffer simultaneously lyses red blood cells and fixes Mf for counting at a later date.

Acknowledgments Many thanks to Brian Chan for proofreading. The protocols in this paper were developed during research funded by the MRC UK, Wellcome Trust, and European Union. References 1. Hoffmann W, Petit G, Schulz-Key H, Taylor D, Bain O, Le Goff L (2000) Litomosoides sigmodontis in mice: reappraisal of an old model for filarial research. Parasitol Today 16:387–389 2. Le Goff L, Lamb TJ, Graham AL, Harcus Y, Allen JE (2002) IL-4 is required to prevent filarial nematode development in resistant but not susceptible strains of mice. Int J Parasitol 32:1277–1284 3. Volkmann L, Bain O, Saeftel M, Specht S, Fischer K, Brombacher F, Matthaei KI, Hoerauf A (2003) Murine filariasis: interleukin 4 and interleukin 5 lead to containment of different worm developmental stages. Med Microbiol Immunol 192:23–31 4. Petit G, Diagne M, Marechal P, Owen D, Taylor D, Bain O (1992) Maturation of the filaria Litomosoides sigmodontis in BALB/c mice; comparative susceptibility of nine other inbred strains. Ann Parasitol Hum Comp 67:144–150 5. Marechal P, Le Goff L, Petit G, Diagne M, Taylor DW, Bain O (1996) The fate of the filaria Litomosoides sigmodontis in susceptible and naturally resistant mice. Parasite 3:25–31

6. Al-Qaoud KM, Taubert A, Zahner H, Fleischer B, Hoerauf A (1997) Infection of BALB/c mice with the filarial nematode Litomosoides sigmodontis: role of CD4+ T cells in controlling larval development. Infect Immun 65(6):2457–2461 7. Hubner MP, Torrero MN, McCall JW, Mitre E (2009) Litomosoides sigmodontis: a simple method to infect mice with L3 larvae obtained from the pleural space of recently infected jirds (Meriones unguiculatus). Exp Parasitol 123:95–98 8. Babayan S, Attout T, Specht S, Hoerauf A, Snounou G, Renia L, Korenaga M, Bain O, Martin C (2005) Increased early local immune responses and altered worm development in high-dose infections of mice susceptible to the filaria Litomosoides sigmodontis. Med Microbiol Immunol 194:151–162 9. Graham AL, Taylor MD, Le Goff L, Lamb TJ, Magennis M, Allen JE (2005) Quantitative appraisal of murine filariasis confirms host strain differences but reveals that BALB/c females are more susceptible than males to Litomosoides sigmodontis. Microbes Infect 7:612–618

Chapter 3 Production of Hymenolepis diminuta in the Laboratory: An Old Research Tool with New Clinical Applications Min Zhang, Amanda J. Mathew, and William Parker Abstract Hymenolepis diminuta, the rat tapeworm, was first described in 1819 by Rudolphi and was studied ­extensively in several laboratories during the mid to latter part of the twentieth century. More recently, the primary use of the organism had been for educational purposes. The organisms require an intermediate insect host to complete their life cycle, making them non-transmissible to other rats or to humans under typical laboratory or educational environments. The organisms effectively colonize rats, but not humans or mice, and are easily maintained in laboratory. They are, with exceedingly rare exceptions, benign (e.g., nonparasitic) in humans, mice, and laboratory rats. Although the benign character of the helminth makes it ideal for educational purposes, the fact that no pathology is associated with colonization has led to decreased interest in the H. diminuta as a model for modern research where efforts are largely motivated by interests in medicine and health. However, more recently work with the “biota alteration” model of inflammatory disease has established that reintroduction of helminths into Western society, a practice often referred to as “helminthic therapy,” is potentially a way of lowering inflammation without compromising immune function. For this effort, the lack of pathology and benign nature of the organism makes H. diminuta an ideal subject for study. In this chapter, we describe production of H. diminuta using l­ aboratory rats and introduction of the organisms into laboratory mice as a model for their effects in humans. Key words Helminthic therapy, Helminth, Biological therapeutic, Inflammation, Anti-inflammatory

1  Introduction Animal models for the study of inflammatory disease are extremely helpful to biomedical research efforts. This is increasingly true as the prevalence of a wide range of inflammatory-related diseases continues to rise in Western society [1–3]. Inflammation-related diseases of Westernization include a broad range of allergic disorders, autoimmune conditions, digestive diseases, and ­ ­neuropsychiatric disorders. An intuitive view is that models aimed at dealing with the root causes of these inflammatory disease will be the most beneficial to medical progress [4]. These root causes include inflammatory diets, sedentary lifestyles, chronic ­psychological stress, and vitamin D deficiency. At the same time, R. Lee Reinhardt (ed.), Type 2 Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1799, https://doi.org/10.1007/978-1-4939-7896-0_3, © Springer Science+Business Media, LLC, part of Springer Nature 2018

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changes in the human biota, the life associated with the ecosystem of the human body, are being recognized as important c­ ontributors to the ever-­increasing prevalence of inflammation-related diseases in Western society [1, 2]. Among the most impactful changes to the biota is the virtual annihilation of helminths from humans [4]. It is now becoming apparent that helminths, ubiquitous symbionts until very recently in human history, are important for immune function and stabilization. With that in mind, animal models to study the use of helminths as therapeutic agents in clinically r­ elevant scenarios are of considerable interest. Hymenolepis diminuta, the rat tapeworm, is now one of the most widely used helminths for therapeutic purposes [5, 6]. However, H. diminuta is not currently approved by any regulatory agency for therapeutic use, and the study of the effects of this ­helminth on humans and even on laboratory animals is in its infancy. Since helminthic therapy effectively alleviates many of the effects of “biota alteration,” one of the primary causes of disease in Western society, it is expected that the study of a wide range of helminths, including H. diminuta, will increase in the foreseeable future. Similar to many rodent models of helminth colonization, H. diminuta exposure leads to increased type 2 cytokines in the ­intestine of both mice and rats. Although IL-4 appears to be the dominant cytokine at low worm burdens in the tolerant rat model, administration of 50 worms leads to a significant increase in IL-13 mRNA and protein production [7]. This increase in cytokine ­correlates with enhanced mucus production, goblet cell h ­ yperplasia, and worm expulsion. Mice are less tolerant to prolonged colonization when exposed to low-dose H. diminuta. Mice ­ ­colonized with five cysticercoids mount a rapid and robust type 2 immune response which leads to clearance of adult worms in both Balb/c and C57BL6 backgrounds [8, 9]. Expulsion in each of these cases is dependent on T cells and is dominated by the ­production of IL-4 and IL-13. Importantly, signaling induced by IL-4 and IL-13 is required for worm expulsion, mucus ­production, and goblet cell hyperplasia as STAT6-deficiency, a key factor in IL-4 receptor signaling, substantially prolongs adult worm ­engraftment [10]. H. diminuta is easily maintained in the laboratory with no ­specialized equipment (see Note 1). The organism is so easily maintained that it is now used in middle school and high school biology classes for educational purposes. Adding further to its utility as a laboratory animal model, the safety profile of the organism is excellent, posing no hazards to humans working with the organisms or to the ­laboratory rats that serve as their primary hosts (see Notes 2 and 3). Unlike most roundworms and some other flatworms, H. diminuta lives exclusively in the lumen of the gut. That is, it does

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not breach the epithelium of the gut, but rather remains in the fecal stream. The organisms have no mechanism by which b ­ reaching the epithelium is possible and essentially “swim” in the intestine [11]. Further, they do not form lesions at the site of attachment in their natural hosts [11]. In humans, helminthic therapy with H. diminuta is ­accomplished by repeated exposure to the cysticercoid life stage of the organisms at 1–6-week intervals [5, 6]. Unlike laboratory rats, neither humans nor mice can host mature, reproducing H. diminuta. For this reason, a mouse model may be the most ­clinically relevant for studying the use of H. diminuta for therapy in humans. In this review, methods are described for (a) ­maintenance in H. diminuta in the laboratory rat and (b) use of mice as a model for the effects of helminthic therapy with H. diminuta in humans (see Notes 4–6).

2  Materials 2.1  Maintaining the Beetles (Tenebrio molitor)

1. Quaker brand, 100% natural whole grain oats: follow the ­directions on the container to store the oats, and discard if mold is visible. 2. Freshly washed organic celery: the celery must be certifiably organic and stored in a refrigerator. Remove thin ends or leafy parts before storing. 3. Nutritional yeast (Bragg Live Food Products, Santa Barbara, CA) at a ratio of approximately 0.06 g nutritional yeast per gram of oats: add only to the nursery (see point 5 below for definition of the nursery). 4. Small plastic containers: reusable food-grade plastic containers are used as housing adult beetles. A typical setup is shown in Fig. 1. These containers are modified by cutting a hole in the lid and gluing a screen mesh onto the lid. In this figure, six batches of beetles are shown (two in the front, one open). See Methods for the definition of a “batch.” 5. Nursery: a container typically larger than that used to contain adult beetles, used to contain mealworms and pupae. It is modified by cutting a hole in the lid and gluing a screen mesh onto the lid. 6. Plastic dome enrichment: these are made from a section of a BPA-free (polypropylene) plastic drinking cup for the beetles in each batch to hide under. The surface of the cup is scored with sandpaper so that the beetles can climb on the plastic. 7. Dehumidifier: is recommended for beetle housing. Our laboratory uses an Eva-Dry EDV-100 petite dehumidifier in each “isolator.”

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Fig. 1 Beetle housing conditions in the laboratory. In this setup, a front opening box is used as an “isolator,” and individual “batches” of beetles loaded with HDCs are kept in food-quality containers modified with wire screens to allow air to circulate in each batch. A dehumidifier is visible in the back, right of the container. The four batch containers and dehumidifier are resting on a mealworm nursery container (bottom of isolator), where mealworms are allowed to pupate and mature to adults

8. A chemical fume hood to store the isolators. 9. Unfitted disposable dust mask is adequate for most purposes; however, fitted respiratory protective equipment may be needed for individuals who are sensitive to dust or mold or who spend considerable time maintaining beetle colonies. 2.2  Maintaining the Rats

1. Sprague Dawley rats (Harlan Sprague Dawley, Indianapolis, IN). 2. A disposable fine tip transfer pipette (Samco Scientific Corp). 3. Maintained in AAALAC-approved barrier facilities at Duke University Medical Center in accordance with institutional guidelines. All animal care and procedures were approved by the institutional animal care and use committee at Duke University (see Note 2).

2.3  Isolating H. diminuta Cysticercoids (HDCs)

1. 0.6% saline solution: distilled water, solid NaCl. 2. A disposable fine tip transfer pipette (Samco Scientific Corp). 3. Sterile petri dishes. 4. Dissecting microscope with 40× magnification.

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3  Methods 3.1  Maintenance of Grain Beetles (Tenebrio molitor) in the Laboratory

1. Build the nursery out of a large plastic container and add the worms and pupae. They are provided with fresh Quaker oats, organic celery, and nutritional yeast seasoning. The oats to yeast ratio is 10:1 in the nursery, with celery as needed. Feeding the beetles a richer diet than oats and celery has reportedly increased the production of beetles but has decreased the therapeutic effect of the HDCs (see Note 7). 2. To prevent the celery from becoming buried in the oat/yeast mix, toothpicks are inserted into the celery. This will also prevent mold from developing (see Notes 8 and 9). 3. Newly hatched beetles are collected over a 2-week period and placed into a new “batch” box, which is labeled. We define one “batch” of HDCs as all HDCs contained in a group of beetles that is “loaded” with HDCs at the same time in the same container. Each batch of beetles consists of between 15 and 70 beetles and is loaded as described below in Subheading 3.2. This batch box will contain roughly 60 grams of fresh Quaker oats. Add the plastic dome enrichment and organic celery. The celery should be at a minimum of 0.23 g/cm2, where the cm2 reflects the area (length by width) of the container. 4. Each “batch” is kept in a separate container, or isolator, as depicted in Fig. 1. The isolator (the large green container in Fig. 1) is a container meant to house multiple batches of beetles and a nursery. Each isolator is outfitted with a dehumidifier and a large front opening for easy access to the batch boxes. 5. The celery in each batch box is changed twice weekly. In addition, the boxes are monitored for pupae or mold during this time.

3.2  Loading Grain Beetles with HDCs

Although the process is straightforward, substantial variation in the number of eggs eaten by individual beetles (and thus the eventual numbers of HDCs per beetle) is observed. It is possible to feed more than 100 eggs to an individual beetle, but the procedure can be adjusted so that an average of between 15 and 70 eggs per beetle are ingested (see Note 10). Since the eggs are produced by the adults living in rats and are present in the rat feces, the procedure involves feeding of rat feces to beetles. 1. To create a “batch” of HDC-loaded beetles, newly hatched (within 5–6 weeks) beetles are selected (see Note 11). Beetles are moved to a new container, or “starvation chamber,” without access to food or a water source (celery) for 2 days. Water and food deprivation ensures that the beetles will eat rat fecal pellets containing H. diminuta eggs.

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2. Harvest fecal pellets from rats colonized with Hymenolepis diminuta. After collection from cages, feces containing Hymenolepis diminuta eggs are stored for no more than 2 days at room temperature prior to use. Do not store feces at 4 °C. 3. Prior to placing the droppings into the starvation chamber, add water dropwise if the droppings appear dry. Drying rat fecal p ­ ellets will kill the eggs. One or more drops of water per pellet are typically added immediately before feeding to the beetles. One pellet will feed a few dozen beetles, and five pellets will feed several hundred beetles. Keep the beetles in the “starvation chambers” with the rat fecal pellets for another 2 days. 4. After feeding for 2 days, the beetles should be removed from any remaining feces and placed in a fresh container with oats and organic celery. 3.3  Maintenance in the Rat

The rats must be maintained with HDCs isolated from grain beetles to ensure production of H. diminuta eggs (see Note 12). For maintenance in a mouse model, see Notes 4–6. 1. The HDCs are isolated from the grain beetles (see Subheading 3.4) and are suspended in 0.6% saline. They are then placed in a disposable “fine tip transfer pipette” (Samco Scientific Corp) as described below. 2. The pipette tip is placed in the rat’s mouth, on the tongue, while gently holding the rat, and the liquid is expelled. The rat is watched carefully during this time to ensure that it swallows. Four to five HDCs are administered per rat. There is generally no need for oral gavage, as the rodents will readily accept the HDCs when fed by an experienced technician. 3. After at least 21 days following the ingestion of HDCs and then monthly thereafter, the feces should be checked for HDC eggs using a standard fecal flotation test. A positive result of a fecal flotation test is shown in Fig. 2, and most veterinarians working in animal research facilities will be able to help with positive identification of tapeworm colonization.

3.4  Isolation of HDCs from Grain Beetles

1. The beetle is placed in a sterile petri dish. The beetle can be placed in the upside-down or upright position for this step of the procedure. 2. The head (or the thorax plus the head, it does not matter) is removed with a swift motion using a clean scalpel or knife. 3. The legs are removed using forceps and dissecting scissors. 4. The wings are removed using forceps and dissecting scissors.

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Fig. 2 Observation of HD eggs by fecal flotation. Rats colonized with H. diminuta have copious amounts of eggs in their fecal material that are readily observed after fecal flotation

Fig. 3 Dissection of HDCs. Although eggs are clearly visible in this photo, HDCs are not discernable without the use of a microscope. The abdomen of a single grain beetle, bottom side up (with legs and wings removed), is shown. In this specimen, the contents of the abdomen have been partially removed and dispersed in the 0.6% saline solution, as described in the Methods

5. The abdomen of the beetle is placed topside (wing side) up in a new sterile petri dish, and about 3 mL of 0.6% saline solution is added (Fig. 3). 6. Using two pairs of tweezers and, if desired, a small knife, the interior of the beetle abdomen is gently scraped out into the saline solution (Fig. 3). This frees the HDCs from the beetle. The HDCs are visible to the naked eye, but will not be

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­ iscernable from other components of the beetle’s abdomen d without a microscope. 7. Once the contents of the beetle’s abdominal cavity are ­suspended in solution on a petri dish, the dish is placed under a dissecting microscope, and the HDCs are harvested. A 20- to 40-fold total magnification (ocular + objective lens) is desirable to obtain a balance between identification of individual HDCs and observation of a broad field. 8. A disposable “fine tip transfer pipette” (Samco Scientific Corp) is used in collection of HDCs.

4  Notes 1. The life cycle of H. diminuta in the laboratory is shown in Fig.  4. Adult, egg-laying helminths are maintained in the ­laboratory using rats as the primary hosts. The eggs, present in the feces of the rats, are consumed by grain beetles, Tenebrio molitor, which serve as the intermediate host in the laboratory. H. diminuta achieve a distinctive, cysticercoid stage in the extraintestinal space of the abdomen of the beetle, which is readily extracted for inoculation of additional laboratory rats. This stage is often referred to as an “HDC” (Hymenolepis diminuta cysticercoids), although the acronym HDC is sometimes used as a general name for H. diminuta by ­ ­individuals using helminthic therapy. In this manuscript, HDC (or HDCs, plural) will refer strictly to the cysticercoids stage of H. diminuta. Mature HDCs have been used for therapeutic purposes in humans and can be used in laboratory mice as a model for its therapeutic effect in humans (Fig. 4). 2. Older beetles can also be loaded, but they will not live as long after they are colonized by the HDCs, so they will be of less utility in further studies. It will take 5–7 weeks for the HDCs to mature once the beetles have ingested the eggs. 3. Results of loading will vary depending on the number of eggs in the pellets and other factors such as the relative humidity, and the procedure will be adjusted to maintain an average colonization rate between 15 and 70 HDCs per beetle ­ ­(average) by increasing or decreasing the number of pellets used per beetle. If the procedure yields more than 70 HDCs per beetle on average, then the number of beetles fed by a single pellet will be increased. If the procedure yields less than 15 HDCs per beetle on average, then the number of beetles per pellet can be decreased. 4. Rats need to be exposed to Hymenolepis diminuta once every few months at most, and sometimes colonization will last for the lifetime of the rat. (Hymenolepis diminuta will live for 4–8

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Fig. 4 Overall schematic of laboratory use of H. diminuta (HD), including maintenance of adults in rats and therapeutic use in humans or in mouse models

months in some rats and longer in others.) This depends in large part upon the breed of the rat, but it can vary for unknown reasons. Thus, colonization should be evaluated periodically, and rats should be recolonized as needed. 5. Rat-to-rat transmission of HDCs is not possible under s­ tandard laboratory conditions, and thus no special housing of the rats is required. The safety and training procedures needed to work with laboratory rodents are well documented and part of ­routine practice in any laboratory. 6. The risk of H. diminuta transmission to humans following exposure is negligible and requires no special safety ­precautions. The effects of ingestion of the larval stage of the organisms appear to be generally beneficial rather than harmful in humans based on sociomedical studies [5, 6], indicating that no p ­ articular safety precautions are warranted for work with H. diminuta. Thus, precautions that need to be taken are p ­ rimarily dictated by precautions that need to be taken when working with its vertebrate and invertebrate primary and ­secondary hosts, respectively.

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7. The primary concern when working with grain beetles is to minimize exposure to particulate antigens. The conditions of cultivation of grain beetles are conducive to growth of yeast/mold. Precautions, such as inserting toothpicks into celery, taken so that the source of food for the beetles (oats) does not come into extensive contact with the source of moisture (­celery) for the beetles. A dehumidifier prevents excess growth of yeast or mold which might increase microbial-derived a­ irborne antigens. 8. In addition, it is recommended that dust be kept to a low level by weekly or twice-weekly cleaning of housing. This is done by dumping the entire nursery into a sieve, allowing the ­droppings to pass through. After the droppings have been removed, the worms and pupae retained in the sieve are returned to the nursery and provided with fresh organic oats, celery, and nutritional yeast seasoning (10:1 ratio of oats to yeast in the nursery, with celery as needed). 9. The methods described are based essentially on methods acquired from individuals producing HDCs for therapeutic purposes in humans. The acquisition of these methods was conducted during the course of IRB-approved sociomedical studies, as described previously [5, 6]. These methods are slightly modified from methods described by Carolina Biological Supply (Greensboro, NC), which sells H. diminuta strictly for educational purposes in both the cysticercoid and egg life stages. Modification of the methodology to enhance the production of HDCs or beetles may decrease the therapeutic effect of the HDCs. In particular, feeding the grain beetles a richer source of nutrition repeatedly yielded improved production of grain beetles but also HDCs with decreased therapeutic impact. Using HDCs that were between 5 weeks and 5 months of age, but not older or younger, was also reported to have the most therapeutic benefit. 10. Unlike laboratory rats, laboratory mice will not readily ingest HDCs administered using the tip of a pipette placed in their mouth. Rather, the mice will do their best to avoid the pipette, and will often bite through the pipette, rendering the pipette ineffective for delivering the HDCs. Oral gavage needles are readily available, but HDCs tend to get hung up in the ­junctions in the needles, making delivery through a standard gavage needle unreliable. For reliable delivery of HDCs to mice, a disposable “fine tip transfer pipette” (Samco Scientific Corp) can be used for delivery, but it must be shielded to ­prevent the mice from biting through the pipette. For this purpose, our laboratory uses a modified 14-gauge IV catheter (Angiocath: Becton, Dickinson and Company, Franklin Lakes, NJ) as shown in Fig. 5 to shield the pipette. To feed the mice, the animals are held by the scruff of the neck, and the shielded pipette is inserted behind the tongue of the mouse with the

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Fig. 5 Device for feeding HDCs to mice. Modification of a 14-gauge IV catheter to shield a pipette is shown. (a) Catheter with needle, (b) catheter modified to make a shield for the pipette, (c) pipette loaded with 20 μL of opaque liquid, (d) pipette loaded with 20 μL of opaque liquid and shielded with a modified catheter, ready for use in feeding HDCs to mice. The asterisk in the diagram indicates the furthest point to which HDCs should be drawn in the pipette. Beyond that point, the HDCs have an increased tendency to stick in the pipette. Of note is the fact that the unshielded pipette, as loaded in diagram c, is appropriate for feeding of rats. The numbers on the scale indicate centimeters

head in the vertical position. After insertion of the shielded pipette, the liquid is immediately dispensed, and the animal is watched closely to ensure that it swallows. 11. Controls in the mouse model should be fed beetle abdomen extracts from beetles that lack HDCs. These controls compensate for the fact that beetle extracts contain both nutritional material and microbial content. Although the microbial content of the extracts (as will all insect-associated bacteria) is benign, it will alter the flora in the mice. Although HDCs can be cleaned to avoid the nutritional and microbial contamination, HDCs in pure saline are excessively “sticky” and difficult to pipette. Thus, either the unpurified (in beetle abdomen extract) HDCs should be used or a carrier protein should be incorporated into the purification medium. 12. H. diminuta initially begin to grow in mice with good e­ fficiency, with most of the HDCs maturing [12]. However, maturation is short lived. Hopkins provides an extensive list of mouse strains that reject H. diminuta, concluding that “all strains” reject the helminths [13]. However, Hopkins also concludes that side-byside studies have not been conducted, so it is difficult to know if there are strain-dependent d ­ ifferences in rejection. Andreassen and colleagues concluded that H. diminuta were expelled from nu/nu mice between days 10 and 20 and were expelled from +/nu mice sooner, at less than 10 days [14].

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References 1. Parker W, Perkins SE, Harker M, Muehlenbein MP (2012) A prescription for clinical immunology: the pills are available and ready for testing. Curr Med Res Opin 28:1193–1202. https:// doi.org/10.1185/03007995.2012.695731 2. Parker W, Ollerton J (2013) Evolutionary biology and anthropology suggest biome reconstitution as a necessary approach toward dealing with immune disorders. Evol Med Public Health 2013:89–103. https://doi. org/10.1093/emph/eot008 3. Bickler SW, DeMaio A (2008) Western diseases: current concepts and implications for pediatric surgery research and practice. Pediatr Surg Int 24(3):251–255 4. Bilbo SD, Wray GA, Perkins SE, Parker W (2011) Reconstitution of the human biome as the most reasonable solution for epidemics of allergic and autoimmune diseases. Med Hypotheses 77(4):494–504. https://doi. org/10.1016/j.mehy.2011.06.019 5. Cheng AM, Jaint D, Thomas S, Wilson J, Parker W (2015) Overcoming evolutionary mismatch by self-treatment with helminths: current practices and experience. J Evol Med 3:235910 6. Liu J, Morey RA, Wilson JK, Parker W (2016) Practices and outcomes of self-treatment with helminths based on physicians’ observations. J Helminthol FirstView 91:1–11 7. Webb RA, Hoque T, Dimas S (2007) Expulsion of the gastrointestinal cestode, Hymenolepis diminuta by tolerant rats: evidence for mediation by a Th2 type immune enhanced goblet cell hyperplasia, increased mucin production

and secretion. Parasite Immunol 29(1):11–21. https://doi.org/10.1111/j.1365-3024.2006. 00908.x 8. McKay DM, Halton DW, McCaigue MD, Johnston CF, Fairweather I, Shaw C (1990) Hymenolepis diminuta: intestinal goblet cell response to infection in male C57 mice. Exp Parasitol 71(1):9–20 9. Palmas C, Bortoletti G, Gabriele F, Wakelin D, Conchedda M (1997) Cytokine production during infection with Hymenolepis diminuta in BALB/c mice. Int J Parasitol 27(7):855–859 10. McKay DM, Khan WI (2003) STAT-6 is an absolute requirement for murine rejection of Hymenolepis diminuta. J Parasitol 89(1):188– 189. https://doi.org/10.1645/00223395(2003)089[0188,SIAARF]2.0.CO;2 11. Arai HP (1980) Biology of the tapeworm Hymenolepis diminuta. Academic Press, New York 12. Hopkins CA, Subramanian G, Stallard H (1972) The development of Hymenolepis diminuta in primary and secondary infections in mice. Parasitology 64(3):401–412 13. Hopkins CA (1980) Immunity and Hymenolepis diminuta. In: Arai HP (ed) Biology of the tapeworm Hymenolepis diminuta. Academic Press, New York, pp 551–614 14. Andreassen J, Hindsbo O, Ruitenberg EJ (1978) Hymenolepis diminuta infections in congenitally athymic (nude) mice: worm kinetics and intestinal histopathology. Immunology 34(1):105–113

Chapter 4 A Mouse Model of Peanut Allergy Induced by Sensitization Through the Gastrointestinal Tract Kelly Orgel and Michael Kulis Abstract Animal models of disease enable the study of the pathology, biomarkers, and treatments for the disease being studied. These models become particularly useful in the study of diseases, such as peanut allergy, that currently have no FDA-approved therapy options. Here, we describe a mouse model of peanut allergy using a peanut extract and cholera toxin that can be applied to both BALB/c and C3H/HeJ mouse strains. Sensitization is induced through the gastrointestinal tract resulting in elevated levels of ­peanut-­specific IgE and anaphylaxis upon challenge with peanut proteins. This model has been used to study the cells and molecules involved in the development of peanut allergy and to evaluate novel ­immunotherapy approaches and the underlying mechanisms of immunotherapy. Potential utilities of this model are numerous and may include studies on microbial influences on peanut allergy and discovery of biomarkers of anaphylaxis. Key words Peanut allergy, Food allergy, Mouse model, Anaphylaxis, IgE

1  Introduction Peanut allergy, a growing public health concern, currently affects about 1% of the US population [1–3]. During an allergic reaction, peanut-specific IgE bound to the high-affinity IgE receptor (FcεRI) on mast cells and basophils is cross-linked by peanut ­allergen [4]. This results in activation of mast cells and basophils, which release histamine, leukotrienes, and other mediators into ­tissues. These mediators are ultimately responsible for the allergic symptoms observed in patients, including hives, angioedema, throat tightness, abdominal pain, and vomiting. Allergic patients are advised to avoid the allergen and carry auto-injectable ­epinephrine in case of accidental exposure [5]. Peanut allergy has drastic implications on the patients’ quality of life [6, 7] and can be life-threatening and even fatal [8, 9], yet there remain no FDA-approved therapy options to treat peanut allergies [10]. Furthermore, there are increasingly more questions R. Lee Reinhardt (ed.), Type 2 Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1799, https://doi.org/10.1007/978-1-4939-7896-0_4, © Springer Science+Business Media, LLC, part of Springer Nature 2018

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surrounding the epidemiology of peanut allergy. Specifically, there is limited knowledge on the causes of food allergies, biomarkers for the disease, and biomarkers for anaphylaxis [11–13]. In order to better understand the disease and to develop new therapeutics, ­animal models of peanut allergy have been developed. The model described here utilizes cholera toxin as an adjuvant to break oral tolerance in either BALB/c or C3H/HeJ mice and sensitize them to peanut through the gastrointestinal tract. Our group has used this model to investigate novel preventive and therapeutic approaches, among other investigations in similar mouse models [14–22].

2  Materials Prepare all reagents with Milli-Q Type 1 ultrapure water when ­possible. All thawed or reconstituted reagents are stored at 4 °C for a maximum of 4 weeks. Other than reagents for ELISAs and ­preparation of peanut extract, only open and use reagents under a sterile biosafety cabinet. Follow all waste disposal instructions when disposing of reagents. The institution’s Division of Laboratory Animal Medicine (DLAM) should be made aware of the use of cholera toxin and informed to not change the cage within 2 days of any treatment with cholera toxin. All animal research should be approved by the Institution’s Animal Care and Use Committee (IACUC). 2.1  Peanut Extract

1. Peanut flour: 12% fat light roast, 50% protein (Golden Peanut Co.). 2. Phosphate-buffered saline (PBS) with 1 mol/L NaCl. 3. Microcentrifuge. 4. 0.4 μM syringe filter. 5. 0.2 μM syringe filter. 6. Bicinchoninic acid assay (BCA). 7. NuPAGE 4–12% Bis-Tris gel and protein gel apparatus (Thermo Fisher).

2.2  Mouse Sensitization

1. Cholera toxin: 1 mg/mL cholera toxin from Vibrio cholerae in ultrapure water. Resuspended aliquot can be stored at 4 °C for 3–4 weeks. 2. 22 g × 1.5″, 1.25 mm straight oral gavage needle. 3. 1 mL syringe. 4. BALB/cJ: female mice (see Note 1). 5. C3H/HeJ: female mice (see Note 1).

A Mouse Model of Peanut Allergy

2.3  Serum Collection, ELISAs for Immunoglobulin Quantification, and Core Temperature Measurements

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1. Microtainer TSS serum separator tubes (Becton Dickenson). 2. 0.05 M carbonate-bicarbonate buffer, pH 9.6 (Sigma). 3. 96-well non-treated polystyrene plate. 4. 2,4-dinitrophenyl hapten conjugated to human serum ­albumin (DNP-HSA): 20 μg/mL in carbonate-bicarbonate buffer (see Note 2). 5. ELISA wash solution: 0.05% Tween 20 in PBS. 6. ELISA blocking solution and antibody diluent: 0.05% Tween 20 and 2% bovine serum albumin fraction V in PBS. 7. Detection antibodies: For specific IgE, 0.5 μg/mL sheep antimouse IgE, 0.5 μg/mL biotinylated donkey anti-sheep IgG, 0.2  μg/mL neutravidin-horseradish peroxidase (HRP). For specific IgG1, 10 ng/mL HRP-conjugated goat anti-mouse IgG1. 8. 3,3′,5,5′-Tetramethylbenzidine (TMB) substrate solution: SureBlue TMB Microwell Peroxidase Substrate (KPL). 9. TMB stop solution (KPL). 10. Microplate colorimetric spectrophotometer. 11. Rectal probe: Small animal probe to measure core body temperature.

3  Methods 3.1  Peanut Extract

1. Extract peanut proteins by mixing peanut flour in a 1:5 (wt:vol) ratio of phosphate-buffered saline (PBS) with 1 mol/L NaCl. 2. Mix solution with stirring bar on magnetic stir plate for 2 h at room temperature while maintaining an alkaline pH (8.5) (see Note 3). 3. Centrifuge solution at 30,000 × g for 45 min at 4 °C. 4. Decant to collect supernatant, and then filter-sterilize it ­sequentially through a 0.4 μM filter followed by a 0.2 μM filter. 5. Determine protein concentration by bicinchoninic acid assay (BCA) using bovine serum albumin (BSA) as the standard (see Note 4). 6. Run collected peanut protein on a NuPAGE gel with a 10–20 μg protein load per well to identify and compare quantities of ­peanut allergens Ara h 1, 2, 3, and 6 between extract ­preparations. Use a molecular weight marker for easy identification of key allergenic protein bands (Fig. 1).

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Fig. 1 SDS-PAGE analysis of peanut extracts. Two peanut extracts are shown side by side with 20 μg of protein loaded per lane. The major peanut allergens, Ara h 1, 2, 3, and 6, are labeled 3.2  Mouse Sensitization

1. Mice can be separated into naïve or sensitized groups. 2. Maintain naïve mice throughout the experiment by treating them with PBS by oral gavage once a week for 4 weeks ­according to the schedule schematic shown in Fig. 2. 3. Sensitize 4-week-old female mice with 200 μL containing 2 mg peanut extract and 10 μg cholera toxin diluted in PBS by oral gavage once a week for 3 weeks. On the fourth week of ­sensitization, gavage mice with 350 μL containing 5 mg peanut extract and 10 μg cholera toxin diluted in PBS.

3.3  Serum Collection and ELISAs for Immunoglobulin Quantification

1. One week after the final sensitization dose, collect 200 μL whole blood by submandibular bleed into BD Microtainer SST serum separator tubes. Allow blood to clot for at least 20 minutes after collection before centrifuging at 8000 RPM for 15 min. Transfer serum layer to 0.7 microfuge tube, and freeze at −20 °C until needed for ELISA. 2. Coat 96 well non-treated polystyrene plate with 20 μg/mL whole peanut extract for sample wells and 20 μg/mL DNP-

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Fig. 2 Schematic of sensitization protocol. Four-week-old female mice are sensitized once weekly for 4 weeks with peanut and cholera toxin by oral gavage. One week later (day 28), mice are bled by submandibular bleed to measure immunoglobulins. The following day, mice can be challenged with peanut extract by intraperitoneal injection

HSA for standard curve wells in carbonate-bicarbonate buffer for 1 h at 37 °C or overnight at 4 °C. 3. Wash plates 3 times with 200 μL ELISA wash solution. 4. Block plates with ELISA blocking and antibody diluent ­solution for 2 h at 37 °C or overnight at 4 °C. 5. Repeat plate washing three times with 200 μL ELISA wash solution. 6. Add serum samples for 1 h at 37 °C. For peanut-specific IgE measurements, use a serum dilution between 1:100 (BALB/c mice) and 1:200 (C3H/HeJ mice). For peanut-specific IgG1 measurements, use a serum dilution between 1:10,000 (BALB/c mice) and 1:20,000 (C3H/HeJ mice). 7. Repeat plate washing three times with 200 μL PBS containing 0.05% Tween 20. 8. For detection of IgE, add 100 μL/well sheep anti-mouse IgE (0.5 μg/mL) for 1 h at 37 °C. Repeat plate washing three times with 200 μL ELISA wash solution. Add 100 μL b ­ iotinylated donkey anti-sheep IgG (0.5 μg/mL), and ­incubate for 1 h at 37 °C. Wash plate three times with 200 μL ELISA wash s­ olution. Add 100 μL/well n ­ eutravidin-horseradish peroxidase (NA-HRP; 0.2 μg/mL) for 1 h at 37 °C. Again, wash plate three times with 200  μL ELISA wash solution. Representative examples of ­peanut-IgE levels pre- and post-­sensitization with peanut extract plus cholera toxin are shown in Fig. 3a. 9. For detection of IgG1, add 100 μL/well HRP-conjugated goat anti-mouse IgG1 used at 10 ng/mL for 1 h at 37 °C. After incubation, wash plate three times with 200 μL ELISA wash solution. 10. Measure HRP activity by blue color development of SureBlue TMB Microwell Peroxidase Substrate solution. To do this, add 100  μL TMB Substrate solution to all wells. Incubate with shaking at room temperature for approximately 10 min.

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Fig. 3 Representative peanut-specific IgE levels and challenge outcomes. (a) Peanut-specific IgE levels in C3H/HeJ and BALB/cJ mice pre- and post-sensitization with peanut extract plus cholera toxin. (b) Changes in body temperatures of peanut (PN)-sensitized C3H/HeJ mice after challenge with 200 μg peanut extract; note that peanut-sensitized mice challenged with an equal volume of PBS do not have decreased body temperature and that naïve (i.e., not sensitized to peanut) mice challenged with peanut extract do not have decreased body temperature. (c) Symptom scores in C3H/HeJ mice; note that only peanut-sensitized mice challenged with peanut exhibit allergic symptoms

11. Add 100  μL of TMB stop solution to each well. Wells that turned blue after TMB addition will turn yellow after the acidic stop solution is added. Avoid causing bubbles in wells as this may interfere with absorbance readings. 12. Read plates on colorimetric spectrophotometer at 450 nm wavelength. 3.4  Intraperitoneal Peanut Challenge

1. Allow animals’ body temperature to rise for 15 min prior to beginning the challenge (see Note 5).

A Mouse Model of Peanut Allergy

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2. Using a rectal thermometer probe, measure and record the body temperatures of the mice. These values will serve as ­baseline temperatures. 3. Inject 200 μL diluted peanut extract in PBS (200 μg for C3H/ HeJ mice and 300 μg for BALB/c mice) into the ­intraperitoneal cavity of each animal. Start a timer after the first mouse has been injected (see Note 6). 4. Measure and record each animal’s body temperature every 15–60 min. Animals experiencing a systemic reaction will have a drop in body temperature, while those experiencing no ­reaction will maintain a constant body temperature (Fig. 3b). 5. During the 30-min body temperature reading, assign a ­symptom score to each animal (Fig. 3c). Symptoms are assigned based on the following scale: 0, no symptoms; 1, scratching around the nose and head; 2, puffiness around the eyes and mouth with reduced activity; 3, labored respiration and/or cyanosis around the mouth and tail; 4, no activity after prodding or tremor and convulsion; and 5, death (see Note 7). 6. If blood samples are required during the challenge, a ­submandibular bleed can be performed at the desired time point (see Note 8). 7. After 60 min, the housing, tunnels, and food can be returned in the cages, and cages can be returned to their location in the housing facility. If a second challenge is needed, the procedure can be repeated after 1 week of resting the animals (see Notes 9 and 10).

4  Notes 1. Female mice are used in this model since male mice do not consistently develop elevated peanut-specific IgE nor experience anaphylaxis upon challenge with peanut extract. Female mice are ordered to arrive at 3 weeks of age. They are allowed to acclimate for 1 week after transfer, and procedures are started at 4 weeks of age. Mice are fed standard chow lacking peanut by the University of North Carolina DLAM personnel and housed under pathogen-free conditions. Mice are housed with four or five animals per cage. 2. Individual peanut allergens (e.g., Ara h 1, 2, 3, and 6) can be assayed and are coated at 5 μg/mL. 3. pH will drop as peanut proteins enter solution. Check pH every 15 min, and adjust to pH 8.5 using 6 M sodium ­hydroxide. For more concentrated solution, mix and adjust pH for longer than 2 h.

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4. Typical concentrations of peanut extract yielded with this ­protocol are ~15–20 mg/mL. 5. Animals that have been sleeping will have a lower body temperature. Removing any housing, tunnels, and food will result in the animals running and their body temperatures rising. If this step is not done, the baseline temperature readings will appear lower and may mask initial decreases during the challenge. 6. The concentrations of peanut provided here are those that we have optimized to see maximal reactions while minimizing animal death. Depending on the level of sensitization of the animals, the dose may be increased or decreased to induce more or less reaction severity. 7. Symptom scores may worsen at later time points, and while we make note of these changes for our own records, we only report one symptom score in the literature and not changes in scores over time. 8. During a systemic reaction, mice experience vasodilation, so obtaining more than 100 μL whole blood can be very difficult. Also, mice tend to experience body temperature decrease following a blood draw, so temperature readings after this ­ point are not useful. 9. Mice should be continuously monitored following challenge until their body temperature stops decreasing. Also, any mouse whose body temperature drops below 30 °C should be ­euthanized for ethical reasons. 10. C3H/HeJ mice that have been sensitized using this model can then undergo immunotherapy to desensitize the animals. References 1. Boyce JA, Assa'ad A, Burks AW, Jones SM, Sampson HA, Wood RA, Plaut M, Cooper SF, Fenton MJ, Arshad SH, Bahna SL, Beck LA, Byrd-Bredbenner C, Camargo CA Jr, Eichenfield L, Furuta GT, Hanifin JM, Jones C, Kraft M, Levy BD, Lieberman P, Luccioli S, McCall KM, Schneider LC, Simon RA, Simons FE, Teach SJ, Yawn BP, Schwaninger JM (2011) Guidelines for the diagnosis and management of food allergy in the United States: summary of the NIAID-sponsored expert panel report. Nutr Res 31(1):61–75. https://doi.org/10.1016/j. nutres.2011.01.001 2. Tang ML, Mullins RJ (2017) Food allergy: is prevalence increasing? Intern Med J 47(3):256–261. https://doi.org/10.1111/ imj.13362

3. Sicherer SH, Munoz-Furlong A, Godbold JH, Sampson HA (2010) US prevalence of selfreported peanut, tree nut, and sesame allergy: 11-year follow-up. J Allergy Clin Immunol 125(6):1322–1326. https://doi. org/10.1016/j.jaci.2010.03.029 4. Ang WX, Church AM, Kulis M, Choi HW, Burks AW, Abraham SN (2016) Mast cell desensitization inhibits calcium flux and aberrantly remodels actin. J Clin Invest 126(11):4103–4118. https://doi. org/10.1172/JCI87492 5. Burks AW (2008) Peanut allergy. Lancet 371(9623):1538–1546. https://doi. org/10.1016/S0140-6736(08)60659-5 6. King RM, Knibb RC, Hourihane JO (2009) Impact of peanut allergy on ­ quality of life, stress and anxiety in the family. Allergy

A Mouse Model of Peanut Allergy 64(3):461–468. https://doi. org/10.1111/j.1398-9995.2008.01843.x 7. Avery NJ, King RM, Knight S, Hourihane JO (2003) Assessment of quality of life in children with peanut allergy. Pediatr Allergy Immunol 14(5):378–382 8. Bock SA, Munoz-Furlong A, Sampson HA (2001) Fatalities due to anaphylactic reactions to foods. J Allergy Clin Immunol 107(1):191– 193. https://doi.org/10.1067/ mai.2001.112031 9. Bock SA, Munoz-Furlong A, Sampson HA (2007) Further fatalities caused by anaphylactic reactions to food, 2001–2006. J Allergy Clin Immunol 119(4):1016–1018. https://doi. org/10.1016/j.jaci.2006.12.622 10. Iweala OI, Burks AW (2016) Food allergy: our evolving understanding of its pathogenesis, prevention, and treatment. Curr Allergy Asthma Rep 16(5):37. https://doi. org/10.1007/s11882-016-0616-7 11. Virkud YV, Burks AW, Steele PH, Edwards LJ, Berglund JP, Jones SM, Scurlock AM, Perry TT, Pesek RD, Vickery BP (2017) Novel baseline predictors of adverse events during oral immunotherapy in children with peanut allergy. J Allergy Clin Immunol 139(3):882–888.e5. https://doi.org/10.1016/j.jaci.2016.07.030 12. Burton OT, Logsdon SL, Zhou JS, Medina-­ Tamayo J, Abdel-Gadir A, Noval Rivas M, Koleoglou KJ, Chatila TA, Schneider LC, Rachid R, Umetsu DT, Oettgen HC (2014) Oral immunotherapy induces IgG antibodies that act through FcgammaRIIb to suppress IgE-mediated hypersensitivity. J Allergy Clin Immunol 134(6):1310–1317.e1316. https:// doi.org/10.1016/j.jaci.2014.05.042 13. Nowak-Wegrzyn A, Sampson HA (2011) Future therapies for food allergies. J Allergy Clin Immunol 127(3):558–573.; quiz 574– 555. https://doi.org/10.1016/j. jaci.2010.12.1098 14. Plundrich NJ, Kulis M, White BL, Grace MH, Guo R, Burks AW, Davis JP, Lila MA (2014) Novel strategy to create h ­ ypoallergenic peanut protein-polyphenol edible m ­ atrices for oral immunotherapy. J Agric Food Chem 62(29):7010–7021. https://doi. org/10.1021/jf405773b 15. Kulis M, Wesley Burks A (2015) Effects of a pre-existing food allergy on the oral

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i­ntroduction of food proteins: findings from a murine model. Allergy 70(1):120–123. https://doi.org/10.1111/all.12519 16. Kulis M, Gorentla B, Burks AW, Zhong XP (2013) Type B CpG oligodeoxynucleotides induce Th1 responses to peanut antigens: modulation of sensitization and utility in a truncated immunotherapy regimen in mice. Mol Nutr Food Res 57(5):906–915. https:// doi.org/10.1002/mnfr.201200410 17. Kulis M, Macqueen I, Li Y, Guo R, Zhong XP, Burks AW (2012) Pepsinized cashew proteins are hypoallergenic and immunogenic and provide effective immunotherapy in mice with cashew allergy. J Allergy Clin Immunol 130(3):716–723. https://doi.org/10.1016/j. jaci.2012.05.044 18. Kulis M, Chen X, Lew J, Wang Q, Patel OP, Zhuang Y, Murray KS, Duncan MW, Porterfield HS, WB A, Dreskin SC (2012) The 2S albumin allergens of Arachis hypogaea, Ara h 2 and Ara h 6, are the major elicitors of anaphylaxis and can effectively desensitize peanut-allergic mice. Clin Exp Allergy 42(2):326–336. https://doi. org/10.1111/j.1365-2222.2011.03934.x 19. Kulis M, Wan CK, Gorentla BK, Burks AW, Zhong XP (2011) Diacylglycerol kinase zeta deficiency in a non-CD4(+) T-cell compartment leads to increased peanut hypersensitivity. J Allergy Clin Immunol 128(1):212–214. https://doi.org/10.1016/j.jaci.2011.02.035 20. Kulis M, Li Y, Lane H, Pons L, Burks W (2011) Single-tree nut immunotherapy attenuates allergic reactions in mice with hypersensitivity to multiple tree nuts. J Allergy Clin Immunol 127(1):81–88. https://doi.org/10.1016/j. jaci.2010.09.014 21. Orgel KA, Duan S, Wright BL, Maleki SJ, Wolf JC, Vickery BP, Burks AW, Paulson JC, Kulis MD, Macauley MS (2017) Exploiting CD22 on antigen-specific B cells to prevent allergy to the major peanut allergen Ara h 2. J Allergy Clin Immunol 139(1):366–369.e362. https:// doi.org/10.1016/j.jaci.2016.06.053 22. Pons L, Ponnappan U, Hall RA, Simpson P, Cockrell G, West CM, Sampson HA, Helm RM, Burks AW (2004) Soy immunotherapy for peanut-allergic mice: modulation of the peanut-allergic response. J Allergy Clin Immunol 114(4):915–921. https://doi.org/10.1016/j. jaci.2004.06.049

Chapter 5 Induction and Characterization of the Allergic Eye Disease Mouse Model Nancy J. Reyes, Rose Mathew, and Daniel R. Saban Abstract Ocular IgE-associated allergy ranges from mild disease (seasonal and perennial allergic conjunctivitis) to more chronic/severe and vision-threatening forms (atopic and vernal keratoconjunctivitis). Whereas mild forms of disease have been studied extensively, less is known about the more chronic forms. Our lab has helped to address this knowledge gap by developing and characterizing an allergen-induced, chronic/ severe, IgE-associated model of ocular allergy referred to as the severe allergic eye disease (AED) model. It is distinct from previously described models that mimic the more mild forms, referred to in the literature as the allergic conjunctivitis (AC) model. The purpose of this method article is to detail the protocol to induce and characterize the AED model and directly compare these mice to the mild AC model. Troubleshooting and implications are also discussed. Keywords Allergic eye disease, AED, Allergic conjunctivitis, T cells, Th2, Allergy, Conjunctivitis, Type 2 immunity, Fibrosis, Dendritic cells

1  Introduction Ocular allergy is a set of specific allergic inflammatory conditions that affect the conjunctiva, the eyelid, and, in severe cases, the c­ornea. Type and severity range from mild seasonal and perennial allergic conjunctivitis (AC) to more severe and chronic such as atopic and vernal keratoconjunctivitis (AKC and VKC, ­respectively) [1]. While these types differ significantly in clinical symptoms, prognosis, and pathobiology, they all share a common IgE-associated etiology [1]. Hence, allergenic CD4+ T helper (Th)2 cells and their cytokines (IL-4, IL-5, IL-13) are central to mediate disease [2–4].

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Though several species, such as the rat and guinea pig, have been used to study ocular allergy, presently the preferred species is the mouse system [1]. Several mouse models exist for studying ocular allergy that are induced by either active or passive ­immunization against a model antigen, such as short rag weed or ovalbumin. Until recently, most mouse models have primarily mimicked the more mild types of ocular allergy akin to AC in people. These models present with classical immediate ­ ­hypersensitivity signs and symptoms that include swelling of the bulbar conjunctiva (i.e., chemosis), superficial conjunctival ­vasodilation (i.e., redness), tearing (Fig. 2), and lid edema [2, 5]. By contrast, AKC and VKC are more chronic and severe ­diseases that have more complex mechanisms and can develop tissue remodeling [6–11], although the pathobiology is less ­ ­understood. To help address current knowledge gaps [3, 12], our lab established a model of chronic ocular allergy, which we refer to as the allergic eye disease (AED) model so as to differentiate it from the classical AC model. Mice with AED present with the more severe clinical manifestations that are present in patients with AKC and VKC [13–15]. At a clinical level, manifestation of ­conjunctival fibrosis, thick mucoid discharge, and eczema of the eyelid are all present in the AED model [5, 16–18] (Fig. 2). On a cellular level, robust eosinophil recruitment occurs in the ­conjunctiva, and lymphangiogenesis is observed in the cornea [16, 19, 20]. Thus, the AED model now enables the study of the chronic eye allergy setting and thereby helps address major ­knowledge gaps in our understanding of the pathobiology. The focus of this methods paper is the induction and analysis of AC and AED using C57BL/6 mice and ovalbumin (OVA) ­antigen, as outlined in Figure 1.

2  Materials 2.1  Chronic AED Model

1. 15 mL or smaller beaker. 2. Stir bar. 3. Stir plate. 4. Ovalbumin: OVA from chicken egg white. 5. Pertussis toxin (PT): Pertussis toxin from Bordetella pertussis. 6. Aluminum hydroxide (Imject Alum). 7. Sterile 1× PBS. 8. 1 mL syringe. 9. 25-gauge needle. 10. C57BL/6 mice (see Note 1).

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Fig. 1 Induction of mild allergic conjunctivitis (AC) and chronic allergic eye disease (AED) models. Allergy is induced in mice by IP injection of immunization mix and 14 days later topical instillation of OVA for 7 consecutive days

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Fig. 2 AED results in more severe clinical disease compared to AC. Clinical scores were assessed at 20 min (to determine immediate hypersensitivity) and 6 h post-challenge (to determine the late-phase reaction). (a) Mice with AED have higher classic allergic clinical responses compared to mice with AC that persisted even 6 h postchallenge. (b) Severe clinical manifestations such as blepharitis (red arrow), mucoid discharge, and corneal epitheliopathy (data not shown) developed in mice with AED but were absent in mice with AC. Data is representative of five independent experiments (n = 5 mice/group/experiment). p   4  °C (see Note 4). After cells are partially thawed, transfer the vial content into the prepared 15 mL tube with complete RPMI 1640 with Benzonase. 4. Centrifuge at 800 × g-force for 7 min. 5. Remove supernatant by decanting or aspiration, and resuspend the pelleted cells in 10  mL of fresh complete RPMI 1640. Benzonase is no longer required as the DNA from nonviable cells will have been degraded during steps 3 and 4. 6. Determine the cell count using a hemocytometer or cell counter, following the manufacturer’s guidelines. 7. Centrifuge the cells at 800 × g-force for 7 min, and resuspend to a final concentration of 4 × 106 cells/mL in complete RPMI 1640. 8. Plate PBMCs in a 24-well plate, adding 0.5 mL per well (i.e., 2 × 106 cells/well). For a cell count of ten million cells, five wells will be plated.

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9. Add 0.5  mL of tree grass (TG) extract suspension to each well. Adding 0.5 mL of extract suspension to 0.5 mL of cell suspension will put the final stimulation concentration of TG extract to 50 μg/mL. 10. Incubate cells in an incubator at 37 °C, 5% CO2, 5% humidity, for 14 days. 11. Every 3–4 days, remove half the volume of medium (0.5 mL) from each well (carefully from the top, without disturbing the cell layer at the bottom of the well) and add 0.5 mL of fresh complete RPMI 1640 with rhIL-2. This should be performed three times over 14 days. 3.2  Coating of 3-Color FluoroSpot Plates

Perform all steps described here in a lamina flow to work in sterile conditions. FluoroSpot plates should be coated the day before assay setup (day 13). 1. Pre-wet IPFL plate by pipetting 50  μL per well of 70% methanol. 2. Discard methanol and immediately wash plate 3× with 150 μL sterile dH2O. 3. Coat plate with 50  μL/well of coating antibody cocktail. A total volume of 5 mL is required to coat 1 plate. 4. Seal plate edges with parafilm (optional, but recommended), and place the plate at 4 °C overnight.

3.3  FluoroSpot Setup

Perform all steps described here in a lamina flow to work in sterile conditions.

3.3.1  Blocking the IPFL Plate Membranes to Prevent Unspecific Binding

1. Decant coating antibody cocktail solution. 2. Wash plate 3× with 100 μL PBS. 3. Add 100 μL of complete RPMI 1640 to each well, and incubate for a minimum of 1 h in a 37 °C incubator. 4. Remove plates and discard cell medium. 5. Add stimulation conditions (Subheading 3.3.2, step 7) and cells (Subheading 3.3.3, step 7) as described below.

3.3.2  Preparation of Stimulation Conditions

1. Prepare 15-mer peptide solutions by adding concentrated peptide stock to complete RPMI 1640 at a concentration of 20 μg/mL (see Note 3). Of note, the peptide solution will be diluted further 1:1 in the FluoroSpot plate due to the addition of cells (see Subheading 3.3.3, step 7); therefore the final concentration of peptide is 10 μg/mL. 2. Prepare timothy grass (TG) extract suspension by adding concentrated extract stock to complete RPMI 1640 at a

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c­ oncentration of 10–100 μg/mL. Of note, the peptide solution will be diluted further 1:1 in the FluoroSpot plate due to the addition of cells (see Subheading 3.3.3, step 7); therefore the final concentration of peptide is 5–25 μg/mL. 3. Prepare PHA solution as a positive control. 4. As a negative control, use cell medium only. 5. Prepare a map of the plate layout to define the positions of each stimulus to be tested. Keep in mind, each stimulation condition will be tested in triplicate, using three neighboring wells. 6. After blocking for a minimum of 1 h, remove the IPFL plate from the 37  °C incubator, and discard the cell medium as described in Subheading 3.3.1, step 4. 7. Add 50  μL of each peptide solution (stimulus) to be tested into three neighboring wells, following the layout specified in the plate map. 3.3.3  Harvesting Cells from Expansion Culture

1. Use a sterile Pasteur pipette, and harvest the wells by carefully pipetting the content of the well up and down to evenly resuspend cells. Transfer the resuspended cells from the well into a clean 15  mL tube, pooling wells from the same donor that have been stimulated with the same stimulus (e.g., TG extract). 2. Centrifuge cells at 800 × g-force for 7 min. 3. Discard the supernatant, and resuspend the cell pellet in 10 mL of fresh complete RPMI 1640 medium. 4. Determine the cell count using a hemocytometer or cell counter, following the manufacturer’s guidelines. 5. Centrifuge cells at 800 × g-force for 7 min. 6. Discard the supernatant, and resuspend the cell pellet for final concentration of 2  ×  106/mL in complete RPMI 1640 medium. 7. Add 50  μL of the cell suspension (50  μL contain 100,000 cells) to each well of the IPFL plate AFTER the stimulation condition has already been added (Subheading 3.3.2, step 7) (see Note 5). 8. Be careful not to cross-contaminate stimulation conditions with the pipette tips while adding the cells. Angle the pipette to avoid touching any well content. 9. After both stimulus and cells have been carefully added, replace the IPFL plate lid, and carefully move the plate back into the incubator for 22–24  h incubation at 37  °C, 5% CO2, 5% humidity. For the clearest reads, avoid disrupting the plate by opening and closing the incubator door for the remainder of the incubation as this may displace cells, pushing them to the edge of the well.

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3.3.4  Color FluoroSpot Development

1. After 22–24 h of incubation, remove the cells by emptying the IPFL plate into a receptacle containing a small amount of bleach. 2. Wash the IPFL plate 5× with 200 μL/well of 10% Tween solution. This step can be performed using an automated plate washer, if available. 3. Prepare detection antibody cocktail. 4. Add 100 μL/well of antibody detection cocktail, and incubate for 2 h at room temperature. 5. Wash the IPFL plate 5× with 10% Tween solution (200 μL/ well). 6. Add 100 mL/well of secondary antibody detection cocktail. Incubate 1 h at room temperature. From this step on, cover the plate to limit light exposure. 7. Wash the plate 5× with 200 μL/well of 10% Tween solution. 8. Empty the plate and add 50 μL/well of Fluorescence enhancer­II and leave the plate for 15 min at room temperature. 9. Empty the plate, and remove residual Fluorescence enhancer­II by firmly tapping the plate against clean paper towels. 10. Remove the underdrain (the soft plastic under the IPFL plate). 11. Leave the IPFL plate in the dark to dry. The plate should be completely dry before analysis. 12. Inspect and count spots using a FluoroSpot reader. Figure 2 shows images of a single well as visualized by a FluoroSpot reader. 13. Store plate in the dark at room temperature.

3.4  Data Analysis

1. Data can be exported from the FluoroSpot reader software directly into Excel for further analysis. 2. The number of spots counted directly translates to the number of cells responsive to the stimulus tested (spot-forming cells, SFC). 3. Single peptides that elicit a response more than twofold above background (SFC induced by cells stimulated with medium alone), with a minimum of 20 SFC, are considered epitopes. 4. Data can be plotted as a bar graph showing the magnitude of reactivity (SFC) in response to each stimulus after subtraction of the background signal (i.e., net SFC) (Fig. 3).

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Fig. 2 Images showing spots resulting from cytokine production by antigen-­ specific cells as visualized by FluoroSpot. Each cell producing a given cytokine in response to antigen stimulation will be detected as one spot in the respective color of the corresponding cytokine

4  Notes 1. PBMC used for allergen T cell epitope identification should be obtained from donors who are truly allergic to the allergen of interest. Allergic sensitization is typically determined by skin prick testing (patient should have a wheal of flare ≥3 mm) or based on allergen-specific IgE titers (patient should have a titer ≥0.35 kU/L). Additionally, a clinical history consistent with the allergy of interest (clinical symptoms, previous diagnosis by medical professional, etc.) is preferable. 2. Cryopreservation can affect cell viability. Thus, it is necessary to count with a good live/dead stain, such as trypan blue. 3. Peptides can be difficult to dissolve into solution using standard buffers. For this reason, peptides may be resuspended at a high concentration in 100% DMSO, a substance in which most peptides are highly soluble. This allows the peptides to be kept at a high stock concentration (e.g., 40 mg/mL), with sufficient room for dilution for the assay to ensure the final DMSO concentration that the cells get exposed to in the assay is low enough to not affect cell viability. Of note, the final DMSO concentration in the assay should not exceed 0.5% of total volume.

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Fig. 3 T cell reactivity to TG extract and individual TG allergen-derived peptides in TG allergic patients. Responses are shown as number of spot-forming cells (SFC) per 106 PBMC, showing the sum of all three cytokines measured (IFNγ, IL-5, and IL-10). Two representative donors are shown

4. Freezing medium contains DMSO, which is toxic for PBMC above a temperature of 4 °C. Therefore, when thawing cells, it is recommended to not fully thaw the whole vial content but leave a small piece frozen before transfer to medium (where the DMSO is diluted to nontoxic concentrations) to avoid cell death and improve viability yield. 5. It is important to add the cells last. If 50 μL of stimulation is added on top of 50 μL of cell suspension, the cells settled on the membrane will be disrupted, and the pressure from the liquid added will displace the cells to the edge of the well. This may result in crescent shapes of spots rather than spots being equally distributed across the entire membrane.

Acknowledgments This work has been supported with federal funds from the National Institute of Allergy and Infectious Diseases, National Institutes of Health, and Department of Health and Human Services, under the grant number U19AI100275.

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References 1. Pawankar R, Canonica GW, Holgate ST, Lockey RF (2012) White book on allergy 2011–2012 executive summary. World Health Organization, Geneva 2. Borghesan F, Bernardi D, Plebani M (2007) In vivo and in  vitro allergy diagnostics: it's time to re-appraise the costs. Clin Chem Lab Med 45(3):391–395. https://doi.org/10.1515/ CCLM.2007.077 3. Akdis M, Verhagen J, Taylor A, Karamloo F, Karagiannidis C, Crameri R, Thunberg S, Deniz G, Valenta R, Fiebig H, Kegel C, Disch R, Schmidt-Weber CB, Blaser K, Akdis CA (2004) Immune responses in healthy and allergic individuals are characterized by a fine balance between allergen-specific T regulatory 1 and T helper 2 cells. J Exp Med 199(11):1567–1575. https://doi.org/10.1084/jem.20032058 4. Powe DG, Huskisson RS, Carney AS, Jenkins D, McEuen AR, Walls AF, Jones NS (2004) Mucosal T-cell phenotypes in persistent atopic and nonatopic rhinitis show an association with mast cells. Allergy 59(2):204–212 5. Salvi SS, Babu KS, Holgate ST (2001) Is asthma really due to a polarized T cell response toward a helper T cell type 2 phenotype? Am J Respir Crit Care Med 164(8 Pt 1):1343–1346. https://doi.org/10.1164/ ajrccm.164.8.2103080 6. Czarnowicki T, Gonzalez J, Shemer A, Malajian D, Xu H, Zheng X, Khattri S, Gilleaudeau P, Sullivan-Whalen M, Suarez-Farinas M, Krueger JG, Guttman-Yassky E (2015) Severe atopic dermatitis is characterized by selective expansion of circulating TH2/TC2 and TH22/ TC22, but not TH17/TC17, cells within the skin-homing T-cell population. J  Allergy Clin Immunol 136(1):104–115.e7. https://doi. org/10.1016/j.jaci.2015.01.020 7. Schulten V, Tripple V, Aasbjerg K, Backer V, Lund G, Wurtzen PA, Sette A, Peters B (2016) Distinct modulation of allergic T cell responses by subcutaneous vs. sublingual allergen-specific immunotherapy. Clin Exp Allergy 46(3):439– 448. https://doi.org/10.1111/cea.12653 8. Bonvalet M, Wambre E, Moussu H, Horiot S, Kwok WW, Louise A, Ebo D, Hoarau C, Van Overtvelt L, Baron-Bodo V, Moingeon P (2011) Comparison between major histocom-

patibility complex class II tetramer staining and surface expression of activation markers for the detection of allergen-specific CD4(+) T cells. Clin Exp Allergy 41(6):821–829. https://doi. org/10.1111/j.1365-2222.2011.03708.x 9. Ellis AK, Frankish CW, O'Hehir RE, Armstrong K, Steacy L, Larche M, Hafner RP (2017) Treatment with grass allergen peptides improves symptoms of grass pollen-­ induced allergic rhinoconjunctivitis. J  Allergy Clin Immunol 140:486–496. https://doi. org/10.1016/j.jaci.2016.11.043 10. Sager N, Feldmann A, Schilling G, Kreitsch P, Neumann C (1992) House dust mite-specific T cells in the skin of subjects with atopic dermatitis: frequency and lymphokine profile in the allergen patch test. J Allergy Clin Immunol 89(4):801–810 11. Wambre E, Van Overtvelt L, Maillere B, Humphreys R, von Hofe E, Ferhat L, Ebo D, Moingeon P (2008) Single cell assessment of allergen-specific T cell responses with MHC class II peptide tetramers: methodological aspects. Int Arch Allergy Immunol 146(2):99– 112. https://doi.org/10.1159/000113513 12. Dillon MB, Schulten V, Oseroff C, Paul S, Dullanty LM, Frazier A, Belles X, Piulachs MD, Visness C, Bacharier L, Bloomberg GR, Busse P, Sidney J, Peters B, Sette A (2015) Different Bla-g T cell antigens dominate responses in asthma versus rhinitis subjects. Clin Exp Allergy 45(12):1856–1867. https:// doi.org/10.1111/cea.12643 13. Oseroff C, Pham J, Frazier A, Hinz D, Sidney J, Paul S, Greenbaum JA, Vita R, Peters B, Schulten V, Sette A (2016) Immunodominance in allergic T-cell reactivity to Japanese cedar in different geographic cohorts. Ann Allergy Asthma Immunol 117(6):680–689. e681. https://doi.org/10.1016/j.anai.2016.10.014 14. Oseroff C, Christensen LH, Westernberg L, Pham J, Lane J, Paul S, Greenbaum J, Stranzl T, Lund G, Hoof I, Holm J, Wurtzen PA, Meno KH, Frazier A, Schulten V, Andersen PS, Peters B, Sette A (2017) Immunoproteomic analysis of house dust mite antigens reveals distinct classes of dominant T cell antigens according to function and serological reactivity. Clin Exp Allergy 47(4):577–592. https:// doi.org/10.1111/cea.12829

Chapter 14 Generation of Allergen-Specific Tetramers for a Murine Model of Airway Inflammation James J. Moon and Marion Pepper Abstract The identification and analysis of allergen-specific CD4+ T cells is critical for understanding how these cells contribute to atopic disease and how to subvert their behavior through immune therapy. The advent of fluorescently labeled soluble tetramers of peptide:MHCII complexes (pMHCII tetramers) has provided investigators with an invaluable means to achieve this goal. Although pMHCII tetramers were first developed over two decades ago, their widespread use has been limited by the technical difficulty of generating these reagents. However, the adoption of various technical innovations from several labs over time has contributed greatly to the increased success in tetramer generation today. Here, we describe a comprehensive protocol for generating pMHCII tetramers using as an example a Derp1:I-Ab tetramer used to study allergen-specific CD4+ T cell responses in murine models of airway inflammation and allergic disease. Key words Allergy, House dust mite, CD4+ T cell, Peptide:MHC complex, Tetramer

1  Introduction Asthma is an inflammatory disease of the airways that affects more than 300 million people worldwide and is rapidly increasing in incidence [1]. Clinically, asthma is characterized by acute, intermittent, and recurrent episodes of inflammation that can be induced by a specific allergen. Allergen-specific CD4+ T cells contribute to this process by producing type 2 cytokines (e.g., IL-4, IL-5, and IL-13) and inducing B cell production of IgE. Both these responses require T cell receptor (TCR) recognition of allergen-­derived peptides bound to MHCII molecules on host antigen-presenting cells. In both murine models of disease and asthmatic patients, allergen peptide:MHCII (pMHCII)-specific CD4+ memory T cells persist in the lungs and lymphoid organs after resolution of inflammation. Upon subsequent exposure to allergen, these cells rapidly drive asthma-induced pathology, but due to the technical challenges of identifying small populations of CD4+ T cells that express allergen pMHCII-specific TCRs, we are R. Lee Reinhardt (ed.), Type 2 Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1799, https://doi.org/10.1007/978-1-4939-7896-0_14, © Springer Science+Business Media, LLC, part of Springer Nature 2018

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just beginning to understand how these Th2 cells differentiate, function, and persist in health and disease. Soluble tetramers of allergen pMHCII provide a powerful state-of-the-art means to overcome the challenges of studying allergen-specific CD4+ T cells. By binding selectively to TCRs with cognate antigen specificity, fluorescently labeled tetramers can be used as staining reagents to directly identify allergen-specific T cells [2]. Here we describe the generation of pMHCII tetramers that can be used to identify both circulating and lungresiding allergen-­specific CD4+ T cells. For an allergen, we have initially focused on the Der p 1 protein (Derp1) of the house dust mite (HDM), Dermatophagoides pteronyssinus. Derp1 is a 30 kDa Group 1 cysteine protease that breaks down epithelial tight junctions and enhances antigen presentation by dendritic cells [3]. Several regions of Derp1 contain epitopes that are highly immunogenic in atopic individuals and certain strains of mice [4, 5]. Using MHCII binding prediction algorithms, we fine-mapped the peptide sequence of an immunodominant epitope in C57BL/6 mice and generated the corresponding I-Ab tetramer [6]. While this protocol specifically describes the generation of this particular Derp1117–131:I-Ab tetramer, such tetramers can be generated for any allergen so long as a well-defined immunogenic epitope in the allergen is known. Because epitope-specific CD4+ T cells are usually present at extremely low frequencies in sampled tissues [7], the use of tetramers is often coupled with magnetic cell enrichment techniques to improve the sensitivity of epitope-specific T cell detection [8, 9]. In our studies, we used tetramer-based cell enrichment in conjunction with intravascular staining and multiparameter flow cytometry to directly analyze allergen-specific T cells in the lymphoid organs, blood, and lungs of mice sensitized with HDM allergen [6]. This has become an instrumental tool in our studies of how Th2 effector and memory cell differentiation is regulated during atopic asthma. The following protocol explains the generation of tetramers using basic laboratory techniques that should enable most immunology labs without access to sophisticated resources to take advantage of this powerful technology. While pMHCI tetramers have been widely available for decades [10], the successful generation of pMHCII tetramers has been stymied by several technical issues including the instability of soluble MHCII heterodimers, the improper folding and glycosylation of MHCII proteins when expressed in bacteria, the difficulty of loading defined peptides into the MHCII binding groove, and the propensity of purified pMHCII to aggregate and precipitate from solution [11]. To overcome these difficulties, we have incorporated into our methods a number of technical innovations developed by several different labs over

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the years. Collectively, they have enabled us to achieve consistent success in the generation of most tetramer species. Our tetramers adopt a design introduced by Teyton and colleagues in which the intracellular domains of the MHCII alpha and beta chains are replaced with acidic and basic Fos-Jun leucine zipper motifs (Fig. 1) [12]. This forces heterodimerization of the extracellular domains of these subunits and increases stability of the soluble protein. To circumvent the challenges of loading peptides into the MHC binding groove, we utilize a strategy developed by Kappler and colleagues in which the peptide antigen is genetically encoded as a fusion product covalently linked to the N-terminus of the MHCII beta chain via a flexible linker [13]. To achieve proper posttranslational processing and high production yields, these constructs are stably expressed as secreted proteins in Drosophila S2 cells. The C-terminus of the alpha chain contains a signal sequence for site-specific biotinylation by E. coli BirA biotin ligase [14], ensuring that pMHCII complexes are in the proper orientation after eventual tetramerization with streptavidin. The BirA ligase is coexpressed along with the pMHCII constructs to mediate efficient in vivo biotinylation of the pMHCII during production, thereby eliminating the need for a distinct biotinylation step [15]. To facilitate purification, the C-terminus of the beta chain contains a 6× polyhistidine sequence to enable nickel column chromatography. Once purified, biotinylated pMHCII complexes are analyzed by Western blot for quantification and quality control and then tetramerized with premium grade streptavidin conjugated to fluorochromes of choice. peptide

flexible linker

MHCII alpha

MHCII beta

acidic leucine zipper

BirA

basic leucine zipper

SA

PE ?@

His

Biotin

Monomer Fig. 1 Design schematic of peptide:MHCII monomer and tetramer

Tetramer

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2  Materials 2.1  Plasmid Cloning

1. Expression plasmids (Addgene): pR I-Abα BirA; pR I-Abβ Derp1-117; pR BirA; pCoPuro. 2. XmaI restriction enzyme. 3. SpeI restriction enzyme. 4. Low-melt agarose. 5. DNA gel extraction kit. 6. DNA ligation kit. 7. Competent cells: DH5α, TOP10, XLI Blue, or equivalent. 8. LB agar plates. 9. LB broth. 10. Ampicillin. 11. AMP+ LB agar plates: LB agar supplemented with 100 μg/ mL ampicillin. 12. DNA miniprep kit. 13. DNA maxiprep kit.

2.2  S2 Cell Culture and Transfection

1. Drosophila S2 cells. 2. Complete S2 medium: Schneider’s Drosophila Medium, 10% heat-inactivated fetal bovine serum, 1% penicillin/streptomycin (100 U/mL each), 50 μg/mL gentamycin, 250 ng/mL fungizone. 3. Calcium Phosphate Transfection Kit (ThermoFisher). 4. Puromycin. 5. Complete S2 medium + Puro: Schneider’s Drosophila Medium, 10% heat-inactivated fetal bovine serum, 1% penicillin/streptomycin (100 U/mL each), 50 μg/mL gentamycin, 250 ng/ mL fungizone, 5 μg/mL puromycin. 6. DMSO. 7. Phosphate buffered saline (PBS). 8. 2× Hanks buffered saline (HBS). 9. Serum-free medium (SFM): Express five serum-free medium, 1% penicillin/streptomycin (100 U/mL each), 50 μg/mL gentamycin, 250 ng/mL fungizone. 10. SFM + Puro: Express five serum-free medium, 1% penicillin/ streptomycin (100 U/mL each), 50 μg/mL gentamycin, 250 ng/mL fungizone, 5 μg/mL puromycin. 11. Pluronic F-68 (ThermoFisher).

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12. 1 M Copper sulfate solution: Dissolve 12.5 g CuSO4 (copper (II) sulfate pentahydrate) in 50 mL dH2O, and pass through sterile 0.2 μm filter. 13. 1 mg/mL D-biotin solution: Dissolve 50 mg D-biotin in 49.5 mL dH2O and pH to 7.0 with 1 N NaOH (this will require approximately 240 μL). Bring volume to 50 mL, and pass through sterile 0.2 μm filter. 14. 3 L spinner or 2 L shaker flasks. 15. 500 mL conical centrifuge tubes. 16. 24 cm round grade 3 filter paper (Whatman). 17. 0.45  μm bottletop vacuum filter. 18. 1 L bottle. 19. 10% sodium azide solution: 10% sodium azide in PBS. 2.3  Protein Purification

1. HisBind Purification Kit (Novagen-EMD Biosciences). 2. HisBind nickel resin (provided in HisBind Purification Kit). 3. 1 × 10 cm glass column. 4. Ring stand. 5. Clamp to attach glass column to ring stand. 6. 500 mL beaker: Waste beaker. 7. 1 L beaker: Collection beaker. 8. 425–600 μm diameter glass beads. 9. 1× charge buffer: 50 mM NiSO4 (provided in HisBind Purification Kit). 10. 1× bind buffer: 500 mM NaCl, 20 mM Tris-HCl, 5 mM imidazole, pH 7.9 (provided in HisBind Purification Kit). 11. Luer-lock stopcock. 12. Small diameter flexible tubing: 1/16 in internal diameter (I.D.) tubing (Tygon). 13. Small diameter flexible tubing: 1/8 in I.D. tubing (Tygon). 14. 10 mL syringe. 15. Hemostat (or similar clamp). 16. PBSA: PBS, 0.05% sodium azide. 17. Octyl β-d-glucopyranoside (Sigma-Aldrich).

(OGP)

detergent

18. 1× elute buffer + 0.2% OGP: 1 M imidazole, 500 mM NaCl, 20 mM Tris–HCl pH 7.9, 0.2% OGP. 19. 1× strip buffer: 500 mM NaCl, 100 mM EDTA, 20 mM Tris– HCl, pH 7.9 (provided in HisBind Purification Kit). 20. Amicon Ultra-15 50 kD concentrating filter (Millipore).

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21. PBSA + 0.2% OGP: PBS, 0.05% sodium azide, 0.2% OGP. 22. 0.2  μm microfuge spin filter (Corning-Costar). 23. Spectrophotometer. 2.4  Protein Purification

1. Streptavidin-R-Phycoerythrin (SA-PE) (ProZyme). 2. Streptavidin-Allophycocyanin (SA-APC) (ProZyme). 3. Tris-glycine acrylamide gel (4–15% or other concentration suitable for 50–100 kD proteins). 4. Protein molecular weight markers. 5. 4× Native PAGE sample buffer: 200 mM Tris pH 6.8, 60% glycerol, 20 mg/mL bromophenol blue. Mix one part to three parts sample prior to gel loading. 6. 5× Native Tris-glycine PAGE running buffer: 125 mM Tris, 1.25 M glycine. Dilute to 1× with dH2O before use. 7. Nitrocellulose membrane and filter paper for Western blot transfer. 8. 10× TBS-Tween 0.1% (TBST): 200 mM Tris pH 7.6, 1.37 M NaCl, 1% Tween-20. Dilute to 1× with dH2O before use. 9. Streptavidin conjugated to horseradish peroxidase (HRP) or fluorescent dye. 10. Enhanced chemiluminescence (ECL) substrate.

3  Methods 3.1  Expression Plasmid Cloning (for New Antigens)

All plasmids required to generate I-Ab Derp1-117 tetramers have been deposited at Addgene. The pR I-Ab beta expression plasmid is designed with convenient XmaI and SpeI restriction sites flanking the peptide sequence to facilitate cloning of new constructs with different peptide antigens. This section describes the cloning of a new construct using complementary oligonucleotide synthesis. 1. Design complementary oligonucleotides for the new peptide insert (Fig. 2). Ideally, new peptide antigens should be designed with a minimal length including just the 9-mer MHCII binding core plus two additional N-terminal residues. No extra C-terminal residues are needed due to the presence of the linker. The ends of the oligos should extend three bases beyond the XmaI and SpeI restriction sites on each side to facilitate enzyme activity. 2. Anneal oligos. Make 100 μM stocks of each oligo in dH2O, and mix 5 μL of each oligo in a 0.2 mL PCR tube.

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substitute new peptide sequence here sense oligo CCCGGGACTGAGGGCNNNNNNNNNNNNNNNNNNNNNNNNNNNNNNNNNGGCTGTGGAGGTACTAGT original sequence (pR I-Ab beta Derp1-117) MHCII binding pocket position -2 -1 1 2 3 4 5 6 7 8 9 XmaI SpeI S P G T E G C Q I Y P P N V N K I G G G G T S G CCCGGGACTGAGGGCTGCCAAATTTACCCACCAAATGTAAACAAAATTGGCGGGGGAGGTACTAGTGGC-3' GGGCCCTGACTCCCGACGGTTTAAATGGGTGGTTTACATTTGTTTTAACCGCCCCCTCCATGATCA signal peptide linker Derp1117-127 peptide sequence GGGCCCTGACTCCCGNNNNNNNNNNNNNNNNNNNNNNNNNNNNNNNNNCCGACACCTCCATGATCA substitute new peptide sequence here

antisense oligo

Fig. 2 Cloning strategy for new antigenic peptide sequence. Complementary oligos are designed to create a double-stranded DNA fragment that will replace the original peptide sequence with a new one. The fragment encompasses flanking XmaI and SpeI restriction sites to facilitate cloning into the pR I-Abβ vector

3. Run the following program in a thermocycler: 95 °C 30 s; step −0.1 °C/s to 4 °C; 4 °C forever. Keep tube on ice after PCR reaction is complete. 4. Run the double-stranded product on a 2% low-melt agarose gel, and purify the band of expected size using a suitable DNA gel extraction kit. 5. Digest the parent vector (pR I-Ab Derp1–117) and new insert fragment with XmaI and SpeI restriction enzymes. 6. Run the digested vector and insert on 1.2% and 2% low-melt agarose gels, respectively, and purify bands of expected sizes. 7. Ligate purified digested vector and insert using a suitable DNA ligation kit. Remember to set up a vector control (no insert). 8. Transform competent cells with ligation reactions and plate out on AMP+ LB agar plates. 9. Miniprep 3–4 colonies using a suitable miniprep kit. Spot replica colonies on a new ampicillin plate for future inoculation of maxiprep cultures. 10. Screen miniprep DNA by sequencing the insert region with the upstream primer pRFOR2 (5′-TGTGCAAAAGAGGTGAATCG-3′). 11. Maxiprep a sequence-verified clone using a suitable maxiprep kit.

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3.2  Drosophila S2 Cell Transfection

Expression of biotinylated pMHCII in Drosophila S2 cells largely follows the DES Drosophila Expression System from ThermoFisher. 1. Using healthy S2 cells in log growth phase, set up 3 × 106 cells in 3 mL Complete S2 medium in 60 mm tissue culture dishes. Incubate overnight at 28 °C. You can place three 60 mm dishes inside a 150 mm dish to provide an additional sterile barrier (see Notes 1–3). 2. Transfect cells using the Calcium Phosphate Transfection Kit. Make sure all reagents are warmed to room temperature prior to transfection. 3. For each transfection, set up a 1.5 mL microfuge tube with the following: pR I-Abα BirA plasmid 9 μg pR I-Abβ (Derp1-117) plasmid 9 μg pR BirA plasmid 9 μg pCoPuro plasmid 1 μg CaCl2 (added last) 240 mM (36 μL of 2 M) Sterile dH2O needed to achieve a total volume of 300 μL. 4. Slowly add mixture dropwise over the course of 1 min into another 1.5 mL microfuge tube containing 300 μL 2× HBS while vortexing. This process produces the fine precipitate that mediates transfection of DNA. 5. Incubate at room temperature for 30–40 min. 6. Slowly drop mixture into cell culture while swirling dish. 7. Incubate 1 day at 28 °C. 8. Replace Complete S2 medium. Without disturbing cells, pipet Complete S2 medium from the dish into a 15 mL centrifuge tube. 9. Carefully rinse cells with 1 mL fresh Complete S2 medium and transfer into the same 15 mL tube. Add 1 mL fresh Complete S2 medium to dish so the cells don’t dry out. 10. Spin down the 15 mL tube at 400 × g-force for 5 min at room temperature, and aspirate and resuspend the cell pellet with 2 mL fresh Complete S2 medium and transfer to the dish to restore volume to 3 mL. 11. Incubate 2 days at 28 °C. 12. Resuspend cells by pipetting and transfer to a T75 flask. Due to evaporation, there will only be about 2.5 mL of cells left per dish. Add 13 mL fresh Complete S2 medium + Puro. From here on, transfected cells should always be kept in puromycin-­ containing media. 13. Incubate 7 days at 28 °C.

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14. Scale up culture for freezing and protein expression. Split cells 1:10 as follows:

(a) Place 0.5 mL of cells into T25 flask containing 4.5 mL of Complete S2 medium + Puro for maintenance. Split this ­culture 1:10 every 7 days until supernatant is successfully harvested from scale-up cultures (see Note 4).



(b) Place 1.5 mL of cells into T75 flask containing 13.5 mL Complete S2 medium + Puro for freeze-down.



(c) Place 5 mL into 2× T225 flasks each containing 45 mL SFM + Puro for scale-up.

15. Incubate cultures at 28 °C. 16. When the T75 freeze-down culture has reached ~107 cells/ mL (usually about 3–5 days), transfer to a 15 mL tube, spin down, and aspirate supernatant down to ~2.5 mL. 17. Add 2.5 mL fresh Complete S2 medium, 0.6 mL sterile DMSO, and suspend cells. This will result in ~3.0 × 107 cells/ mL in 45% fresh complete S2 medium, 45% conditioned complete S2 medium, and 10% DMSO. 18. Aliquot ~1 mL to each of five labeled cryotubes. 19. Freeze down and transfer to liquid nitrogen for long-term storage. 3.3  pMHCII Expression

1. When the T225 SFM culture reaches 5–10 × 106 cells/mL (about 5–7 days), transfer the whole cell culture (100 mL) into a sterile 3 L spinner flask or 2 L shaker flask. Rinse remaining cells in the T225 flasks with 1.0 L prewarmed SFM + Puro and pool together in the new flask (see Note 5). 2. Add 10 mL Pluronic F-68 to a final concentration of 0.1% (see Note 6). 3. Incubate at 28 °C with constant spinning or shaking at 125 rpm. Lower speeds (~100 rpm) are okay too, but the cell densities may be slightly lower. 4. When the culture reaches 1.0–1.5 × 107 cells/mL (about 5–7 days), induce expression by adding 880 μL of 1 M copper sulfate solution and 2.2 mL of 1 mg/mL D-biotin solution for a final concentration of 800 μM CuSO4 and 2 μg/mL D-biotin. 5. Incubate 14 days at 28 °C, spinning or shaking at 125 rpm. 6. Transfer and spin down cells in 500 mL conical centrifuge tubes at ~1000 × g-force for 10 min in a tabletop centrifuge. 7. Transfer supernatant to new 500 mL conical tubes and spin down again to remove residual cells.

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8. Filter supernatant through a piece of 24 cm round grade 3 filter paper folded into a funnel set up over a 0.45 μm bottletop vacuum filter, draining into a 1 L bottle. 9. Add 5 mL of 10% sodium azide solution to achieve a final concentration of 0.05%, and store at 4 °C. If aggregates form in the supernatant over time, filter it again before proceeding to purification. 3.4  pMHCII Purification

Soluble monomeric pMHCII complexes are purified from cell culture supernatants via nickel affinity chromatography targeting the 6× His epitope on the C-terminus of the MHCII beta chain. This section describes a protocol adapted from the HisBind Purification Kit from Novagen-EMD Biosciences. Alternatively, purification by immunoaffinity chromatography may be used to achieve higher purity product (see Note 7). 1. Add 8 mL of well mixed HisBind nickel resin slurry to a 1 × 10 cm glass column clamped on a ring stand with a 500 mL waste beaker placed below. The resin is supplied as a 50% suspension slurry, so 4 mL of packed resin will remain in the column after the liquid portion has drained away (see Note 8). 2. Add about 1/4 inch layer of 425–600 μm diameter glass beads to the top of the resin bed to protect the top surface from disruption as fluids are added. 3. Wash the column by adding 12 mL (three column bed volumes) dH2O and let drain. 4. Charge the column by adding 20 mL (five column bed volumes) of 1× charge buffer and letting it drain. The color of the resin should turn from white to blue. 5. Add 12 mL (three column bed volumes) of 1× bind buffer to the column and let it drain. The blue color will fade a little, but overall the column should still be blue. 6. Affix a closed Luer-lock stopcock to the bottom of the column. Add a few mL of culture supernatant to the top of the resin, at first by slowly running it down the inside of the column to prevent disruption of the resin. Secure the cap on top of the column. 7. Place the bottle containing the culture supernatant on a shelf a few feet above the top of the column. Set up a length of small diameter flexible tubing that will connect the bottle of culture supernatant to the cap on top of the column. Rather than sticking the tubing directly into the bottle of supernatant, you can connect the end to a 1 mL pipet (with cotton removed) and then insert the pipet into the bottle. This will allow for better draining of the bottle. We recommend using 1/16 in I.D. tubing with short lengths (~2 in) of 1/8 in I.D. tubing at

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each end to act as couplings for attachment to the pipet at one end and the nipple on the column cap at the other end. Make sure that the lowest point in the tubing loop hangs below the bottom of the column. This safety loop will prevent the column from running dry after the bottle has emptied. 8. Prime the tubing by using a 10 mL syringe to draw fluid from the bottle through the entire length of tubing, and then clamp the tubing about an inch from the end with a hemostat. Remove the syringe, and attach the end of the tubing to the nipple on the column cap. Reattach the column cap to the top of the column. 9. Unclamp the tubing and open the stopcock. The fluid should start dripping from the bottom of the column. Use the stopcock to adjust the flow rate to about 2 mL per minute. Let the column drip overnight, collecting the flow-through fraction in a new 1 L bottle. The safety loop should prevent the resin from drying out once the supernatant bottle has emptied. 10. Elevate the tubing so that the remaining supernatant passes through the column. Close the stopcock, and remove the cap and tubing from the column. Transfer the column on the ring stand to a bench in the lab. The column should be bluish green with some hint of brown impurities mixed in. 11. Wash the column by adding 8 mL (two column bed volumes) PBSA, collecting the flow-through into a 15 mL tube for backup. 12. Add 16 mL (four column bed volumes) of 1× elute buffer +0.2% OGP to the top of the column. Collect eluted sample into a new 15 mL tube. 13. Regenerate the column by adding 12 mL (three column bed volumes) 1× strip buffer and let it drain. The resin should change in color from blue to white. Close the stopcock, and add another 2 mL to the top of the column. Replace the cap on the column, and store the column at 4 °C. 14. If the sample contains precipitated material, pass through a 0.2 μm spin filter before moving on. 15. Add the entire sample (~16 mL) to the upper chamber of an Amicon Ultra-15 50 kD concentrating filter and spin at ~1500 × g-force until the volume is reduced to ~1 mL. To prevent overspinning, start out conservatively with a 5 min spin and then adjust subsequent spin times accordingly. 16. Discard the flow-through in the lower chamber and add 15 mL PBSA + 0.2% OGP to the upper chamber. Spin down to ~1 mL again. 17. Repeat twice for a total of three buffer exchanges which will reduce the imidazole concentration to well below 1 mM.

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18. Transfer sample to a 0.2 μm microfuge spin filter and spin at >16,000 × g for 1 min to remove any aggregates. 19. Measure the OD280 on spectrophotometer (try a 1:10 dilution first) of the sample, and calculate the pMHCII sample concentration. concentration = OD280 × dilution factor/extinction coefficient (final units in mg/mL) The extinction coefficient for pMHCII is ~1.0 cm−1(mg/ mL)−1 (i.e., an OD280 of 1.0 corresponds to a concentration of 1.0 mg/mL). 3.5  Titration of Biotinylated pMHCII

Because the sample still contains a significant amount of impurities and the biotinylation process is not 100% efficient, your concentration based on the OD280 reading will be an overestimate of the true concentration of biotinylated pMHCII. The following titration assay will empirically determine how much of your pMHCII sample should be mixed with streptavidin-fluorochrome conjugates (SA-PE or SA-APC) to create fully armed tetramers (see Notes 9 and 10). 1. Prepare 35 μL of pMHCII sample at 200 μg/mL in PBSA. 2. Prepare a twofold dilution series starting with 2 μM SA-PE or SA-APC. Start with 30 μL of 2 μM, and mix 15 μL of this stock with 15 μL of PBSA to make a 30 μL of 1 μM dilution. Repeat to make a 1:2 serial dilution series in PBSA extending down to 125 nM. 3. Add 5 μL of the 200 μg/mL pMHCII stock (1 μg total) to each tube and then 5 μL of each of the SA-PE (or SA-APC) dilutions as shown in Table 1. Add 5 μL PBSA to the first and last samples to bring the volume of all samples to 10 μL. 4. Incubate for 30 min at room temperature. 5. Run the samples on a 4–15% PAGE gel under non-denaturing conditions. Load molecular weight markers or pMHCII from a previous prep in a separate lane for reference. 6. Transfer the proteins to a nitrocellulose membrane. 7. Analyze biotinylated pMHCII by western blotting with a streptavidin probe and enhanced chemiluminescence (ECL) or fluorescence based detection. 8. Lane #1 should show a single strong band at about 80 kD, corresponding to biotinylated pMHCII (Fig. 3). Lane #2, which contains the maximum dose of SA-PE, should present no band because all of the biotinylated pMHCII should be bound to SA-PE. The 80 kD band will eventually reappear as the SA-PE concentration falls and can no longer occupy all of

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Table 1 Streptavidin titration of biotinylated pMHC Total amount of Tube pMHC

Volume of pMHC to be added

Total amount of SA-PE

Volume of SA-PE to be added

1

1 μg

5 μL of 200 μg/mL

None

0

2

1 μg

5 μL of 200 μg/mL

10 pmol

5 μL of 2 μM

3

1 μg

5 μL of 200 μg/mL

5 pmol

5 μL of 1 μM

4

1 μg

5 μL of 200 μg/mL

2.5 pmol

5 μL of 500 nM

5

1 μg

5 μL of 200 μg/mL

1.25 pmol

5 μL of 250 nM

6

1 μg

5 μL of 200 μg/mL

0.625 pmol

5 μL of 125 nM

7

None

0

10 pmol

5 μL of 1 μM

Fig. 3 Representative Western blot of biotinylated pMHC monomer following titration with streptavidin. Titrated mixtures of pMHC monomer and SA-PE from Table 1 were blotted with a streptavidin-HRP probe and visualized by enhanced chemiluminescence. The indicated band represents free biotinylated pMHC that was not saturated by SA-PE

biotinylated pMHCII. The SA-PE amount where the biotinylated pMHCII band just reappears defines the saturation point. If no saturation point is observed, then repeat the assay making adjustments to the amounts of pMHCII loaded in each lane. Be conservative with your assumptions so that you always error on the side of underestimating the true concentration of biotinylated pMHCII. 9. To calculate the concentration of biotinylated pMHCII in your sample, first multiply the molar amount of SA-PE in the saturation point by a factor of 4 to determine the molar amount of biotinylated pMHCII that was loaded in that lane. Divide this amount of biotinylated pMHCII by the

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5 μL volume in which it was delivered to determine the concentration of biotinylated pMHCII in the “200 μg/mL” diluted stock of pMHCII monomer prepared in Subheading 3.5, step 1. Then multiply this concentration by the dilution factor to calculate the concentration of biotinylated pMHCII in the original sample. 3.6  pMHCII Tetramerization

It is important to mix at least 4 moles of biotinylated pMHCII for each mole of SA-PE (or SA-APC) to ensure that the resulting tetramer is fully multimerized. For making calculations, use the concentration of biotinylated pMHCII empirically determined in the previous section (Subheading 3.5) and the concentration of SA-PE specified in the data sheet for each production lot from the manufacturer. 1. Calculate how many moles of tetramer to generate (see Notes 11 and 12). 2. Determine the volume of SA-PE that is needed by dividing the desired molar amount of tetramer by the concentration of the SA in the SA-PE. 3. Determine the molar amount of biotinylated pMHCII that is needed by multiplying the desired molar amount of tetramer by 4 and then again by a safety factor of 2 to ensure that there is an excess of pMHCII to SA-PE. To determine the volume of biotinylated pMHCII needed, divide this molar amount by the empirically determined concentration of biotinylated pMHCII from the previous titration step (Subheading 3.5) (see Note 13). 4. Mix the calculated volumes of SA-PE and biotinylated pMHCII together, and incubate at room temperature for 30 min. 5. Spin the tetramerized product through a 0.2 μm microfuge spin filter to remove aggregates. 6. Measure the OD566 (try a 1:20 dilution first), and calculate the PE concentration. concentration = OD566 × dilution factor/extinction coefficient (final units in μM). The extinction coefficient for PE is 1.96 cm−1 μM−1 (i.e., an OD566 of 1.96 corresponds to a concentration of 1.0 μM). The extinction coefficient for APC at 650 nm is 0.70 cm−1 μM−1 (i.e., an OD650 of 0.70 corresponds to a concentration of 1.0 μM). 7. Ideally, there should be exactly one molecule of SA conjugated to each molecule of PE in the SA-PE stock. However, this stoichiometry is usually slightly off. The data sheet for each lot of SA-PE should specify the individual concentrations of SA and PE. Convert the measured PE concentration of your sample to that of SA by multiplying by the [SA]:[PE] ratio obtained

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from the data sheet. Because SA constitutes the core of the tetramer, the concentration of SA should be the actual concentration of the tetramer. 8. Dilute the concentration of tetramer to 1.0 μM, and store at 4 °C in the dark. Do not freeze. 9. The final staining concentration for cells is usually 10 nM, but every production lot of tetramer should be titrated in a pulldown experiment on mice immunized with the corresponding antigen (usually peptide + CFA immunized by subcutaneous injection and assessed 7 days after immunization). 10. After determining how much pMHCII is needed to make an ideally sized batch of tetramer, freeze down appropriately sized aliquots of your remaining pMHCII at −80 °C for future batches.

4  Notes 1. In general, S2 cells are grown at 28 °C without CO2. Maintain humidity in the incubator with a tray of water. 2. Optimal medium volumes for tissue culture vessels for S2 cells are: T25 5 mL T75 15 mL T225 50 mL 2 L shaker 1 L 3. A T25 flask of parental cells in 5 mL complete medium should be continuously maintained, split no more than 1:10 every 3–4 days until all planned transfections are completed. 4. Transfected S2 cell cultures can be maintained indefinitely by splitting them 1:10 in complete medium + drug about once per week. S2 cells require a soluble autocrine growth factor for optimal growth, so any split beyond 1:10 will result in a substantial slowdown in growth and possible loss of the culture. Because the cells do not do well at low densities, subcloning is very difficult. 5. Avoid the use of shaker flasks with baffled sides, as these will increase the generation of bubbles and foam which can damage cells. 6. Pluronic F-68 is a surfactant that is added to the shaker or spinner culture to prevent cell shearing. It may also inhibit aggregation of secreted pMHCII in the media. 7. This protocol describes pMHCII purification via nickel column chromatography because this process is easy to set up with commercially available kits and does not pose a high risk

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for pMHCII precipitation. However, the purity of the resulting pMHCII is low with large amounts of contaminants from the S2 cell culture. While these contaminants do not significantly impact the function of the final tetramer reagents, they complicate calculations of pMHCII protein concentration and are aesthetically unpleasing due to their visible brown hue. IF desired, size exclusion chromatography (S-300 Sepharose or similar) may be used to remove most of these contaminants, which are much larger than pMHCII complexes. Alternatively, immunoaffinity chromatography may be used to achieve much greater purities of pMHCII. We have found that the I-Ab-­ specific antibody clone Y3P, but not M5/114, works very well for this purpose. However, an associated risk with immunoaffinity purification is that the pMHCII may sometimes precipitate from solution during elution with the low or high pH buffers that are necessary to disrupt antibody-antigen binding. The tendency for pMHCII to precipitate varies for different constructs and can only be reliably predicted by trial and error. 8. The HisBind resin has a binding capacity of about 8 mg protein per mL. 4 ml of resin should therefore be capable of binding 32 mg of protein, which is above the 3–30 mg yields that are typical of 1 L cultures. If a yield of more than 25 mg is achieved, it may be worth running the supernatant through a second time to harvest any protein that was missed due to saturation of the column. 9. The quality of tetramers is highly dependent on the quality of the streptavidin reagents used. We use SA-PE and SA-APC from ProZyme, which have worked very well compared to other vendor sources and come with detailed specifications including relative concentrations of SA and fluorochrome in each production lot. 10. Note that tetramer concentrations will only be as accurate as the listed concentration of SA-PE from the manufacturer, so it is best to titrate your sample to the exact tube of streptavidin that you will be using to tetramerize your pMHCII. 11. The ideal concentration of the final tetramer is 1 μM, which is used at 1:100 for staining at a final concentration of 10 nM. The testing of most cell samples will therefore only require 1–2 μL of reagent, so 500 μL of tetramer would be sufficient for a few hundred tests and would represent a reasonable amount of tetramer to make in a single batch such that most of it is used before getting too old. 500 μL of tetramer at 1 μM corresponds to 500 pmol, but because some tetramer is lost during tetramerization, you may wish to start off making somewhere between 500 pmol and 1.0 nmol tetramer.

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12. The shelf life of tetramers varies considerably for different MHC and antigens, but a good arbitrary expiration date is 1 year. 13. When tetramerizing pMHCII, always err on the side of excess pMHCII. The affinity of monomeric pMHCII for TCR is several orders of magnitude lower than that of tetramer, so excess free pMHCII in the final reagent is not a concern. References 1. Locksley RM (2010) Asthma and allergic inflammation. Cell 140(6):777–783 2. Nepom GT (2012) MHC class II tetramers. J Immunol 188(6):2477–2482 3. Chua KY et al (1988) Sequence analysis of cDNA coding for a major house dust mite allergen, Der p 1. Homology with cysteine proteases. J Exp Med 167(1):175–182 4. Yssel H et al (1992) T cell activation-inducing epitopes of the house dust mite allergen Der p I. Proliferation and lymphokine production patterns by Der p I-specific CD4+ T cell clones. J Immunol 148(3):738–745 5. Hoyne GF, Callow MG, Kuo MC, Thomas WR (1994) Inhibition of T-cell responses by feeding peptides containing major and cryptic epitopes: studies with the Der p I allergen. Immunology 83(2):190–195 6. Hondowicz BD et al (2016) Interleukin-­ 2-­ dependent allergen-specific tissue- resident memory cells drive asthma. Immunity 44(1):155–166 7. Jenkins MK, Moon JJ (2012) The role of naive T cell precursor frequency and recruitment in dictating immune response magnitude. J Immunol 188(9):4135–4140 8. Moon JJ et al (2007) Naive CD4(+) T cell frequency varies for different epitopes and predicts repertoire diversity and response magnitude. Immunity 27(2):203–213

9. Legoux FP, Moon JJ (2012) Peptide:MHC tetramer-based enrichment of epitope-specific T cells. J Vis Exp 68:e4420 10. Altman JD et al (1996) Phenotypic analysis of antigen-specific T lymphocytes. Science 274(5284):94–96 11. Vollers SS, Stern LJ (2008) Class II major histocompatibility complex tetramer staining: progress, problems, and prospects. Immunology 123(3):305–313 12. Scott CA, Garcia KC, Carbone FR, Wilson IA, Teyton L (1996) Role of chain pairing for the production of functional soluble IA major histocompatibility complex class II molecules. J Exp Med 183(5):2087–2095 13. Kozono H, White J, Clements J, Marrack P, Kappler J (1994) Production of soluble MHC class II proteins with covalently bound single peptides. Nature 369(6476):151–154 14. Beckett D, Kovaleva E, Schatz PJ (1999) A minimal peptide substrate in biotin holoenzyme synthetase-catalyzed biotinylation. Protein Sci 8(4):921–929 15. Yang J, Jaramillo A, Shi R, Kwok WW, Mohanakumar T (2004) In vivo biotinylation of the major histocompatibility complex (MHC) class II/peptide complex by coexpression of BirA enzyme for the generation of MHC class II/tetramers. Hum Immunol 65(7):692–699

Chapter 15 The Generation and Use of Allergen-Specific TCR Transgenic Animals Manon Vanheerswynghels, Wendy Toussaint, Martijn Schuijs, Leen Vanhoutte, Nigel Killeen, Hamida Hammad, and Bart N. Lambrecht Abstract The generation of allergen-specific TCR transgenic animals allows for the characterization of allergen-­ specific T-cell responses in vivo and in vitro and is a powerful tool to study adaptive immunity to allergens. Here we describe an approach starting from the isolation of antigen-specific T-cell hybridomas and using PCR, flow cytometric, and co-culture methods to obtain antigen-specific MHC class II-restricted CD4+ TCR transgenic mice on the Rag2−/− background. Key words Hybridoma, T-cell fusion, T-cell receptor, Allergen, Transgenic

1  Introduction The generation of allergen-specific T-cell receptor (TCR) transgenic animals has made a tremendous impact on the field of immunology and has greatly enhanced the possibilities to investigate the role of antigen-specific T cells in (allergic) disease [1–3]. Recognition of the TCR in the context of MHC-peptide ligands is critical in developmental selection and peripheral activation of T cells. A greater understanding of the TCR-ligand interaction as well as the activation and differentiation of T cells in vivo is essential for our understanding of pathology during disease. The advent of TCR transgenic mice has proven to be a powerful tool that enables researchers to phenotypically monitor a cohort of T cells with defined antigen specificity. In this way, various TCR transgenic mouse models have been instrumental for elucidating the process of thymic selection, central and peripheral T-cell tolerance, and T-cell responsiveness to different antigens or allergens [2, 4–8]. To obtain antigen-specific clones, T-cell hybridomas were generated by fusing T-cell lymphoma cell line BW5147 with in vitro restimulated CD4+ spleen cells of C57BL/6 mice that received the R. Lee Reinhardt (ed.), Type 2 Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1799, https://doi.org/10.1007/978-1-4939-7896-0_15, © Springer Science+Business Media, LLC, part of Springer Nature 2018

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specific antigen in vivo. Antigen-specific clones were single cell sorted and expanded, and a single clone was used for RNA isolation and cDNA synthesis using random hexamer primers. cDNA was amplified using a PCR screening panel, and the amplified partial Vα and Vβ products were sequenced and used to determine the full-length TCR. Next, antigen-specific hybridoma cDNA was used to amplify the full-length Vα- and Vβ-chain with a proofreading enzyme using full-length primers. The amplified bands were cloned in the TCR expression vectors p428 and p783, co-injected in an equimolar concentration of 1 ng/μL into the pronuclei of fertilized C57BL/6 Rag2−/− oocytes, and transferred to a foster mother. Founders were screened for TCR functionality by flow cytometry, and offspring was genotyped by PCR with primers specific for the Vα- and Vβ-transgene. Below, we describe a detailed protocol used for the generation of novel mouse models with MHC class II-restricted CD4+ TCR transgenic T cells.

2  Materials 2.1  Immunization

1. Antigen formulated on aluminum hydroxide: dilute the antigen in sterile PBS in a 15 mL conical tube. Shake the aluminum hydroxide (Imject Alum, Thermo Scientific) well before use, and add it dropwise to the antigen in a 1/20 dilution while gently vortexing. Vortex the solution for 1 h prior to injection. Inject 500 μL intraperitoneally with a 26-gauge needle (see Note 1).

2.2  T-Cell Fusion [9]

1. BW5147 lymphoma cell line.

2.2.1  Culture of the BW5147 Fusion Partner

2. DMEM10: DMEM supplemented with 10% fetal calf serum (FCS), nonessential amino acids (NEAA, 100× diluted from commercial stock), 50 μg/mL gentamycin, 50 μM beta-­ mercaptoethanol, and 1 mM sodium pyruvate (all Gibco). 3. 25cm2 culture flask.

2.2.2  In Vitro Restimulation of Lymph Node Cells

1. 70 μm cell strainer. 2. 96-well round bottom plate. 3. HBSS (Gibco). 4. RPMI5: RPMI supplemented with 5% FCS, 50 μg/mL gentamycin, 50 μM beta-mercaptoethanol, and 2 mM GlutaMAX (all Gibco). 5. Peptide solution: prepare 100 μL peptide solution per well. Dilute the peptide to 10 μg/mL in RPMI5 (final concentration of 5 μg/mL in the well).

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1. MACS buffer: prepare a solution of 0.5% bovine serum albumin (BSA) and 2 mM EDTA in PBS. 2. CD4+ T-cell isolation kit, Miltenyi Biotech or equivalent.

2.2.4  T-Cell Fusion

1. Two 400 mL beakers and two 600 mL beakers. 2. DMEM (Gibco). 3. 50% polyethylene glycol (PEG), commercial solution of 50% (Roche). 4. DMEM20: DMEM supplemented with 20% FCS, NEAA (100× diluted from commercial stock), 50 μg/mL gentamycin, 50 μM beta-mercaptoethanol, and 1 mM sodium pyruvate (all Gibco). 5. 96-well flat bottom plate.

2.2.5  Culture of the Fused T Cells

1. 2× HAT medium: dilute 50× hypoxanthine/aminopterin/thymidine supplement (Gibco) to 2× with DMEM20 medium. Store up to 2 months at 4 °C. 2. 1× HT medium: dilute 100× hypoxanthine/thymidine supplement (Gibco) to 1× in DMEM20. Store up to 2 months at 4 °C. 3. 24-well plate. 4. DMEM20: DMEM supplemented with 20% FCS, NEAA (100× diluted from commercial stock), 50 μg/mL gentamycin, 50 μM beta-mercaptoethanol, and 1 mM sodium pyruvate (all Gibco). 5. DMEM10: DMEM supplemented with 10% FCS, NEAA (100× diluted from commercial stock), 50 μg/mL gentamycin, 50 μM beta-mercaptoethanol, and 1 mM sodium pyruvate (all Gibco).

2.3  Screening of the T-Cell Clones 2.3.1  Culture of Bone Marrow-Derived Dendritic Cells [10]

1. 70% ethanol. 2. RPMI (Gibco). 3. Mortar and pestle. 4. 70 μm cell strainer. 5. Osmotic lysis buffer: prepare a solution of 155 mM NH4Cl, 1 mM KHCO3, 100 μM EDTA in water. Adjust the pH to 7.1–7.4 using HEPES. Sterilize using a 0.22 μm bottle top filter. 6. 100 mm petri dish. 7. RPMI5: RPMI supplemented with 5% FCS, 50 μg/mL gentamycin, 50 μM beta-mercaptoethanol, and 2 mM GlutaMAX (all Gibco).

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8. Recombinant mouse GM-CSF: dilute GM-CSF to a final concentration of 20 ng/mL in RPMI5. 9. PBS. 2.3.2  Initial Screening

1. 96-well round bottom plate. 2. DMEM10: DMEM supplemented with 10% FCS, NEAA (100× diluted from commercial stock), 50 μg/mL gentamycin, 50 μM beta-mercaptoethanol, and 1 mM sodium pyruvate (all Gibco). 3. Peptide solution: prepare 100 μL peptide solution per well. Dilute the peptide to 4 μg/mL in DMEM10 (final concentration of 1 μg/mL). 4. Mouse IL-2 ELISA ready-set-go (eBioscience). 5. Microtiter plate reader.

2.3.3  Confirmation Screening

1. Material and buffers as described in Subheading 2.3.2, initial screening, except for the peptide solution. 2. Peptide solution: dilute the peptide to 4 μg/mL in DMEM10 (final concentration of 1 μg/mL in the well), and prepare a 1 in 2 serial dilution. Do this by each time transferring half of the tube content to a new tube with an equal volume of DMEM10, for five steps in a row. Prepare 50 μL per sample (see Note 14). 3. Anti-mouse CD3 antibody, anti-mouse CD4 antibody, and anti-mouse CD69 antibody. 4. Fixable live/dead staining (e Bioscience). 5. Fc-blocking reagent. 6. FACS buffer: prepare a solution of 0.25% BSA, 1 mM EDTA, and 0.05% NaN3 in PBS. 7. Compensation beads (e Bioscience). 8. Flow cytometer.

2.3.4  Evaluation of the TCR Subtype

1. Mouse Vβ TCR screening panel (Becton Dickinson). 2. Anti-mouse Vα2 antibody, anti-mouse Vα3.2 antibody, anti-­ mouse Vα8.3 antibody, anti-mouse Vα11.1,11.2 antibody, anti-mouse CD3 antibody, and anti-mouse CD4 antibody. 3. Material and buffers as described in Subheading 2.3.3 confirmation screening (see items 4–8).

2.3.5  Freezing of Peptide-Specific T-Cell Hybridomas

1. Freezing medium: DMEM supplemented with 20% FCS, NEAA (100× diluted from commercial stock), 50 μg/mL gentamycin, 50 μM beta-mercaptoethanol, 1 mM sodium pyruvate (all Gibco) and 10% DMSO (Invitrogen).

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2. Freezing container. 3. Cryovials. 4. −70 °C freezer. 5. −150 °C freezer or liquid nitrogen. 2.4  Generation of Monoclonal Cells

1. Anti-mouse CD3 antibody, anti-mouse CD4 antibody, antibody against the corresponding Vα and Vβ. 2. Material and buffers as described in Subheading 2.3.3 confirmation screening (see items 4–7). 3. FACS ARIA cell sorter or equivalent. 4. DMEM20: DMEM supplemented with 20% FCS, NEAA (100× diluted from commercial stock), 50 μg/mL gentamycin, 50 μM beta-mercaptoethanol, and 1 mM sodium pyruvate (all Gibco). 5. 96-well round bottom plate. 6. 24-well plate. 1. Material and buffers as described in Subheading 2.3.3 confirmation screening.

2.5  Confirmation and Selection of the Monoclonal T-Cell Clones

2. Material and buffers as described in Subheading 2.3.5 freezing of peptide-specific T-cell hybridomas.

2.6  Cloning

1. RNAse-free 1.5 mL tubes and tips.

2.6.1  RNA Preparation

2. TriPure reagent (Roche). 3. Chloroform. 4. Isopropanol. 5. 75% ethanol. 6. RNAse-free water.

2.6.2  cDNA Preparation

1. SensiFAST cDNA synthesis kit (Bioline) or equivalent.

2.6.3  Determination of the T-Cell Receptor Subtype by PCR

1. Fast taq DNA polymerase and the corresponding buffer. 2. Deoxynucleotide triphosphate (dNTP). 3. TCR subtype primers (see Tables 1 and 2). 4. Thermal cycler. 5. 1× TBE buffer: dilute 10× TBE buffer (Roche) to 1× in water. 6. 2% agarose gel: dissolve 2% agarose in 1× TBE buffer and add 1/10,000 SYBR Safe DNA gel stain (Invitrogen). Boil and pour the solution in a gel casting system. 7. Power source.

TGGATGGTTTGAAGGACAGTG

CTGTTTATCTCTGCTGACCGG

ACGAAGGACAAGGATTCACTGT

CTGGAGGACTCAGGCACTTACT

TRAV01

TRAV02

TRAV03

TRAV04

Vα4

GGTACCCGACTCTTTTCTGGT

ACCCTTTCAGAAGATGACTTCC

TTTAAAGTCCCAAAGGCCAA

TCCTGAAAGTCATTACGGCTG

AGAGCCTCAAGGGACAAAGAG

AGACTCCCAGCCCAGTGACT

ACATCAGAGAGCCGCAACC

CCCTGCCCAGCTAATCTTAAT

CTGCAGCTGAGATGCAAGTATT

TCCTATGGTGGATCCATTTACC

TGGACAGAAAACAGAGCCAA

TRAV06

TRAV06-4

TRAV06-5

TRAV06-6

TRAV06-7

TRAV07

TRAV07-5

TRAV08

TRAV09

TRAV09-1

TRAV10/05-2

Vα3

Vα1

Vα13

TRAV05

Vα11

Vα5

Anti-mouse Vα11.1/11.2

TCR Vα antibodies

Flow cytometry

CTCAAGTACTATTCCGGAGACCCAGTGGTT Anti-mouse Vα3.2

CAGCAGAGCCCAGAATCCCTC

GGAAGCAGCAGAGGTTTTGAAGCTACATAC

CGTTCAAATATGGAAAGAAAGCAGACCCAA

AATGGGAGGTTAAAGTCAACATTCAATTCT

AAGGTTTTCTCAAGTACGGAAATAAACGAA

GTGGCATCTCTGTTTATCTCTGCTGACCGG

IMGT subgroup Primer sequence

Primer sequence

IMGT subgroup

Vα12

PCR TCR panel [12]

PCR TCR panel [11]

Overview of the correspondence between different nomenclatures in the TCR Vα PCR panels and TCR Vα antibodies

Table 1

188 Manon Vanheerswynghels et al.

CAGGCAAAGGTCTTGTGTCC

ACGCCACTCTCCATAAGAGCA

GCTCTTTGCACATTTCCTCC

TGCAGTTATGAGGACAGCACTT

CTGCAGTTATGAGAACAGTGCTT

CCAGACGATTCGGGAAAGTA

TTCCATCGGACTCATCATCAC

AACCTGAAGAAATCCCCAGC

GGAAGACGGAAGATTCACAGTT

ACGCTCCTAATAGACATTCGCT

GTTCCTCTTCAGGGTCCAGA

CACCAGCAGGTTCTGGGTTC

TRAV11

TRAV12

TRAV13

TRAV141/14-­2

TRAV14-3

TRAV15

TRAV16

TRAV17

TRAV19

TRAV20

TRAV21

TRAC

CAGGCAGAGGGTGCTGTCC

CGGCACATTGATTTGGGAGTC

NJ109 NJ110

GGCCCCATTGCTCTTGGAATC

NJ108

AGTATGGCTTTCCTGGCTATTGCCTCTGAC

CGACAAACGTCTTCTTCTACTGCAAAAGAG

ACCTTCTTCAATAAAAGGGAGAAAAAGCTCa Anti-mouse Vα2

CTGACATCCACCACAGTCACTAAGGAACGT

CAAAGAGCTGCGACGTTCCTT

Vα2 in this panel corresponds to both TRAV14-1, TRAV14-2, and TRAV14-3.

a

TCR Vα antibodies

Flow cytometry

ACAGACAACAAGAGGACCGAGCACCAAGGG Anti-mouse Vα8.3

Vα9

Vα6

Vα7

Vα2

Vα10

Vα8

CTGGTTGACCAAAAAGACAAAACGTCAAAT

IMGT subgroup Primer sequence

Primer sequence

IMGT subgroup Vα14

PCR TCR panel [12]

PCR TCR panel [11]

Allergen-Specific TCR Transgenic Mice 189

GTGGCAGTTTTGCATTCTGTGCC

CCACTTGCAGCCTCAGCTCCG

GGCCGTTTGTCTCCTGGTGGC

TGTGGCCGAGTCATCAGGCTT

GGAAGCAGGACACACAGGACCCA

AGTCCTCCAGTGCTGTGGGTTGG

GGCTCCTAAGCTGTGTGGCCT

TACACAGCAGAGTCCTCCGGCTC

AGCTGCAGGCTTCTCCTCTATGT

AACGACTGGGCACCGTCTCA

TGTTACAGACATGGGACAGAATGTCA

GCCAAGCTGCTGGCATACATAGT

TGCTCTGCTGTATGGCCCTTTGT

ACAGTGCAGAGTCCTTTGGCTC

GCAGTGCTGCATTTTGTTTCCTGC

TGTCTGGGAGGGAGAGGCCAA

CCTCACTAGTGCTGAATTCTCCCAA

Vβ1 F

Vβ1 R

Vβ2 F

Vβ2 R

Vβ3 F

Vβ3 R

Vβ4 F

Vβ4 R

Vβ5 F

Vβ5 R

Vβ6 F

Vβ6 R

Vβ7 F

Vβ7 R

Vβ8 F

Vβ8 R

Vβ9 F

TRBV05

TRBV04

TRBV03

TRBV02

TRBV01

IMGT subgroup

Primer sequence

IMGT subgroup

ACGGTGCCCAGTCGTTTTAT

TAAACGAAACAGTTCCAAGGC

TCACTCTGAAAATCCAACCCA

ATGGACAATCAGACTGCCTCA

ACACGGGTCACTGATACGGA

Primer sequence

PCR TCR panel designed in house [11]

PCR TCR panel designed in the house

Anti-mouse Vβ10b

Anti-mouse Vβ4

Anti-mouse Vβ2

TCR panel

Flow cytometry

Overview of the correspondence between different nomenclatures in the TCR Vβ PCR panels and TCR Vβ screening panel for flow cytometry

Table 2

190 Manon Vanheerswynghels et al.

ACATGGAGAAGGAGCCCTCGG

TGGTGGAATCACCCAGACACCT

GCTGGCCCAGAAGTACATGGAGG

GCCACGGACACCAGGCACTTC

AGGTGCTCTGTCGAGTCCCGC

CTCGCTGATTCTGCCTGGGGC

AGTCCTCCAGTTCCAAGGCACTCA

CTTCCCTGACCCCGCCTGGAA

CGGCAGAGTCCTCTAGCTCCAAGG

CTGGGGCACCACCCTGCTTTC

CCTCTAGGTCCAAGGCACTCAGG

TGGGCTCCAGGCTCTTTCTGGT

TGCGTTGTTCTGGTGGCCTTGT

GGCTCCAGGCTCTTCTTCGTGC

TGGCTTGGTCTGGAGGCCTTGT

TGGGCTCCAGACTCTTCTTTGTGGT

TGGAGGCCTTGTACCCATCAGGG

GCACCAGGCTTCTTGGCTGGG

GGCTGTGTCGCCCTGCTTTG

TGGGCATCCAGACCCTCTGTTG

Vβ9 R

Vβ10 F

Vβ10 R

Vβ11 F

Vβ11 R

Vβ12-1 F

Vβ12-1 R

Vβ12-2 F

Vβ12-2 R

Vβ12-3 F

Vβ12-3 R

Vβ13-1 F

Vβ13-1 R

Vβ13-2 F

Vβ13-2 R

Vβ13-3 F

Vβ13-3 R

Vβ14 F

Vβ14 R

Vβ15 F

TRBV15

TRBV14

TRBV13-3

TRBV13-2

TRBV13-1

TRBV12-2

TRBV12-1

IMGT subgroup

Primer sequence

IMGT subgroup

CGCAGCAAGTCTCTTATGGAA

GCGACACAGCCACCTATCTC

TGGCTTCCCTTTCTCAGACA

GGCTACCCCCTCTCAGACAT

CCAGAACAACGCAAGAAGACT

AGATAAAGGAAACCTGCCCAG

GGATTCCTACCCAGCAGATTC

Primer sequence

PCR TCR panel designed in house [11]

PCR TCR panel designed in the house

(continued)

Anti-mouse Vβ12

Anti-mouse Vβ13

Anti-mouse Vβ8.1b

Anti-mouse Vβ8.2b

Anti-mouse Vβ8.3

Anti-mouse Vβ5.1a

Anti-mouse Vβ5.2a

TCR panel

Flow cytometry

Allergen-Specific TCR Transgenic Mice 191

TGCTGGCACACAGATACACAGC

GGCCCCCAGGCTCCTTTTCTG

CCGCTGAGTCCTGGGGTTGC

TGTGATCTTCTGTCTTCTTGCAGCC

AGGCCTGCAGAGCCAATGTAG

GGTGTCACCACGAACCTAAGATACA

TTCCAGATCTGCTGGCCCCAC

TGCTGGGTAACCCTTTGTCTCCT

CGGCCATCTCGTTCTTCTGGGC

ACTGCTTCTATTACTTCTGGGGCCT

AGGATAAGTTGGGATGACTGATGGGA

TGCAAACTTTTCTACTGTGTGCCCT

GCTTGCTGGACTGGATCTCTATGGT

TGTGGCTTCTCTGGTATGTAGCCC

AGCTGCAGGGTGAGCTTAAGGGA

Vβ15 R

Vβ16 F

Vβ16 R

Vβ17 F

Vβ17 R

Vβ18 F

Vβ18 R

Vβ19 F

Vβ19 R

Vβ20 F

Vβ20 R

Vβ21 F

Vβ21 R

Vβ22 F

Vβ22 R

TRBV21

TRBV20

TRBV19

TRBV17

TRBV16

IMGT subgroup

Primer sequence

IMGT subgroup

AAAATGCCCTGCTAAGAAACC

TTCCCATCAGTCATCCCAAC

GAAGGCTATGATGCGTCTCG

TGAGAAGTTCCAATCCAGTCG

ATAGATGATTCAGGGATGCCC

Anti-mouse Vβ6

Anti-mouse Vβ9

Anti-mouse Vβ11

TCR panel

PCR TCR panel designed in house [11]

PCR TCR panel designed in the house Primer sequence

Manon Vanheerswynghels et al.

Table 2 (continued) Flow cytometry

192

GGGTGCACGGCTCATTTGCTA

GTGAGCTGGAGGGGCACACAG

TGGGTGCAAGACTGCTCTGCTG

ACAGACTGCTGGCACAGAGCTACA

GTGATCACTTCGCAGGAAAACTTGA

AGAGCTGCTGGCACAGAAGTG

GGCTACAAGGCTCCTCTGTTACAC

CTGGATGTCCCTTTTCAGGGATACA

GCCAAGTTCATGCATTGTCTGGCCT

AGGCAGGAGTGTGGAGGGCT

CCAGTCTCTCAAGATGTGTGGTCCT

GCACACATGCCTGGTCGATGCT

AGGCTCATCTCTGCTGTGGTGCT

CCCGCTTCTTCCGTGAGACCC

ACATTCCTGCTACTTCTTTGGAGCC

AGGTTTGGCCGGCTGATTGGA

CTCTCCTTGCCTTTCTCCTGGGC

GTCGTCCTTCGGCCTGGAAGC

TGGCCAGGGGCTTCTTCCCT

GGAACTGCACTTGGCAGCGGA

Vβ23 F

Vβ23 R

Vβ24 F

Vβ24 R

Vβ25 F

Vβ25 R

Vβ26 F

Vβ26 R

Vβ27 F

Vβ27 R

Vβ28 F

Vβ28 R

Vβ29 F

Vβ29 R

Vβ30 F

Vβ30 R

Vβ31 F

Vβ31 R

Cb1and2 F

Cb1and2 R

TRBV31

TRBV30

TRBV29

TRBV26

TRBV24

TRBV23

IMGT subgroup

Primer sequence

IMGT subgroup

TTCATCCTAAGCACGGAGAAG

GGACAAGTTTCCAATCAGCCG

AAAGGATACAGGGTCTCACGG

AGTGTCCTTCAAACTCACCTT

GCATCCTGGAAATCCTATCCT

CAGCCTGGGAATCAGAACG

Primer sequence

PCR TCR panel designed in house [11]

PCR TCR panel designed in the house

(continued)

Anti-mouse Vβ14

Anti-mouse Vβ7

Anti-mouse Vβ3

Anti-mouse Vβ17a

TCR panel

Flow cytometry

Allergen-Specific TCR Transgenic Mice 193

TGGTGGCAGACAAGACCCCTT

GGCCAGAGGGCTCACCCAAAC

TGGCCATCAGCACCAGGCCAC

Cb1 R

Cb2 F

Cb2 R

b

a

The MR9-4 antibody reacts with the Vβ5.1 and 5.2 TCR The MR5-2 antibody reacts with the Vβ8.1 and 8.2 TCR

GTCAGCACGGACCCTCAGGC

Cb1 F

TRBC1

IMGT subgroup

Primer sequence

IMGT subgroup

TGCAATCTCTGCTTTTGATGGCTC

Primer sequence

PCR TCR panel designed in house [11]

PCR TCR panel designed in the house

Table 2 (continued)

TCR panel

Flow cytometry

194 Manon Vanheerswynghels et al.

Allergen-Specific TCR Transgenic Mice 2.6.4  Subcloning the TCR in the Teasy Vector

195

1. Phusion® high-fidelity DNA polymerase (New England Biolabs) or equivalent. 2. Material and buffers for PCR described in Subheading 2.6.3 determination of the T-cell receptor subtype by PCR. 3. Agarose Gel DNA Extraction Kit (Roche) or equivalent. 4. pGEM®-T Easy Vector System (Promega). 5. Deoxyadenosine triphosphate (dATP). 6. Fast taq DNA polymerase and the corresponding buffer. 7. T4 DNA ligase and 10× T4 ligase buffer (Promega) or equivalent. 8. Bacterial LB/agar plates with ampicillin and blue/white selection: dissolve 10.3 g LB broth and 7.5 g agar in 500 mL Milli-­Q water and autoclave. When cooled to approximately 37 °C, add 100 μg/mL ampicillin, 0.5 mM IPTG, and 80 μg/ mL X-gal (stock dissolved in DMSO), and pour plates (10– 15 mL per 100 mm petri dish). 9. DH10B electrocompetent cells. 10. Electroporation system. 11. 0.2 cm cuvettes. 12. SOC medium (Invitrogen) or equivalent. 13. LB liquid medium: dissolve 10.3 g LB broth in 500 mL Milli-­Q water and autoclave. When cooled to approximately 37 °C, add 100 μg/mL ampicillin. 14. High pure plasmid isolation kit (Roche) or equivalent. 15. Restriction enzymes for restriction analysis and the corre sponding restriction buffer. 16. 1× TBE buffer: dilute 10× TBE buffer (Roche) to 1× in water. 17. 2% agarose gel: dissolve 2% agarose in 1× TBE buffer, and add 1/10,000 SYBR Safe DNA gel stain (Invitrogen). Boil and pour the solution in a gel casting system. 18. Power source.

2.6.5  Cloning of the TCR in CD4 Expression Vector

1. Genopure plasmid maxi kit (Roche) or equivalent. 2. SalI and XhoI restriction enzyme and the corresponding restriction buffer. 3. Agarose Gel DNA Extraction Kit (Roche) or equivalent. 4. T4 DNA ligase and 10× T4 ligase buffer (Promega) or equivalent. 5. Material, cells, and buffers for transformation as described in Subheading 2.6.4 subcloning of the TCR in the Teasy vector (see items 7–13). 6. Material and buffers for restriction analysis as described in Subheading 2.6.4 subcloning of the TCR in the Teasy vector (see items 14–17).

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2.6.6  Preparation of the Injection Fragments

1. Genopure plasmid maxi kit (Roche) or equivalent. 2. NotI restriction enzyme and the corresponding restriction buffer. 3. Agarose Gel DNA Extraction Kit (Roche) or equivalent. 4. Material and buffers for restriction analysis as described in Subheading 2.6.4 subcloning of the TCR in the Teasy vector (see items 14–17). 5. Injection buffer: prepare a solution of 10 mM tris, pH 7.5, 0.1 mM EDTA, and 100 mM NaCl in ultrapure water. Sterile filter this solution. 6. Eppendorf centrifuge.

2.7  Evaluation of the TCR Mice

1. Material and buffers for PCR described in Subheading 2.6.3 determination of the T-cell receptor subtype by PCR.

2.7.1  Genotyping

1. 10 mM EDTA: dilute 0.5 M commercial EDTA in PBS (20 mL EDTA per 1 L PBS).

2.7.2  Analysis of the Presence of T Cells in Peripheral Blood

2. PBS. 3. Lympholyte cell separation medium (Cedar lanes). 4. Anti-mouse CD3 antibody, anti-mouse CD4 antibody, and anti-mouse CD69 antibody. 5. Material and buffers as described in Subheading 2.3.3 confirmation screening (see items 4–8).

2.7.3  Evaluation of TCR Specificity in a DC-T-Cell Co-culture

1. Material and buffers as described in Subheading 2.3.1 culture of bone marrow-derived dendritic cells. 2. Material and buffers as described in Subheading 2.3.3 confirmation screening (see items 4–8). 3. 70 μm cell strainer. 4. Osmotic lysis buffer: prepare a solution of 155 mM NH4Cl, 1 mM KHCO3, 100 μM EDTA in water. Adjust the pH to 7.1–7.4 using HEPES. Sterilize using a 0.22 μm bottle top filter. 5. PBS. 6. MACS buffer: prepare a solution of 0.5% BSA and 2 mM EDTA in PBS. Sterilize using a 0.22 μm bottle top filter. 7. CD4+ T-cell isolation kit, Miltenyi Biotech or equivalent. 8. CellTrace CFSE cell proliferation kit for flow cytometry (eBioscience), or equivalent. Prepare a solution of 50 μM CFSE by 1/100 dilution of the CFSE stock in RPMI. 9. RPMI. 10. RPMI5: RPMI supplemented with 5% FCS, 50 μg/mL gentamycin, 50 μM beta-mercaptoethanol, and 2 mM GlutaMAX (all Gibco).

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11. Peptide solution: dilute the peptide to 4 μg/mL in RPMI5 (final concentration of 1 μg/mL in the well), and prepare a ½ serial. Do this by each time transferring half of the tube content to a new tube with an equal volume of RPMI5, for five steps in a row. Prepare 50 μL per sample (see Note 14, use RPMI5 instead of DMEM10). 12. 96-well round bottom plate.

3  Methods 3.1  Immunization

1. Inject 10–15 mice intraperitoneally with 10 μg antigen formulated on aluminum hydroxide, twice with a 1-week interval (days 1 and 8). Ten days after the last injection, inject the mice on 3 consecutive days (days 18, 19, and 20) with an intranasal injection of 10 μg of non-formulated antigen. Sacrifice the mice 1 day after the last injection (day 21) (see Notes 1 and 2).

3.2  T-Cell Fusion

All centrifugation steps are performed in a cooled centrifuge, for 7 min at 400 × g, unless specified otherwise. Cells are cultured in a humidified incubator at 37 °C and 5% CO2. All products that come into contact with the cells should be sterile. Sterile working procedures should be applied.

3.2.1  Culture of the BW5147 Fusion Partner

1. Start-up BW5147 cells by rapid thawing in a 37 °C water bath. Resuspend cells in 9 mL of pre-warmed DMEM10 medium in a 15 mL conical tube and centrifuge cells at room temperature. 2. Resuspend cells in 1 mL DMEM10. Count cells and dilute to 2 × 105 cells/mL in DMEM10 medium. Transfer 10 mL of the cell suspension into a 25cm2 culture flask and culture for 2 weeks. 3. Passage cells every 2 or 3 days by seeding at 2 × 105 cells/mL or 105 cells/mL, respectively. Cell concentrations should not exceed 106 cells/mL. 4. Seed cells 1 day prior to T-cell fusion at 3 × 105 cells/mL. For optimal results, cell viability should be over 90%.

3.2.2  In Vitro Restimulation of Lymph Node Cells

1. Isolate mediastinal lymph nodes from immunized mice. Obtain a single-cell suspension by mashing all lymph nodes through a 70 μm cell strainer using 1 mL HBSS, and rinse the cell strainer with 2 mL HBSS. Centrifuge cell suspension, and resuspend the cell pellet in 10 mL RPMI5 (see Note 3). 2. Count cells and dilute to a concentration of 3 × 106 cells/mL in RMPI5. Seed 100 μL (3 × 105 cells) per well in a 96-well round bottom plate. Add 100 μL antigen-specific peptide

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per well at a final concentration of 5 μg/mL in RMPI5 (see Note 4). 3. Culture cells for 4 days without passaging. 3.2.3  CD4+ T-Cell Isolation

1. Harvest restimulated lymph node cells, and transfer cells to a 50 mL conical tube, centrifuge, and resuspend cells in 10 mL MACS buffer. 2. Pre-warm centrifuge to room temperature for T-cell fusion. 3. Isolate CD4+ T cells using Miltenyi Biotech CD4+ T cells isolation kit, according to manufacturer’s instructions (see Note 5).

3.2.4  T-Cell Fusion

1. Pre-warm in a 37 °C warm water bath:

(a) Two 400 mL and two 600 mL beakers all with 75 mL water inside



(b) Four 50 mL tubes with DMEM inside



(c) 2 mL aliquot of 50% poly ethylene glycol (PEG)



(d) One 50 mL tube with DMEM20

2. Harvest BW5147 cells and wash cells twice using 40 mL DMEM. Centrifugation steps are performed at room temperature. Resuspend cells in 10 mL DMEM (see Note 6). 3. Wash the enriched T cells twice using 10 mL DMEM. Centrifugation steps are performed at room temperature. Resuspend cells in 1 mL DMEM in a 15 mL conical tube. 4. Count BW5147 and enriched T cells, and add an equal number of BW5147 cells to the enriched T-cell suspension. 5. Centrifuge 5′ at 800 × g at room temperature and aspirate the supernatant. 6. Place the 400 mL beakers with warm water (37 °C) in the 600 mL beakers with warm water (37 °C) in the laminar flow, in order to mimic two warm water baths. Place the tube with cells and medium in separate baths. 7. Slowly and dropwise add 1 mL of pre-warmed PEG using a 1 mL micropipette to the cells during 1 min while gently stirring the pellet with the pipet (see Note 7). 8. Slowly and dropwise add 2 mL of pre-warmed DMEM to the cells with a 2 mL pipet, while gently stirring. Continue stirring for a total of 2 min (see Note 8). 9. Slowly and dropwise add an additional 7 mL of pre-warmed DMEM with a 10 mL pipet over a 2–3 min period while gently stirring. 10. Centrifuge 5′ 800 × g, at room temperature.

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11. Aspirate supernatant and vigorously add 10 mL of DMEM20 to the pellet, as to release the pellet completely. However, do not pipet the suspension up and down. 12. Dilute cells between 105 and 106 cells/mL by gently adding the appropriate volume of DMEM20 while stirring, based on the total number of T cells used for fusion. 13. Plate the cells in a 96-well flat bottom plate, by pipetting two drops (~100 μL) of cell suspension per well with a 10 mL pipet and culture for 1 day (see Note 9). 3.2.5  Culture of the Fused T Cells

1. After 1 day, gently add 100 μL of 2× HAT medium to each well (see Note 10). 2. After 6 or 7 days, remove the top half of the medium of each well using a multichannel pipet and gently add 100 μL of fresh 2× HAT medium to each well. 3. Monitor cells daily; cells should be expanded when medium becomes slightly yellow and expansion of the cells is clearly visible under the microscope (about 50% of the bottom of the well is covered with cells). Expand the cells by reseeding them in a 24-well plate containing 1 mL of 1× HT plating medium. 4. After 1 day, add an additional 1 mL of 1× HT plating medium. 5. Duplicate cultures of clones that are growing well, by transferring half of the cells to a fresh 24-well plate and adding 1 mL of fresh DMEM20 medium. In this way, one plate can be used for screening experiments, and one for culture. From now on, cells should be able to grow in lower FCS conditions and can thus be cultured in DMEM10 medium. 6. Monitor cells daily, and maintain them by reseeding 25% of the cells in DMEM10 in case the medium turns yellow (see Note 11).

3.3  Screening of the T-Cell Clones 3.3.1  Culture of Bone Marrow-Derived Dendritic Cells

1. Isolate both femurs from one mouse and sterilize in 70% ethanol for 5 min. Aspirate the ethanol and rinse the femurs twice with 10 mL of RPMI. Crush the bones in 10 mL RPMI in a mortar with a pestle and repeat twice; pass the suspension over a 70 μm cell strainer and centrifuge. 2. Lyse the erythrocytes by resuspending the cell pellet in 2 mL osmotic lysis buffer and incubate for 4 min on ice. Add 10 mL of RPMI and centrifuge the cell suspension. 3. Count and plate 3 × 106 cells per 100 mm petri dish in 10 mL RPMI5 with 20 ng/mL recombinant GM-CSF. Culture the cells for 8–10 days. On day 3, add 10 mL of RPMI5 with 20 ng/mL recombinant GM-CSF. On days 6 and 8, replace the top half of the medium by 10 mL RPMI5 with 20 ng/mL recombinant GM-CSF.

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4. Harvest the cells by resuspending with a 10 mL pipet, and rinse the plate with 10 mL ice-cold PBS. Centrifuge and count the cells. 3.3.2  Initial Screening

1. For every individual clone, start a co-culture by seeding 50 μL of bone marrow dendritic cells (2 × 105 c/mL) in 2 wells of a 96-well round bottom plate (see Note 12). Add 50 μL of 4 μg/mL peptide solution to one well (final concentration of 1 μg/mL) and 50 μL of medium to the second well. Resuspend the hybridoma clone with a 1 mL micropipette, transfer 100 μL of cells to each well, and culture overnight. All steps are performed in DMEM10. 2. After 1 day, centrifuge the plates and collect the supernatant. Test the supernatant in an IL-2 ELISA. Clones that are specific for the peptide will show high IL-2 production in the presence of the peptide. Clones not responding to antigen can be discarded (see Note 13).

3.3.3  Confirmation Screening

1. In order to validate and asses the specificity of the clones, a similar co-culture experiment is performed. Start a co-culture by seeding six wells with 50 μL of bone marrow dendritic cells (2 × 105 c/mL) in a 96-well round bottom plate (see Note 12). Count and add 100 μL of the hybridoma cells at 106 cells/ mL. Prepare a serial dilution of the peptide in 50 μL and add to the different wells (include a medium control) (see Note 14). All steps are performed in DMEM10. 2. Culture overnight and quantify IL-2 production in supernatant by ELISA (see Note 13). 3. Additionally, evaluate increased expression of CD69, on CD3+CD4+ T cells, after peptide stimulation using flow cytometry. In short, prepare a 50 μL staining mix per sample including all required antibodies in the recommended dilution, Fc- blocking reagent and live/dead stain in PBS (see Note 15). Prepare compensation beads and add single stain reactions, containing each individual antibody and the live/dead stain separately, in order to set the voltages on the flow cytometer. Centrifuge and resuspend cells in 50 μL staining buffer, and incubate cells and compensation beads at 4 °C in the dark. After 30 min, wash and resuspend samples in 150 μL FACS buffer and analyze on a flow cytometer. 4. Clones with the highest specificity, e.g., the lowest peptide concentration that still induces upregulation of CD69 and production of IL-2, can be selected (see Fig. 1).

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Fig. 1 Co-culture of a hybridoma clone with bone marrow-derived dendritic cells, in the presence or absence of antigen-specific peptide. Upregulation of CD69 (a) and induction of IL-2 (b) of the hybridoma clone after co-­culture in the presence of peptide 3.3.4  Evaluation of the TCR Subtype

1. Evaluate the selected clones for their TCR variable domain by flow cytometry (as described above), using the mouse Vβ TCR screening panel (Becton Dickinson) and commercially available TCR Vα antibodies (see Note 16).

3.3.5  Freezing of Peptide-Specific T-Cell Hybridomas

1. Select one clone based on all available screening data, and freeze all other specific clones as backup.

3.4  Generation of Monoclonal Cells

2. Collect T-cell hybridomas from their respective wells and transfer to a 15 mL conical tube (see Note 17). Centrifuge the cells, aspirate the supernatant, and resuspend the pellet in 1 mL of freezing medium (10% DMSO in DMEM20). Transfer the cell suspension into a cryovial and store it in a freezing container at −70 °C. After 1 day cells can be transferred to liquid nitrogen for further storage. 1. Stain cells from the selected clone with anti-CD3, anti-CD4, and, if available, anti-TCR Vα and -Vβ (as described before). 2. Single cell sort stained cells using fluorescence-activated cell sorting (FACS) in a 96-well round bottom plate with 200 μL DMEM10/well (see Note 18). 3. Culture sorted cells and feed weekly by replacing the top half of the medium with 100 μL/well DMEM10. 4. Expand growing cells of wells that become slightly yellow and where expansion of the cells is clearly visible under the microscope, by resuspending and transferring the cells to a 24-well plate with 1 mL of DMEM10. 5. Cells can be cultured as described before. Passage cells every 2 or 3 days by seeding at 2 × 105 cells/mL or 105 cells/mL, respectively. Cell concentrations should not exceed 106 cells/mL.

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3.5  Confirmation and Selection of the Monoclonal T-Cell Clones

1. Screen monoclonal T-cell clones for their peptide specificity, by performing a co-culture using a peptide concentration range, as described before at “second confirmation screening” (see Subheading 3.3.3). 2. Select one clone with high peptide specificity, preferably with a TCR subtype to which antibodies are available. This clone will be used for cloning of the T-cell receptor. 3. Freeze cells of the selected clone as described in “freezing of peptide-specific T-cell hybridomas”(see Subheading 3.3.5). Prepare multiple vials of 106—2 × 106 cells. Preferentially, some peptide-specific clones can be frozen as a backup. 4. In addition, freeze cell pellets of the selected clone for RNA isolation by pelleting 106 cells by centrifugation. Aspirate supernatant and snap freeze the cell pellet on dry ice or in liquid nitrogen, finally store the pellets at −70 °C.

3.6  Cloning 3.6.1  RNA Preparation

All centrifugation steps are performed in a cooled Eppendorf centrifuge. Cool centrifuge to 4 °C prior to the experiment. RNA isolation should be performed using RNAse-free materials and reagents. 1. Resuspend the cell pellet in 1 mL of TriPure reagent and vortex extensively. Add 200 μL chloroform, and vortex and incubate for 3 min at room temperature. Centrifuge for 15 min at 12,000 × g. 2. Carefully pipet the upper transparent layer into a new tube. Add 500 μL isopropanol, and vortex and incubate for 5 min at room temperature. Centrifuge for 10 min at 12,000 × g. 3. Remove the supernatant and add 1 mL of 75% ethanol. Vortex and centrifuge for 5 min at 7500 × g. 4. Remove supernatant and air-dry pellet. Resuspend pellet in 20 μL RNAse-free water.

3.6.2  cDNA Preparation

1. Prepare cDNA using random hexamer primers, following the manufacturer’s instructions, using 1 μg RNA.

3.6.3  Determination of the T-Cell Receptor Subtype by PCR

1. Perform a PCR using a panel of TCR subtype primers (see Tables 1 and 2), with 0.1 μL of cDNA per reaction as a template (following the manufacturer’s instructions). The fusion partner BW5147 will give a positive reaction for TRAV20 and TRBV12-­ 1. Other positive reactions are derived from the allergen-specific TCR and should be sequenced (see Note 19). Determine the correct Vα and Vβ subtype by blasting the sequence using the IMGT database (www.imgt.org). Design full-length primers in order to amplify the complete Vα and Vβ including a piece of the 5′ UTR. The primers should contain SalI or XhoI restriction sites to allow subcloning the Vα

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and Vβ into the TCR expression vectors p783 and p428, respectively. 3.6.4  Subcloning the TCR in the Teasy Vector

1. Amplify the Vα and Vβ by performing a proofreading PCR using 2 μL cDNA as a template and their full-length primers (following the manufacturer’s instructions). Gel purify the amplicons using a gel isolation kit (following the manufacturer’s instructions). 2. A-tail and subclone the amplicons using the T easy vector system (Promega). Transform the ligation reaction in high-­ efficiency competent cells applying blue-white selection screening for positive clones (following the manufacturer’s instructions). 3. Grow the positive clones overnight in 5 mL liquid culture (LB + 100 μg/mL ampicillin) at 37 °C while shaking and isolate DNA using a mini prep kit (following the manufacturer’s instructions). 4. Confirm correct clones by using restriction analysis and sequencing.

3.6.5  Cloning of the TCR in CD4 Expression Vector

1. Prepare a maxi prep from one positive TCR Vα and one TCR Vβ clone, following manufacturer’s instructions. 2. Excise the inserts from the T easy vector by SalI of XhoI digestion. Incubate 10 μg of DNA in 1× restriction buffer with 1 μL (10 units) of SalI or XhoI enzyme in a 50 μL reaction, for a minimum of 3 h at 37 °C. Gel purify the inserts (following the manufacturer’s instructions). 3. Linearize the CD4 expression vector p428 and p783 (see Fig. 2) with SalI. Incubate 10 μg of vector with 1 μL (10 units) of SalI enzyme in a 50 μL reaction, for a minimum of 3 h at 37 °C. Gel purify the vectors (following the manufacturer’s instructions). 4. Determine the concentration of isolated DNA and prepare a 10  μL ligation reaction of maximum 100 ng total DNA, including insert and vector in a 3:1 molar ratio and 1 μL (1 Unit) T4 DNA in 1× T4 DNA ligase buffer. The Vα should be cloned in the p783 and the Vβ in the p428 expression vector. Incubate for 30 minutes at room temperature, followed by overnight at 4 °C (see Note 20). Transform the ligation reaction in high-­efficiency competent cells (following the manufacturer’s instructions). 5. Positive colonies can be determined using a colony PCR. Single colonies are transferred to a 96-well PCR plate containing 25  μL of Milli-Q water/well. Use 5 μL in a colony PCR using the CD4 forward and reverse primer (CTCCTCGA CCCACTTCTGATG and GCTCAGATTCCCAACCAACAA, respectively) in combination with the reverse primer from the

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Fig. 2 Both TCR expression vectors p428 and p783 contain the mouse CD4 promoter and the mouse CD4 enhancer followed by a piece of mouse CD4 genomic sequence, a cloning site, and an SV40 polyA signal. The genomic sequence in both vectors contains mouse CD4 exon I, intron I containing different deletions in each vector followed by exon II up to the translation start site. The intron in plasmid p783 contains the transcriptional silencer element that is normally responsible for terminating CD4 expression in CD8 lineage cells. Vα cDNA is cloned into the p783 and Vβ cDNA into the p428 expression vector. Injection fragments are excised from the pNNO3 backbone by NotI digestion and injected into fertilized oocytes a

b

1230 bp F1

R1

800 bp

1230 bp F1

R2

F1

R2

Insert in forward orientation 200 bp

630 bp

600 bp

400 bp

R1

R1

Insert in reverse orientation 30 bp

200 bp

400 bp

600 bp

30 bp

Fig. 3 Example of a colony PCR to evaluate the presence and orientation of the insert. A colony PCR is performed using CD4 forward (F1) and reverse (R1) primers in combination with the reverse primer from the TCR primer panel corresponding to the insert (R2). F1 and R1 are located on the TCR expression vectors ~200 bp upstream and ~30 bp downstream of the TCR cloning site SalI. In this example, R2 is located 600 bp in the insert. In correct orientation (a), the colony PCR will result in a product of ~800 bp (~200 bp upstream the insert +600 bp of insert). In reverse orientation (b), the colony PCR will result in a product of ~630 bp (600 bp of insert + ~30 bp downstream the insert). An additional product of ~1230 bp might be amplified in both orientations. In colonies where the vector re-ligated, not harboring any insert at all, a ~230 bp product is amplified

TCR primer panel corresponding to the insert. Analysis of the colony PCR on a 2% agarose gel will provide information on the presence and orientation of the insert (Fig. 3). Positive colonies are transferred to 5 mL liquid culture (LB + 100 μg/ mL ampicillin) and grown overnight at a 37 °C while shaking. Colonies are viable for a limited time and should be transferred to the medium within 8 h. 6. Isolate DNA using a mini prep kit (following the manufacturer’s instructions) and confirm correct clones by using restriction analysis and sequencing.

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1. Prepare a maxi prep from one positive TCR Vα and one TCR Vβ clone, following the manufacturer’s instructions. 2. Prepare the injection fragments by NotI digestion of the construct. Incubate 20 μg of maxi prep DNA in 1× restriction buffer with 2 μL (10 units) of NotI restriction enzyme in a 100 μL reaction and incubate for a minimum of 3 h at 37 °C. Gel purify the fragments (following the manufacturer’s instructions). 3. Determine the concentration of isolated DNA and confirm the fragments by restriction analysis. Incubate 100 ng of the fragments in 1× restriction buffer with 0.5 μL (5 units) of restriction enzyme in a 20 μL reaction, for a minimum of 3 h at 37 °C. Analyze on an agarose gel. 4. Prepare an equimolar solution of 1 ng/μL of both TCR injection fragments (TCR Vα and TCR Vβ) in 0.22 μm pre-filtered injection buffer. Centrifuge for 30 minutes at 14,000 × g in a cooled centrifuge. Transfer the supernatant into a 1.5 mL tube and use this for microinjection in fertilized Rag2−/− oocytes.

3.7  Evaluation of the TCR Mice 3.7.1  Genotyping

3.7.2  Analysis of the Presence of T Cells in Peripheral Blood

1. Perform a PCR using primers from the TCR PCR panel corresponding to the construct (TCR Vα and Vβ). Analyze PCR products on a 2% agarose gel for presence of the transgene. Breed the transgenic founders with rag2−/− mice to obtain germline transmission and characterize (3.7.2. and 3.7.3.) each line to select the best line for use in further experpiment. Since the mice were generated via microinjection, the construct will be integrated multiple copy, randomly into the mouse genome. Each transgenic founder will have a different integration site and copy number and should be characterized separately as an independent line. Microinjection was performed in Rag2−/− oocytes. As Rag2-­ deficient animals fail to generate mature T or B lymphocytes, mature T cells will only be present in the peripheral blood when the inserted constructs provide a functional T-cell receptor. Therefore, assessing the presence of mature T cells in the peripheral blood will provide evidence of a functional TCR transgenic mouse (see Fig. 4). 1. Collect blood of transgenic animals in a 1.5 mL tube containing 100 μL of 10 mM EDTA and add PBS to acquire a total volume of 800 μL. Pipet 400 μL lympholyte on the bottom of the tube, allowing it to form a layer under the diluted blood. Centrifuge at 600 ×g for 20 min at room temperature, without brake (see Note 21). 2. Collect the interface with a 200 μL pipet and transfer to a new 1.5 mL tube. Add FACS buffer to acquire a total volume of 1.2 mL and centrifuge. Stain cells with anti-CD3 and anti-CD4, as described before, and determine the presence of T cells.

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Fig. 4 Evaluation of the presence of CD3+CD4+ T cells in the peripheral blood of TCR transgenic (a), wild-type (b) and rag2−/− (c) mice, analyzed by flow cytometry. Since TCR transgenic mice were generated by microinjection in rag2−/− oocytes, mature T cells will only be present when the inserted constructs provide a functional T-cell receptor

3.7.3  Evaluation of TCR Specificity in a DC-T-Cell Co-culture

1. One week prior to the experiment, prepare bone marrow dendritic cells, as described above. 2. Isolate the spleen and lymph nodes from a transgenic animal (see Note 22). 3. Obtain a single-cell suspension by mashing the spleen and lymph nodes through a 70 μm cell strainer in 1 mL PBS and rinse with 2 mL PBS. Centrifuge the cell suspension and lyse the erythrocytes by resuspending the cell pellet in 1 mL osmotic lysis buffer, incubate for 4 min on ice, and centrifuge after adding 10 mL of PBS. 4. Isolate CD4+ T cells using Miltenyi Biotech CD4+ T cells isolation kit, according to manufacturer’s instructions. 5. Wash the enriched CD4+ T cells twice in RPMI and label the cells with CFSE (see Note 23). Resuspend the cells in 1 mL of RPMI, dropwise add 110 μL of 1/100 pre-diluted CFSE (5 μM final concentration), while gently vortexing the cell suspension and incubate at 37 °C (see Note 24). After 10 min stop the CFSE staining by adding 10 mL of RPMI5, wash the cells twice in RPMI5 and count (see Note 25). 6. Start a co-culture by seeding 50 μL of bone marrow-derived dendritic cells (2 × 105 cells/mL) in 6 wells of a 96-well round bottom plate. Add 100 μL of the enriched CD4+ T cells at 106 cells/mL. Prepare a 1 in 2 serial dilution of the TCR-specific peptide in 50 μL and add to the different wells (include a medium control), starting at 1 μg/mL of peptide (see Note 14). All steps are performed in RPMI5. Culture the cells for 4 days.

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Fig. 5 Proliferation of CD3+CD4+ TCR transgenic T cells after a co-culture with bone marrow-derived dendritic cells in the absence (a) or presence of peptide in decreasing doses (b–f). Antigen-specific peptide was present in the culture at 1 μg/mL (b), 0.2 μg/mL (c), 0.04 μg/mL (d), 0.008 μg/mL (e), 0.0016 μg/mL (f). CFSE content is decreased by half each time cells divide, resulting in one peak per generation of proliferating cells

7. Evaluate T-cell proliferation by assessing the CFSE profiles of CD3+CD4+ T cells by flow cytometry, as described before (see Note 26) (see Fig. 5).

4  Notes 1. As an example, for an antigen with a stock concentration of 200  μg/mL, 50 μL of antigen is diluted with 425 μL PBS, followed by dropwise addition of 25 μL aluminum hydroxide. Mice are injected intraperitoneally with 500 μL of this formulation. 2. The optimal immunization protocol is dependent on the antigen and should be optimized. 3. Activated T cells can be isolated from draining lymph nodes or spleen, depending on the route of antigen administration. 4. Depending on the peptide, incubation time and peptide dose can be different. This should be optimized before. The pep-

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tide used can be a known immunogenic peptide, or ­immunogenicity can be predicted using IEDB database, and confirmed by in vitro experiments. 5. CD4+ T-cell isolation is described using the CD4+ T cell isolation kit from Miltenyi Biotech. However, isolation can also be performed by FACS or any other magnetic cell separation protocol. 6. It is important to wash the cells in medium without FCS or HEPES, as PEG will precipitate proteins, and HEPES can be toxic for the cells. 7. Pipetting the cells should be avoided from this step onwards, to avoid disrupting the fragile cell contacts. 8. Slow addition of the wash medium is necessary to gradually dilute the PEG, without lysing the cells. Small cells clumps will be visible. 9. Avoid using the outer wells of the plate. 10. HAT is added as a selection marker. It will kill the HAT sensitive unfused BW5147 cells, while fused cells will become HAT insensitive and survive. 11. Some clones will grow faster than others. Therefore, clones that become confluent very quickly after feeding can be reseeded at a lower density. 12. Here, we describe the use of bone marrow-derived dendritic cells as antigen-presenting cells. Alternatively, the syngeneic spleen cells can be used. In this case, plate 50 μL of the syngeneic spleen cells (2 × 106 c/mL). 13. Cytokine ELISA performed according to the manufacturer’s instructions. 14. Preparation of a serial dilution with peptide. Prepare a stock of 4 μg/mL (final concentration 1 μg/mL) and dilute 1/2 in DMEM10 for several steps. Mix 100 μL of peptide stock with 100 μL of DMEM10 (final concentration of 0.5 μg/mL), mix well, and transfer 100 μL to a second tube with 100 μL DMEM10 (final concentration 0.25 μg/mL); repeat till the appropriate amount of dilutions is obtained. The immunogenicity of the peptide used will determine the concentrations that need to be tested; therefore the example provided above is merely an indication of the required concentrations. 15. A maximum of 106 cells is stained in 50 μL. Increase the staining volume accordingly if the cell number is higher. 16. Selecting clones that express TCR domains to which an antibody is available provides advantages for future detection of TCR transgenic cells in mice.

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17. Ideally, cells are reseeded 1 day prior to freezing. Cells are then in log phase and grow exponentially, increasing recovery after freezing. 18. Single clones can also be obtained by other techniques such as limiting dilution. 19. There are different TCR screening panels available, and the nomenclature can be misleading. The numbering used in the different panels is not the same, and this should be taken into account when analyzing the TCR subtype. An overview of the correspondence between nomenclatures can be found in Tables 1 and 2. 20. p428 is a good choice for a TCR Vβ chain because of a deletion in a critical silencer region of the CD4 promotor, it is already expressed from double-negative thymocyte stage onwards. Consequently, a Vβ chain driven from this promotor gives a critically early and continued signal for T-cell development of the cells carrying the transgene. p783 is a good choice for expression of a class II-restricted Vα chain because it is not expressed in double-negative thymocytes (due to reinsertion of the CD4 silencer region) and thus will not contribute to a signal that would favor the unwanted formation of aberrant gamma-delta-like cells, which are often a complication in TCR transgenic mice. On top, p783 is silenced in CD8 cells and thus will not favor their development. 21. Alternatively, peripheral blood can also be analyzed directly after several erythrocyte lysis steps with osmotic lysis buffer. However, we experience that T cells are very fragile at this stage and can suffer from the repeated osmotic lysis steps. Isolating the lymphocytes via a gradient as described in the protocol increases the recovery of these cells. 22. Before performing this assay, make sure that there is sufficient transgenic offspring, as to ensure the transgenic line. 23. It is important that no FCS is present in the medium, as this markedly reduces the CFSE labeling. 24. A maximum of 50 × 106 cells is labeled in 1 mL; increase the volume of RPMI and CFSE accordingly if the cell number is higher. 25. The presence of FCS in the medium for washing of the cells is essential, as FCS stops the labeling, eliminating the toxic effects of CFSE. 26. It is important to include a live/dead marker in the staining, as a lot of nonproliferating cells die during the culture and should be excluded from the analysis.

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References antigen-­specific T cells in thymus by restricting 1. Coquet JM, Schuijs MJ, Smyth MJ, Deswarte MHC molecules. Nature 335(6192):730–733 K, Beyaert R, Braun H, Boon L, Hedestam Gunilla BK, Nutt Steven L, Hammad H, 7. Shimonkevitz R, Colon S, Kappler JW, Marrack Lambrecht BN (2015) Interleukin-21-­ P, Grey HM (1984) Antigen recognition by producing CD4+ T cells promote type 2 H-2-restricted T cells. II. A tryptic ovalbumin immunity to house dust mites. Immunity peptide that substitutes for processed antigen. 43(2):318–330 J Immunol 133(4):2067–2074 2. Plantinga M, Guilliams M, Vanheerswynghels 8. Barnden MJ, Allison J, Heath WR, Carbone M, Deswarte K, Branco-Madeira F, Toussaint FR (1998) Defective TCR expression in transW, Vanhoutte L, Neyt K, Killeen N, Malissen genic mice constructed using cDNA-based B, Hammad H, Lambrecht Bart N (2013) alpha- and beta-chain genes under the control Conventional and monocyte-derived CD11b+ of heterologous regulatory elements. Immunol dendritic cells initiate and maintain T helper 2 Cell Biol 76(1):34–40 cell-mediated immunity to house dust mite 9. Wiley J & Sons (2009) Current protocols in allergen. Immunity 38(2):322–335 immunology. In: Kruisbeek Ada M. Generation 3. Dullaers M, Schuijs MJ, Willart M, Fierens of mouse T cell hybridomas, Vol 1, chapter K, Van Moorleghem J, Hammad H, 3.14 Lambrecht BN (2016) House dust mite- 10. Inaba K, Inaba M, Romani N, Aya H, Deguchi driven asthma and allergen-specific T cells M, Ikehara S, Muramatsu S, Steinman RM depend on B cells when the amount of (1992) Generation of large numbers of deninhaled allergen is limiting. J Allergy Clin dritic cells from mouse bone marrow cultures Immunol 140(1):76–88 supplemented with granulocyte/macrophage 4. Kisielow P, Bluthmann H, Staerz UD, colony-stimulating factor. J Exp Med Steinmetz M, von Boehmer H (1988) 176(6):1693–1702 Tolerance in T-cell-receptor transgenic mice 11. Hermansson A, Ketelhuth D, Strodthoff D, involves deletion of nonmature CD4+8+ thyWurm M, Hansson E, Nicoletti A, Paulsson-­ mocytes. Nature 333(6175):742–746 Berne G, Hansson G (2010) Inhibition of T 5. Sha WC, Nelson CA, Newberry RD, Kranz cell response to native low-density lipoprotein DM, Russell JH, Loh DY (1988) Positive and reduces atherosclerosis. J Exp Med negative selection of an antigen receptor on T 207(5):1081–1093 cells in transgenic mice. Nature 12. Wang Q, Malherbe L, Zhang D, Zingler K, 336(6194):73–76 Glaichenhaus N, Killeen N (2001) CD4 6. Kisielow P, Teh HS, Bluthmann H, von promotes breadth in the TCR repertoire. Boehmer H (1988) Positive selection of J Immunol 167:4311–4320

Chapter 16 Using Cytokine Reporter Mice to Visualize Type-2 Immunity In Vivo Mark Dell’Aringa and R. Lee Reinhardt Abstract Type-2 cytokine production plays a critical role in the context of type 2 immunity and allergic inflammation. Interleukin-4 (IL-4) and IL-13 are key modulators of the cell-mediated and humoral immune hallmarks most commonly associated with type-2 immune responses. However, production of these cytokines by lymphocytes and their tissue localization has been difficult to detect in vivo. As such, the field has relied heavily on ex vivo restimulation and in vitro differentiation assays to understand type-2 cytokine biology. Although these studies have greatly informed our understanding of type-2 cytokine regulation, it is becoming increasingly clear that the data does not always provide a true accounting of the complexity of type-2 immune cell biology in vivo. Described below is a protocol used to detect IL-4-competent and protein-­ producing cells in the lung and lymph nodes of mice after infection with a helminth. Importantly, this protocol has also been used to successfully identify reporter expression and cell function in vivo using various other cytokine-reporter systems. Key words Interleukin-4, T helper 2 cells (Th2), T follicular helper cells (Tfh), Helminth, Tyramide signal amplification, Immunohistochemistry

1  Introduction Type-2 immunity results from the hosts response to parasitic worm infection, a process commonly observed in developing regions of the world where over 1.5 billion individuals are infected with intestinal helminths [1]. In urban centers, type-2 inflammation more commonly presents as atopy, allergy, and asthma [2, 3]. Clearance of helminths and allergic pathology are both mediated by T helper 2 (Th2) cells. Th2 cells promote helminth expulsion and allergic pathology by producing cytokines, such as IL-4 and IL-13, that are critical for mucus production, goblet cell hyperplasia, smooth muscle contractility, and innate immune cell recruitment to the site of colonization [4–7]. In addition to the type-2 immune hallmarks orchestrated by Th2 cells in peripheral tissues, specific humoral hallmarks, such as the increased production of immunoglobulin

R. Lee Reinhardt (ed.), Type 2 Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1799, https://doi.org/10.1007/978-1-4939-7896-0_16, © Springer Science+Business Media, LLC, part of Springer Nature 2018

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(Ig)E and high-affinity IgG1 are also associated with type-2 immune responses. Affinity maturation and isotype class-switching of B cells to IgE and IgG1 is heavily reliant on interactions of germinal center (GC) B cells with IL-4-producing T follicular helper (Tfh) cells [8–11]. The ability to accurately detect IL-4- and IL-13-expressing cells and to identify their precise location within affected organs are critical steps toward a better understanding of type-2 immunity. As such, detection of IL-4-expressing Th2 and Tfh cells in vivo provides an opportunity to understand key aspects of type-2 immunity that cannot be explored in an in vitro or ex vivo setting. The generation of two IL-4 reporter mice, IL44get and IL4KN2, has significantly enhanced our ability to understand the biology of IL-4-expressing cells in vivo [12, 13]. Briefly, the IL44get mouse has an internal ribosomal entry site (IRES)-enhanced green fluorescent protein (GFP) gene knocked-in downstream of the stop codon of the endogenous IL-4 locus. Upon the transcription of the il4 locus in these mice, a bicistronic mRNA transcript is generated allowing both IL-4 and GFP to be expressed. IL-4-expressing cells, often referred to as IL-4-competent cells, are identified by their robust expression of GFP. The IL4KN2 mouse has the endogenous il4 locus replaced with a human CD2 (huCD2) reporter. In these mice, IL-4 protein-producing cells can be identified by surface expression of huCD2. Human CD2 expression correlates with recent IL-4 protein production in these mice [12, 14]. By using mice expressing both IL-4 reporters, a distinction can be made between cells that are competent for IL-4 (GFP+) and cells that are actively producing IL-4 (huCD2+). These mice have provided novel insight into previously unappreciated aspects of type-2 immune cell biology. Described below is a protocol used to stain for GFP and huCD2 for detection of IL-4-expressing and IL-4-producing Th2 and Tfh cells using immunohistochemistry. The GFP expression from IL44get mice in most cases is not bright enough to be detected by conventional fluorescent microscopes and is quenched following tissue fixation. Therefore, to detect IL44get-competent cells, tissue sections are fixed and stained with anti-GFP antibodies, which are further amplified using tyramide signal amplification (TSA) to increase the signal-to-noise ratio [15]. This procedure is demonstrated on IL44get-fixed lung (Fig. 1) and mediastinal lymph node (Fig. 2) tissue sections obtained from Nippostrongylus brasiliensis-­ infected IL44get mice. To detect IL-4-producing cells using IL4KN2 reporter mice, sections are stained with anti-huCD2 and amplified using TSA. We show tyramide amplified huCD2 staining of fresh-­ frozen lymph node sections (Fig. 2) from helminth-infected IL4KN2 mice. Importantly, with minor modifications, this protocol can be adapted for use with other type-2 cytokine reporter mice [4, 16–18].

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Fig. 1 Detection of IL-4 expressing CD4+ T cells in the lung of N. brasiliensis-infected mice. IL44get mice were infected with 500 (L3) N. brasiliensis worms, and lungs were harvested and prepared for histology 9 days postinfection. (a) Lungs were stained for CD4 (red) and GFP/IL-4 (green). A composite picture is shown on the far right. The bottom panel shows images acquired from a wildtype mouse  (reporter-negative control). All images were acquired at 100× magnification (10x objective)

2  Materials 2.1  Mice

1. IL44get reporter mice: to assess IL-4 mRNA expression, read out as GFP. 2. IL4KN2 reporter mice: to assess IL-4 protein production, read out as huCD2. 3. C57BL/6 mice: serve as negative controls for GFP and huCD2 reporters.

2.2  Infection of IL-4 Reporter Mice with N. brasiliensis

1. 26-gauge (G) needle. 2. 1 mL syringe. 3. 0.9% saline in distilled water. 4. L3 N. brasiliensis larvae. 5. 50 mL conical tubes. 6. 2 mL Eppendorf tubes. 7. Isoflurane anesthesia machine. 8. 70% ethanol.

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Fig. 2 Imaging of IL-4 expressing and IL-4 producing T cells in the lymph node of N. brasiliensis-infected mice. IL44get or IL4KN2 mice were infected with 500 (L3) N. brasiliensis worms. The lung-draining lymph node was harvested and prepared for histology 9 days later. (a) Representative 50× images (5x objective) of the entire lymph node. Lymph nodes were stained for CD4 (red), GFP/IL-4 (green), and IgD (blue). A composite image is shown. A WT control for negative GFP staining is shown on the far right. (b) Representative 10× images of the lymph node showing a germinal center as negative IgD staining within the B-cell follicle. A WT control of GFP staining is shown on the right. (c) A 100× image (10x objective) of fresh-frozen IL4KN2 lymph node is shown. The lymph node was stained for CD4 (red), huCD2/IL-4 (green), and IgD (blue). A WT control for huCD2 staining is shown in a composite on the far right

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1. Forceps and scissors for dissection. 2. Suture thread. 3. 18-gauge catheter needle for lung inflations. 4. Phosphate buffered saline (PBS). 5. 4% paraformaldehyde solution: 2 mM MgCl2, 1.25 mM EGTA with a final pH of 7.5 in PBS. 6. Optimal cutting temperature (O.C.T.) freezing media. 7. 10 × 10 × 5 mm cryomolds for freezing lymph nodes. 8. 15 × 15 × 5 mm cryomolds for freezing lung tissue. 9. 2-methylbutane. 10. Dry ice. 11. 50 mL conical tubes. 12. 30% sucrose in PBS. 13. Acetone.

2.4  Cutting Tissue Sections

1. Positively charged slides for applying tissue sections. 2. Cryostat. 3. Optimal cutting temperature (O.C.T.) freezing media. 4. Small paintbrushes (size 0–4). 5. Low- or high-profile microtome blades (depending on the cryostat model).

2.5  Staining Tissue Sections

1. PBS. 2. 50 mL conical tube. 3. Normal IgG rat serum. 4. Purified anti-CD16/CD32 antibody to block Fc receptors. 5. TNB blocking buffer: 0.1 M Tris–HCl, 0.15 M NaCl, and 0.5% blocking reagent (supplied in tyramide amplification kits) in H2O, pH 7.5. 6. Fc block: anti-CD16/CD32 antibodies (0.5 mg/mL stock). 7. 1% H2O2/0.1% sodium azide solution: 1% H2O2, 0.1% sodium azide in PBS. 8. Avidin- and biotin-blocking kit to block endogenous biotin. 9. Purified polyclonal rabbit anti-GFP antibody (1 mg/mL stock; see Note 1). 10. Donkey anti-rabbit antibody conjugated to biotin (1 mg/mL stock). 11. Antihuman CD2 antibody conjugated to biotin (0.5 mg/mL stock). 12. Horseradish peroxidase (HRP) conjugated to streptavidin.

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13. Fluorescein tyramide amplification kit. 14. Anti-CD4 antibody conjugated to biotin (0.5 mg/mL stock). 15. Anti-B220 antibody conjugated to biotin (250 μg/mL stock). 16. Alexa Fluor 555 tyramide amplification kit. 17. Biotin tyramide amplification kit. 18. DyLight 649 conjugated to streptavidin (0.5 mg/mL stock). 19. Vectashield to prevent photobleaching of stained tissue sections. 20. Cover slips (22 × 40 mm or 22 × 50 mm). 21. 4′,6-diamidino-2-phenylindole (DAPI) stock solution: 1 mg/ mL DAPI in ddH2O. 22. DAPI Working Solution: 0.5 μg/mL in PBS. 2.6  Imaging and Analysis of Tissue Sections

1. Fluorescent microscope. Specifications of the microscope used in this protocol: inverted Zeiss 200M with 175-W xenon lamp in a DG4 lamp housing (Sutter Instruments), motorized stage (Applied Scientific Instruments, Inc.) for tiling and montage acquisition, Lambda 10-2 filter wheel control unit (Sutter Instruments), Sensicam CCD camera (Cooke), Cy3, Cy5, FITC, and DAPI filter cubes, 5× and 10× air objectives were used. 2. FIJI (ImageJ) for image analysis.

3  Methods 3.1  Infection of IL44get Mice with N. brasiliensis

3.2  Extraction and Fixation of the Mediastinal Lymph Node

1. Harvest third-stage N. brasiliensis larvae (L3) from cultured fecal material of N. brasiliensis-infected rats. 2. Dilute the larval worms in saline to a concentration of 500 L3 worms per 200 μL, and inject mice subcutaneously in the rear flank using a 26 G needle (see Note 2). IL4KN2 lymph nodes should not be fixed but frozen immediately after removal from the mouse. Only perform fixation steps for IL44get tissues. 1. Euthanize mice according to institutional and IACUC guidelines. 2. Carefully open the chest cavity of the mouse by cutting upwards on both sides of the rib cage (see Note 3). 3. Gently push the lungs aside, and remove the connective tissue surrounding the lymph node. Once most of the connective tissue has been removed, excise the lymph node by placing the forceps underneath the node and pulling gently upward. Carefully dissect excess fat from the lymph node (see Note 4).

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4. Place the lymph node in a 50 mL conical tube containing 2 mL of 4% PFA (see Note 5). 5. After 2 h remove the lymph node from the PFA, and place it in a new 50 mL conical with 2 mL of PBS to wash the tissue. 6. After 4–6 h, place the lymph node in a new 50 mL conical with 2 mL of 30% sucrose and incubate overnight at 4 °C. 7. The next day make a dry ice bath by placing several pieces of dry ice into a small container with 2-methylbutane. 8. Fill a small 10 × 10 × 5 mm cryomold with O.C.T. and place the lymph node cortical dome side down into the O.C.T. media and gently submerge the tissue to ensure it is completely covered with O.C.T. Freeze the lymph node in O.C.T. by placing them cryomold into the dry ice bath. 9. Store the frozen tissues at −80 °C until ready to section. 3.3  Extraction and Fixation of Lung Tissue

1. Once the mediastinal lymph node has been removed, remove the thymus and heart. 2. Carefully clear excess fat and connective tissue in the chest cavity and neck to reveal the trachea. Make a small incision into the upper portion of the trachea (see Note 6). 3. Attach a catheter needle cover to a 5 mL syringe filled with 4% PFA. Carefully place the catheter needle cover and 5 mL syringe gently into the opening of the trachea, and tie it securely with suture thread. 4. Inject the PFA into the trachea to inflate the lung. Once the lung is properly inflated, remove the syringe and quickly tighten the suture thread to prevent PFA from leaking out of the lung and/or trachea. 5. Fill a 50 mL conical with 20 mL of 4% PFA. Place the inflated lung into this conical, and allow it to fix for 3 h. Turn conical upside-down during incubation to allow lung to fully submerge in PFA. 6. Remove the lung from the PFA, and cut the trachea to remove the suture wire. Wash the lung with PBS to remove excess PFA. Place the lung into a 50 mL conical filled with 20 mL of PBS. 7. After 1 h place the lung into a new 50 mL conical with 20 mL of PBS and repeat once. Incubate the lung in PBS overnight at 4 °C. 8. The next day, place the lung into a 50 mL conical with 20 mL of 10% sucrose at 4 °C. After 1 h, move the lung to a new 50 mL conical with 20 mL of 30% sucrose, and incubate for 1 h at 4 °C.

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9. Then place the lung into a 50 mL conical of 30% sucrose/ O.C.T. media (1:1). Let the lung incubate overnight at 4 °C. 10. Fill a 15 × 15 × 5 mm cryomold with O.C.T. and place a lobe of the lung inside the mold. Freeze the lung using a dry ice, 2-methylbutane bath and immediately store at −80 °C. 3.4  Sectioning Tissues on the Cryostat

1. Remove the frozen tissues from the −80 °C freezer and place inside the cryostat to allow the tissues to warm up to −20 °C. 2. Freeze the O.C.T tissue block to the chuck with O.C.T. 3. Cut the tissues into 8 μM sections, roll them flat onto the stage using the anti-roll bar or with a paintbrush, then collect them onto room-temperature positively charged slides. 4. IL4KN2 tissues should be lightly fixed/dehydrated in acetone for 10 min after they are transferred to the slides. Do not perform this step for the PFA-fixed IL44get tissues. 5. Store slides at −80 °C until ready to stain the sections.

3.5  Staining IL44get the Lung and Lymph Node Tissue Sections

All steps are performed at room temperature. Once fluorescent antibodies or tyramide are added to slides all incubation steps should be done in a covered humidified chamber. Staining lung and lymph node sections for CD4 and IL-4 (GFP): 1. Remove tissue sections from −80 °C, and allow them to equilibrate to room temperature for about 10 min. 2. Once the sections have dried, use a PAP pen to outline the tissues. This will allow you to put liquid on the slide to submerge the tissue without the liquid running off the slide. Never let tissue dry from this point forward. 3. Place the slides into a Coplin jar filled with 1% H2O2/0.1% sodium azide. Incubate for 30 min. Dislodge bubbles that form on the tissues by slowly agitating the slides in the Coplin jar. 4. Wash the slides gently by lightly running PBS over the tissue using a 6 mL transfer pipette, and then place the slides in a Coplin jar filled with PBS. 5. Place the slides into a humidified chamber (see Note 7). Incubate slides with 1% rat serum and 1% Fc block in TNB buffer to block nonspecific binding. Add enough blocking solution to the slide to fill the region outlined with the PAP pen and submerge the tissues. Incubate for 30 min. 6. Wash the slides with PBS as described previously in step 4 (and for all upcoming steps requiring a PBS wash). Use avidin solution to block endogenous biotin. Incubate for 20 min. 7. Wash with PBS. Incubate with biotin solution for 20 min to bind up excess avidin.

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8. Wash with PBS. Dilute polyclonal rabbit ant-GFP antibody 1:2000 in TNB and stain IL44get tissues for 1 h. 9. Wash with PBS. Incubate tissues for 30 min with biotinylated donkey anti-rabbit diluted 1:500 in TNB. 10. Wash with PBS. Incubate tissues for 25 min with streptavidin conjugated to horseradish peroxidase (HRP) diluted 1:200 in TNB. 11. Wash with PBS. Stain the slides with FITC-conjugated tyramide at 1:100 in PBS (or amplification diluent supplied in the kit) for 8 min (see Note 8). 12. Wash the slides with PBS immediately after staining with FITC-conjugated tyramide. 13. Repeat steps 3, 6, and 7 to quench peroxidase activity and to block biotin and avidin (see Note 9). 14. Wash the slides and incubate with biotinylated anti-CD4 at 1:250 in TNB buffer for 1 h. 15. Wash with PBS. Incubate the slides with streptavidin conjugated to HRP at 1:200 in TNB and incubate tissues for 25 min. 16. Wash the slides and incubate with Alexa Fluor 555 conjugated to tyramide at 1:200 in PBS for 8 min. Immediately wash with PBS. 17. Wash all the slides and repeat steps 3, 6, and 7 for the lymph node tissues only. Lung tissues should be incubated with DAPI working solution for 5 min. 18. Wash the lung tissues with PBS for a final time. Apply two to three drops of Vectashield onto a cover slip and place it carefully onto the slide (see Note 10). The lung tissues are now ready to be imaged. Staining lymph node sections for IgD: 19. Wash lymph node tissues with PBS and stain with biotinylated anti-IgD at 1:250 in TNB for 1 h. 20. Wash the lymph node tissues with PBS and stain with streptavidin conjugated to HRP diluted at 1:200 in TNB for 25 min. 21. Wash again with PBS and stain the lymph node tissues with biotinylated tyramide at a concentration of 1:100 in PBS for 8 min. 22. Wash with PBS and stain lymph node tissues with streptavidin conjugated to DyLight 649 diluted at 1:500 in TNB. Incubate for 30 min. 23. Wash with PBS, and stain the lymph node tissues with DAPI working solution for 5 min. After the DAPI stain, wash the lymph node tissues with PBS a final time, and apply a coverslip with Vectashield to the slide as described previously. The lymph node tissues are now ready to be imaged.

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3.6  Staining IL-4KN2 Lymph Node Sections

1. Remove tissue sections from −80 °C and allow them to equilibrate to room temperature for approximately 15 min. 2. Once the sections have dried, use a PAP pen to outline the tissues. This will allow you to put liquid on the slide to submerge the tissue without the liquid running off the slide. 3. Place the slides into a Coplin jar filled with 1% H2O2/0.1% sodium azide for 30 min. This will quench the endogenous peroxidase activity. Dislodge any bubbles that form on the tissues by agitating the slides in the Coplin jar. 4. Wash the slides gently by lightly running PBS over the tissue using a 6 mL transfer pipette and place the slides in a Coplin jar filled with PBS. 5. Place the slides into a humidified chamber (see Note 7). Incubate slides with 1% rat serum in Fc block to block nonspecific antibody binding. Add enough blocking solution to the slide to fill the region outlined with the PAP pen and submerge the tissues. Incubate for 30 min. 6. Wash the slides with PBS as described previously in step 4 (and for all upcoming PBS wash steps). Use avidin solution to block endogenous biotin. Incubate for 20 min. 7. Wash with PBS. Incubate with biotin solution for 20 min to soak up excess avidin. 8. Wash with PBS. Incubate the tissues for 1 h in a 1:250 dilution of biotinylated antihuman CD2 antibody in TNB. 9. Wash with PBS. Incubate tissues for 25 min with streptavidin conjugated to horseradish peroxidase (HRP) diluted 1:200 in TNB. 10. Wash with PBS. Stain the slides with FITC-conjugated tyramide at 1:100 in PBS for 8 min (see Note 8). 11. Wash the slides with PBS immediately after staining with FITC-conjugated tyramide. 12. Repeat steps 3, 6, and 7 to quench peroxidase activity and to block biotin and avidin (see Note 9). 13. Wash the slides and incubate with biotinylated anti-CD4 at 1:250 in TNB buffer for 1 h. 14. Wash with PBS. Incubate the slides with streptavidin conjugated to HRP 1:200 in TNB and incubate tissues for 25 min. 15. Wash the slides and incubate with Alexa Fluor 555 conjugated to tyramide at 1:200 in PBS for 8 min. Immediately wash with PBS. 16. Wash all the slides and repeat steps 3, 6, and 7. 17. Wash with PBS and stain with biotinylated anti-IgD at a concentration of 1:250 in TNB for 1 h.

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18. Wash the tissues with PBS, and stain with streptavidin conjugated to HRP diluted at 1:200 in TNB for 25 min. 19. Wash again with PBS, and stain the tissues with biotinylated tyramide at a concentration of 1:100 in PBS for 8 min. 20. Wash with PBS, and incubate tissues with streptavidin conjugated to DyLight 649 diluted at 1:500 in TNB for 30 min. 21. Wash with PBS and stain the tissues with 1 μg/mL DAPI in TNB for 5 min. After the DAPI stain, wash the lymph node tissues a final time with PBS. Apply two to three drops of Vectashield to a coverslip, and place the coverslip onto the slides. The IL4KN2 lymph node tissues are now ready to be imaged. 3.7  Image Acquisition and Analysis

1. Power up microscope and associated equipment according to your institution’s instructions. 2. Place an IL44get or IL4KN2 tissue slide under the microscope objective. Using the DAPI filter locate the tissue and focus appropriately. 3. Using the acquisition software, find the optimal exposure times for each channel you are imaging (see Note 11). Set the images to your desired brightness settings (see Note 12). 4. Take images at the desired magnifications in each channel (see Note 13). For this experiment, we acquired images using 5× and 10× objectives. Ensure the images taken are representative of the entire tissue. 5. Analyze images using your software of choice.

4  Notes 1. Polyclonal anti-GFP antibodies have worked better than monoclonal antibodies in our hands. 2. The worms will settle quickly in solution and need to be thoroughly mixed in between each injection. 3. Be sure not to puncture the lungs when opening the rib cage. You will need to subsequently inflate the lungs with 4% PFA; a damaged lung will make inflation very difficult. 4. Do not crush or squeeze the lymph node. Inflamed lymph nodes can be very fragile and are prone to rupturing. Only grasp the lymph node from underneath the tissue, and be gentle when cleaning excess fat off the lymph node. 5. Fresh PFA solution is optimal for fixation of tissues. PFA older than 1 week will begin to degrade and work less efficiently. Fixation to mediate light cross-linking of proteins is required

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to ensure cytoplasmic GFP is not washed away during the staining protocol. 6. To inflate the lungs, the trachea must remain intact. Do not cut entirely through the trachea, but only make a small incision to open the tracheal lumen. 7. A humidified chamber is used to prevent drying of the tissue sections. This can be accomplished with readily available materials: we cover the bottom of a large plastic Tupperware container with moist paper towels, 1 mL pipettes are then placed on top of the towels, and the slides rest suspended above the moisture upon two of the 1 mL pipettes while incubating. 8. It is important to wash immediately with PBS after incubating the slides with tyramide. Over-incubation with tyramide can lead to nonspecific background staining. 9. Only perform these blocking steps again if you are planning on doing additional staining steps. If your subsequent stains do not require tyramide amplification or horseradish peroxidase, the hydrogen peroxide incubation step can be skipped. If the additional stains do not require a biotinylated antibody, then additional avidin and biotin blocks can be skipped as well. 10. Vectashield is a mounting medium that helps to prevent photobleaching of fluorescently stained tissues. 11. Exposure times will vary. However, it is important that within each experiment, exposure settings are maintained between samples. Using different exposure times will bias data and any subsequent quantification if comparing different groups. 12. When comparing images, it is important to keep brightness and contrast settings the same. 13. Be sure not to move the objective when acquiring images in multiple channels at the same position.

Acknowledgments This research was funded in part by NIH grant R01AI119004 (R.L.R). References 1. Hotez PJ, Brindley PJ, Bethony JM, King CH, Pearce EJ, Jacobson J (2008) Helminth infections: the great neglected tropical diseases. J Clin Invest 118(4):1311–1321. https://doi. org/10.1172/JCI34261 2. Lambrecht BN, Hammad H (2014) The immunology of asthma. Nat Immunol 16(1):45–56. https://doi.org/10.1038/ ni.3049

3. Locksley RM (2010) Asthma and allergic inflammation. Cell 140(6):777–783. https:// doi.org/10.1016/j.cell.2010.03.004 4. Liang HE, Reinhardt RL, Bando JK, Sullivan BM, Ho IC, Locksley RM (2012) Divergent expression patterns of IL-4 and IL-13 define unique functions in allergic immunity. Nat Immunol 13(1):58–66. https://doi. org/10.1038/ni.2182

Tissue Detection of Cytokines Using Cytokine Reporter Mice 5. Grunig G, Warnock M, Wakil AE, Venkayya R, Brombacher F, Rennick DM, Sheppard D, Mohrs M, Donaldson DD, Locksley RM, Corry DB (1998) Requirement for IL-13 independently of IL-4 in experimental asthma. Science 282(5397):2261–2263 6. Corry DB, Folkesson HG, Warnock ML, Erle DJ, Matthay MA, Wiener-Kronish JP, Locksley RM (1996) Interleukin 4, but not interleukin 5 or eosinophils, is required in a murine model of acute airway hyperreactivity. J Exp Med 183(1):109–117 7. Fallon PG, Jolin HE, Smith P, Emson CL, Townsend MJ, Fallon R, McKenzie AN (2002) IL-4 induces characteristic Th2 responses even in the combined absence of IL-5, IL-9, and IL-13. Immunity 17(1):7–17 8. Haynes NM, Allen CD, Lesley R, Ansel KM, Killeen N, Cyster JG (2007) Role of CXCR5 and CCR7 in follicular Th cell positioning and appearance of a programmed cell death gene-­ 1high germinal center-associated subpopulation. J Immunol 179(8):5099–5108 9. Reinhardt RL, Liang HE, Locksley RM (2009) Cytokine-secreting follicular T cells shape the antibody repertoire. Nat Immunol 10(4):385– 393. https://doi.org/10.1038/ni.1715 10. Zaretsky AG, Taylor JJ, King IL, Marshall FA, Mohrs M, Pearce EJ (2009) T follicular helper cells differentiate from Th2 cells in response to helminth antigens. J Exp Med 206(5):991– 999. https://doi.org/10.1084/ jem.20090303 11. King IL, Mohrs M (2009) IL-4-producing CD4+ T cells in reactive lymph nodes during helminth infection are T follicular helper cells. J Exp Med 206(5):1001–1007. https://doi. org/10.1084/jem.20090313

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12. Mohrs K, Wakil AE, Killeen N, Locksley RM, Mohrs M (2005) A two-step process for cytokine production revealed by IL-4 dual-reporter mice. Immunity 23(4):419–429 13. Mohrs M, Shinkai K, Mohrs K, Locksley RM (2001) Analysis of type 2 immunity in vivo with a bicistronic IL-4 reporter. Immunity 15(2):303–311 14. Scheu S, Stetson DB, Reinhardt RL, Leber JH, Mohrs M, Locksley RM (2006) Activation of the integrated stress response during T helper cell differentiation. Nat Immunol 7(6):644–651 15. Reinhardt RL, Khoruts A, Merica R, Zell T, Jenkins MK (2001) Visualizing the generation of memory CD4 T cells in the whole body. Nature 410(6824):101–105 16. Nussbaum JC, Van Dyken SJ, von Moltke J, Cheng LE, Mohapatra A, Molofsky AB, Thornton EE, Krummel MF, Chawla A, Liang HE, Locksley RM (2013) Type 2 innate lymphoid cells control eosinophil homeostasis. Nature 502(7470):245–248. https://doi. org/10.1038/nature12526 17. Neill DR, Wong SH, Bellosi A, Flynn RJ, Daly M, Langford TK, Bucks C, Kane CM, Fallon PG, Pannell R, Jolin HE, McKenzie AN (2010) Nuocytes represent a new innate effector leukocyte that mediates type-2 immunity. Nature 464(7293):1367–1370. https://doi. org/10.1038/nature08900 18. Huang Y, Guo L, Qiu J, Chen X, Hu-Li J, Siebenlist U, Williamson PR, Urban JF Jr, Paul WE (2015) IL-25-responsive, lineage-­negative KLRG1(hi) cells are multipotential ‘inflammatory’ type 2 innate lymphoid cells. Nat Immunol 16(2):161–169. https://doi. org/10.1038/ni.3078

Chapter 17 Live Imaging of IL-4-Expressing T Follicular Helper Cells in Explanted Lymph Nodes Mark Dell’Aringa, R. Lee Reinhardt, Rachel S. Friedman, and Jordan Jacobelli Abstract The generation of class-switched, high-affinity, antibody-producing B cells plays a critical role in the ­establishment of type 2 immunity to intestinal helminths as well as in the pathogenesis of allergy and asthma. The generation of these high-affinity, antibody-producing B cells occurs in germinal centers (GC) and relies on interactions with follicular dendritic cells (FDCs) and T follicular helper (Tfh) cells. One ­critical mediator produced by Tfh cells in GCs is interleukin-4 (IL-4). Tfh-derived IL-4 drives class ­switching to type 2 antibody isotypes IgE and IgG1 and is required for high-affinity IgG1 production. In vivo detection of IL-4-expressing Tfh cells is required to better understand the role of these cells during the GC response. Detection of IL-4-expressing cells has been greatly improved by the generation of the IL-44get reporter mice, which read out IL-4 expression as green fluorescent protein (GFP). Much has been learned from these mice with regard to type 2 immunity using flow cytometry and immunohistochemistry. However, these methods do not allow the study of cellular behavior and interactions in real time. In ­contrast, multi-­photon microscopy allows for deep tissue imaging and tracking of multiple cell types in intact tissues over time. Here, we describe a protocol for in vivo detection of IL-4-expressing Tfh cells in an explanted popliteal lymph node by multi-photon microscopy. The dynamics of Tfh cell motility and their interactions with FDC networks in the GCs were analyzed. Key words T follicular helper cell, Follicular dendritic cell, Germinal center, IL-4, Two-photon microscopy, Multi-photon microscopy, Fluorescent reporter mice

1  Introduction The incidence of allergic and asthmatic disease is rapidly ­increasing worldwide. It is estimated that hundreds of millions of individuals suffer from allergic symptoms, with 300 million people being afflicted with asthma [1]. Allergic and asthmatic

Rachel S. Friedman and Jordan Jacobelli contributed equally to this work. R. Lee Reinhardt (ed.), Type 2 Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1799, https://doi.org/10.1007/978-1-4939-7896-0_17, © Springer Science+Business Media, LLC, part of Springer Nature 2018

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responses are largely the result of detrimental type 2 immune responses to ­normally inert environmental antigens. A hallmark of allergic and asthmatic responses is the presence of ­immunoglobulin (Ig) E and IgG1. IgE and IgG1 are antibody isotypes that are produced by subsets of B cells that have ­undergone maturation in response to antigen and cytokines in lymphoid tissues. The majority of B-cell isotype switching occurs in germinal centers (GC). The m ­ aturation, selection, and s­ urvival of GC B cells are tightly regulated by their interactions with ­follicular dendritic cells (FDCs) and CD4+ T ­follicular helper cells (Tfh cells) [2–4]. In addition, cytokine ­production by Tfh cells plays a key role in the GC B-cell m ­ aturation process. One essential cytokine produced by Tfh cells is IL-4. In most ­physiologic settings, Tfh cells are the predominant source of IL-4 protein within the lymph node and GC [3, 5, 6]. The in vivo detection of IL-4-expressing Tfh cells is important to better understand the role, location, and dynamics of Tfh cell function. The ability to detect IL-4-expressing Tfh cells has been greatly aided by the generation of IL44get reporter mice [7]. IL44get reporter mice allow for visualization of IL-4 competent cells through the production of green fluorescent protein (GFP). Although these mice have been used successfully in both flow ­cytometric and static imaging studies, such methods are limited in their ability to provide important data on the dynamics of IL-4expressing cells in living tissues [2, 3, 7, 8]. As such, little is known about their cellular interactions and motility during allergic responses. Multi-photon microscopy allows for imaging and ­tracking of live cells over time in intact tissues, and therefore, this technology provides an opportunity to assess true cellular behavior in real time. Various studies have been performed using ­multi-photon microscopy to investigate the cellular dynamics and interactions of GC B cells, Tfh cells, and FDCs [9–11]. Here, we describe how to use the IL44get reporter mice ­combined with multi-photon microscopy to image live, IL44getexpressing cells in explanted popliteal lymph nodes. We imaged GFP+ IL44get cells in an endogenous setting using a homozygous IL44get mouse (IL44get/4get), to maximize the GFP fluorescent s­ ignal. The B-cell follicle and follicular dendritic cells (FDCs) were ­delineated by fluorescently labeled B cells and antigen-antibody complexes, respectively (Fig. 1). Due to the high cell density of GFP+ cells in homozygous IL44get/4get mouse, it is not always ­possible to reliably track individual GFP-expressing cells in the endogenous setting to analyze motility properties. To enhance the likelihood of successful tracking of GFP+ IL44get cells, we also ­ performed a transfer experiment using IL44get/+ OT-II T cells (Fig.  2). The protocol and experiments described below can be adapted to investigate the motility and interaction dynamics of IL-4-expressing cells in vivo using multi-photon microscopy.

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Fig. 1 Detection of GFP+ IL44get/4get cells in an endogenous setting. (a) Schematic describing the experimental set up. IL44get/4get mice were immunized with OVA precipitated in alum on day 0. Six days postimmunization, VPD-stained naïve B cells were transferred and FDCs were labeled using PE immune complexes. On day 7, mice were euthanized, and the popliteal lymph node was harvested for multi-photon analysis. (b) Representative multi-photon images of VPD-stained B cells (blue), PE immune complexes (red), and GFP+ IL44get/4get cells (green) taken at 870 nm. A composite image is shown and potential GC regions are circled. Scale bar = 50 μm

2  Materials 2.1  Mice

1. IL-44get reporter mouse. 2. C57BL/6 mouse to obtain naïve B cells for subsequent staining and transfer (see Subheadings 3.2 and 3.3). 3. C57BL/6 mouse to serve as a negative control for GFP ­expression during microscopy. 4. OT-II mouse crossed to IL44get reporter mouse as donor for the transfer experiment.

2.2  Immunization to Induce Tfh Differentiation and Germinal Center Formation

2.3  Isolation of Naïve B Cells for Subsequent Labeling and Transfer

1. Ovalbumin. 2. Alum adjuvant (Imject; Thermo Fisher Scientific). 3. 1 mL syringe with 28G needle for injection. 4. Phosphate buffered saline (PBS). 5. Isoflurane and anesthesia machine. 1. 70% ethanol. 2. 3 mL syringe plunger. 3. 80 μM filter mesh.

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Fig. 2 Transfer of IL44get/+ OT-II T cells allows for more reliable tracking and analysis. (a) Schematic of the ­experimental set up. Briefly, heterozygous IL44get/+ OT-II T cells were transferred into a naïve C57BL/6 recipient on day-1. On day 0, mice were immunized with OVA precipitated in alum. On day 6, VPD-stained B cells were transferred, and FDCs were marked with IC PE. (b) Representative multi-photon image of PE immune complexes marked FDCs and GFP+ IL44get/+ T cells. Images were taken at 910 nm to maximize GFP signal since the ­transferred cells only contained one allele of the 4get reporter, resulting in a very dim signal. The second image shows successful tracking of GFP+ IL44get/+ cells using Imaris software. Scale bar = 50 μm. (c) Quantifications of speed (μm/min) and displacement over time (μm/min) distributions of tracked GFP+ IL44get/+T cells

4. Dissection tools (forceps and scissors). 5. Fetal calf serum (FCS). 6. Phosphate buffered saline (PBS). 7. FACS buffer: (2% FCS in PBS). 8. Cell isolation buffer: (2% FCS in PBS with 1 mM EDTA). 9. Complete Roswell Park Memorial Institute (RPMI) 1640 media: (10% FCS, 100 U/mL penicillin, 100 μg/mL ­streptomycin, and 0.3 mg/mL l-glutamine). 10. ACK lysis buffer: (150 mM NH4Cl, 10 mM KHCO3, 0.1 mM Na2EDTA in water). 11. B-cell negative selection kit and isolation magnets (STEMCELL Technologies). 12. 50 mL conical tubes. 13. 15 mL conical tubes. 14. 5 mL polystyrene tubes. 15. Refrigerated centrifuge.

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16. Violet proliferation dye (VPD, Becton Dickinson) or desired dye for labeling cells. 17. Hank’s Buffered Saline Solution (HBSS) with calcium and magnesium. 18. Trypan blue. 19. Cell counter. 20. 1 mL syringe with 28G needle for tail vein injection. 2.4  In Vivo Labeling of Follicular Dendritic Cells

1. B-Phycoerythrin (PE). 2. Purified Anti-PE antibodies (Rockland). 3. 1 mL syringe with 28G needle for intraperitoneal and ­subcutaneous injections. 4. PBS.

2.5  Imaging of Explanted Lymph Node and Data Analysis

1. 70% ethanol. 2. Forceps and scissors for dissection. 3. 6-well plate. 4. Plastic cover slips cut into small pieces. 5. Dissecting microscope. 6. Vetbond (3 M) glue to adhere lymph node to cover slips. 7. Warmed RPMI (without phenol red). 8. Tank of carbogen (95% O2 + 5% CO2). 9. Peristaltic pump. 10. Vacuum trap. 11. HPLC filter for bubbling gas in media. 12. Tubing (for carbogen and peristaltic pump).

2.6  Multi-photon Microscope and Imaging Software

The microscope we used is a dual IR laser Olympus FV1000MPE multi-photon microscope with the following specifications: 1. Microscope body: Olympus BX61 upright with motorized XY-stage and objective Z-drive. 2. Primary IR laser: Spectra-Physics MaiTai HP DeepSee with pre-chirp compensation. 3. AOM-regulated laser power output capable of ­depth-­dependent modulation. 4. Confocal scanner with two sets of galvanometers scanning mirrors. 5. Four non-descanned photomultiplier tube detectors. 6. 25× high 1.05 NA water immersion Olympus objective. 7. Custom-dedicated fluorescent emission filter cubes.

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8. Vibration isolation air table. 9. Fluoview FV10-ASW software version 3.0. 10. Warner instruments environmentally controlled flow chamber for maintenance of tissue temperature and oxygenation. Warner PH-1 stage, JG-23 chamber, In-line Heater/Cooler SC-20, Dual Channel Bipolar Temperature Controller CL-200A. 11. Imaging software for data analysis (Imaris or Volocity).

3  Methods 3.1  Immunization of Mice

1. On day 0 of the experiment, dilute 100 μg of ovalbumin in PBS to a volume of 20 μL. Add 20 μL of alum and mix the solution for 5 min by pipetting gently up and down (see Note 1). 2. Anesthetize mice using isoflurane in an approved anesthesia machine according to established institutional and IACUC guidelines. 3. Using a 28-gauge needle, inject IL44get mice subcutaneously in the footpad with 40 μL of ovalbumin/alum mixture.

3.2  Preparation of Single-Cell Suspensions for B-Cell Isolation

1. Euthanize C57BL/6 mice according to institutional and IACUC guidelines. 2. Using sterile, aseptic technique, gently remove peripheral lymph nodes (popliteal, inguinal, axillary, brachial, and mesenteric) from the mouse and place them in a 50 mL conical with 2 mL of 2% FCS in PBS. Remove the spleen from the mouse and place in a separate 50 mL conical with 2% FCS. 3. Prepare single-cell suspensions of lymph node and splenic cells. Briefly, pour the lymph nodes and spleen into separate 3 cm petri dishes, and crush them using the back of a plunger from a 3 mL syringe. Add 5 mL of 2% FCS to each petri dish, and filter the lymph node and splenic cells, using an 80 μm mesh, into separate 15 mL conical tubes. Add an additional 5 mL of 2% FCS into each dish and filter into the appropriate tubes. Centrifuge the cells at 400 × g for 5 min to pellet the cells. Remove the supernatant. Dilute the lymph node and splenic cells in 1 mL cell isolation buffer (these cells are ready for selection). 4. Proceed with B-cell-negative selection according to ­manufacturer’s instructions. We used a B-cell-negative selection kit from STEMCELL Technologies. Similar kits are also ­available from Miltenyi and Dynabeads (Thermo Fisher). 5. Count the total number of cells in the negatively selected ­fraction using Trypan blue exclusion and a hemocytometer. 6. Purified B cells will be stained with VPD (Becton Dickinson) in the subsequent step.

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3.3  Stain Purified B Cells with Violet Proliferation Dye

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1. Resuspend previously selected B cells at 1 × 107/mL in HBSS with calcium and magnesium (see Note 2) in a 15 mL conical. 2. Add VPD at 1 μM final concentration and mix well by vortexing. 3. Incubate the cells for 30 min at 37 °C. 4. Add 1× the volume of 100% FCS and incubate for 2 min at room temperature. 5. Wash the cells twice with complete RPMI (cRPMI). Centrifuge the cells for 5 min at 400 × g. Remove the supernatant after each spin. Finally, wash the cells once with PBS, centrifuging again for 5 min at 400 × g. Resuspend cell pellet in PBS such that you have 1.5 × 107 cells per 200 μL. 7. Transfer 1.5 × 107 VPD-stained B cells into IL44get or C57BL/6 mice by intravenous injection into the lateral tail vein.

3.4  In Vivo Labeling of Follicular Dendritic Cells Using Immune-­ Complexed (IC) Phycoerythrin (PE)

3.5  Preparation of Lymph Nodes for Imaging

1. Dilute 2 mg of anti-PE antibody in 300 μL PBS. 2. Inject 2 mg anti-PE antibody intraperitoneally into desired mice using a 28-gauge needle. 3. Dilute 10 μg of PE in 40 μL of PBS and inject subcutaneously into the immunized footpad. Injection of PE should occur approximately 8 h after injection of the anti-PE antibody. 1. Euthanize the recipient mice according to IACUC and institutional guidelines. 2. Extract the popliteal lymph nodes and place into a six well dish filled with 4 mL of room temperature RPMI (no phenol red). Gently remove the lymph node by placing your forceps under it. Do not crush, squeeze, or pull on the lymph node during dissection (see Note 3). 3. Using a dissection microscope, carefully clean the fat off the cortical side of the lymph nodes. Be sure not to damage the lymph nodes or allow the lymph nodes to dry out (see Note 4). 4. Cut a plastic cover slip into small 5 mm square pieces and place a small amount of Vetbond on one of the squares. Remove excess glue, leaving only a residual film on the cover slip. 5. Remove residual media from the lymph node, without allowing it to dry out, by moving the lymph node around a dry petri dish. Quickly place the lymph node cortical side up on the cover slip. The Vetbond will secure the lymph node to the cover slip. Immediately, submerge your cover slip and lymph node face down in RPMI without phenol red to cure the remaining V ­ etbond present on the cover slip and to ensure the lymph node does not dry out.

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3.6  Imaging Explanted Lymph Node with Multi-photon Microscope

1. Turn on the microscope and start the software. 2. Place RPMI without phenol red media in a 37 °C water bath to warm. Turn on the carbogen tank and place the HPLC filter attached to the carbogen tank into the RPMI media. Place the tubing leading from your peristaltic pump into your RPMI as well. Start the peristaltic pump and allow a slow but steady stream of warmed RPMI medium to fill your flow chamber. Fit the vacuum apparatus and tubing on to the flow chamber to prevent media overflow. 3. Turn on the heating apparatus and set it so that the temperature remains between 35.5 °C and 36.5 °C with the objective ­submerged. Place the temperature probe into the media-filled flow chamber. Continue to monitor the temperature ­throughout the duration of the experiment (see Note 5). 4. Secure the lymph node to the stage by turning off your ­peristaltic pump and removing the media from the flow chamber. Put a very small amount of vacuum grease onto the glass portion of the stage and place the coverslip with the lymph node securely onto the grease. Turn the peristaltic pump back on once the lymph node has been secured to the stage. 5. Tune the laser to 870 nm and set the laser power accordingly (see Note 6). 6. Starting at the top of the lymph node, acquire 1.5 × 1.5 mm tiled Z-stack of approximately 300 μm depth. Use this ­composite image of the lymph node to guide you to a field of view c­ ontaining a germinal center to perform more extensive time-lapse imaging. 7. Navigate to your desired location and take a 60 μm Z-stack over a 30-min period with intervals of 30 s. A 3 μm Z-distance will allow all cells to be imaged in at least one z-plane. A minimum XY resolution of 1 μm/pixel will allow cell–cell interaction to be analyzed. 8. Analyze images and videos using your software program of choice (Imaris or Volocity).

3.7  Transfer of IL44get OT-II T Cells for Subsequent Multi-photon Microscopy

1. Euthanize an IL44get OT-II mouse according to IACUC and institutional guidelines. 2. Prepare single-cell suspensions from the spleen and brachial, axillary, and inguinal lymph nodes as described in Subheading 3.2. 3. Proceed with T-cell-negative selection according to manufacturer’s guidelines. We used a kit purchased from ­ STEMCELL, but they can also be obtained from Miltenyi. 4. Count cells using trypan blue exclusion on a hemocytometer. Dilute cells in PBS to a concentration of 5 × 105 cells per 200 μL.

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5. Transfer 5 × 105 cells into a naïve C57BL/6 mouse by ­intravenous injection in the lateral tail vein on day -1. 6. On day 0 immunize mice with 100 μg OVA precipitated in alum as described in Subheading 3.1. 7. On day 6 stain isolated naïve B cells with VPD as described in Subheadings 3.2 and 3.3 and transfer into the immunized recipient. 8. On day 6 mark FDCs using PE immune complex method detailed in Subheading 3.4. 9. Euthanize the recipient mouse on day 7 according to institutional and IACUC guidelines. Extract the popliteal lymph node and prepare for imaging as outlined in Subheading 3.5. 10. Image the explanted popliteal lymph node as described in Subheading 3.6 (see Note 7). 11. Analyze acquired images and videos using your software program of choice (Imaris or Volocity). Track your cells of interest and quantify cell motility statistics such as speed and displacement over time (Fig. 2c) (see Note 8).

4  Notes 1. Ovalbumin must be mixed well with alum prior to ­immunization in order to properly adsorb and precipitate with the alum. Pipetting up and down for 5 or more minutes is highly ­recommended to ensure a strong immunization. 2. It is important to use HBSS that has calcium and magnesium. Using HBSS that is not supplemented with calcium and ­magnesium will result in poor staining and cell recovery with the violet proliferation dye. 3. Disrupting the architecture of the lymph node will cause your cells to stop migrating within the lymph node. Be sure not to crush, squeeze, or tear the lymph node when extracting the lymph node from the mouse, removing excess fat from the lymph node, or fixing the lymph node to the cover slip. 4. When removing fat or manipulating the lymph node, ensure that the lymph node does not dry out. If the lymph node dries out, it can cause reduced cellular motility as well as increased autofluorescence in subcapsular cells, such as macrophages. 5. Physiologic lymphocyte motility peaks between these ­temperatures. Falling outside this temperature range will result in reduced movement of your cells that is independent of your experimental conditions [12]. 6. The laser power and voltages used for each experiment will depend on the microscope and may vary between experiments.

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Typically, it is a good idea to start with a low laser power and high detector power. If the laser power is too high, it could lead to an enhanced rate of photobleaching and may damage your tissue. 7. Since our transferred OT-II T cells only had one 4get allele, we acquired the images shown for our IL44get transfer experiment at 910 nm in order to maximize GFP+ excitation. Reliable imaging of VPD does not occur at 910 nm, and we could not image VPD B cells, PE immune complexes, and GFP ­simultaneously. Use of homozygous IL44get OT-II T cells would allow for better imaging of GFP at 870 nm. Successful ­detection of VPD, PE immune complexes, and GFP can occur at this wavelength. Alternatively, a two IR laser system can allow for simultaneous imaging of these fluorescent labels by exciting at both 810 nm and 910 nm. 8. A large variety of statistics are available to assess cell dynamics and motility. Speed and displacement over time are just examples. Additionally, quantifications of cellular interactions can be ­performed to determine if two cell types interact, how often they may interact, and whether the interactions lead to changes in motility.

Acknowledgments This research was funded in part by NIH grants R01AI125553 (J.J.), R01DK111733 (R.S.F), and R01AI119004 (R.L.R). The content of this work is solely the responsibility of the authors and does not necessarily represent the official views of the NIH. References 1. Pawankar R, Canonica GW, Holgate ST, Lockey RF (2012) Allergic diseases and asthma: a major global health concern. Curr Opin Allergy Clin Immunol 12(1):39–41. https://doi.org/10.1097/ ACI.0b013e32834ec13b 2. Liang HE, Reinhardt RL, Bando JK, Sullivan BM, Ho IC, Locksley RM (2012) Divergent expression patterns of IL-4 and IL-13 define unique functions in allergic immunity. Nat Immunol 13(1):58–66. https://doi. org/10.1038/ni.2182 3. Reinhardt RL, Liang HE, Locksley RM (2009) Cytokine-secreting follicular T cells shape the antibody repertoire. Nat Immunol 10(4): 385–393. https://doi.org/10.1038/ni.1715

4. Wang X, Cho B, Suzuki K, Xu Y, Green JA, An J, Cyster JG (2011) Follicular dendritic cells help establish follicle identity and p ­ romote B cell retention in germinal centers. J Exp Med 208(12):2497–2510. https://doi. org/10.1084/jem.20111449 5. King IL, Mohrs M (2009) IL-4-producing CD4+ T cells in reactive lymph nodes during helminth infection are T follicular helper cells. J Exp Med 206(5):1001–1007. https://doi. org/10.1084/jem.20090313 6. Zaretsky AG, Taylor JJ, King IL, Marshall FA, Mohrs M, Pearce EJ (2009) T follicular helper cells differentiate from Th2 cells in response to helminth antigens. J Exp Med 206(5): 991–999. https://doi.org/10.1084/jem. 20090303

Multi-photon Imaging of IL-4 Expressing Cells in Lymph Nodes 7. Mohrs M, Shinkai K, Mohrs K, Locksley RM (2001) Analysis of type 2 immunity in vivo with a bicistronic IL-4 reporter. Immunity 15(2):303–311 8. Bao K, Carr T, Wu J, Barclay W, Jin J, Ciofani M, Reinhardt RL (2016) BATF modulates the Th2 locus control region and regulates CD4+ T cell fate during antihelminth immunity. J Immunol 197(11):4371–4381. https://doi. org/10.4049/jimmunol.1601371 9. Victora GD, Schwickert TA, Fooksman DR, Kamphorst AO, Meyer-Hermann M, Dustin ML, Nussenzweig MC (2010) Germinal center dynamics revealed by multiphoton microscopy with a photoactivatable fluorescent reporter. Cell 143(4):592–605. https://doi. org/10.1016/j.cell.2010.10.032

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10. Hauser AE, Junt T, Mempel TR, Sneddon MW, Kleinstein SH, Henrickson SE, von Andrian UH, Shlomchik MJ, Haberman AM (2007) Definition of germinal-center B cell migration in vivo reveals predominant intrazonal circulation patterns. Immunity 26(5):655–667. https://doi.org/10.1016/j. immuni.2007.04.008 11. Allen CD, Okada T, Tang HL, Cyster JG (2007) Imaging of germinal center selection events during affinity maturation. Science 315(5811):528–531 12. Miller MJ, Wei SH, Parker I, Cahalan MD (2002) Two-photon imaging of lymphocyte motility and antigen response in intact lymph node. Science 296(5574):1869–1873

Chapter 18 Imaging Precision-Cut Lung Slices to Visualize Leukocyte Localization and Trafficking Miranda R. Lyons-Cohen, Hideki Nakano, Seddon Y. Thomas, and Donald N. Cook Abstract Pulmonary dendritic cells (DCs) are potent antigen-presenting cells that can activate both naïve and memory/effector T cells. However, very little is known of how movements and localization of DCs contribute to these events. To study this, we have developed new procedures that combine precision-cut lung slices with cell staining using fluorescently tagged antibodies to detect individual cell types. In this chapter, we describe these methods in detail and show how they can be used to study the localization of not only DCs but also other leukocytes of interest, as well as structural cells within the lung. Key words Live-cell imaging, Precision-cut lung slices, Lung, Dendritic cells, Macrophages, Antigen-­ presenting cells, Localization, Confocal microscopy, Lymphatics, Epithelial cells, Airway epithelium

1  Introduction Dendritic cells (DCs) span the interface between innate and adaptive immunity and have several unique features that distinguish them from other cell types. They acquire proteins from the extracellular environment, and digest them into peptides that are presented on the cell surface in context of major histocompatibility complex (MHC) class II [1]. Pulmonary DCs, which are located within the airway epithelium, lung parenchyma, and alveolar spaces, acquire inhaled antigens and migrate to regional mediastinal lymph nodes (LNs) to activate CD4+ T cells, thereby initiating adaptive immune responses to those inhaled proteins [2]. Despite the well-established ability of DCs to migrate to regional LNs, not all DCs appear to do this. Indeed, many antigen-bearing DCs remain in the lung for several months after acquiring antigen [3, 4]. It seems likely that these resident DCs can interact with memory/effector T cells that reenter the lung from the circulation, but

R. Lee Reinhardt (ed.), Type 2 Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1799, https://doi.org/10.1007/978-1-4939-7896-0_18, © Springer Science+Business Media, LLC, part of Springer Nature 2018

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it is unclear whether the functions of resident DCs are similar or different than those of DCs that migrate to regional LNs. Flow cytometry is a powerful technique for quantitative analysis of cell surface proteins on a variety of immune cell types, but it provides no information on the precise localization and migration patterns of these cells into, within, and out of the lung. Likewise, in vitro chemotaxis assays can assess the ability of cells to respond to chemotactic factors in vitro, but these assays do not reproduce the complex, three-dimensional environment of the intact lung. Here, we describe how precision-cut lung slices (PCLS) can be generated, stained with fluorescently tagged antibodies, and visualized by confocal microscopy to study the location and movement of individual cell types within the lung. This procedure has multiple advantages, including retention of the three-dimensional architecture of the lung slices, the ability to visualize an area as large as a cross section of an entire lobe, as well as the opportunity to use video microscopy to visualize cell movements and interactions in the lung slices at 37 °C. We have used this protocol to successfully visualize the localization of a wide variety of immune cell types, including different types of dendritic cells, macrophages, neutrophils, T cell and B cells, as well as structural cells such as lymphatic, endothelial, and epithelial cells.

2  Materials 2.1  Lung Preparation

1. Sodium pentobarbital is dissolved and diluted with water: 250 mg/kg body weight. 2. Cork board or polystyrene foam base. 3. 1 mL syringe. 4. 1.5 in., 20-gauge (G) needle. 5. Polyethylene tubing (BD Diagnostic Systems): 0.86 mm inside diameter, 1.27 mm outside diameter. 6. Phosphate buffered saline (PBS): (Mg−, Ca−) pH 7.2–7.4. 7. 2% agarose: 2% low melting point agarose (GeneMate Sieve GQA); warmed to 40 °C. 8. 40 °C water bath. 9. Refrigerator or cold room (4–8 °C).

2.2  PCLS Preparation

1. Compresstome™ VF-300 (Precisionary Instruments) or equivalent instrument with metal cooling block, plunger, and metal syringe. 2. Double-edge stainless razor blade (Electron Microscopy Sciences). 3. Super glue (Krazy Glue): All-purpose, no-run gel.

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4. Forceps. 5. Thin spatula or paintbrush. 6. 24-well plate. 2.3  PCLS Staining

1. PBS containing 2% fetal bovine serum (FBS). 2. Complete RPMI-10 medium (cRPMI-10): RPMI1640 containing 10% FBS and 10 mM Hepes. 3. Normal mouse serum (Jackson ImmunoResearch). 4. Normal rat serum (Jackson ImmunoResearch). 5. Fc blocker: anti-CD16/32 mAb (clone 2.4G2) purified mAb or hybridoma culture supernatant. 6. Antibody combinations designed to detect specific cell types [5]. Dendritic cell panel: CD324/E-cadherin—Alexa Fluor 488 (AF488) (clone DECMA-1) CD88/C5Ra1—Phycoerythrin (PE) (clone 20/70) CD11c/alpha X integrin—Brilliant Violet 605 (BV605) (clone N418) CD103/alpha E integrin—Allophycocyanin (APC) (clone 2E7) Structural cell panel: CD90.2/Thy1.2—APC (clone 53-2.1) CD324/E-cadherin—AF488 (clone DECMA-1) CD31/PECAM-1—PE (clone MEC13.3) Myeloid cell panel: CD324/E-cadherin—AF488 (clone DECMA-1) CD88/C5Ra1—PE (clone 20/70) CD11c/alpha X integrin—BV605 (clone N418) CD172a/Sirp1a—APC (clone P84) 7. Antibody cocktail for static imaging: Antibodies usually work well at 1–5 μg/mL, but optimal concentation should be determined by reseachers. Antibodies added to 1 mL PBS containing 2% FBS, 50 μL normal mouse serum, 50 μL normal rat serum, and Fc blocker (5 μg/mL mAb or 100 μL supernatant). Antibody cocktail for live-cell imaging: 1–5 μg/mL mAbs added to 1 mL cRPMI-10 containing 50 μL normal mouse serum, 50 μL normal rat serum, and Fc blocker (5 μg/mL mAb or 100 μL supernatant).

2.4  Confocal Microscopy of PCLS

1. 35-mm glass-bottom microwell dishes (MatTek Corporation). 2. 12 mm coverslip (Carolina Biological Supply). 3. ProLong Gold Antifade Mountant (Thermo Fisher Scientific).

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4. Confocal microscope: e.g., Zeiss 880 multiphoton laser-­ scanning microscope. 5. Plan-Apochromat 20×/0.8 M27 objective lens (Carl Zeiss). 6. ZEN: image processing software (Carl Zeiss). 2.5  Live-Cell Imaging of PCLS

1. Cold room (4–8 °C). 2. Incubator, 37 °C. 3. Matrigel: growth factor reduced, basement membrane matrix, Phenol Red-free, LDEV-free (Corning). 4. Nunc Lab-Tek chambered cover glass (8 slots) (Thermo Fisher Scientific). 5. Bare platinum wire: wire bent into 5–8 mm long “L” shape (World Precision Instruments).

3  Methods 3.1  Lung Preparation

1. Euthanize mouse by intraperitoneal injection of 5 mg sodium pentobarbital. Pin the animal to a polystyrene foam base. 2. Open the abdominal cavity and cut the inferior vena cava to drain blood away from the lungs. Carefully puncture the diaphragm to allow the rib cage to expand. 3. Remove salivary glands and muscle to expose the upper trachea. Make a small incision on the anterior side of the thickest band of cartilage using fine forceps or scissors, being careful not to cut all the way though the trachea. The incision should be just large enough to allow a 20-G needle to pass though (Fig. 1). 4. Insert a 20-G needle into a section of the polyethylene tubing, leaving approximately 1 cm of tubing beyond the end of the needle. Cut the tubing at an angle of approximately 45° to bevel the end, and attach the needle to a 1-mL syringe. Load the syringe with 0.8 mL of warm (40 °C) 2% agarose, immediately place the end of the tubing into the incision in the trachea, and slowly inject the agarose into the lungs. The lungs should be seen inflating within the chest. Without moving the needle or syringe, tape the syringe to the polystyrene foam base to ensure the needle remains in the trachea (Fig. 1) (see Note 1). 5. Place the mouse, with base and syringe still inserted in the trachea, at 4 °C for at least 10 min to allow the agarose to completely solidify. If necessary, the animal can be kept at 4 °C for up to several hours. 6. Carefully excise the agarose-inflated lungs, and place them in a 35-mm dish with ice-cold PBS containing 2% FBS. Keep on ice. 7. Any lobe of the lung can be used to make lung slices, but we prefer the right superior lobe. Ensure that the interior part of

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Fig. 1 Lung inflation preparation. (a) A 1-mL syringe attached to 20-G needle with beveled polyethylene tubing slid onto the tip has been inserted into the trachea, agarose injected, and the syringe taped in place. (b) Higher magnification image of the hole cut on the anterior side of the thickest band of cartilage that is large enough to insert a 20-G needle. (c) Higher magnification image of fully inflated lungs in the chest cavity

the lung (where it connects to trachea) faces down in the dish. The lobe used and its orientation on the plunger will determine the size of vessels and airways that will be visible. The orientation described here is ideal for visualization of large airways and parenchyma (Fig. 2a) (see Note 2). 3.2  PCLS Preparation

1. To prepare lung slices, use a Compresstome, metal cooling block, plunger and metal syringe, or similar, according to the manufacturer’s instructions. Place a small drop of super glue onto the plunger and spread the glue in a circular motion (Fig. 2b) (see Note 3). 2. Immediately after covering the plunger with super glue, gently grasp the excised lung lobe with forceps with the trachea side down, dab it on a Kimwipe paper to remove excess liquid, and carefully place it on top of the plunger. Trim off any extra tissue extending beyond the edge of the plunger (Fig. 2c). 3. Move the plunger down so that the sides of the metal syringe move up and over the tissue, creating a well with the lung glued at the bottom, no more than several centimeters deep. Tape around the bottom of the metal syringe to hold this position in place (Fig. 2d). 4. Carefully pour 2% agarose (40 °C) into the well until the agarose just covers the top of the lobe.

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Fig. 2 Orientation of lung attachment onto plunger. Diagram of the preferred orientation of the lung glued onto the plunger. (a, b) The lung, resting trachea side down in a dish, is dabbed dry, then placed gently onto the wet super glue on the plunger in the same orientation. (c) The lobe is trimmed as necessary when hanging over the edge of the plunger. (d) The plunger is moved down to create a space to add agarose. Four to six mm from the top of the tissue to the syringe top is optimal

5. Surround the metal syringe with the ice-cold chilling block, and cool the lobe and 2% agarose for 1–2 min. A shorter cooling time than this might cause disintegration of the agarose during slicing, which could lead to unevenly cut PCLS. 6. Load the specimen syringe into the tissue slicer and fill the buffer tank with ice-cold PBS. Align the step motor drive with the specimen syringe and remove the piece of tape from the syringe. Turn the switch to the fast-forward (FF) position until the step motor drive just touches the back of the plunger. Turn off the motor when the drive touches the plunger. 7. Align the fresh blade with the specimen syringe. Set the Compresstome settings to desired slice thickness (120–150 μm, continuous cutting; oscillation, 9; and speed, 3–4). 8. Fill buffer tank with cold PBS, and turn on the power. 9. During slicing, PCLS separate from agarose and fall into the buffer tank. Using a thin spatula or paintbrush, collect the PCLS one at a time as they fall into the buffer tank, and place them in a 24-well plate containing ice-cold PBS (2% FBS). It is important to keep the slices in the order that they are cut to maintain consistency between experiments. Although individual tissue samples are different due to age, gender, and weight, the tenth slice usually yields reproducibly sized airways.

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1. Design a panel of fluorescently tagged antibodies based on the cell types of interest, taking into account the potential for fluorescent spectral overlap and the detection capabilities of the microscope. Fluorescence spectral viewer tools such as FluoroFinder (https://fluorofinder.com) and BioLegend Fluorescence Spectra Analyzer (http://www.biolegend.com) are useful to design panels. 2. Depending on the method, prepare the antibody cocktail for static imaging or live imaging with a desired final concentration, which needs to be optimized for each individual antibody (usually 1–5 μg/mL). Transfer the antibody cocktail solution to a 35-mm dish, and keep on ice in the dark until ready to use. 3. Choose a PCLS having an anatomical area of interest, such as large airways or periphery of the lung, and place in a 35-mm dish containing 1 mL antibody cocktail. 4. Stain in the dark, on ice, rocking slowly for 20–60 min. A shorter incubation time is recommended for live-cell imaging. 5. Remove the antibody solution from the dish, and rinse the slices twice with 1 mL ice-cold PBS containing 2% FBS for static PCLS imaging, or cold cRPMI-10 for live-cell imaging. Proceed to either Subheading 3.4 for confocal microscopy of static PCLS imaging or Subheading 3.5 for live-cell video imaging.

3.4  Confocal Microscopy of PCLS

1. Pipette 100 μL PBS containing 2% FBS onto a 35-mm round glass-bottom dish. Using a spatula or paintbrush, place the stained PCLS onto the drop, gently manipulate the slice until it is flat and spread out. Remove the PBS with a pipette and a Kimwipe paper (Fig. 3a, b). The slice should lie as flat as possible. The above procedures should be done quickly to avoid photobleaching. 2. Place 15 μL of room temperature ProLong Gold antifade mounting medium onto the slice, and gently place a round glass coverslip into the well of the plate (Fig. 3). 3. When viewing under a confocal microscope using ZEN software, use the “tile” tool to locate and mark the edges of the tissue in the “convex hull” setting. 4. While setting the z-stack range, verify at multiple areas of the slice, especially along the edges, that the set ranges are appropriate. The z-range will likely need to be greater than the thickness of the tissue itself because it is difficult to get the tissue to lay completely flat. For example, with a 150-μm-thick PCLS, the z-stack range will need to be set from 150 to 200 μm to accommodate curves in the slice. Depending on the z-stack range and the size of the tissue, each full-lung scan will take

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a

Lung slice

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Pipette tip

PBS drop

Glass well in the center Glass-bottom dish

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Glass cover slip Mounting medium

Lung slice

Fig. 3 Flattening of lung slice for static imaging. (a) PBS (2% FBS, 100 μL) is placed onto a 35-mm glass-­ bottom dish. A lung slice is placed onto the drop using a spatula or paintbrush and gently manipulated until it spreads out. (b) Remove the PBS with a pipette and a Kimwipe paper. The slice should not have any wrinkles or bumps. (c) Place 15 μL mounting medium onto the slice and gently place a round glass coverslip into the well of the plate. The lung slice should be as flat as possible at this point

between 7 and 12 h. Each lung is unique and the settings must be adjusted for every experiment. 3.5  Live-Cell Imaging of PCLS

1. Pipette 50 μL cRPMI-10 into one slot of a Lab-Tek chambered cover glass. Use a spatula to place the PCLS onto the medium, gently manipulating it until the slice is flat and spread out. Remove the 50 μL medium using a pipette and a Kimwipe paper (Fig. 3a, b). It is acceptable if the edges of the slice are laying vertically against the wall of the chamber. 2. Using forceps, carefully drop “L”-shaped platinum wire weights on top of the slice. We recommend using two weights that form a square with the four sides of the two “L” -shaped wires to be used as a viewing window under the microscope (Fig. 4a, b) 3. In a cold room, mix 150 μL of Matrigel with 50 μL cRPMI­10 in a microtube. Use pre-cooled pipette tips for this procedure, or the Matrigel will solidify in the tip. This ratio of Matrigel and medium (3:1) creates a gel dense enough to hold the PCLS down and maintains cell viability. 4. In the cold room, gently pipette 100 μL Matrigel-medium mixture on top of the PCLS, making sure that the gel goes on top of the lung slice and that the slice does not float up on top of the gel. The gel should work as a weight, anchoring the PCLS down onto the bottom of the chamber (Fig. 4b, c). 5. Carefully transfer the chambered cover glass to a 37 °C incubator and let the Matrigel-medium solidify for ~10 min.

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L-shape platinum wire

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b

Matrigel-medium mixture Medium

Chambered coverglass bottom

c

Matrigel-medium ; mixture

Platinum wire

Chambered coverglass

Lung slice

Fig. 4 Flattening of lung slice for live-cell imaging. (a) After placing a PCLS on a drop of cRPMI-10 in a chambered cover glass, the medium is removed with a pipette and a Kimwipe paper as depicted in Fig. 3a, b. Two L-shaped platinum wire weights are carefully placed on top of the slice to form a square imaging window. (b) Matrigel-medium mixture is carefully pipetted directly on top of the lung slice. (c) Final PCLS set-up for live-cell imaging. The Matrigel and wire weights are anchoring the PCLS down, and the slice should lie flat at this point. Once solidified, the Matrigel holds PCLS down throughout imaging

6. Transfer the entire chambered cover glass onto the pre-warmed stage (37 °C, 7.5% CO2) of a confocal microscope. Sample should be maintained at 37 °C and supplied with 7.5% CO2 throughout imaging. 7. Set the z-stack range and tile range based on the desired interval between frames. 1 × 1 tile and 30 z-stacks will generally result in 1 frame/2.5 min. Ensure the autofocus function is activated before starting the experiment to prevent x–y and z-drift.

4  Notes 1. After injection, the lungs will be larger and inflated within the chest. If only one lobe expands, the needle has been inserted too deeply (past the bronchus), and it will be necessary to pull it out slightly before proceeding. This part of the procedure needs to be done relatively quickly, before the agarose starts to solidify.

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2. Mice have one large left lobe and four smaller lobes on the right. For PCLS imaging experiments that involve introduction of allergens into the lungs by oropharyngeal or intranasal aspiration, we use the right superior lobe due to a consistent and dispersed exposure to inhaled reagents. However, other lobes also can be studied as long as the same lobe is used when comparing strains or treatments. 3. Super glue should not touch the sides of the metal syringe. If the super glue touches the sides of the syringe, it will glue the syringe and plunger together.

Acknowledgments We thank Michael Sanderson, Jun Chen, and Keiko Nakano for their advice and help with preparing lung slices and Jeff Tucker and Erica Scappini for microscopic analysis. This work was supported by the Intramural Research Program of the National Institutes of Health and the National Institute of Environmental Health Sciences. References nodes. J Exp Med 205(12):2839–2850. 1. Lambrecht BN, Hammad H (2009) Biology of https://doi.org/10.1084/jem.20081430 lung dendritic cells at the origin of asthma. Immunity 31(3):412–424. https://doi. 4. Julia V, Hessel EM, Malherbe L, Glaichenhaus org/10.1016/j.immuni.2009.08.008 N, O'Garra A, Coffman RL (2002) A restricted subset of dendritic cells captures airborne anti 2. Steinman RM (2007) Lasker Basic Medical gens and remains able to activate specific T cells Research Award. Dendritic cells: versatile conlong after antigen exposure. Immunity trollers of the immune system. Nat Med 16(2):271–283 13(10):1155–1159. https://doi. org/10.1038/nm1643 5. Lyons-Cohen MR, Thomas SY, Cook DN, Nakano H (2017) Precision-cut mouse lung 3. Jakubzick C, Bogunovic M, Bonito AJ, Kuan slices to visualize live pulmonary dendritic cells. EL, Merad M, Randolph GJ (2008) Lymph-­ J Vis Exp (122):e55465. https://doi. migrating, tissue-derived dendritic cells are org/10.3791/55465 minor constituents within steady-state lymph

Chapter 19 Study of IgE-Producing B Cells Using the Verigem Fluorescent Reporter Mouse Zhiyong Yang, James B. Jung, and Christopher D. C. Allen Abstract Immunoglobulin E (IgE) is the least abundant antibody isotype in mammalians, yet it plays a critical role in allergy and asthma. IgE-producing (IgE+) B cells are rare and difficult to detect, which have hindered progress to understand their generation and differentiation. Recently developed new fluorescent IgE reporter mice have enabled better understanding of the biology of IgE+ B cells. We here describe the usage of the Verigem IgE reporter mouse to study IgE+ B cells and plasma cells by flow cytometry and microscopy. Key words IgE, B cells, IgE reporter, Verigem, Flow cytometry, Microscopy, Imaging, Germinal center, Plasma cells

1  Introduction Immunoglobulin, or antibody, is an important component of humoral immunity. Mammals produce five major isotypes of antibodies – IgM, IgD, IgG, IgE, and IgA. Among these isotypes, IgE is the least abundant isotype, ranging from undetectable to micrograms per milliliter in the serum [1]. Despite its low abundance, IgE plays a critical role in allergy and asthma [2, 3]. Understanding how IgE-producing (IgE+) B cells are produced and differentiated under normal and pathophysiological conditions is crucial for developing new therapies for IgE-mediated allergic diseases. Upon activation by binding of their surface B cell receptors (BCR) to cognate antigens, naïve B cells may undergo class-switch recombination from IgM/IgD to other isotypes of BCR. Some of these activated B cells may differentiate into short-lived plasma cells (PCs) extrafollicularly [4]. Alternatively, activated B cells can form or enter germinal centers (GCs), where class-switch recombination, affinity maturation, and differentiation into memory B cells or long-lived PCs take place [5]. Due to the technical difficulties to directly study IgE+ B cells, further explained below, it has been R. Lee Reinhardt (ed.), Type 2 Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1799, https://doi.org/10.1007/978-1-4939-7896-0_19, © Springer Science+Business Media, LLC, part of Springer Nature 2018

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unclear whether IgE+ B cells follow the same route of activation and differentiation as B cells expressing other BCR isotypes or display unique characteristics in these processes. As IgE is the least abundant antibody isotype, the IgE+ PCs that produce these antibodies, and their precursor IgE+ B cells, are accordingly very rare. An even more confounding fact is that in addition to IgE+ B cells, several types of immune cells are also coated with IgE. For example, some B cells serve as reservoir for IgE through their surface low-affinity IgE receptor CD23 [6]. Basophils, mast cells, and dendritic cells in humans also absorb IgE through high-affinity IgE receptor, FcεRI, on their surface [7]. Thus it is almost impossible to identify genuine IgE+ B cells by conventional direct surface staining with anti-IgE antibodies. To overcome these challenges, special staining methods for flow cytometry analysis have been applied or developed (reviewed in [8]). For example, an acid wash method was employed to strip IgE noncovalently bound to CD23 [9, 10]. Two techniques have been developed recently to stain intracellular IgE, which is exclusively present in IgE-producing B cells [11, 12]. Around the same time, at least three reporter mouse strains were generated for tracking IgE+ B cells with fluorescent protein reporters (reviewed in [8]). Two of the reporter strains, M1′/GFP and CεGFP, placed the IRES-GFP cassette 3′ of the last exon of membrane IgE, M2 [13, 14]. The third IgE reporter mouse strain, Verigem, was generated in our laboratory [12]. In the Verigem reporter, a T2A-Venus cassette is placed immediately before the stop codon of the IgE M2 exon (see Fig. 1). T2A is a type of 2A peptide that allows separation of two or more proteins encoded in a single mRNA into individual proteins during the process of translation [15], and Venus is a brighter variant of yellow fluorescent protein [16]. Therefore, all B cells expressing the membrane IgE BCR in homozygous Verigem reporter mice will be labeled by Venus. The abovementioned IgE reporters can be used for flow cytometric characterization of IgE+ B cells by directly tracking the fluorescent reporters [12–14]. Furthermore, the fluorescent reporters can also be used as a readout of IgE+ B cells in immunohistochemistry and immunofluorescence imaging, bypassing the interference of other immune cells decorated with surface IgE [12, 13]. A unique advantage of these IgE reporters is that they can be used for live cell imaging in intact tissues, making the monitoring of interaction of IgE+ B cells and other cells possible [12, 13]. In this chapter, we describe detailed protocols for using of the Verigem reporter to track IgE+ B cells by flow cytometry and microscopy. These protocols may also be suitable for the analysis of the other reporter mice.

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Fig. 1 Diagram of the genomic allele, mRNA, and protein of the Verigem reporter. The relative location of exons and polyadenylation sites (pA) of the IgE genomic locus are shown (a). The coding sequence of T2A-Venus is fused in frame to the coding sequence of the membrane IgE M2 exon. Also shown are one FRT site placed immediately 3′ of the coding sequence of Cε4s exon (the last exon for secreted IgE) and one loxP site placed immediately 3′ of the Venus coding sequence. The coding sequences for membrane IgE, T2A, and Venus will be transcribed and spliced into single mRNA molecules (b), but the IgE BCR and Venus will be separated into individual proteins during translation at the T2A site (c)

2  Materials 2.1  Materials for Cell Culture, Immunization, and Flow Cytometry Analyses

1. 5–3/4″ Pasteur pipettes. 2. 100 mm diameter, 60 mm diameter Petri dishes. 3. Complete DMEM medium (cDMEM): DMEM high glucose medium with 10% fetal bovine serum (FBS), 10 mM HEPES, 1× penicillin/streptomycin/l-glutamine. 4. DNase I (10 mg/mL). 5. Complete RPMI growth medium (cRPMI): RPMI 1640 medium with 10% FBS, 10  mM HEPES, 1× penicillin/ streptomycin/l-glutamine, 50 μM β-mercaptoethanol. 6. Recombinant murine interleukin-4 (IL-4). 7. Anti-CD40 (FGK-45, Miltenyi Biotec, 2 mg/mL). 8. U-bottom 96-well tissue culture plate. 9. U-bottom 96-well assay plate. 10. FACS buffer: 1× PBS, 2% FBS, 1 mM EDTA, 0.1% sodium azide (omit sodium azide for CD138 staining [17]). 11. TruStain fcX (anti-mouse CD16/32, clone 93) antibody (BioLegend). 12. Peanut agglutinin (PNA), biotinylated. 13. A fluorescent streptavidin conjugate such as Qdot 605 streptavidin conjugate.

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14. Fixable Viability Dye eFluor 780 (Thermo Fisher). 15. Propidium iodide (PI) in H2O (1 mg/mL). 16. DAPI (4′,6-diamidino-2-phenylindole) in H2O (1 mM). 17. Alum adjuvant (Alhydrogel, Accurate Chemical and Scientific). 18. 4-hydroxy-3-nitrophenylacetyl conjugated to chicken gamma globulin (NP-CGG, ratio 30–39, Biosearch Technologies). 19. Tuberculin syringe with permanently attached needle, 1/2 cc, 27G 1/2. 20. Swinging bucket centrifuge that can accommodate 15/50 mL tubes and plates. 21. Tissue culture incubator. 22. Flow cytometer. 2.2  Materials for Preparation and Microscopic Imaging of Tissue Sections

1. 1× phosphate buffered saline (1× PBS). 2. 4% w/v paraformaldehyde (PFA) in 1× PBS. The PFA must be methanol-free; typically this is prepared from pure powder or from sealed ampules (such as from Electron Microscopy Sciences). To avoid degradation, 4% PFA/PBS should be frozen in aliquots at ≤−20 °C and thawed freshly for each use. 3. 30% w/v sucrose in 1× PBS. 4. 1000 mL pipette tip box lid. 5. Dry ice pellets. 6. Ethanol, 200 proof. 7. Optimal cutting temperature (O.C.T.) compound. 8. Cryomold 10 × 10 × 5 mm (biopsy size). 9. Cryostat. 10. Good quality microtome blades (Tissue-Tek Accu-Edge High Profile Microtome Blades). 11. Superfrost Plus Microscope Slides or other treated slides optimized for tissue adherence. 12. Acetone. 13. Humidity chamber for slide staining. 14. Glass container with rack to hold slides (we typically use containers that hold 300–500 mL volume). 15. Section staining buffer: 1× PBS, 1% normal mouse serum, and 0.1% bovine serum albumin. However, if rat anti-mouse IgG1 antibody will be used, normal mouse serum should be ­substituted with normal rat serum, since the former contains free mouse IgG1. 16. Kimwipes tissue paper or equivalent. 17. Fluoromount-G Scientific).

mounting

medium

(Thermo

Fisher

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18. Slip-Rite coverslips, 24 × 55. 19. Rabbit anti-GFP tag polyclonal antibody (Thermo Fisher Scientific). 20. Additional staining antibodies (see Notes 10, 12). 21. Clear nail polish.

3  Methods Flow cytometry can be used for characterization of IgE+ B cells generated in vitro or in vivo. Some general notes are listed in Notes section (see Note 1). 3.1  Flow Cytometric Analysis of IgE+ B Cells Generated by In Vitro Cell Culture

1. Euthanize a Verigem reporter mouse, and harvest the spleen into a 15 mL conical tube with 5 mL cDMEM medium on ice. Mash the spleen in medium on a 40  μm cell strainer in a 100 mm diameter Petri dish using the bulb of a 3 cc syringe. Pipette the cell suspension three times through a 5–3/4″ Pasteur pipette to break up tissue particles. Then pass the cell suspension through the cell strainer again to remove unbroken cell aggregates. Centrifuge the cell suspension at 400 × g-force at 4 °C for 5 min. 2. Remove the supernatant and add 10 μL DNase I to the cell pellet, and then resuspend the cell pellet in 1 mL cDMEM by pipetting slowly and carefully using a P1000 pipette. 3. Count the cells. The splenocyte suspension can be used directly for cell culture or can be used for B cell purification before setting up the culture (see Note 2). Adjust the final cell concentration to 1 million cells per mL in cRPMI. 4. Add 100 μL cRPMI with 2× final concentration of IL-4 and anti-CD40 to each well of a U-bottom 96-well plate (see Note 3). Then add 100 μL cRPMI containing 100,000 splenocytes or purified B cells.  5. Place the culture in a humidified tissue culture incubator at 37 °C with 5% CO2. 6. After culture for desired amount of time (see Note 4), centrifuge the 96-well plate with cultured cells at 700 × g-force at 4 °C for 2 min. 7. Dump the culture media into a sink by thrusting the plate downward quickly while holding the plate firmly. Hold the plate horizontally, and tap against your palm gently until the cell pellet is fully resuspended in the residual liquid. To block non-specific binding of antibodies to Fc receptors, add 25 μL FACS buffer with 1:50 diluted TruStain fcX™ to each well. Incubate on ice for 10 min.

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8. Add 200 μL FACS buffer to each well. Centrifuge the 96-well plate at 700 × g-force at 4 °C for 2 min. Dump the supernatant  and resuspend the cell pellet as in step 7. Add 25  μL FACS buffer with diluted antibodies, such as anti-IgM, antiIgD, anti-B220, anti-CD138, anti-IgG1, and anti-IgE (see Note 4). Incubate on ice for 20–60 min. 9. Add 200 μL FACS buffer to each well. Centrifuge the 96-well plate at 700 × g-force at 4 °C for 2 min, and then dump the supernatant. Wash the cell pellet again with 200 μL FACS buffer/well. 10. Resuspend the cell pellets in 50 μL FACS buffer with 1:3500 DAPI or 1:2000 PI. 11. Load the samples for flow cytometer analysis (Fig. 2a, b; see Note 5). 3.2  Flow Cytometric Analysis of IgE+ B Cells Generated In Vivo (See Note 6)

Normally IgE+ B cells are nearly undetectable in naïve mice maintained in a clean-specific pathogen-free facility. But these cells can be induced either by immunization with an antigen or by infection with a parasite. Here we describe using NP-CGG immunization as a model to induce IgE+ B cells in vivo. 1. Dissolve lyophilized NP-CGG in sterile water to the concentration of 2.5  mg/mL.  The reagent may be sonicated for 5 min to facilitate dissolving it. The NP-CGG solution can be aliquoted and frozen for future use. 2. Prepare NP-CGG and alum mixture for immunization. For injection at each site, mix 5 μL NP-CGG, 1.5 μL 10× PBS, and 8.5 μL water. Then add 15 μL alum. Mix by tapping or inverting the container. 3. Inject 30 μL NP-CGG:alum mixture subcutaneously using a tuberculin syringe to each site. 4. After immunization for desired amount of time (see Note 7), sacrifice immunized mice according to approved institutional protocol. Collect draining lymph nodes (LNs) of each mouse into a 1.5 mL Eppendorf tube with 1.2 mL cDMEM. Store samples on ice until all the samples have been collected. 5. Mash the draining LNs from each animal on a 70  μm cell strainer in a 60 mm Petri dish using the bulb of a 1 cc syringe. Pipette the cell suspension two times through a 5–3/4″ ­Pasteur pipette to break tissue particles. Then pass the cell suspension through the cell strainer twice to remove unbroken cell aggregates. Measure cell concentration of each sample. 6. Add 3 × 106 cells to each well of a round bottom 96-well plate. If necessary, add FACS buffer to bring the volume of each sample to about 200 μL. Centrifuge the plate at 700 × g-force

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Fig. 2 Flow cytometric characterization of IgE+ B cells generated in vitro or in vivo from Verigem reporter and control mice. (a, b) B cells from WT control or homozygous Verigem mice were cultured in vitro with anti-CD40 (250 ng/mL) and murine IL-4 (25 ng/mL) for 4 days. (c, d) Cell suspensions from draining LNs of WT control or homozygous Verigem mice 7  days after subcutaneous immunization with NP-CGG in alum adjuvant were stained and analyzed by flow cytometry. Cells in panels a and c were gated on live singlets; cells in panel b were gated on live singlets that are IgD−, IgM−, and CD19+. Cells in panel d were pooled from the draining LNs of five homozygous Verigem mice and were pre-gated as GC B cells (live singlets that are CD138−B220highCD 19+CD38lowGL7highIgDlow), and PCs (live singlets that are CD138+B220−CD38lowIgDlow) were further gated according to their expression of Venus and surface IgG1 (panel d, left), and histograms of surface IgE staining on these gated populations are further plotted (panel d, right)

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at 4 °C for 2 min. Dump the supernatant, and then resuspend cell pellet of each sample in 50  μL FACS buffer with 1:50 diluted TruStain fcX. If desired, to identify GC B cells, PNA biotin (diluted 1:2000) can be added at the same time as the TruStain fcX, since the PNA reagent is a lectin and not an antibody. Alternatively, a biotin-conjugated antibody can be added directly to the samples after incubation with TruStain fcX for 10 min, allowing time for the Fc receptors to become blocked. Incubate on ice for 20–60 min. 7. Add 200 μL FACS buffer to each well. Centrifuge the 96-well plate at 700 × g-force at 4 °C for 2 min, and then dump the supernatant. Wash the cell pellet again with 200 μL FACS buffer/well. 8. Resuspend cell pellet of each sample in 50 μL FACS buffer, with a diluted fluorescent conjugate of streptavidin (such as 1:400 Qdot 605 streptavidin conjugate) and other appropriately diluted antibodies for cell surface staining. If desired, 1:500 diluted eFluor 780 viability dye may be included in the antibody mixture to label dead cells (we now routinely use this reagent to exclude nonviable cells even for surface staining and have found it can be added in our normal FACS buffer at the same time as antibodies). Incubate on ice for 20–60 min. 9. Add 200 μL FACS buffer to each well. Centrifuge the 96-well plate at 700 ×g-force at 4 °C for 2 min, and then dump the supernatant. Wash the cell pellet again with 200 μL FACS buffer/well. 10. Resuspend the cell pellet in 25 μL FACS buffer. If no fixable viability dye was included in step 8, then DAPI or PI should be included at this step (see Note 1). Combine staining replicates in one tube. 11. Load the samples for flow cytometer analysis (Fig. 2c, d). 3.3  Staining of IgE+ B Cells in LN Cryostat Sections 3.3.1  Preparation of Whole LNs (See Note 8)

1. Remove LN(s) and place in a 1.5–5.0 mL tube containing 4% PFA/PBS, pre-chilled to 4 °C. 2. Rotate the tubes on a rotator at 4 °C for 2 h. 3. Remove LNs from PFA and wash three times with 1× PBS.  Allow LNs to soak in PBS for at least 5  min for each wash. 4. Transfer washed LNs to new tubes containing 30% sucrose/ PBS solution pre-chilled to 4 °C. 5. Rotate the tubes on a rotator at 4 °C overnight. 6. Transfer the LNs to a cryomold and fill with O.C.T. compound, avoiding bubbles. 7. Freeze the tissue rapidly. We prepare a dry ice pellet and ethanol bath in the lid of a 1000 mL pipette tip box. The ­cryomold

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should be held or positioned such that it is floating in the bath during the freezing process (avoid letting the ethanol reach the unfrozen tissue/OCT). Once frozen, you may leave the block temporarily on dry ice while preparing other samples and then absorb excess ethanol with a Kimwipes and transfer to a −80 °C freezer for long-term storage. 3.3.2  Cryostat Sectioning of Frozen LNs

1. Section frozen specimen in a cryostat with the chamber temperature set at −22 to −20 °C. If the system has the ability to separately adjust the object temperature, set this to about 3 °C warmer than the chamber temperature, which will assist with capturing the sections (we typically use an object temperature of −18 °C and a chamber temperature of −21 °C). The sections should be 5–10  μm thick (we typically cut 7  μm sections). 2. Mount LN sections on microscope glass slides (can typically mount ten or more sections on a single slide); allow to air-dry for at least 1 h. 3. Place slides with mounted tissue in a glass chamber with pre-­ chilled 4 °C acetone for 10 min. 4. After removing from acetone, allow slides to air-dry for at least 1  h before staining. Slides may alternatively be stored at −80  °C with desiccant (avoid getting the tissue wet). Best results are obtained by directly proceeding to the staining steps.

3.3.3  Staining Sectioned Tissue

1. Use a scriber to etch the glass around a region containing all the tissue on each slide (see Note 9). 2. Hydrate the tissue by placing glass slides with tissue in a glass chamber with 1× PBS for at least 15 min. 3. In the meantime, prepare primary antibody master mixture (see Note 10) containing rabbit anti-GFP antibody at a dilution of 1:400 (see Note 8) in section staining buffer. Each slide will need 350 μL of section staining buffer. 4. Remove each glass slide one at a time from the glass chamber containing PBS, run the edge of the slide along the glass chamber to remove excess fluid and dry the back of the slide on a paper towel, quickly dry a region around the tissue with a piece of folded Kimwipes (see Note 11), place the slide in humidity chamber, and quickly add the primary stain solution (it is important for the tissue not to dry out during this process). 5. Stain for 2–3 h at room temperature. 6. Prepare the secondary antibody mixtures (see Notes 10 and 12) in section staining buffer.

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7. Flick off the staining solution onto a paper towel, and wash each slide by placing it on a rack in a glass chamber with an excess of PBS.  Allow at least 3  min to wash each slide (see Note 13). 8. Remove one slide at a time from the PBS wash container, run the edge of the slide along the glass chamber to remove excess fluid and dry the back of the slide on a paper towel, quickly dry a region around the tissue, place slide in humidity chamber, and quickly add the secondary stain solution (again, it is imperative that the tissue does not dry out during this process). Repeat for additional slides. 9. Stain for 2–3 h at room temperature in the dark (covering the humidity chamber with foil is adequate for light protection). 10. Flick off the staining solution and wash slides as described in step 7. 11. Remove one slide at a time from the PBS wash container, run the edge of the slide along the glass chamber to remove excess fluid and dry the back of the slide on a paper towel, quickly dry a region around the tissue, and then add mounting media and coverslip carefully avoiding bubbles. (You may store the slides in a box at 4 °C in the dark overnight if needed). Prior to imaging, let the mounting media begin to solidify by leaving the slides exposed to air at room temperature, protected from light, for 1–2 h. You may seal the coverslips using clear nail polish. Proceed to step 12 or store the slides in a box at 4 °C in the dark until ready to image. Best results are obtained when imaging within 24–48 h after staining. 12. Collect images on a suitable microscope. Care must be taken in the adjustment of exposure times and display levels (see Fig. 3). 3.4  Live Microscopy of the Verigem Reporter

The Verigem reporter mice may also be used for live imaging. Two-photon laser scanning microscopy is the preferred method because this enables visualization deep within living tissues with minimal phototoxicity. In live imaging, since the tissue is not fixed, Venus fluorescence is detectable in both IgE+ GC B cells (Venuslo) and in IgE+ PCs (Venushi). Since detailed protocols have been provided elsewhere for the preparation of explanted tissues [18] and intravital two-photon microscopy [19], we simply provide the following advice for visualization of the Verigem reporter by two-­ photon microscopy. 1. A sensitive detector, such as a gallium arsenide phosphide (GaAsP) detector, will facilitate the detection of IgE+ GC B cells with weak Venus fluorescence. A rather high detector voltage may be needed to visualize weak Venus fluorescence, and this voltage may result

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Fig. 3 Representative microscopy of the Verigem reporter in LN  cryostat sections. Heterozygous Verigem reporter mice were immunized subcutaneously with NP-KLH in alum adjuvant, and LNs were collected 7 d later. (a) Example of a GC (IgDlow region) with IgE+ B cells detected by anti-GFP staining (red). (b) Demonstration of the difference in sensitivity between anti-GFP staining and Venus fluorescence and the importance of the display adjustments to detect IgE+ PCs (Venushigh) versus IgE+ GC B cells (Venuslow). All four panels show the same image with either anti-GFP staining (red, upper panels) or with Venus fluorescence (green, lower panels). The image display levels are adjusted in a standard fashion to show the full range of signal detected (left panels) versus an exaggerated narrow range making the red/green signal appear bright (right panels). Note the GC B cells (e.g., see the yellow arrowhead) can only be detected by anti-GFP staining on the right panels with the bright adjustment of display levels. While PCs can be directly detected by either Venus fluorescence or anti-GFP staining (inset c) with normal display levels, the GC B cells cannot be visualized by Venus fluorescence but can be visualized by the anti-GFP staining with the bright adjustment of display levels (inset d). Within the GC, the bright display adjustment of the Venus channel instead reveals autofluorescent cells, such as tingible body macrophages (e.g., see the white arrowhead in (b, lower right panel)). The bright adjustment of display levels also results in oversaturation of the PCs; thus it may be difficult to properly visualize PCs and GC B cells with the same display settings. Anti-GFP was detected in the far red channel (see Note 10)

in oversaturation of IgE+ PCs with bright Venus fluorescence. Given the pronounced green/yellow autofluorescence of some cell types, it is recommended to include a Verigem-negative control mouse tissue in the experiment to validate that the observed signal originates from Venus fluorescence.

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2. The use of a 525/50 filter with high efficiency transmission characteristics will maximize signal to noise. This filter would also detect green (e.g., GFP) signal. To distinguish GFP from Venus, narrow band filters can be used, such as 510/20 for GFP and 535/30 for Venus. However, note that the use of narrow band filters will greatly reduce the Venus signal. 3. We have determined Venus has two distinct two-photon excitation peaks near 940 nm and 1020 nm (unpublished observations). In our experience, the 1020 nm peak gives the strongest excitation on systems with suitable optical properties. However, we have observed that some pre-compensation (“pre-chirp”) devices, found on many two-photon laser setups, may interfere with laser excitation in the 1020 nm range. Some microscope optics, such as the objective, may also lose transmission efficiency above 1000  nm. We routinely use a Coherent Ultra II Ti:Sapphire laser without a pre-compensation device and an objective optimized for transmission up to 1300 nm, and we consistently observe the best excitation at 1020  nm. Overall, the two-photon excitation of Venus can occur over a broad spectrum between 890 nm and 1040 nm, and thus other wavelengths may be preferred for optimal coexcitation of other fluorophores. Another important consideration is that second harmonic generation, such as from collagen in the lymph node capsule, will appear at half the excitation wavelength. This means that excitation at 1020 nm will result in second harmonic generation at 510 nm, which will appear in the typical green channel (typically the same channel as the Venus signal), which may be undesirable in some cases.

4  Notes 1. General notes for flow cytometry analysis: (1) Tissues should be collected in media pre-chilled to 4 °C and stored on ice. Samples should be kept on ice throughout the staining procedure and before flow cytometric analysis, and centrifuged at 4  °C. (2) Dead cells in the samples should be excluded for further analysis. Dead cells can be identified by washable viability dyes such as Fixable Viability Dye eFluor 780 or by DNA-binding dyes DAPI or PI. The eFluor 780 dye can be included during the last cell surface staining step (this reagent can be added together with other antibodies to surface markers), whereas DAPI or PI should be added after the c­ ompletion of staining procedures. (3) The information of reagents used for flow cytometry staining can be found in our previous publication [20]. (4) The surface membrane IgE expression on

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IgE+ B cells from Verigem reporter mice is about two- to threefold higher than normal. This could be due to the insertion of an FRT site and/or T2A-Venus coding sequence into the IgE genomic locus. Although our previous study found little difference between wild-type (WT) IgE+ B cells and Verigem IgE+ B cells in terms of their differentiation and kinetics [12], readers are still advised to keep in mind that Verigem IgE+ cells have a higher level of membrane IgE than WT IgE+ cells. 2. We have found that comparable numbers of IgE+ B cells can be induced from purified B cells or from total splenocytes [12], and IgE+ B cells generated either way demonstrated the same propensity for antigen-independent PC differentiation [20]. Thus, efforts in purifying B cells can be saved if the in vitro culture is only going to be analyzed by flow cytometry. However, if the effect of other immune cells could potentially interfere with your assay, purified B cells should be used for culture. 3. IgE+ B cells can be generated from naïve B cells activated with LPS and/or anti-CD40 together with IL-4 in in vitro culture. We have found that anti-CD40 plus IL-4 is more potent than LPS plus IL-4 in inducing IgE+ B cells. Additionally, LPS from different batches can vary dramatically in inducing B cell class switching and differentiation (unpublished observations). The generation and differentiation of IgE+ B cells is relatively insensitive to the amount of IL-4 (ranging from 10 to 100 ng/ mL) but is sensitive to the amount of anti-CD40 (≥62.5 ng/ mL) [20]. Therefore the concentration of anti-CD40 should be tested empirically under different experimental settings. 4. IgE+ and IgG1+ cells will start to appear when naïve B cells have been cultured for 3 days, and peak numbers are observed when cells have been cultured for 4 days. We routinely stain in  vitro cultured cells with anti-IgM, anti-IgD, anti-B220, anti-CD138, anti-IgG1, and anti-IgE. Anti-B220 staining can help gate out B220− cells of non-B cell lineages. Some of the cultured B cells will differentiate into PCs, and CD138 is a marker for these cells (PCs will be stained more weakly for B220 but will still appear positive compared to non-B cells). Note that a recent study reported that sodium azide negatively affects CD138 staining and thus sodium azide should be omitted from buffers for optimal staining of this marker [17]. Staining with anti-IgE antibody for cells expressing Venus is optional but can help to corroborate the specificity of the Verigem reporter. Note that in step 3.1.8, the initial washing step after Fc block, prior to antibody staining, is optional, but is included in the protocol to minimize the staining volume and thus reduce the amount of antibodies needed. 

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5. Contrary to mouse B cells expressing other isotypes of BCR, IgE+ PCs express higher surface IgE BCR than IgE+ B cells. Accordingly, the IgE+ PCs carrying the Verigem reporter express much higher Venus than IgE+ B cells. The resultant fluorescence of Venus in IgE+ PCs B cells is so strong that it can exceed the detection range of flow cytometry with standard detector settings. This is true for IgE+ PCs generated in  vitro and in  vivo. Thus it may be necessary to lower the voltage of the “green channel” by 5–10% to make sure that the Venus fluorescence falls within the range of detection of the cytometer. 6. Examples of flow cytometry of B cells from in  vitro culture and cells from draining LN are presented in Fig. 2 to make the following points. (1) Control animals should be included for in  vivo experiments. Although expression of Venus in the Verigem reporter correlates tightly with IgE BCR expression, there are several types of cells, such as eosinophils, that have autofluorescence in the green channel. While there are almost no cells that appear to be Venus+ from the WT B cells cultured in  vitro (Fig.  2a), autofluorescent cells are readily detected from the analysis of tissue samples. For example, immunized WT control and Verigem reporter mice have a similar percentage of IgDhigh cells that appear to be Venus+ (Fig. 2c) in lymph nodes. However, only the sample from the Verigem reporter mouse has Venus+ cells in IgDlow population. Therefore Venus fluorescence alone is not sufficient for specifically identifying IgE+ B cells. Inclusion of non-Verigem control animals in the experiment and staining of additional surface markers [12] will render more confidence in identifying genuine IgE+ B cells. (2) It is advisable to test the FACS settings for detecting the Verigem reporter with cultured Verigem cells before using the mice for in  vivo studies. Under optimal in  vitro culture conditions, the abundance of IgE+ cells can be comparable to that of IgG1+ cells (Fig.  2b). However, even at the peak of in vivo response, the number of IgE+ B cells is usually about 100-fold less than that of IgG1+ B cells (Fig. 2d). Therefore the technical conditions for assessing the Verigem reporter can be most easily tested through in vitro cell cultures to generate a large frequency of IgE+ B cells. (3) Flow cytometry analysis of in vivo IgE response should start with sufficient amount of cells. Since IgE+ cells are still rare even at the peak of responses to immunization or parasite infection, we usually start with four wells of cells from each sample for one staining scheme, with three million cells/well. These wells are then pooled for flow cytometric analysis. About 30–50% of cells will be retained at the completion of staining. As an example, in Fig. 2d, out of about 22 million events pooled from 5 Verigem reporter

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mice 7 days after immunization, there are only 895 Venus+ GC B cells and 237 Venus+ PCs. Therefore, it is very important to start the staining with a sufficient amount of cells to avoid stochastic effects. (4) Some B cells expressing other isotypes of BCRs may also be Venus+. For example, all B cells generated during in vitro cell culture from the Verigem reporter have a background shift in the Venus channel comparing to the WT control (Fig. 2b), which may be due to low-level translation of the germline transcript (unpublished observations). There are also small fractions of IgG1+ GC B cells and PCs generated in vivo from Verigem reporter mice that are weakly Venus positive (Fig. 2d). These Verigemlow cells that also express other isotypes of BCR could be transitional cells that recently switched to IgE, as suggested by surface IgE staining. Alternatively, they could have switched to IgE in the nonproductive IgH allele, as was described with the CεGFP reporter mouse [14], although translation of this allele is not expected [8]. The frequency of Verigemlow cells expressing other isotypes seems to vary depending on immunization conditions and time points. We are actively investigating the identity of these cells. Nevertheless, this again underscores the importance of including non-Verigem controls and additional cell surface markers (such as IgG1) to establish whether bona fide IgE+ B cells are being detected. 7. The IgE response in the immunization model described here will peak at around 7  days after immunization and quickly decline over time. In other models, such as Nippostrongylus brasiliensis infection, the peak IgE response may be closer to 10–14 days after immunization [12]. 8. Fixation with PFA is essential to preserve Venus (YFP) inside the cells during sectioning. However, the fluorescence of Venus from IgE+ B cells with Verigem reporter will be greatly diminished after fixation by PFA, which precludes the visualization of IgE+ GC B cells (see Fig. 3). Thus we include rabbit anti-GFP antibody in the primary antibody staining solution. As a YFP derivative, Venus is recognized by this anti-GFP antibody. It is important to include LNs from an immunized WT mouse as a negative control. In some cases we have observed that non-specific staining can occur. This non-specific staining is particularly prominent at the peak of the extrafollicular PC response, as some PCs will encode antibodies that cross-react with the reagents used for staining (this is particularly notable in our experience in mice immunized with antigens containing NP and with the use of Cy/Alexa dye fluorophores). 9. Prior to the hydration step, we use an etching pen (diamond scriber) to etch a region around the tissue(s). This makes it easier to dry a region around the tissue prior to adding the staining

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mix since the tissue becomes difficult to see after rehydration. The dried region around the tissue keeps the staining solution contained and enclosed for the duration of the staining step. 10. Suggested additional complementary staining reagents are PNA biotin to identify GC B cells, rat anti-mouse CD35 (BD Biosciences, clone 8C12) to identify follicular dendritic cells, rat anti-mouse CD138 (BD Biosciences, clone 281-2) to identify PCs, rat anti-mouse IgD (BioLegend, clone 11-26c.2a) or goat anti-mouse IgD (Fc) (Nordic-MUbio) to detect follicular B cells, rat anti-mouse IgE biotin (BD Biosciences, clone R35–118), rat anti-mouse IgE PE (BioLegend, clone RME-1), and rat anti-mouse IgG1 (BD Biosciences, clone A85-1). Some of the aforementioned reagents can be used when directly conjugated to fluorophores if the signal has sufficient intensity; alternatively, purified antibodies can be detected with fluorescently conjugated secondary antibodies. We typically obtain polyclonal F(ab′)2 secondary reagents, made in donkeys, that are conjugated to AMCA, Cy3, or Alexa Fluor 647 from Jackson ImmunoResearch and are absorbed against many species (must be absorbed against mouse), such as Cy3 F(ab′)2 donkey anti-rat IgG. Good streptavidin conjugates that can be paired with biotin staining are streptavidin-Cy3 (Jackson ImmunoResearch) and streptavidin-­ Alexa Fluor 647 (Thermo Fisher Scientific). 11. A folded Kimwipes or paper towel is sufficient for the drying step. For hydration and washing steps, we dip our slides in glass containers of 400–500 mL PBS. 12. For detection of anti-rabbit GFP antibody staining, we have found good success using donkey anti-rabbit Alexa Fluor 488 (Jackson ImmunoResearch) and donkey anti-rabbit Alexa Fluor 647 (Jackson ImmunoResearch). Detection in the far red channel (such as with Alexa Fluor 647) gives the least background due to low tissue autofluorescence in the far red region of the visible light spectrum. Detection in the far red channel also allows simultaneous detection of Venus fluorescence (see Fig.  3). A dilution of 1:300 for both reagents is sufficient. 13. The duration of washes may affect staining results. We aim to achieve a 3–5 min wash time for each slide. This is accomplished by staggering the start time for putting the slides in the wash container. For example, add one slide to the wash container, wait 1 min, add the next slide to the wash container, and repeat until four slides have been added (at this time, the first slide will have been washed for about 3 min). Now remove the first slide, and proceed to the next staining step as described in Subheading 3.3.3, step 8. When you have added the antibody stain to this slide, put another slide in the wash ­container, and then remove

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the slide that has been washing for the longest time, and then follow Subheading 3.3.3, step 8 again. Continue repeating by adding one slide and removing one slide from the wash container about every 1 min until all slides have been washed for about 3–5 min each.

Acknowledgments These protocols were developed for research projects supported by the UCSF Sandler Asthma Basic Research Center, the UCSF Cardiovascular Research Institute, the Weston Havens Foundation, and grants DP2HL117752 and R01AI103146 from the National Institutes of Health. C.D.C.A. is a Pew Scholar in the Biomedical Sciences, supported by The Pew Charitable Trusts. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health or of the Pew Charitable Trusts. References 1. Dullaers M, De Bruyne R, Ramadani F, Gould HJ, Gevaert P, Lambrecht BN (2012) The who, where, and when of IgE in allergic airway disease. J Allergy Clin Immunol 129(3):635–645. https://doi.org/10.1016/j.jaci.2011.10.029 2. Graham MT, Nadeau KC (2014) Lessons learned from mice and man: mimicking human allergy through mouse models. Clin Immunol 155(1):1–16. https://doi.org/10.1016/j. clim.2014.08.002 3. Gould HJ, Sutton BJ (2008) IgE in allergy and asthma today. Nat Rev Immunol 8(3):205– 217. https://doi.org/10.1038/nri2273 4. MacLennan IC, Toellner KM, Cunningham AF, Serre K, Sze DM, Zuniga E, Cook MC, Vinuesa CG (2003) Extrafollicular antibody responses. Immunol Rev 194:8–18. https:// doi.org/10.1034/j.1600-065X.2003.00058.x 5. Allen CD, Okada T, Cyster JG (2007) Germinal-center organization and cellular dynamics. Immunity 27(2):190–202. https:// doi.org/10.1016/j.immuni.2007.07.009 6. Cheng LE, Wang ZE, Locksley RM (2010) Murine B cells regulate serum IgE levels in a CD23-dependent manner. J  Immunol 185(9):5040–5047. https://doi. org/10.4049/jimmunol.1001900 7. Shin JS, Greer AM (2015) The role of Fc epsilon RI expressed in dendritic cells and monocytes. Cell Mol Life Sci 72(12):2349–2360. https:// doi.org/10.1007/s00018-015-1870-x

8. Yang Z, Robinson MJ, Allen CD (2014) Regulatory constraints in the generation and differentiation of IgE-expressing B cells. Curr Opin Immunol 28:64–70. https://doi. org/10.1016/j.coi.2014.02.001 9. Katona IM, Urban JF Jr, Scher I, Kanellopoulos-­ Langevin C, Finkelman FD (1983) Induction of an IgE response in mice by Nippostrongylus brasiliensis: characterization of lymphoid cells with intracytoplasmic or surface IgE.  J Immunol 130(1):350–356 10. Erazo A, Kutchukhidze N, Leung M, Christ AP, Urban JF Jr, Curotto de Lafaille MA, Lafaille JJ (2007) Unique maturation program of the IgE response in  vivo. Immunity 26(2):191–203. https://doi.org/10.1016/j. immuni.2006.12.006 11. Wesemann DR, Magee JM, Boboila C, Calado DP, Gallagher MP, Portuguese AJ, Manis JP, Zhou X, Recher M, Rajewsky K, Notarangelo LD, Alt FW (2011) Immature B cells preferentially switch to IgE with increased direct Smu to Sepsilon recombination. J  Exp Med 208(13):2733–2746. https://doi. org/10.1084/jem.20111155 12. Yang Z, Sullivan BM, Allen CD (2012) Fluorescent in  vivo detection reveals that IgE(+) B cells are restrained by an intrinsic cell fate predisposition. Immunity 36(5):857–872. https://doi.org/10.1016/j. immuni.2012.02.009

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13. Talay O, Yan D, Brightbill HD, Straney EE, Zhou M, Ladi E, Lee WP, Egen JG, Austin CD, Xu M, Wu LC (2012) IgE(+) memory B cells and plasma cells generated through a germinal-­ center pathway. Nat Immunol 13(4):396–404. https://doi.org/10.1038/ni.2256 14. He JS, Meyer-Hermann M, Xiangying D, Zuan LY, Jones LA, Ramakrishna L, de Vries VC, Dolpady J, Aina H, Joseph S, Narayanan S, Subramaniam S, Puthia M, Wong G, Xiong H, Poidinger M, Urban JF, Lafaille JJ, Curotto de Lafaille MA (2013) The distinctive germinal center phase of IgE+ B lymphocytes limits their contribution to the classical memory response. J Exp Med 210(12):2755–2771. https://doi. org/10.1084/jem.20131539 15. de Felipe P, Luke GA, Hughes LE, Gani D, Halpin C, Ryan MD (2006) E unum pluribus: multiple proteins from a self-­processing polyprotein. Trends Biotechnol 24(2):68–75. https:// doi.org/10.1016/j.tibtech.2005.12.006 16. Nagai T, Ibata K, Park ES, Kubota M, Mikoshiba K, Miyawaki A (2002) A variant of

yellow fluorescent protein with fast and efficient maturation for cell-biological applications. Nat Biotechnol 20(1):87–90. https:// doi.org/10.1038/nbt0102-87 17. Wilmore JR, Jones DD, Allman D (2017) Protocol for improved resolution of plasma cell subpopulations by flow cytometry. Eur J  Immunol 47(8):1386–1388. https://doi. org/10.1002/eji.201746944 18. Matheu MP, Parker I, Cahalan MD (2007) Dissection and 2-photon imaging of peripheral lymph nodes in mice. J  Vis Exp 7:265. https://doi.org/10.3791/265 19. Murooka TT, Mempel TR (2012) Multiphoton intravital microscopy to study lymphocyte motility in lymph nodes. Methods Mol Biol 757:247–257. https://doi. org/10.1007/978-1-61779-166-6_16 20. Yang Z, Robinson MJ, Chen X, Smith GA, Taunton J, Liu W, Allen CD (2016) Regulation of B cell fate by chronic activity of the IgE B cell receptor. eLife 5:e21238. https://doi. org/10.7554/eLife.21238

Chapter 20 Chromatin Preparation from Murine Eosinophils for Genome-Wide Analyses Carine Bouffi, Artem Barski, and Patricia C. Fulkerson Abstract Dynamic gene expression is a major mechanism that directs hematopoietic lineage commitment and differentiation. Recent advances have revealed an association between chromatin signatures and functional genetic sequences such that these chromatin signatures can be used to predict regulatory elements such as enhancers that may direct lineage differentiation. Our understanding of the genetic elements that regulate eosinophil development is very limited, likely due to the technical challenges in working with a rare complex cell. Herein, we describe protocols to sort mature eosinophils from the bone marrow of mice and to prepare chromatin that can be used for ChIP studies for genome-wide mapping of histone marks and transcription factors in mature eosinophils. Comprehensive epigenomic profiling during critical stages in eosinophil development will ultimately aid in defining the gene regulatory networks necessary to regulate eosinophil production. Key words Chromatin immunoprecipitation (ChIP), Gene regulation, Epigenetics

1  Introduction Eosinophils participate in homeostatic and pathologic processes primarily through their ability to store and rapidly release an abundance of mediators, including cytokines, growth factors, and cytotoxic proteins [1, 2]. In the peripheral blood, eosinophils normally represent less than 5% of white cells, and eosinophilia is generally characterized as mild (450–1500 eosinophils per microliter), moderate (1500–5000 eosinophils per microliter), or severe (greater than 5000 eosinophils per microliter). In general, blood eosinophilia results from enhanced eosinophilopoiesis (i.e., increased production of mature cells from eosinophil lineage-committed progenitors [EoPs] in the bone marrow). Dynamic gene expression is a major regulatory mechanism that directs hematopoietic lineage commitment and differentiation [3, 4]. Gene expression control is achieved, in part, via access of the transcriptional machinery to gene regulatory elements, such as promoters and enhancers. R. Lee Reinhardt (ed.), Type 2 Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1799, https://doi.org/10.1007/978-1-4939-7896-0_20, © Springer Science+Business Media, LLC, part of Springer Nature 2018

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Modifications to chromatin, a complex formed between genomic DNA and histone proteins, can profoundly influence gene expression via alterations in DNA accessibility [5]. Technological advances in recent years have revealed an association of specific patterns of histone modifications (i.e., chromatin signature or fingerprint) with functional genetic sequences such that these chromatin signatures can be used to predict regulatory elements such as enhancers that may direct lineage differentiation [6–9]. Our understanding of the genetic elements that regulate eosinophil development is very limited. We previously identified genome-wide epigenome changes that occur at critical stages in eosinophil development—eosinophil-­ lineage commitment and eosinophil maturation. We performed chromatin immunoprecipitation coupled with massively parallel sequencing (ChIP-seq) to identify genetic regulatory elements that are active during homeostatic eosinophil production in the bone marrow [10]. Our studies provided the first demonstration of stage-specific epigenomic signatures in the eosinophil lineage [10]. Previously, molecular assays involving chromatin required large numbers of purified cells (often as many as 10–50 × 106), which made studies focused on transcriptional regulation in rare cell populations, like eosinophils, very challenging and expensive to perform. In addition, eosinophils contain many highly charged granule proteins and enzymes that cleave DNA and RNA [11]; thus, isolation of quality chromatin from these complicated cells can be challenging. With our collaborators, we have developed a methodology to perform ChIP-seq reproducibly with ~0.5 × 106 eosinophils. Herein, we describe protocols to sort mature eosinophils from the bone marrow of mice and to prepare chromatin that can be used for ChIP studies for genome-wide mapping of histone marks and transcription factors in mature eosinophils.

2  Materials 2.1  Equipment

1. Sterile mortar and pestle. 2. 70 μm cell strainer. 3. 50 mL conical tubes. 4. 15 mL conical tubes. 5. 1.5 mL microcentrifuge tubes. 6. FACS tubes: 5 mL polystyrene round bottom tubes. 7. Table top centrifuge. 8. Rocking platform. 9. Rotating platform.

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Table 1 Control and sample staining for mature murine eosinophils

Tube

Anti-CCR3

Anti-Siglec-F

Live/ dead stain

Unstained FMO1—Siglec-F gating

X

FMO2—CCR3 gating

X X

FMO3—Live/dead gating

X

X

Sample

X

X

X

X

10. Sonicator. 11. Gel electrophoresis equipment. 2.2  Eosinophil Purification

1. Red blood cell lysis buffer (8.3 g/L ammonium chloride in 0.01 M Tris–HCl buffer). 2. PBS. 3. Fetal bovine serum (FBS). 4. Collection buffer (2% FBS in PBS). 5. FACS buffer (PBS with 0.5% BSA and 2 mM EDTA). 6. Sorting media (Iscove’s Modified Dulbecco’s Media with 20% FBS). 7. Fluorescent antibodies in FACS buffer (see Table 1). 8. Cell viability stain (diluted in PBS per manufacturer’s direction).

2.3  Chromatin Preparation and Immunoprecipitation

1. 5 M sodium chloride (NaCl). 2. 10% sodium dodecyl sulfate (SDS) solution: 10% (w/v) SDS in ddH2O. 3. 10% sodium deoxycholate (NaDOC) solution: 10% (w/v) NaDOC in ddH2O. 4. 10% Triton-X solution: 10% (w/v) Triton-X in PBS. 5. Proteinase K. 6. 2.5 M glycine solution: 2.5 M glycine in PBS. 7. Glycerol. 8. TE buffer with 0.1% SDS. 9. 10× cross-linking solution: 8.8% formaldehyde, 0.1 M NaCl, 1 mM EDTA, 0.5 M EGTA, 50 mM HEPES buffer in ddH2O.

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10. L1 buffer: 50 mM HEPES buffer, 140 mM NaCl, 1 mM EDTA, 10% glycerol, 0.5% NP-40, 0.25% Triton-X in ddH2O. 11. L2 buffer: 200 mM NaCl, 1 mM EDTA, 0.5 mM EGTA, 10 mM Tris, pH 8.0 in ddH2O . 12. Protease inhibitor cocktail (10×): Mixture of AEBSF (104 mM), aprotinin (80 μM), bestatin (4 mM), E-64 (1.4 mM), leupeptin (2 mM), and pepstatin A (1.5 mM) in DMSO. Diluted 1:10 in L1, L2, and sonication buffers for use per manufacturer’s instructions. 13. Sonication buffer: TE + 0.1% SDS in ddH2O. 14. Chromatin QC solution: 0.3 M NaCl, 2 μL of 10% SDS, 0.5  μL of RNase A, and 2.5 μL of proteinase K to sterile ddH2O (final volume 40 μL). 15. Agarose. 16. Chromatin immunoprecipitation (ChIP) buffer: 150 mM NaCl, 0.1% NaDOC, 1% Triton-X, 10% protease inhibitor cocktail in TE buffer with 0.1% SDS. 17. Magnetic protein A beads: magnetic beads with recombinant protein A covalently coupled to the surface for immunoprecipitation.

3  Methods 3.1  Eosinophil Purification by FACS

Murine eosinophils can be purified from whole bone marrow cells via cell sorting. Whole bone marrow cells (~10–15 × 106 per mouse) are collected from crushed femurs and tibiae (see Note 1). Mature eosinophils comprise 1–3% of whole bone marrow cells at homeostasis (see Note 2) and are identified by surface expression of Siglec-F and CCR3 (see Fig. 1). 1. Collect whole bone marrow cells from dissected and cleaned murine femurs and tibiae that were crushed in a sterile mortar and pestle containing collection buffer (see Note 3). 2. Push the collected cells through a 70-μm strainer into a 50 mL conical tube to remove debris and create a single-cell suspension. 3. Lyse red blood cells using lysis buffer per manufacturer’s instructions. Centrifuge cells at 500 × g-force at 4°C, discard supernatant, and resuspend pellet in 2 mL collection buffer. Count cells in the suspension. 4. Centrifuge cells at 500 × g-force at 4°C, discard supernatant, and then resuspend cell pellet in 500 μL FACS buffer for every 100 × 106 live cells and transfer cell suspension to a 5 mL polystyrene round bottom tube (FACS tube).

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Fig. 1 Eosinophil gating strategy. A representative gating strategy to identify mature eosinophils from naive murine bone marrow is shown and starts with pregating on viable single cells and then further gating on cells that coexpress Siglec-F and CCR3. Percentage of parent gate is shown

5. Transfer 50 μL of the total cell suspension to another FACS tube for staining controls. Add 150 μL of FACS buffer to this cell suspension, mix gently, and transfer 50 μL of this diluted cell suspension to three FACS tubes for a total of four tubes for staining controls (see Table 1). Save the remaining 450 μL of cell suspension for sorting. 6. Add fluorescent antibodies to the cell suspensions in FACS tubes for staining controls and sample cells to be sorted (see Note 4). 7. Incubate cells at room temperature for 15–20 min in the dark (to protect antibodies conjugated with fluorochromes from the light), and then add cell viability stain for an additional 5 min. 8. Centrifuge at 500 × g-force for 5 min. Discard the supernatant. 9. Wash cell pellets by resuspending cells with 500 μL FACS buffer and centrifuge at 500 × g-force for 5 min. 10. Resuspend pelleted bone marrow cells to be sorted at a density of 20 × 106 cells per mL of FACS buffer. Cell pellets for staining controls can be resuspended in a final volume of 100– 200 μL FACS buffer. 11. Live mature murine eosinophils are identified from whole bone marrow suspensions via surface expression of Siglec-F and CCR3 (see Fig. 1). Sort cells directly into 5 mL sorting media in a 15 mL conical tube that has been coated with the sorting media by gently inverting the tube several times (see Note 5). 3.2  Chromatin Fixation

1. Make note of the final volume of sorted eosinophils in the collection tube (see Note 6). 2. Add freshly made 10× cross-linking solution directly to the collection tube containing sorted murine eosinophils, and mix gently by inversion (see Note 7).

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3. Incubate the cells on ice for 8 min. 4. To stop the cross-linking reaction, add one-twentieth of the final volume of a 2.5 M glycine solution to each tube. Place the tube on a rocking platform at a slow/medium speed at room temperature for 5 min. 5. Centrifuge at 2000 × g-force for 10 min at 4°C. Discard the supernatant. 6. Gently resuspend the cell pellet in 1 mL ice-cold PBS, and transfer the suspension into a fresh 1.5 mL microcentrifuge tube. 7. Centrifuge cells at 2000 × g-force for 5 min at 4°C, and carefully decant and discard the supernatant. 8. Resuspend the cell pellet in 1 mL ice-cold PBS. 9. Repeat step 7. 10. Centrifuge the cell pellet again at 3000 × g-force for 5 min to dry the pellet. Remove and discard any residual liquid in the tube. 11. Freeze and store at −80°C until ready for use (see Note 8). 3.3  Nuclei Preparation

1. Prepare L1 and L2 buffers. Add protease inhibitor cocktail to each buffer. Keep L1 buffer on ice and L2 buffer at room temperature until ready to use. 2. Thaw the frozen cell pellet by incubation on ice. 3. Resuspend the cell pellet in 1 mL of L1 buffer with added protease inhibitor cocktail. Incubate on a rotating platform at 4 °C for 10 min. 4. Centrifuge at 5000 × g-force for 5 min at 4 °C. 5. Carefully remove and discard the supernatant using a micropipette or vacuum. 6. Resuspend the cell pellet in 1 mL of L2 buffer with added protease inhibitor cocktail. 7. Incubate on a rotating platform at room temperature for 10 min. 8. Centrifuge at 5000 × g-force for 5 min at 4 °C. Carefully remove and discard the supernatant using a micropipette or vacuum. 9. Wash the pelleted nuclei by adding 0.5 mL of sonication buffer to lift the pellet up from the bottom of the tube without resuspending into a suspension (see Note 9). Do not disrupt the pellet.

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The sonication intensity, length, and number of cycles needed for optimal chromatin shearing should be determined experimentally for each cell type and cell density and will vary between machines. 1. Resuspend the pelleted nuclei in 130 μL of sonication buffer with added protease inhibitor cocktail. 2. Transfer the nuclei suspension into a small vial or microcentrifuge tube for sonication. The suspension should be loaded into vials without bubbles for proper and reproducible sonication. 3. Sonicate the nuclei to generate sheared chromatin that is between 200 and 500 base pairs in size. The length, intensity, and number of cycles will need to be determined experimentally as it will vary by machine (see Note 10). 4. Collect the chromatin suspension into a 1.5 mL microcentrifuge tube and transfer 10 μL for quality control testing. 5. Add 5 μL of 100% glycerol (5% glycerol final concentration) to the rest of the suspension (~100 μL) and store at −80 °C until ready for use.

3.5  Chromatin Quality Control Testing

1. To the microcentrifuge tube with the chromatin for quality control, add 40 μL of chromatin QC solution to the 10 μL of sheared chromatin. 2. Incubate the quality control chromatin at 37 °C for 60 min. Incubate at 65 °C for an additional 4–5 h up to overnight (see Note 11). 3. Run the chromatin sample on an agarose gel (1%) to determine the size of sheared chromatin (see Fig. 2). Sheared chromatin should be between 200 and 500 base pairs.

3.6  Chromatin Preparation for Immunoprecipitation

1. If sonication looks appropriate, thaw the chromatin sample on ice. 2. Add 100 μL of ChIP buffer to the sample. 3. Centrifuge at 20,000 × g-force for 10 min at 4°C.

Fig. 2 Chromatin quality control testing. The size of sheared chromatin from two independent experiments is shown on 1% agarose gel. The left panel shows a smear of DNA between 200 and 500 base pairs. This chromatin was used for chromatin immunoprecipitation. The right panel shows uneven shearing, and this chromatin was not used for further experiments

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4. To preclear the chromatin solution, add 5 μL protein A-coated magnetic beads (same beads will be used for the immunoprecipitation) to the sample and incubate between 30 and 60 min at 4°C on a rotating rack. 5. Put tubes on magnet for 1 or 2 min, and collect the negative fraction that contains the pre-cleared chromatin. Adjust total volume up to 200 μL with ChIP Buffer. 6. Keep 2 μL of the negative fraction (or 1% of total) for input control. 7. Transfer 198 μL of chromatin suspension into tube. Chromatin suspension can be stored with added glycerol (final 5% glycerol content) at −80°C until needed for future ChIP studies (see Note 12).

4  Notes 1. For the highest quality of chromatin, the eosinophils need to be sorted from fresh whole bone marrow preparations that have not undergone any bead-mediated depletion or enrichment. It has been our experience that cells that have been incubated with beads yield poor quality and amount of chromatin. 2. We typically sort eosinophils from the pooled bone marrow of ten mice (~100 × 106 total cells) to typically yield around 1 × 106 mature eosinophils. 3. Alternatively, the femurs can be flushed with collection buffer and pooled prior to pushing thru a cell strainer. Total cell yield is smaller but collection is faster. 4. Each antibody lot and viability dye should be titrated for optimal staining. We typically use the antibodies directed against Siglec-F (PE-conjugated) and CCR3 (FITC-conjugated) at 1:200 dilutions. We have found that the viability dyes are very potent and use at a dilution of 1:3000. We make a 1:1000 dilution in PBS of the original stock and add one-third of the final volume to the cell suspension in FACS buffer. 5. We use fluorescence minus one (FMO) controls to identify and gate cells when multiple fluorochromes are used in a given panel. To set the gates for sorting, we use the unstained initially, then FMO3 to set the gate for live cells, and then FMO1 and FMO2 to set gates for Siglec-F and CCR3-positive cells, respectively. We sort cells using a 70 μm nozzle on the FACSAria. Coat the 15 mL conical tube with the sorting media by rotating and inverting the tube after adding 5 mL sorting media. This prevents the sorted cells from sticking to the side of the tubes.

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6. Since the cross-linking buffer is added at 10×, the total volume that the sorted cells are suspended in needs to be recorded. 7. One-tenth of the final volume of the cross-linking buffer is added to the cells suspended in the sorting media. For example, if the cells are sorted in 1 mL, then 0.11 mL of the 10× cross-linking buffer would be added to cells directly. The final concentration of formaldehyde is 0.8% and should not exceed 1%. 8. The fixed cell pellets can be stored indefinitely at −80°C. We have successfully performed ChIP on pellets stored for up to 6 months. 9. If the pellet dissolves in the buffer, centrifuge the suspension at 5000 × g for 10 min to repellet the nuclei and move on to shearing. 10. In our laboratory, we use a Covaris sonicator to shear the chromatin with the nuclei suspended in small vials. The settings that we have determined to be optimal when starting with ~1 × 106 mature eosinophils are 10% duty cycle, 5 intensity (equals 105 pulse intensity), and 200 burst for 45 s. 11. A PCR machine can be used for the heating cycles, but we recommend that the last cycle be set for 15 °C to avoid precipitating the SDS in the solution. 12. We typically yield ~1.5 μg chromatin from 1 × 106 sorted eosinophils. We use ~1–1.5 μg of chromatin for every ChIP. The ratio of chromatin to antibody needs to be determined experimentally as the optimal ratio depends on the antibody used for the immunoprecipitation. References 1. Lee J, Rosenberg HF (2013) Eosinophils in health and disease, 1st edn. Elsevier/Academic Press, London; Waltham, MA 2. Rosenberg HF, Dyer KD, Foster PS (2013) Eosinophils: changing perspectives in health and disease. Nat Rev Immunol 13(1):9–22. https://doi.org/10.1038/nri3341 3. Kee BL (2011) A comprehensive transcriptional landscape of human hematopoiesis. Cell Stem Cell 8(2):122–124. https://doi. org/10.1016/j.stem.2011.01.006 4. May G, Soneji S, Tipping AJ, Teles J, McGowan SJ, Wu M, Guo Y, Fugazza C, Brown J, Karlsson G, Pina C, Olariu V, Taylor S, Tenen DG, Peterson C, Enver T (2013) Dynamic analysis of gene expression and genome-wide transcription factor binding during lineage specification of multipotent progenitors. Cell Stem Cell 13(6):754–768. https://doi. org/10.1016/j.stem.2013.09.003

5. Maston GA, Evans SK, Green MR (2006) Transcriptional regulatory elements in the human genome. Annu Rev Genomics Hum Genet 7:29–59. https://doi.org/10.1146/ annurev.genom.7.080505.115623 6. Wang Z, Zang C, Rosenfeld JA, Schones DE, Barski A, Cuddapah S, Cui K, Roh TY, Peng W, Zhang MQ, Zhao K (2008) Combinatorial patterns of histone acetylations and methylations in the human genome. Nat Genet 40(7):897–903. https://doi.org/10.1038/ ng.154 7. Heintzman ND, Stuart RK, Hon G, Fu Y, Ching CW, Hawkins RD, Barrera LO, Van Calcar S, Qu C, Ching KA, Wang W, Weng Z, Green RD, Crawford GE, Ren B (2007) Distinct and predictive chromatin signatures of transcriptional promoters and enhancers in the human genome. Nat Genet 39(3):311–318. https://doi.org/10.1038/ng1966

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8. Hon GC, Hawkins RD, Ren B (2009) Predictive chromatin signatures in the mammalian genome. Hum Mol Genet 18(R2):R195–R201. https://doi. org/10.1093/hmg/ddp409 9. Barski A, Cuddapah S, Cui K, Roh TY, Schones DE, Wang Z, Wei G, Chepelev I, Zhao K (2007) High-resolution profiling of histone methylations in the human genome. Cell 129(4):823–837. https://doi.org/10.1016/j. cell.2007.05.009

10. Bouffi C, Kartashov AV, Schollaert KL, Chen X, Bacon WC, Weirauch MT, Barski A, Fulkerson PC (2015) Transcription factor repertoire of homeostatic eosinophilopoiesis. J Immunol 195(6):2683–2695. https://doi. org/10.4049/jimmunol.1500510 11. Marichal T, Mesnil C, Bureau F (2017) Homeostatic eosinophils: characteristics and functions. Front Med (Lausanne) 4:101. https://doi.org/10.3389/fmed.2017.00101

Chapter 21 A Sensitive and Integrated Approach to Profile Messenger RNA from Samples with Low Cell Numbers Sandy Lisette Rosales, Shu Liang, Isaac Engel, Benjamin Joachim Schmiedel, Mitchell Kronenberg, Pandurangan Vijayanand, and Grégory Seumois Abstract Transcriptomic profiling by RNA sequencing (RNA-Seq) represents the preferred approach to measure genome-wide gene expression for understanding cellular function, tissue development, disease pathogenesis, as well as to identify potential biomarkers and therapeutic targets. For samples with small cell numbers, multiple methods have been described to increase the efficiency of library preparation and to reduce hands-on time and costs. This chapter reviews our approach, which combines flow cytometry and the most recent high-resolution techniques to perform RNA-Seq for samples with low cell numbers as well as for single-cell samples. Our approach reduces technical variability while increasing sensitivity and efficiency. Thus, it is wellsuited for large-scale gene expression profiling studies with limited samples for basic and clinical studies. Key words RNA-Seq, Single-cell RNA-Seq, Low-input RNA, Transcriptomics, Smart-Seq2

1  Introduction Large-scale transcriptomic profiling studies constitute an important first step to understanding diverse biological phenomena such as tissue development, cellular differentiation and functionality, and disease pathogenesis [1–3]. They also represent a promising tool for development of diagnostics and therapeutics [4]. However, such studies can be limited by sample availability and heterogeneity, especially in immunology [5]. The heterogeneous and complex cellular composition of tissue samples can confound analyses by increasing the biological noise and masking subtle but biologically relevant changes in mRNA expression. Diverse methods have been used to address these issues, for example, using more accessible sources of

Sandy Lisette Rosales and Shu Liang contributed equally to this work. R. Lee Reinhardt (ed.), Type 2 Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1799, https://doi.org/10.1007/978-1-4939-7896-0_21, © Springer Science+Business Media, LLC, part of Springer Nature 2018

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cells, such as blood, or using ex vivo culture to expand rare cell populations derived from tissue samples [6, 7]. However, multiple reports have shown that the data derived by these alternative methods do not always reflect the exact processes that occur in vivo. Recently developed protocols in RNA sequencing for samples with low cell number or single-cell hold great potential for exploring biological systems with unprecedented resolution [8]. Among these protocols, Smart-Seq2 has shown significant advantages, such as increased efficiency of reverse transcription and library preparation, as well as reduced hands-on time and costs [9]. Smart-­Seq2 has been recently appreciated as the most reproducible and sensitive method for low-input RNA-Seq and single-cell RNA-Seq [10]. The protocol consists of an oligo-based Poly-A tailed mRNA capture followed by a high-fidelity reverse transcription using locked nucleic acids (LNA)-oligos to capture full-length transcripts. mRNA complementary (c)DNA strands are then amplified using a PCR-based process and sequencing adaptors integrated into the cDNA fragments in a single step using Tn5 transposase technology [9]. Our work explores the merits of both bulk RNA-Seq and single-­ cell RNA-Seq in revealing disease-associated patterns or changing paradigms in development of rare immune cell subsets [2, 3]. To overcome problems arising from tissue paucity and heterogeneity, we designed a flow cytometry-based method for isolating pure populations from dispersed tissue samples, coupled with the highly sensitive, medium-throughput RNA-Sequencing Smart-­ Seq2 protocol. The procedure allows for the performance of both bulk RNA-Sequencing and single-cell RNA-Seq assays from the same cell sorting experiment. We have successfully applied our protocol to multiple research projects. In the study by Engel et al., we performed RNA-Seq analysis at the bulk and single-cell levels in three thymic invariant natural killer T cell (iNKT) subsets [3]. Analysis showed an extensive and unexpected diversity of global gene expression levels revealing unique cell type-specific functional molecular patterns. We have also applied our protocol to investigate qualitative differences in T helper 2 (Th2) cells from subjects with allergic asthma and allergic rhinitis. Although both diseases share clinical and pathological features characterized by an exaggerated Th2-type inflammation, allergic rhinitis patients do not develop asthma, suggesting a divergence in disease mechanisms. Analysis of circulating Th2 cells isolated from both disease groups and from healthy subjects revealed differentially expressed genes involved in cell survival, metabolic pathways, and activation persistence [2]. In this chapter, we provide a detailed step-by-step description of the entire procedure from cell sorting to final library quantification for either bulk or single-cell RNA-Seq. It is an integrated, multi-sample, and highly sensitive approach. Figure 1 ­displays an overview of critical steps, quality control steps, and hands-on time estimations for the planning of experiments.

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2  Materials 2.1  Long-Term Common Stock Solutions (See Note 1)

1. 1 M Tris pH 8.0*. 2. 0.5 M EDTA pH 8.0*. 3. TE buffer 1*: 10 mM Tris pH 8.0; 1 mM EDTA pH 8.0, store at 4 °C. 4. TE buffer 2*: 1 mM Tris pH 8.0, 100 μM EDTA pH 8.0, store at 4 °C. 5. TE buffer 3*: 10 mM Tris pH 8.0, 100 μM EDTA pH 8.0, store at 4 °C.

2.2  Common Reagents and Specific Laboratory Equipment

1. Molecular biology grade water (DNase, RNase, protease, endotoxin—free; referred as ultrapure water). 2. RNase AWAY solution (see Note 2). 3. 70% and 80% ethanol solutions (freshly prepared). 4. Recombinant RNase inhibitor, store at −20 °C. 5. Ethanol 200 proof, anhydrous 99.5%. 6. 10 mM dNTP, store at −20 °C. 7. Collecting tube: 1.5 mL Axygen Maxymum recovery PCR tubes (Fisher Scientific). 8. Plate 1: 96-well semi-skirted PCR plates (BioRad) (see Note 3). 9. Plate 2: Hard-shell thin-wall 96-well skirted PCR plates. 10. 0.2 mL RNase-free PCR 8-tube strips. 11. Plate 3: MicroAmp Fast 96-Well Reaction Plate (Life Technologies, see Note 4). 12. qPCR MicroAmp optical adhesive film (Life Technologies). 13. MicroAmp Clear Adhesive Film (Life Technologies). 14. 96-well magnetic device (Axygen). 15. Repeater micropipette. 16. Thermomixer. 17. Real-time quantitative PCR system. 18. Capillary DNA-RNA electrophoresis equipment (Advance Analytical Fragment analyzer or Agilent Bioanalyzer).

2.3  Specific Reagents and Buffers 2.3.1  Sample Collection from Size-Limited Samples

For samples with more than 5000 cells: 1. TRIzol LS, store at room temperature (see Note 5). 2. 1.5 mL sterile tubes: RNase-free, individually wrapped. For single-cell or low cell number collection: 1. 2× low-input cell lysis buffer (LI-LB)*: 0.2% Triton X-100 (vol/vol) (see Note 6).

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2.3.2  RNA Extraction

1. miRNeasy micro kit (Qiagen)—contains the following: miRNeasy MinElute spin columns and RWT and RPE buffers. The RPE and RWT buffers have to be reconstituted with 100% ethanol. 2. RNase-free DNase I kit (Qiagen)—contains the following: RNase-free DNase I, RNase-free Buffer RDD, and RNase-free water. 3. DNAse mix: 1:8 vol:vol ratio of reconstituted RNase-free DNase I and RNase-free Buffer RDD. 4. Chloroform. 5. 0.5 mL Nunc Cryobank vials (Thermo Fisher Scientific).

2.3.3  RNA Quality and Quantity Measurement

1. SuperScript III First-Strand Synthesis System for RT-PCR (Invitrogen), store at −20 °C. 2. Shaved ice from −80 °C freezer. 3. Annealing mix: 5 μM OligodT(20) primers, 10 ng/μL of random hexamers, and 2.4 U/μL of RNase-OUT for a total volume of 3.5 μL per sample (see Note 7). 4. SuperScript III reverse transcription mix: 1× strand buffer, 5 mM MgCl2, 10 mM DTT, 0.5 mM dNTP, 1.2 U/μL of RNase-OUT, and 10 U/μL SuperScript III enzyme for a total volume of 5.6 μL per sample. 5. Human β2m housekeeping gene primers: 25 nM FWD: 5′-CTG CCG TGT GAA CCA TGT GAC TTT-3′; 25 nM REV: 5′-TGC GGC ATC TTC AAA CCT CCA TGA-3′ (see Notes 8 and 9). 6. SYBR Green FAST universal 2× qPCR master mix (Roche). 7. Housekeeping gene qPCR mix: 1× SYBR Green Master Mix, 0.31 μM for each primers, complete to 7.5 μL with ultrapure water. 8. Applied Biosystems QuantStudio 6 Flex Real-Time PCR System.

2.3.4  cDNA Synthesis by High-Fidelity Reverse Transcription

1. 5 M betaine (see Notes 1 and 10). 2. SuperScript II reverse transcriptase (RT) kit: includes RT enzyme and 25 mM MgCl2 solution (Life Technologies), store at −20 °C. 3. Template switching oligo (TSO): 5′-AAG CAG TGG TAT CAA CGC AGA GTA CAT rGrG+G-3′, HPLC purified (Exiqon); dilute with TE buffer 1–100 μM (see Notes 8 and 9). 4. Oligo-dT30VN: 5′-ACA AGC AGT GGT ATC AAC GCA GAG TAC T(30)VN-3′, HPLC purified (Integrated DNA Technologies); dilute with TE buffer 1–100 μM (see Notes 8 and 9).

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5. Reverse transcription mix (RT-mix): 10 U/μL SuperScript II RT, 1 U/μL RNase Inhibitor, 1× SuperScript II first strand buffer, 5 mM DTT, 1 M betaine, 0.6 mM MgCl2, 1 μM template switching oligo (TSO). 2.3.5  cDNA Amplification

1. SYBR Green dye, store at −20 °C. 2. 50× SYBR Green solution = 1:1000 dilution of stock. 3. 50× Rox reference dye (Invitrogen), store at −20 °C. 4. KAPA HiFi HotStart ready mix kit, store at −20 °C. 5. Oligo ISPCR: 5′-AAG CAG TGG TAT CAA CGC AGA-3′, HPLC purified (Integrated DNA Technologies); dilute with TE buffer 1–100 μM (see Notes 8 and 9).

2.3.6  cDNA Size Selection

1. Agencourt AMPure XP beads, store at 4 °C (Beckman Coulter).

2.3.7  cDNA Quantification

1. Bacteriophage lambda DNA (Life Technologies). Sonicate and dilute up to 40 ng/μL with TE buffer 1 (see Note 11). 2. Quant-iT PicoGreen dsDNA reagent (Life Technologies), store at −20 °C. 3. Prepare PicoGreen solution by diluting Quant-iT PicoGreen dsDNA reagent 500-fold with TE buffer 3 (see Note 12). 4. Plate 4: 96-well plate, flat bottom, black for fluorescence measurements. 5. Top-read 96-well plate reader as, for example, SpectraMax M2 Multi-Mode Microplate Reader.

2.3.8  cDNA Library Preparation

1. Nextera XT DNA library preparation kit: 96 reactions, store at −20 °C. 2. Nextera XT index kit: 96 indexes, store at −20 °C. 3. Tagmentation mix: Illumina Nextera XT reagents amplicon tagmentation buffer and tagmentation DNA mix in a 1:2 ratio, respectively. 4. Agencourt AMPure XP beads (Beckman Coulter), store at 4 °C.

3  Methods Before starting any bench work, UV sterilize instruments (pipettes, racks, etc.) for at least 15 minutes, and clean bench and equipment with 70% ethanol solution and RNase AWAY solution (see Note 2).

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3.1  Sample Collection from Size-­ Limited Samples

Depending on the cell number and the scientific objectives, we suggest three distinct approaches to sort cells (see Fig. 1). For sample sizes ranging between 5000 and 200,000 cells, start from Subheading 3.1.1. For single cells or cell numbers less than 5000 cells, proceed directly to Subheading 3.1.2 (see Note 13).

3.1.1  Sample Collection for Samples with More than 5000 Cells

If the size of the cell population of interest ranges between 5000 and 200,000 cells, we suggest performing the sorting directly into TRIzol LS lysis buffer. Proceed with the following steps: 1. Directly sort cells into 750 μL of TRIzol LS in RNase-free individually wrapped 1.5 mL tubes. 2. During the sort, either after every 5 min of sorting or after every 10,000 cells sorted, pause sorting, close tubes, and vortex for about 15 s. Place tubes back on the sorting device and resume sorting (see Note 14). 3. After sorting, vigorously pipette each sample to lyse cells properly. 4. Fill up the volume to 1 mL with ultrapure water, vortex for 30 s, incubate at room temperature for 1 min, and then briefly spin and store samples at −80 °C (see Note 15). Once all samples are collected, proceed with the RNA extraction (see Subheading 3.2).

3.1.2  Single-Cell or Low Cell Number Collection

1. For less than 5000 cells, we recommend sorting a fixed cell number for all cell populations of interest into a fixed volume of complete LI-LB (see Note 16). Sort cells directly into 0.2 mL RNase-free tubes. For single cells, we recommend sorting cells directly in 96-well plates with 4 μL of complete LI-LB (Plate 1). 2. Dispense the appropriate volumes (see Table 1, Note 17) of complete LI-LB into 0.2 mL PCR tubes or 4 μL into each plate well for single-cell sorts. 3. After sort collection, promptly close tubes or seal plates firmly with adhesive seal. Vortex with care for 15 s and spin at 3000 × g-force for 2 min. 4. Samples are ready for cDNA synthesis or can be stored at −80 °C until needed.

3.2  RNA Extraction

RNA extraction is performed with Qiagen miRNeasy micro kits (see Note 18). We have slightly modified the manufacturer’s protocol to increase extraction yield. 1. Add 100 μL (1:5 vol:vol) of chloroform to 500 μL of cells lysed in TRIzol LS and close tube securely. Vortex the tube for 30 s.

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Only for Trizol samples

Flow cytometry sorting Time

5,000 > 200,000 cells

0.5 to 4h

Sort in 1.5 ml tubes with Trizol LS

2 h/16 samples

RNA extraction

2h

Single cell

Sort in 0.2 ml tubes with LI-LB i.e. 400 cells in 8 µL

Sort in 96 well with 4µL LI-LB

mRNA AAAAAAAAAAAAAAA

QC

Quantity: B2m qPCR Quality : RIN

3.5 h

Reverse Transcription (RT)

4h

Cycle determination (CtD)

IS-PCR…TrGrG+G

LNA-TSO oligo

QC

NVTTTTT…IS-PCR

OligodT-30VN

ISPCR…TrGrG+G ISPCR…….C-C-C---

AAAAAA NVTTTTTTT…

IS-PCR primers GGG CCC

cDNA Amplification

2h 1.5 h

< 5,000 cells

Full length cDNA

AAAAAA TTTTTT

Tn5 + Illumina adaptors

Quantification: Pico Green Quality: Bioanalyzer

1h

Tagmentation

1h

index primers Amplification

2h

Final Library DNA purification

Tagmentation: Fragmentation and adaptor insertion GGG CCC

GGG CCC AAAAAA TTTTTT

**

QC

24 h

Sequencing Single-end 50 bp reads

Quality: Bioanalyzer

Illumina Index primers

**

Amplification with Index primers

Quantification: PicoGreen

1.5 h

AAAAAA TTTTTT

** ** Final Library

** **

** **

** **

Sequencing

Read 1 ** **

Read 2 ** **

Fig. 1 Overview of the method. The different sorting methods depending on the sample availability for bulk or single-cell RNA-Seq are indicated on the top of the schematic. All the major steps of the procedure including quality control steps and timing are shown on the left for easy planning of the experiment. In parallel with the procedure, we display a schematic of the molecular process

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Table 1 Low-input RNA lysis buffer

LI-LB Mix for 48 samples

Stock concentration

Lysis buffer

Volume for single cell samples (μL) 191.5

RNAse inhibitor

40 U/μL

10.1

dNTP

10 mM

100.8

Molecular grade water

100.8

Aliquot per sample (μL)

8

All components needed to prepare the low-input RNA lysis buffer are listed with stock concentrations as well as volumes needed to prepare buffer for 48 samples

2. Incubate at room temperature for 2–3 min. Centrifuge for 15 min at 12,000 × g-force at 4 °C. 3. Transfer the upper aqueous phase to a fresh 1.5 mL collecting tube (see Note 19). 4. Measure volume and add 1.5× volumes of 100% ethanol to the tube. Mix thoroughly by pipetting (see Note 20). 5. Place up to 700 μL of sample, including any precipitate, into a miRNeasy MinElute spin column. 6. Place the loaded miRNeasy MinElute spin column in a 2 mL tube (from Qiagen kit). Close the lid and centrifuge at 10,000 × g-force for 30 s at room temperature. 7. Reload the flow-through into the miRNeasy MinElute column, close the lid, and centrifuge at 10,000 × g-force for 30 s at room temperature. 8. Discard the flow-through by pipetting. Repeat steps 5–8 until all of the sample (and flow-through) has been put through the column. 9. Add 350 μL of Qiagen RWT buffer into the column. Spin at 10,000 × g-force for 30 s at room temperature. Discard the flow-through. 10. Prepare DNase mix (see Subheading 2.3.2). Dispense 80 μL of DNase mix on every column. Incubate for 15 min at room temperature. 11. Add 500  μL of RWT buffer to the RNeasy MinElute spin column, close lid, and spin the column at 10,000 × g-force for 30 s at room temperature. 12. Pipette the flow-through and reload it into the RNeasy MinElute column for a second time. Close lid and spin the

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column at 10,000 × g-force for 30 s at room temperature. Discard the flow-through by pipetting. 13. Pipette 500  μL of Qiagen RPE buffer onto the RNeasy MinElute spin column, close lid, and centrifuge for 30 s at 10,000 × g-force at room temperature. Discard the flow-­ through by pipetting. 14. Add 500  μL of freshly prepared 80% ethanol to the RNeasy MinElute spin column, close the lid, and centrifuge for 2 min at 10,000 × g-force. Discard the flow-through and the collection tube. 15. Place the RNeasy MinElute spin column in a new 2 mL collection tube (provided with the kit). 16. Open the lid of the spin column and centrifuge at full speed (≈17,000 × g-force) for 5 min at room temperature to dry the column matrices. Discard flow-through and 2 mL collection tubes. 17. Prepare TE buffer 2 and prewarm to 60 °C. 18. Place the RNeasy MinElute spin column in a fresh 1.5 mL collecting tube, and add 16 μL of warmed TE buffer 2 directly to the center of column matrix. Close the lid gently, and centrifuge for 1 min at full speed to elute the RNA. 19. Pipette the RNA flow-through back into the column a second time, and centrifuge at full speed for another minute. 20. Collect RNA in Nunc Cryobank cryogenic vials and store at −80 °C. 3.3  RNA Quality and Quantity Measurement

All the following steps describe how to determine the quantity and quality of total RNA extracted from samples sorted into TRIzol LS (see Notes 21 and 22). 1. Thaw SuperScript III reagents on ice. Keep all reagents on ice. 2. Transfer 1.5 μL (10%) of total RNA from each sample to a Plate 1. 3. Prepare annealing mix and SuperScript III reverse transcription mix for all samples and for the standard RNA sample (see Subheading 2.3.3, Tables 2 and 3, Note 23). 4. Dispense 3.3 μL of annealing mix in sample wells containing the 1.5 μL of total RNA. 5. Place the plate for 5 min at 65 °C on a thermoblock to allow the unfolding of RNA secondary structures. 6. Immediately cool down samples using shaved ice from an −80 °C freezer for at least 1 min before proceeding to reverse transcription. These steps allows OligodT(20) primers and random hexamers to bind efficiently to unfolded RNAs.

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Table 2 Reverse transcription oligo annealing mix

Oligo Mix for 48 samples

Stock concentration

Volume for TRIzol LS or low input samples (μL)

Oligo dT (20)

50 μM

25.2

Random Hexamers

50 ng/μL

50.4

RNAse OUT

40 U/μL

15.12

Molecular grade water

88.2

totRNA per sample (μL)

1.5

All components needed to prepare the reverse transcription oligo mix are listed with stock concentrations as well as volumes needed to prepare the mix for 48 samples

Table 3 Reverse transcription mix for single-cell samples

RT Mix for 48 samples

Stock concentration

Volume for TRIzol LS and low input samples (μL)

10× First strand Buffer

10×

50.4

MgCl2

25 mM

100.8

DTT

100 mM

50.4

dNTP

10 mM

25.2

RNAse OUT

40 U/μL

15.1

SuperScript III RT

200 U/μL

25.2

Oligo bound RNA per sample (μL)

5

All components needed to prepare the reverse transcription mix for single-cell samples are listed with stock concentrations as well as volumes needed to prepare the mix for 48 samples and one standard sample

7. Dispense 5.3 μL of SuperScript III mix to both sample well and in the standard RNA well. Vortex and pulse-spin the plate. 8. Run the following program on a thermoblock to synthesize cDNA: 25 °C for 10 min, 50 °C for 50 min, 85 °C for 5 min, and 4 °C on hold. 9. Take samples out of the thermoblock, and add 15 μL of ultrapure water to every 10 μL cDNA sample. Use 2.5 μL (10%) of diluted cDNA for the β2m qPCR quantification experiment (see Notes 23 and 24).

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Table 4 qPCR master mix

qPCR Master Mix for 48 samples

Stock concentration

Volume for TRIzol LS and low input samples (μL)

SybrGreen Master mix



588

Fwd Primers

5 μM

70.6

Rev Primers

5 μM

70.6

Molecular grade water

153

cDNA template per sample (μL)

2.5

All components needed to prepare the qPCR master mix are listed with stock concentrations as well as volumes needed to prepare the mix for 48 samples and standard dilutions

10. Regarding the standard samples with known concentration, dilute cDNA to reach 6.4 ng/μL, and prepare a 7-point 1:4 standard serial dilution. Use 2.5 μL of each dilution for a qPCR reaction in a Plate 3 (see Note 23). 11. Prepare housekeeping gene qPCR mix (see Note 25 and Table 4). 12. Dispense 7.5  μL of housekeeping gene qPCR mix to standards and sample wells in a Plate 3. Vortex and pulse-spin plate. 13. Perform qPCR program: 95 °C for 10 min; 40 cycles, 95 °C for 15 s; and 65 °C for 1 min. 14. Based on Ct values, determine the RNA sample quantity and concentration, and run approximately 1 ng (but no more than 10%) of the sample on a capillary DNA-RNA electrophoresis equipment following manufacturer’s recommendations. Determine the RNA integrity number (RIN) as well as the ratios between the 28S- and 18S-rRNA peaks (see Note 26; Fig. 2a, b). 3.4  cDNA Synthesis by High-Fidelity Reverse Transcription

All the following steps are common for all samples independently of the collection method. All reactions are performed at room temperature unless otherwise noted. The following steps are adapted from the Smart-Seq2 method described by Picelli et al. [9]. For convenience, as minor changes are necessary in the procedure depending on the method chosen (bulk RNA-Seq from TRIzol LS, bulk-low input in LI-LB, or single-cell RNA-Seq), we have clearly annotated these changes to the appropriate method throughout the steps (see Note 27).

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a 60 Fluorescence (a.u.)

40

18S rRNA 5S & small RNAs

28S rRNA RIN : 9.5 28S/18S ratio:1.7

20 0

20

0 25

b

Good RIN>7.5 28S/18S ratio>1

Degraded RNA Bad RIN : 1 28S/18S ratio:0.05

10

200

1000 4000 Size (bp)

512

128

R2(>5000cells) =0.85 20 %

Total

b

Unique

> 20 % > 20 %

mRNA Intronic Intergenic

Total

Housekeeping genes B2M

GAPDH

Unique

mRNA Intronic Intergenic

CD4 Naive T cells phenotypic genes

YWHAZ

SELL SELE

CD4

GATA3

Set2

Set1

mRNA expression (rpkm)

Chr 15

c

1.6 kb Chr 12

Housekeeping genes ACTB

7.9 kb Chr 1

7.2 kb Chr 12

1.3 kb Chr 10

5.0 kb

CD8 T cells phenotypic genes IL2RG CXCR4

CD52

mRNA expression (rpkm)

20 single cell RNA-Seq

B2M

1 kb Chr 8

Chr 15

0.4 kb Chr 7

0.2 kb Chr 1

0.1 kb Chr 2

0.2 kb Chr X

0.5 kb

Fig. 5 Quality control post-sequencing. (a) Graphs show different types of mappability percentage for around 10–15 million single-ended 50 bp sequencing reads to the reference genome (hg19) for two independent sets of bulk RNA-Seq libraries (left) and single-cell RNA-Seq (right). Percentage rates shown are for a total number of reads (total), for reads mapping only to one and unique genomic location, reads mapping to coding regions (mRNA), reads mapping to intronic regions, reads mapping to intergenic regions compared to the total number of reads passing sequencing filters. The red dashed lines indicate the threshold values for elimination. (b and c) UCSC genome browser tracks showing sequencing RNA profiles along different gene loci, for two independent sets of 8 bulk CD4 T cell RNA-Seq libraries (b) and 20 CD8 T cells single-cell RNA-Seq (at higher resolution) (c). Rpkm, reads per kilobase per million mapped

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46. We usually perform 50 bp single-end read sequencing and aim to generate 15 million mapping reads per sample. This protocol achieves greater than 80% mapping to reference genome and uniquely mapped reads. As an example, Fig. 5a shows the fraction of reads that map with coding, intronic, and intergenic regions. Figure 5b, c shows example of consistency of RNA-Seq results achieved with this procedure. It illustrates read profiles distribution along different gene loci for two independent sets of bulk RNA-Seq libraries for CD4 T cells (b) and 20 CD8 T single-cell RNA-Seq (at higher resolution). UCSC genome browser tracks; Rpkm, reads per kilobase per million mapped. References 1. Costa V, Aprile M, Esposito R, Ciccodicola A (2013) RNA-Seq and human complex diseases: recent accomplishments and future perspectives. Eur J Hum Genet 21(2):134–142. https://doi.org/10.1038/ejhg.2012.129 2. Seumois G, Zapardiel-Gonzalo J, White B, Singh D, Schulten V, Dillon M, Hinz D, Broide DH, Sette A, Peters B, Vijayanand P (2016) Transcriptional profiling of Th2 cells identifies pathogenic features associated with asthma. J Immunol 197(2):655–664. https:// doi.org/10.4049/jimmunol.1600397 3. Engel I, Seumois G, Chavez L, Samaniego-­ Castruita D, White B, Chawla A, Mock D, Vijayanand P, Kronenberg M (2016) Innate-­ like functions of natural killer T cell subsets result from highly divergent gene programs. Nat Immunol 17(6):728–739. https://doi. org/10.1038/ni.3437 4. Byron SA, Van Keuren-Jensen KR, Engelthaler DM, Carpten JD, Craig DW (2016) Translating RNA sequencing into clinical diagnostics: opportunities and challenges. Nat Rev Genet 17(5):257–271. https://doi.org/10.1038/ nrg.2016.10 5. Zhao Y, Simon R (2010) Gene expression deconvolution in clinical samples. Genome Med 2(12):93. https://doi.org/10.1186/gm214 6. Cai C, Langfelder P, Fuller TF, Oldham MC, Luo R, van den Berg LH, Ophoff RA, Horvath S (2010) Is human blood a good surrogate for brain tissue in transcriptional studies? BMC Genomics 11:589. https://doi. org/10.1186/1471-2164-11-589 7. Mohamad-Fauzi N, Ross PJ, Maga EA, Murray JD (2015) Impact of source tissue and ex vivo

expansion on the characterization of goat mesenchymal stem cells. J Anim Sci Biotechnol 6(1):1. https://doi. org/10.1186/2049-1891-6-1 8. Proserpio V, Lonnberg T (2016) Cutting-­edge single-cell genomics and modelling in immunology. Immunol Cell Biol 94(3):224. https://doi.org/10.1038/icb.2015.117 9. Picelli S, Faridani OR, Bjorklund AK, Winberg G, Sagasser S, Sandberg R (2014) Full-length RNA-seq from single cells using Smart-seq2. Nat Protoc 9(1):171–181. https://doi. org/10.1038/nprot.2014.006 10. Ziegenhain C, Vieth B, Parekh S, Reinius B, Guillaumet-Adkins A, Smets M, Leonhardt H, Heyn H, Hellmann I, Enard W (2017) Comparative analysis of single-cell RNA sequencing methods. Mol Cell 65(4):631– 643. e634. https://doi.org/10.1016/j. molcel.2017.01.023 11. Seumois G, Vijayanand P, Eisley CJ, Omran N, Kalinke L, North M, Ganesan AP, Simpson LJ, Hunkapiller N, Moltzahn F, Woodruff PG, Fahy JV, Erle DJ, Djukanovic R, Blelloch R, Ansel KM (2012) An integrated nano-scale approach to profile miRNAs in limited clinical samples. Am J Clin Exp Immunol 1(2):70–89 12. Wright ES, Vetsigian KH (2016) Quality filtering of Illumina index reads mitigates sample cross-talk. BMC Genomics 17(1):876. https://doi.org/10.1186/ s12864-016-3217-x 13. Illumina (2017) Effects of index misassignment on multiplexing and downstream analysis. Illumina Technical Note

Chapter 22 An Integrated and Semiautomated Microscaled Approach to Profile Cis-Regulatory Elements by Histone Modification ChIP-Seq for Large-Scale Epigenetic Studies Diana Youhanna Jankeel, Justin Cayford, Benjamin Joachim Schmiedel, Pandurangan Vijayanand, and Grégory Seumois Abstract Chromatin immunoprecipitation followed by sequencing (ChIP-Seq) is the preferred approach to map histone modifications and identify cis-regulatory DNA elements throughout the genome. Multiple methods have been described to increase the efficiency of library preparation and to reduce hands-on time as well as costs. This review describes detailed steps to perform cell fixation, chromatin shearing, immunoprecipitation, and sequencing library preparation for a batch of 48–96 samples with small cell numbers. The protocol implements a semiautomated platform to reduce technical variability and improve signal-to-noise ratio as well as reduce hands-on time, thus allowing large-scale epigenetic studies of clinical samples with limited cell numbers. Key words ChIP-Seq, H3K27ac, IP-Star, Tagmentation

1  Introduction Genome-wide profiling of histone modifications in DNA regions by chromatin immunoprecipitation followed by sequencing (ChIP-­ Seq) represents the preferred method to identify cis-regulatory DNA elements (active enhancers, promoters, silencers, insulators) that are playing important roles in gene regulation and cellular development [1–4]. However, large-scale ChIP-Seq experiments in clinical samples, besides from being technically challenging, are often limited by the quantity of cells or tissue of interest [5]. In addition, the heterogeneous cellular composition of clinical samples can confound analyses and mask significant changes in gene regulation. To overcome these hurdles, a number of microscaling

Diana Youhanna Jankeel and Justin Cayford contributed equally to this work. R. Lee Reinhardt (ed.), Type 2 Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1799, https://doi.org/10.1007/978-1-4939-7896-0_22, © Springer Science+Business Media, LLC, part of Springer Nature 2018

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techniques have been reported [5–7]. We developed a sensitive and robust microscaled ChIP-Seq assay to profile histone modification marks for as little as 10,000 cells [8, 9]. We applied the method to a translational research project in which we profiled H3K4me2 marks in three types of circulating CD4+ T cells (naive, TH1 and TH2 memory T cells) directly isolated from blood samples of a cohort of healthy individuals and asthmatic patients. Looking at epigenetic changes between cell types and disease groups, we have identified a number of new active and poised promoters and enhancers, new potential transcription factor binding sites, and functional SNPs that could play a role in T cell development and asthma pathogenesis [9, 10]. In murine cells, we used H3K27ac-­ ChIP-­Seq assay along with RNA-Seq to characterize the different subtypes of developing NKT cells present in the thymus [8]. More recently, a new method called “ChIPmentation” has been described by Schmidl et al. [11]. It combines chromatin immunoprecipitation with a single-step integration of sequencing adaptors using Tn5 transposase technology, increasing library preparation efficiency and reducing hands-on time. In this review, we detail an integrated, high-sensitive, and semiautomated approach to perform every step of the procedure (cell fixation, chromatin shearing, immunoprecipitation, and library preparation) for up to 48 samples with very low cell numbers (10,000–100,000 cells). This approach reduces technical variability and hands-on time and thus is suited for large-scale epigenetic studies. Figure 1a displays an overview of the entire procedure. It describes all critical steps, quality control (QC) steps, and an estimation of hands-on time. For clarity purposes, this chapter was subdivided into six subsections corresponding to cells fixation, chromatin shearing, chromatin immunoprecipitation, library preparation by tagmentation of DNA fragments, amplification with barcoded adaptors, library purification and size selection, as well as the description of DNA quantification using a PicoGreen assay. For each section, materials and methods are described.

2  Materials 2.1  Cell Fixation (See Note 1)

1. 37% formaldehyde. 2. 5 M NaCl*. 3. 0.5 M EDTA pH 8.0*. 4. 0.5 M EGTA pH 8.0*. 5. 1 M HEPES pH 7.5*. 6. 2.5 M glycine*. 7. Phosphate-buffered saline (PBS) pH 7.5.

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8. Complete cell culture medium pH 7.5: Dulbecco’s Modified Eagle’s Medium (DMEM) complemented with 5% fetal bovine serum and 2% human serum (for human cells). 9. Short-term 10× cell fixation buffer: 11% formaldehyde solution, 100 mM NaCl, 1 mM EDTA, 0.5 mM EGTA, 50 mM HEPES, pH 7.5 completed with nuclease-free ultrapure water. Store at room temperature. 10. 1.5 mL Axygen Maxymum recovery tubes. 11. Rotating platform. 12. Liquid nitrogen. 2.2  Chromatin Shearing by Sonication (See Note 1)

1. Dry ice. 2. 1 M Tris–HCl pH 8.0*. 3. 0.5 M EDTA pH 8.0*. 4. 10% sodium dodecyl sulfate (SDS)*. 5. 1 M sodium butyrate (NaBu). 6. Protease Inhibitor Cocktail 200× (PI) (Sigma Aldrich). 7. Short-term complete lysis buffer: 50 mM Tris–HCl pH 8.0, 10 mM EDTA, 0.25% SDS in nuclease-free ultrapure water. Right before use, add 1 M NaBu (to reach 20 mM) and 200× Protease Inhibitor Cocktail (to reach 1×). 8. 0.6 mL Axygen Maxymum recovery tubes. 9. Bioruptor Pico. 10. 0.2 mL 8-tube strips. 11. PureLink RNase A 20 mg/mL (Invitrogen). 12. Proteinase K Solution 20 mg/mL, RNA grade (Thermo Fisher Scientific). 13. Thermomixer. 14. Agarose. 15. 50× Tris–acetate–EDTA (TAE) buffer*: 2 M Tris base, 1 M acetate, and 50 mM EDTA, pH 8.5–9.0. 16. 6× loading dye buffer. 17. SYBR Gold nucleic acid gel stain (Invitrogen). 18. Electrophoresis display. 19. UV transilluminator with a camera. 20. 0.5 mL Nunc Cryobank vials (Thermo Fisher Scientific).

2.3  Automated Chromatin Immunoprecipitation for Histone Modifications

All buffers for this series of steps are obtained from the True MicroChIP Kit (Diagenode): 1. tC1 buffer.

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2. Complete tC1 buffer: right before use, add 20 μL of protease inhibitor cocktail 200× and 80 μL of 1 M NaBu to 4 mL of tC1 buffer. 3. tBW1 buffer. 4. tW1 buffer. 5. tW2 buffer. 6. tW3 buffer. 7. tW4 buffer. 8. tE1 buffer. 9. Protease inhibitor (PI) 200× (Sigma Aldrich). 10. 1 M sodium butyrate (NaBu). 11. ChIP grade antibody: for example, H3K27ac (Diagenode). 12. 30 mg/mL Protein A Dynabeads (Invitrogen). 13. 200  μL 8-tube strips + cap strips for SX-8G IP-Star (Diagenode). 14. 2 mL microtubes for SX-8G IP-Star Compact (Diagenode). 15. Medium reagent containers for SX-8G IP-Star Compact (Diagenode). 16. Tips (bulk) for SX-8G IP-Star (Diagenode). 17. SX-8G IP-Star Compact automated platform (Diagenode). 18. 8-channel micropipette (10–100 μL). 2.4  Library Preparation by Transposase Integration of Illumina Library Adaptors (See Note 1)

1. 1 M Tris–HCl pH 8.0*. 2. 25 mM MgCl2*. 3. N,N-dimethylformamide. 4. DNA tagmentation enzyme from Nextera Kit (Illumina). 5. Short-term tagmentation buffer: 10 mM Tris–HCl pH 8.0, 5 mM MgCl, 10% N,N-dimethylformamide, 1:24 (vol:vol) of DNA tagmentation enzyme. Keep on the ice. 6. 8-tube strip magnet (Diagenode). 7. Metallic 96-well rack. 8. ChIP Buffer tC1 from True MicroChIP Kit (Diagenode). 9. Tips (bulk) for SX-8G IP-Star (Diagenode). 10. 200  μL 8-tube strips + cap strips for SX-8G IP-Star Compact (Diagenode). 11. 2 mL microtubes for SX-8G IP-Star Compact (Diagenode). 12. SX-8G IP-Star Compact automated platform (Diagenode). 13. PureLink RNase A 20 mg/mL (Thermo Fisher Scientific). 14. Proteinase K Solution 20 mg/mL, RNA grade (Thermo Fisher Scientific).

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15. Thermomixer. 16. 8-channel multichannel pipette (10 to 100 μL). 2.5  Purification and Amplification of the Tagmented DNA Fragments (See Note 1)

1. 8-tube strips magnet (Diagenode). 2. ChIP DNA Clean & Concentrator column-based Kit (Zymo Research), including washing solution to be reconstituted with ethanol, 200-proof. 3. Ethanol, 200-proof, anhydrous 99.5%. 4. 1.5 mL collection tubes. 5. 1 M Tris–HCl pH 8.0*. 6. 0.5 M EDTA pH 8.0*. 7. 10× Kapa HiFi HotStart Ready Mix (Kapa Biosystems). 8. SYBR Green dye. 9. 50× Rox dye. 10. Nextera index primers kit (Illumina). 11. Common CtD reaction mix, per sample: 0.275 μL of 2 Nextera index primers (25 μM), 2.75 μL of preheated 2× KAPA HiFi HotStart Ready Mix, 0.11 μL of 1:1000 diluted SYBR Green dye, and 0.11 μL of ROX passive dye. Complete the volume to 4 μL with nuclease-free water. 12. Common Amp reaction mix, per sample: 27.5 μL of preheated 2× KAPA HiFi Hot start ready mix, and complete the volume to 31 μL with nuclease-free water. Then add 2.5 μL of 2 Nextera index primers (25 μM) to each sample. 13. 0.2 mL PCR 8-tube strips with individual cap. 14. qPCR plate and seal: 96-well PCR plate, low profile, skirted; qPCR MicroAmp Optical Adhesive Film (Life Technologies). 15. TE buffer 1: 1 mM Tris–HCl pH 8.0 and 1 μM EDTA (see Note 2)*. 16. Thermomixer. 17. Real-time quantitative PCR system.

2.6  Purification of DNA Post-­ Amplification, Size Selection, and Quantification (See Note 1)

1. AMPure XP beads solution (Beckman Coulter). 2. Magnet for 96-well plate (Axygen). 3. Ethanol washing solution: 100 mL of 80% ethanol solution. 4. Plate 1: 96-well semi-skirted PCR plates (BioRad), higher volume capacity. 5. Plate 2: 96-well hard-shell thin-wall 96-well skirted PCR plates. 6. MicroAmp Clear Adhesive Film (Life Technologies). 7. 1 M Tris–HCl pH 8.0*.

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8. 0.5 M EDTA pH 8.0*. 9. TE buffer 1: 1 mM Tris–HCl pH 8.0, 1 μM EDTA (see Note 2)*. 10. TE buffer 2: 10 mM Tris–HCl pH 8.0, 1 μM EDTA (see Note 2)*. 11. 40 ng/μL of sonicated standard DNA: sonicate Lambda phage DNA (Life Technologies) to obtain fragments at length ranging between 300 and 600 bp. Aliquot and store at −20 °C. 12. Quant-iT PicoGreen dsDNA Reagent (Thermo Fisher Scientific). 13. 96-well plate, flat bottom, black for fluorescence measurements.

3  Methods 3.1  Cell Fixation

To perform good-quality ChIP experiments, DNA and histones need to be cross-linked using formaldehyde. The following steps describe how cells are fixed in a 1% formaldehyde solution, washed, and spun to obtain a cell pellet that can be subsequently snap frozen in liquid nitrogen and stored at −80 °C for up to a year (see Fig. 1a and Note 3). The following steps of this procedure occur at room temperature. 1. Bring cell suspension concentration to 1–2 × 106 cells/mL of complete cell culture medium in either a 15 mL tube if less than 10 mL of cell suspension, or a 50 mL tube for 10–30 mL per tube. If less than 1 million cells, use 0.5 mL of complete cell culture medium in a 1.5 mL tube. 2. Prepare the appropriate amount of 10× cell fixation buffer: total volume of cell suspension (at 1–2 million cells per mL)/10. 3. Place the 2.5 M glycine solution at room temperature and PBS on ice, and have a large bucket of ice to accommodate all tubes after fixation. Also, prepare a container with liquid nitrogen (see Note 4). 4. Vortex cell suspension at medium speed and add, drop by drop, 1:10 (vol:vol) of 10× cell fixation buffer. 5. Place tubes on a rotating platform at low rpm, and incubate the tubes for 10 min at room temperature (see Note 5). 6. After incubation, vortex the tubes at medium speed, and stop the reaction by adding 1:20 (vol:vol) of 2.5 M glycine solution, invert the tubes twice, and place them on ice for at least 5 min. Perform the following steps at 4 °C or on ice. 7. Spin tubes at 800 × g-force for 5 min at 4 °C, discard supernatant, and resuspend the pellets with 5 mL of ice-cold PBS. Incubate on ice for 2 min.

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Automated Multi-Sample Micro-Scaled ChIP-Seq Assay

a Sample Collection and Processing

Sonication

Preparation of cells

ChIP & Library Preparation

Cell lysis using 0.25% SDS buffer

FACS Sorting

Day 1 Place the chromatin Hands on (500ng), beads, time: 1.5 and Ab in the hours IPStar machine

Sonication using Bioruptor

DNA Tagmentation

Cell Fixation Checking fragment size using DNA gel electrophoresis

Hands on time: 3 hours Total time: 5 hours

Washes using IP Star machine

Day 2 Hands on time: 3.5 hours

DNA decrosslinking Check concentration using PicoGreen

Store Chromatin at -80 °C

DNA purification using Zymo columns

Hands on time: 3 hours

Perform CtD

Total time: 6 hours

b

Amplification

48 ChIP Samples/week (3 ChIP runs of 16 samples per run) for sheared chromatin Mon

Tue

Wed

Start Day1 of ChIP A

Start Day2 of ChIP A

Start Day3 of ChIP A

Start Day1 of ChIP B

Start Day2 of ChIP B

Start Day3 of ChIP B

Start Day1 of ChIP C

Start Day2 Start Day3 of ChIP C of ChIP C

Start Day4 of ChIP A/ B/C (could be done on Fri too) Total Hands on Time: 6.5 hours

5 hours

7.5 hours

Thu

Day 3 Hands on time: 2.5 hours

Store samples at -20 °C

Fri Purify samples using AmpureXP beads

6 hours

Size selection using Ampure XP beads

2.5 hours

Quantification & median size analysis

Day 4 Hands on time:5 hours

Pooling and sequenicng

Fig. 1 Overview of the method. (a) Flow chart connecting all major steps of the procedure including quality controls and timing for easy planning of the experiment. (b) Diagram illustrates ChIP-tagmentation procedure schedule for 48 samples over a week. The sequencing would take place on the following week

8. Spin tubes at 800 × g-force for 5 min at 4 °C, discard the supernatant, and carefully resuspend the pellets with 1 mL of ice-cold PBS. Transfer sample to precooled 1.5 mL tubes (see Note 6). 9. Spin the tubes for 5 min at 1200 × g-force at 4 °C, and remove as much of the supernatant as possible without affecting the integrity of the cell pellet.

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10. Snap freeze the pellets in liquid nitrogen, and store in −80 °C freezer (see Note 7). 3.2  Chromatin Shearing

This protocol is set up for the preparation of pellets containing 0.3–3 million cells (see Note 8). It is optimized for the use of the Bioruptor Pico (Diagenode) (see Note 9). 1. About 20 min before starting the protocol, switch the Bioruptor on to cool the water to 4 °C. 2. Pre-warm the sonicator unit by performing 3 cycles of 16 s ON/32 s OFF twice using balancing tubes only (see Note 10). 3. Take out the pellets from the freezer and keep them on dry ice. Do not allow thawing of the pellet before adding the lysis buffer (see Note 11). 4. Add 70 μL of short-term complete lysis buffer, kept at room temperature (RT), to the pellet, and allow it to thaw for 1 min. 5. Carefully resuspend the pellet for 1 min. Use a 200 μL tip keeping the end of the tip very close to the bottom of the tube to create pressure on cell flow (see Note 12). 6. Allow cells to lyse for 1 more minute at room temperature, and then put the sample on ice. From this step, keep the samples on ice (or at 4 °C). 7. To proceed with the sonication, place the samples symmetrically into the tube holder, and fill any gaps with balancing tubes. 8. Place the samples on the rack in the chilled water bath, and let them incubate for 1 min. 9. Perform sonication for x cycles (depending on cell type) with the settings 16s ON/32s OFF (see Note 13). 10. Take the tube holder out of sonicator after every three cycles, and place it on ice. 11. Carefully vortex and pulse-spin the tubes to collect the samples at the bottom of the tubes (see Note 14). 12. Spin the samples at maximum speed (>14,000 × g-force) for 15 min at 4 °C (see Note 15). 13. Transfer supernatant (approx. 70 μL) into fresh low-binding 0.6 mL tubes and keep on ice. 14. To assess the sonication efficiency, take out 1–7 μL (up to 10% of total volume) of supernatant from the sonicated samples (see Note 16), and transfer to fresh 0.2 mL PCR tubes, called QC tubes. 15. Make up the volume to 10 μL in QC tubes with short-term complete lysis buffer, and add 1 μL of RNase A. Incubate the sample at 37 °C for 30 min on a thermomixer with shaking (800 rpm).

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16. Add 1  μL of proteinase K to QC tubes containing 11 μL sample, and incubate at 65 °C for 2 h on a thermomixer with shaking (800 rpm). 17. Take out 2 μL of the decrosslinked sample from the QC tubes for quantification by PicoGreen assay (see Subheading 3.6, steps 31–41). 18. Determine the chromatin concentration for each sample. Based on the initial cell numbers, estimate sonication efficiency (see Note 17). 19. Mix the rest of the sample (10 μL) with 2 μL of 6× loading dye buffer, and load the sample on a 1.2% agarose—1× TAE gel. Run electrophoresis for 1 h at 70 V in 1× TAE buffer. Stain the gel with SYBR Gold dye (1:20,000) in 1× TAE buffer for 20 min, wash it twice with 1× TAE buffer for 10 min, and read it using a UV transilluminator. 20. If quantity results given by PicoGreen measurements and gel analysis indicate successful sonication, then proceed with the preparation of chromatin stocks aliquots for storage (see Fig. 2b). 21. Spin the sample tube again at maximum speed for 15 min at 4 °C. 22. Measure the volume using the pipette, and dilute the samples to set the chromatin concentration to 25 ng/μL (see Note 18). 23. Store all aliquots of sheared chromatin at −80 °C. 3.3  Automated Chromatin Immunoprecipitation for Histone Modifications

This protocol is designed to use the automated ancillary liquid handler SX-8G IP-Star from Diagenode (see Fig. 1a, b and Note 19). Every ChIP reaction sample will contain 500 ng chromatin (20  μL at 25 ng/μL) of sheared DNA equivalent to around 100,000 cells (see Note 19). The following steps describe the preparation of the different 8-tube strips required to set up the automated platform as illustrated in Fig. 3a. 1. Take out 16 chromatin aliquots from −80 °C freezer, and place them on ice to allow the chromatin to thaw slowly. After the chromatin tubes are thawed, vortex them briefly and pulse-spin. 2. Take two 8-tube strips and label them appropriately. To avoid a mix-up in samples, color (or number) the left side and right sides of both strips (see Note 20). 3. Chromatin 8-tube strips preparation: Pipette 100 μL of complete tC1 buffer (supplemented with protease inhibitor and NaBu) into each tube of two 0.2 mL 8-tube PCR strips. 4. Transfer 20 μL of each chromatin sample to the 8-tube strips containing 100 μL of complete tC1 buffer.

Diana Youhanna Jankeel et al.

a

b

Number of cycles

70%

100

B Cell line

500

DL 1 2 3 4 5 6 7 8 9 10 11 12 DL

500

500 100

100

c

Number of samples (12 cycles)

100

10%

128

Fluoresence

c

6

64 R2=0.62

32 16

0

8 6 2 4 pre-amplification (ng)

½ intensity

8 10 12 14 16 18 20 22

≈15 + 1,2Ct

Large fraction

Fluorescence (a.u.)

post-amplification (ng)

b

Percentage of PSS (%)

d

back

Log2 quamntity (ng)

a

Final library 85% 65%) are not located between 100 and 500 bp (see Fig. 2b). Remediation steps will be to sonicate for a few more cycles. Repeat the QC steps as described. Alternatively, repeat the sonication from a new pellet. 18. Based on the results from PicoGreen, we suggest diluting the sample to 25 ng/μL (= 500 ng in 20 μL) with complete lysis buffer (containing proteinase inhibitors and NaBu). We suggest that for each sample, aliquot the chromatin into 3 (or more) labeled Nunc storage cryovials with 20 μL each; pipette any leftovers into an additional Nunc storage cryovial. 19. We optimized the procedure and labor workflow for up to 48 samples organized into three rounds of 16 ChIP reactions per week. For technical purposes, in context of large-scale project with more than 16 samples, we suggest to run 14 samples (100,000 cells = 500 ng each = 20 μL aliquot), 1 technical duplicate control (a second aliquot of 20 μL), and 1 permanent sample used for all rounds of ChIP to control for batch effects (e.g., chromatin from a related cell line). We estimate this step will require 1.5 h of hands-on time. 20. Color code suggestion: 1–8 and 9–16; color the left side of the first strip tube blue (#1), the right side of both strip tubes in red (#8 and 16), and the left side of the second strip tube green (#9); this will reduce frequent samples mix-ups. 21. Optional: if planning to use INPUT (non IP’d samples counterparts): add 90 μL of complete tC1, and take out 10 μL of

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sample. Store at −20 °C. Proceed to Subheading 3.4 step 12 (see Note 31). 22. Numbers are indicative; the concentration of antibody per ChIP experiment has to be validated by specific titration experiments. 23. We consider that all samples receive the same amount of antibody; if planned differently, then the volumes have to be recalculated accordingly. 24. tW4 needs to be added to position “A” instead of tE1 buffer (to avoid elution from the beads before tagmentation reaction). The program has the following settings: Ab coating, 2 h, 4 °C, middle; IP reaction, 10 h, 4 °C, middle; washes, 5 min, 4 °C, middle. This program takes around 18 h to complete. 25. The hands-on estimated time is 1 h and a total time of around 3 h for the completion of the experiment (see Fig. 1a, b). 26. The N,N-dimethylformamide is a very unstable reagent that has to be kept in the flammable/hazardous cabinet at room temperature. Handle with care! 27. Some of the beads can stick to the side of the tubes, don’t mix, pipette slowly out the supernatant from the bottom of the tubes. 28. Tagmentation is a very time-sensitive reaction; even if we controlled that the tagmentation reaction was not happening at 4 °C, it is reasonable to act promptly. 29. We have validated the tagmentation time and set it up optimally for 3 min as others have also shown [11]. 30. Select the wash program in IP-Star (3-min washes). The buffers used are tE1 in position A, tC1 buffer in positions C&D, and tW4 in positions E&F—this program takes 2 h and 18 min to complete. The machine only has an 8 samples version; contact Diagenode to get the 16 samples. 31. Optional: if planning to use INPUT (non IP’d samples counterparts), add 90 μL of tE1 to the 10 μL inputs samples and then follow steps. 32. The hands-on estimated time is 3 h; the entire procedure will take around 5 h. 33. Others suggested to using AMPure XP beads for this cleanup step [11]. However, in our hands we observed a lot of variability and loss of material; hence we suggest using a column-­ based purification. 34. We do not know what is the composition of the elution buffer included in the kit; we therefore suggest to use out TE buffer 1 that will stabilize DNA fragment for storage but won’t interfere with downstream steps.

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35. Use a unique pair of indices for all samples; those samples will not go through sequencing. 36. All volumes are increased by 1.1 to account for volume error during dispensing. 37. Optional: Add a melt curve. We use the qPCR StepOne equipment from Life Technologies, and it requires the use of a passive dye to normalize fluorescence signals. 38. All combinations of index primer pairs have to be unique for sequencing purposes; choose wisely, respect color balance, and refer to Illumina website. 39. Amplified product can be stored for a few days at −20 °C without DNA purification. 40. The hands-on estimated time is 4 h for a total of a 6-h procedure. The following steps are written for a batch of 3 × 16 set of samples. If there are not multiple rounds of ChIP planned, then those steps can be completed on the same day without storage at −20 °C. 41. The plate being used needs to have wells with larger capacities (at least 300 μL) instead of the regular 0.2 mL PCR plate. 42. We advise to perform those steps during early stages; after becoming familiar with the method, those steps could be abandoned. 43. During this step, the fragments >1000 bp will be captured on beads; smaller fragments will stay in solution as shown in Fig. 3. AMPure XP beads to samples volume ratios have to be validated. 44. Now, all wells in the plate should have a bead pellet. 45. At this step, only the fragments with a size length ranging between 200 and 1000 bp will be captured with the beads. The smaller fragments will stay in solution and be discarded, as shown in Fig. 3. 46. Plates can be stored at −20 or −80 °C for long-term storage. 47. We suggest 28 μL to account for pipetting variance when performing the downstream steps. 48. The volumes shown are for one PicoGreen 96-well plate as shown in Fig. 2d; quantity for 24 samples will be measured per plate. For a regular set of 96 samples, you will have to prepare 4 PicoGreen plates. For consistency, it’s best to prepare one series of DNA standard points for all plates. 49. This is equivalent to 2 μL of dye per mL of TE buffer 2. 50. It is very important to add the drop of samples in the liquid and not on the side the wells; the measurement can be affected if so. You can protect the plate from light with aluminum foil (recommended).

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Chr 15

PATL2 B2M

Chr 10

IL15RA

IL2RA

Chr 12

GAPDH (locus)

Batch 2

H3K27ac Enrichment (rpkm)

Batch 1

Genes

25 kb

11 kb

30 kb

Fig. 4 Example of consistency of the method. Examples of consistency for H3K27ac ChIP-Seq enrichment for two different batches of 10 CD4 T cells samples (100,000 cells ChIP-tagmentation each) along different genomic coordinates corresponding to three gene loci with UCSC genome tracks. rpkm, reads per kilobase per million mapped

51. We should expect no more than 20% loss. If more than a 20% loss is measured, then verify your bead ratios or elution steps. 52. We usually perform 50 bp single-end read sequencing and aim to generate 15 million mapping reads per sample. Figure 4 shows as an example of consistency for H3K27ac ChIP-Seq enrichment for two different batches of 10 CD4 T cells samples (100,000 cells ChIP-tagmentation) along different genomic coordinates corresponding to three gene loci with UCSC genome tracks, rpkm: reads per kilobase per million mapped.

Acknowledgments We thank the Vijayanand lab members for technical help and constructive discussions and Dr. Sharron Squazzo from Diagenode for technical assistance with the Bioruptor Pico and SX-8G IP-Star Compact machine and protocols. This work was supported by NIH grants (P.V.): NIH R24 AI108564, NIH U19 AI118626, NIH R01 HL114093, NIH R01 AI121426 and NIH S10 OD016262S10.

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References 1. Andersson R, Gebhard C, Miguel-Escalada I, Hoof I, Bornholdt J, Boyd M, Chen Y, Zhao X, Schmidl C, Suzuki T, Ntini E, Arner E, Valen E, Li K, Schwarzfischer L, Glatz D, Raithel J, Lilje B, Rapin N, Bagger FO, Jorgensen M, Andersen PR, Bertin N, Rackham O, Burroughs AM, Baillie JK, Ishizu Y, Shimizu Y, Furuhata E, Maeda S, Negishi Y, Mungall CJ, Meehan TF, Lassmann T, Itoh M, Kawaji H, Kondo N, Kawai J, Lennartsson A, Daub CO, Heutink P, Hume DA, Jensen TH, Suzuki H, Hayashizaki Y, Muller F, Forrest ARR, Carninci P, Rehli M, Sandelin A (2014) An atlas of active enhancers across human cell types and tissues. Nature 507(7493):455–461. https://doi.org/10.1038/nature12787 2. Creyghton MP, Cheng AW, Welstead GG, Kooistra T, Carey BW, Steine EJ, Hanna J, Lodato MA, Frampton GM, Sharp PA, Boyer LA, Young RA, Jaenisch R (2010) Histone H3K27ac separates active from poised enhancers and predicts developmental state. Proc Natl Acad Sci U S A 107(50):21931–21936. https://doi.org/10.1073/pnas.1016071107 3. Ernst J, Kheradpour P, Mikkelsen TS, Shoresh N, Ward LD, Epstein CB, Zhang X, Wang L, Issner R, Coyne M, Ku M, Durham T, Kellis M, Bernstein BE (2011) Mapping and analysis of chromatin state dynamics in nine human cell types. Nature 473(7345):43–49. https://doi. org/10.1038/nature09906 4. Roadmap Epigenomics C, Kundaje A, Meuleman W, Ernst J, Bilenky M, Yen A, Heravi-Moussavi A, Kheradpour P, Zhang Z, Wang J, Ziller MJ, Amin V, Whitaker JW, Schultz MD, Ward LD, Sarkar A, Quon G, Sandstrom RS, Eaton ML, Wu YC, Pfenning AR, Wang X, Claussnitzer M, Liu Y, Coarfa C, Harris RA, Shoresh N, Epstein CB, Gjoneska E, Leung D, Xie W, Hawkins RD, Lister R, Hong C, Gascard P, Mungall AJ, Moore R, Chuah E, Tam A, Canfield TK, Hansen RS, Kaul R, Sabo PJ, Bansal MS, Carles A, Dixon JR, Farh KH, Feizi S, Karlic R, Kim AR, Kulkarni A, Li D, Lowdon R, Elliott G, Mercer TR, Neph SJ, Onuchic V, Polak P, Rajagopal N, Ray P, Sallari RC, Siebenthall KT, SinnottArmstrong NA, Stevens M, Thurman RE, Wu J, Zhang B, Zhou X, Beaudet AE, Boyer LA, De Jager PL, Farnham PJ, Fisher SJ, Haussler D, Jones SJ, Li W, Marra MA, McManus MT, Sunyaev S, Thomson JA, Tlsty TD, Tsai LH, Wang W, Waterland RA, Zhang MQ, Chadwick LH, Bernstein BE, Costello JF, Ecker JR, Hirst M, Meissner A, Milosavljevic A, Ren B,

Stamatoyannopoulos JA, Wang T, Kellis M (2015) Integrative analysis of 111 reference human epigenomes. Nature 518(7539):317– 330. https://doi.org/10.1038/nature14248 5. Furey TS (2012) ChIP-seq and beyond: new and improved methodologies to detect and characterize protein-DNA interactions. Nat Rev Genet 13(12):840–852. https://doi. org/10.1038/nrg3306 6. Dahl JA, Collas P (2008) A rapid micro chromatin immunoprecipitation assay (microChIP). Nat Protoc 3(6):1032–1045. https:// doi.org/10.1038/nprot.2008.68 7. van Galen P, Viny AD, Ram O, Ryan RJ, Cotton MJ, Donohue L, Sievers C, Drier Y, Liau BB, Gillespie SM, Carroll KM, Cross MB, Levine RL, Bernstein BE (2016) A multiplexed system for quantitative comparisons of chromatin landscapes. Mol Cell 61(1):170–180. https://doi.org/10.1016/j. molcel.2015.11.003 8. Engel I, Seumois G, Chavez L, Samaniego-­ Castruita D, White B, Chawla A, Mock D, Vijayanand P, Kronenberg M (2016) Innate-­ like functions of natural killer T cell subsets result from highly divergent gene programs. Nat Immunol 17(6):728–739. https://doi. org/10.1038/ni.3437 9. Seumois G, Chavez L, Gerasimova A, Lienhard M, Omran N, Kalinke L, Vedanayagam M, Ganesan AP, Chawla A, Djukanovic R, Ansel KM, Peters B, Rao A, Vijayanand P (2014) Epigenomic analysis of primary human T cells reveals enhancers associated with TH2 memory cell differentiation and asthma susceptibility. Nat Immunol 15(8):777–788. https:// doi.org/10.1038/ni.2937 10. Schmiedel BJ, Seumois G, Samaniego-­ Castruita D, Cayford J, Schulten V, Chavez L, Ay F, Sette A, Peters B, Vijayanand P (2016) 17q21 asthma-risk variants switch CTCF binding and regulate IL-2 production by T cells. Nat Commun 7:13426 11. Schmidl C, Rendeiro AF, Sheffield NC, Bock C (2015) ChIPmentation: fast, robust, lowinput ChIP-seq for histones and transcription factors. Nat Methods 12(10):963–965. https://doi.org/10.1038/nmeth.3542 12. Pchelintsev NA, Adams PD, Nelson DM (2016) Critical parameters for efficient sonication and improved chromatin immunoprecipitation of high molecular weight proteins. PLoS One 11(1):e0148023. https://doi. org/10.1371/journal.pone.0148023

Chapter 23 Library Preparation for ATAC-Sequencing of Mouse CD4+ T Cells Isolated from the Lung and Lymph Nodes After Helminth Infection Laura D. Harmacek, Preeyam Patel, Rachel Woolaver, R. Lee Reinhardt, and Brian P. O’Connor Abstract Although conventional methods such as MNase-seq, DNase-seq, and ChIP-seq have been used effectively to assess chromatin and locus accessibility at the genome level, these techniques generally require large numbers of input cells. As such, much of what we understand in terms of epigenetic regulation and locus accessibility in CD4+ T cell subsets comes from in vitro culture systems, which allow for the production of large numbers of polarized T cells. However, obtaining such numbers directly ex  vivo from tissues of ­individual mice is difficult. Here we describe a method combining cytokine reporter mice and Assay for Transposase-Accessible Chromatin with high-throughput sequencing (ATAC-seq) to identify genome wide locus accessibility in a small number of cytokine-expressing CD4+ T cells. This method takes you from cell isolation to library generation and quality control to query. Because the Il4 and Ifng loci are reciprocally regulated in polarized CD4+ T cell subsets (Th1 vs. Th2), we investigated the ability of this approach to identify transposase integration in both IL-4- and IFN-γ-expressing CD4+ T cells isolated directly from the lung and lymph nodes after helminth infection. Key words ATAC-seq, Tn5 transposase, CD4+ mouse T cells, NGS library preparation, Lung and lymph node T cell prep

1  Introduction ATAC-seq is a robust new method to query the chromatin ­structure of cells [1], with 174 publications in the past year, and 435 ­publications in the past 5 years utilizing this technique according to PubMed. Genomic organization within cells is complex, and for genes to be transcribed the chromatin structure needs to be ­accessible to transcription factors, chromatin modifying enzymes, and other transcriptional machinery. The more accessible the ­chromatin, the faster transcriptional changes might take place if the necessary signals are received and proteins are present. This is ­especially important in hematopoietic cells, which travel in the R. Lee Reinhardt (ed.), Type 2 Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1799, https://doi.org/10.1007/978-1-4939-7896-0_23, © Springer Science+Business Media, LLC, part of Springer Nature 2018

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bloodstream potentially encountering many different ­microenvironments. ATAC-seq has been used to understand the dynamic chromatin structure in hematopoietic cells [2], resting CD4 T cells [3, 4] and in various disease states [5]. This method uses a modified Tn5 transposase to insert known short DNA sequence tags into open and accessible genome. In brief, a crude nuclear extract is made from 50,000 cells. Then, cells are treated for a predetermined incubation time with the Tn5 transposase. DNA is column purified from the cells and then PCR amplified based on the max fluorescence intensity. The resulting individually barcoded libraries are size selected and quality controlled on a ­fragment analyzer to assess concentration and size profile. The method presented here is optimized for mouse CD4+ T cells. The CD4+ T cells used were isolated from the lung and lungdraining lymph nodes of individual cytokine reporter mice ­following Nippostrongylus brasiliensis infection. This helminth model drives a prototypic type 2 immune response in the m ­ ediastinal lymph nodes, lungs, and small intestine of infected animals [6]. Importantly, both IL-4-expressing and interferon-gamma (IFNγ)-expressing CD4+ T cells can be isolated from the infected ­tissues using this model [7]. Specifically, CD4+ T cells expressing IL-4 from Il44get mice and IfngGreat were isolated based on their ­expression of GFP (IL-4) and YFP (IFN-γ), respectively [8, 9]. To validate the method, locus accessibility was analyzed at Il4 and Ifng loci of wild-type (reporter-negative) and cytokine-reporter positive cells.

2  Materials 2.1  Cell Isolation and Sorting

1. Il44get cytokine reporter mice: Green fluorescent protein (GFP) marks IL-4 mRNA expression.

2.1.1  Mice

2. IfngGreat cytokine reporter mice: Yellow fluorescent protein (YFP) marks IFN-γ mRNA expression. 3. C57BL/6 mice serve as negative controls for cytokine reporter mice.

2.1.2  Helminths

1. L3 larvae of Nippostrongylus brasiliensis.

2.1.3  Extraction and Preparation of Lung and Mediastinal Lymph Nodes for Cell Sorting

1. Forceps for dissection. 2. Scissors for dissection. 3. 50 and 15 mL conical tubes. 4. 5 mL FACS tubes. 5. 60 mm petri dishes. 6. Razor blades. 7. 80 micron nylon mesh. 8. 50 micron nylon mesh.

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9. 1× phosphate-buffered saline (PBS). 10. Complete RPMI 1640 (cRPMI): 10% fetal bovine serum (FBS), 100 units(U)/mL penicillin and streptomycin (P/S), 2  mM  l-glutamine, 50  μM 2-mercaptoethanol dissolved in RPMI 1640. 11. FACS sorting buffer: 2% FBS, 100 U/mL P/S in PBS. 12. ACK (Ammonium-Chloride-Potassium) Red Blood Cell (RBC) lysis buffer: 0.15 M NH4cl, 10 mM KHCO3, 0.1 mM disodium ethylenediaminetetraacetic acid (EDTA) in water (pH 7.2–7.4). 13. Lung digestion solution: 250 μg/mL collagenase XI, 100 μg/ mL Liberase TM (Roche), 200 mg/mL DNAse I, 1 mg/mL hyaluronidase in RPMI 1640. 14. FACS sorting buffer  +  4′,6-diamidino-2′-phenylindole ­dihydrochloride (DAPI): 0.5 mg/mL DAPI, 2% FBS, 100 U/ mL P/S in PBS. 15. Purified, unlabeled anti-mouse CD16/32 Fc block (TruStain fcX). 16. Antibodies for flow cytometry (conjugated fluorochrome; clone): CD3 (PE-Cy7; PE145-2C11), CD4 (Alexa fluor 647; RM4-5), CD8α (PerCP-Cy5.5; 53-6.7), B220 (PerCP-Cy5.5; RA3-6B2), CD11b (PerCP-Cy5.5; M1/70), CD11c (PerCP-­ Cy5.5; N418), NK1.1 (PerCP-Cy5.5; PK136), TER119 (PerCP-Cy5.5). 2.1.4  Cell Lysis, Transposition Reaction, and Cleanup

All solutions, mixes, and buffers are made with UltraPure Molecular Biology Grade water. Since the end result is NGS sequencing, make sure to use excellent molecular biology technique. Wipe down work area and pipettes with 70% ethanol, use filtered pipette tips, and do not cross contaminate solutions. 1. Cold 1× PBS (see Note 1). 2. Lysis buffer: 10 mM Tris Cl, pH 7.4, 10 mM NaCl, 3 mM MgCl2, 0.1% (v/v) molecular biology-grade IGEPAL CA-630 (see Table 1; Note 2). 3. Transposase Master Mix: TD 2× reaction buffer from Illumina Nextera kit, TDEnzyme1 (Nextera Tn5 Transposase from Illumina Nextera kit (see Table 2). 4. UltraPure Molecular Biology Grade water. 5. Warm water bath set at 37 °C. 6. Qiagen MinElute PCR Purification Kit (Qiagen): Qiagen MinElute columns, PB buffer, PE buffer, Elution buffer (EB). 7. Refrigerated centrifuge (set to 4 °C).

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Table 1 Lysis buffer for ATAC-seq Lysis buffer for ATAC-seq

Stock

Final

1 mL

Tris Ph7.4

1 M

10 mM

10 μL

MgCl2

1 M

3 mM

3 μL

NaCl

5 M

10 mM

2 μL

IGEPAL CA-630

100%

0.10%

1 μL

UltraPure water

984 μL

Store at 4 °C for up to 1 week

Table 2 Transposase Master Mix

2.1.5  QPCR and Library Amplification

Transposase Master Mix

Volume

TD (2× reaction buffer from Nextera kit)

25 μL

TDEnzyme1 (Nextera Tn5 Transposase from Nextera kit)

2.5 μL

UltraPure Molecular Grade Water

22.5 μL

1. PCR Primer 1 [custom-synthesized) [1] (see Note 3). 2. Barcoded PCR Primer Ad2.1 (custom-synthesized) and more barcoded adapter primers depending on pooling and ­sequencing strategy [1] (see Note 4). 3. NEBNext High-Fidelity 2× PCR Master Mix. 4. 100× SYBR green I. 5. Single PCR tubes. 6. 96-well plate for qPCR reactions. 7. qPCR and PCR machines. 8. Qiagen MinElute PCR Purification Kit: Qiagen MinElute ­columns, PB buffer, PE buffer, Elution buffer (EB).

2.1.6  Library Size Selection, Cleanup, and QC

1. Agencourt AMPure XP beads (Beckman Coulter). 2. Freshly made 70% ethanol. 3. Strong 1.5 mL tube magnet. 4. High Sense Bioanalyzer DNA kit. 5. Agilent Bioanalyzer machine (see Note 5). 6. UltraPure 1 M Tris–HCl (pH 8.0), diluted to 10 mM. 7. LoBind 1.5 mL tubes.

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3  Methods 3.1  Cell Isolation from Lung and Mediastinal Lymph Nodes

1. Subcutaneously infect mice with 500 N. brasiliensis L3 larvae at the base of the tail and euthanize mice 9 days later. 2. Harvest mediastinal lymph nodes and place them into a 50 mL conical with FACS sorting buffer. Place the tube on ice until all organs are harvested. 3. Gently excise lungs from the plural cavity and rinse with PBS. Place lungs into a 50 mL conical with 2 mL of cRPMI 1640. Keep organ tube on ice until ready for digestion. 4. Individually dissociate mediastinal lymph nodes in a 60  mm petri dish with 2  mL of PBS using the flat end of a 5  mL syringe plunger. Pass cells through an 80 micron nylon filter, collect cells in a 15 mL conical, and wash the petri dish in a total volume of 10 mL of PBS. Centrifuge 15 mL conical at 500 × g (g-force) for 5 min. Resuspend cell pellet in 1 mL of in FACS sorting buffer and place on ice. 5. Remove lungs from the 50 mL conical and place them into a 60 mm petri dish. Finely mince lung tissue with a razor blade. 6. To this petri dish, add 5 mL of lung digestion solution. Tissue is incubated at 37°C for 30 min (see Note 6). 7. The digested lung is passed through an 80 micron nylon filter into a 50 mL conical. The digestion mixture is quenched with 10  mL of FACS sorting buffer and centrifuged at 500  ×  g (g-force) for 5 min. 8. Following removal of the supernatant, resuspend the lung cell pellet in 2 mL ACK RBC lysis buffer, and incubate for 2–4 min at room temperature. 9. ACK RBC lysis buffer is quenched with the addition of 5 mL of cold PBS and centrifuged at 500 × g (g-force) for 5 min. 10. Decant supernatant and resuspend lung cells in 2 mL of FACS sorting buffer. Cell suspension is placed on ice.

3.2  Cell Staining and FluorescentActivated Cell Sorting

1. Wash single cell suspensions obtained from the lung (50 mL conical) and lymph nodes (15 mL conical) with an additional 10 mL of PBS, and spin at 500 × g (g-force) for 5 min. 2. Gently resuspend cell pellets in 50  μL (lymph nodes) or 500 μL (lung) of a 1:100 dilution of Fc Block in FACS sorting buffer, and incubate at 4 °C for 15 min. 3. Incubate cell suspension with an additional 50  μL (lymph nodes) or 500 μL (lung) of a 1:200 dilution of fluorescently conjugated antibodies (noted in the Materials section) diluted in FACS sorting buffer at 4 °C for 30 min on ice, and then wash two times with 10 mL of PBS.

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Fig. 1 Gating scheme for cell sorting of GFP− and GFP+ CD4+ T cells from the lymph node and post-sort purity of sorted populations. Single cell suspensions of lymph node cells from Il44get mice were prepared and stained for flow cytometry as described. Successive gates were drawn for sorting CD4+ T cells that are GFP− and GFP+ as followed: (a) lymphocytes, (b, c) single cells, (d) live (DAPI−), (e) CD3+ (CD8−, B220−, CD11c−, Ter119−, NK1.1−), (f) CD3+ CD4+ T cells, (g) CD4+ GFP− or GFP+ T cells. Post-sort purity for ungated (h) GFP− and (i) GFP+ CD4+ T cells. Numbers in the graphs represent percentage of gated populations from parent populations

4. Resuspend final cell pellet in FACS sorting buffer + DAPI, and pass cells through a 40 micron nylon filter into a new 5 mL FACs tube immediately before cell sorting is performed. 5. Lymphocytes were identified by forward and side scatter. DAPI+, PerCP-Cy5.5+ cells were excluded in a “dump” gate (Fig.  1). CD3+CD4+ T cells were then gated from the ­remaining cells, and 50,000–150,000 GFP/YFP− or GFP/ YFP+ cells from the lymph nodes or lung populations were sorted into separate 5 mL FACS tubes (Fig. 1) (see Note 7). 3.3  Cell Transposase Reaction

Tn5 transposase enzyme incubation times may vary based on cell type and exposure. Test incubation times with each new cell type. A range of incubation times (30 min, 40 min, 50 min) were tested on 50,000 cells (Fig.  2). In this example and for the cell types sorted in this protocol, 40 min incubation time yielded the most periodicity in both the higher molecular weight range and the lower molecular weight range. Fifty minutes was too long as the higher molecular weight fragments are reduced in abundance, and most of the fragments are smaller. The methods below were used to test these incubation times (see Note 8). 1. Place 50,000 flow sorted CD4+ GFP/YFP +/− cells in a 1.5 mL tube and wash in 50 μL cold PBS. Spin samples in a prechilled (4 °C) centrifuge at 500 × g (g-force) for 5 min (see Note 9).

Assaying Chromatin Dynamics of In Vivo-Isolated Helper T Cells [FU]

a

[FU] 200

100

100

0

0 35

150 300

500 1000 [FU] 150

10380[bp]

333

b

35

150 300

500 1000

10380[bp]

c

100 50 0 35

150 300

500 1000

10380[bp]

Fig. 2 Optimizing Tn5 transposase incubation time on a given cell type: These traces are representative of three Tn5 transposase incubation time points taken on a mouse cell type and made into ATAC-seq libraries. (a) 30 min incubation, (b) 40 min incubation, and (c) 50 min incubation

2. Resuspend cells in 50  μL fresh cold lysis buffer and spin at 500 × g (g-force) for 10 min at 4 °C. 3. Remove lysis buffer, and resuspend cells in 50 μL Transposase Master Mix, pipetting ten times. 4. Incubate cells in a 37 °C water bath for predetermined amount of time (e.g., 40 min). 5. Purify DNA in a Qiagen MinElute column found in the MinElute PCR Purification Kits. To do this, add 250  μL (5×) PB binding buffer to the 50 μL transposase reaction. 6. Apply to column and spin. Add 750  μL wash buffer to the ­column. Spin and then discard wash. 7. Spin 1 additional minute to dry membrane. 8. Elute in 22 μL elution buffer (EB) (see Notes 10 and 11). 3.4  qPCR Amplification

This step allows you to find the max fluorescence intensity of each sample and then calculate the number of PCR amplification rounds necessary to maximize library efficiency while reducing PCR duplicates. Include a no template control where you add master mix alone and no DNA to ensure primers or master mix are not contaminated. 1. Run a qPCR reaction on 2 μL or 10% of each sample by qPCR following Table 3 (see Note 12). 2. Set qPCR instrument with the cycles as follows: 1 cycle: 5 min 72 °C, 30 s 98 °C; 30 cycles: 10 s 98 °C, 30 s 63 °C, 1 min 72 °C.

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Table 3 qPCR Master Mix qPCR Master Mix

Volume

Nuclease-free H2O

4.41 μL

25 μM PCR Primer 1 “noMX”

0.25 μL

25 μM PCR Primer/Adapter 2.1 (or any other 2.× primer)

0.25 μL

100× SYBR green I

0.09 μl

NEBNext High-Fidelity 2× PCR Master Mix

5 μL

Add DNA individually to each aliquoted master mix

Volume

1-2 μL transposed DNA (10% of the total volume)

2 μL

Table 4 PCR amplification Master Mix PCR amplification Master Mix

Volume

25 μM PCR Primer 1 “no MX”

1.5 μL

NEBNext High-Fidelity 2× PCR Master Mix

25 μL

Nuclease-free H2O (up to 50 μL)

4 μL

Add barcoded primers individually

Volume

25 μM Barcoded PCR Primer/Adapter 2

1.5 μL

Add DNA individually

Volume

All μL from original transposased DNA

~18–20 μL

3. To calculate the number of cycles needed (N), plot linear Rn versus cycle, not the ΔRn versus cycle, and determine the cycle number that corresponds to one-third of the maximum ­fluorescent intensity. Refer to Fig. 3. If N falls between two cycles, round down to the nearest integer. This is referred as N cycles in the next step (see Notes 13 and 14). 3.5  PCR Amplification

1. To amplify transposed DNA fragments, combine the following in a 0.2 mL PCR tube according to Table 4 (see Notes 15 and 16). 2. Amplify transposase fragments with the following PCR thermal cycler as follows (see Note 17):

(a) 1 cycle: 5 min 72 °C, 30 s 98 °C

(b) N cycles: 10 s 98 °C, 30 s 63 °C, 1 min 72 °C. 3. Purify DNA in a Qiagen MinElute column and elute in 22 μL elution buffer (EB). See Subheading 3.3, step 5.

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Assaying Chromatin Dynamics of In Vivo-Isolated Helper T Cells

Amplification Plot 15,000,000 14,000,000 13,000,000 12,000,000

Max=14,500,000 Max=12,900,000

11,000,000 10,000,000 9,000,000

Rn

8,000,000 7,000,000 6,000,000 5,000,000 4,000,000 3,000,000 2,000,000 1,000,000 0 -1,000,000

2

4

6

8

10

12

14

16

n=9 Cycles n=12 Cycles

18

20

22

24

26

28

Cycle

Fig. 3 Calculating number of PCR amplification cycles: Representative amplification plot demonstrating the correct number of cycles to perform on two individual ATAC-seq samples. To calculate the additional number of cycles needed (N), plot linear Rn versus cycle and determine the cycle number that corresponds to one-third of the maximum fluorescent intensity 3.6  Library Size Selection and Cleanup:

1. Add ~38 μL nuclease-free water to the sample to bring up to 50 μL total. 2. Add 25  μL (0.5× volume) of AMPure XP well-mixed RT beads to each sample (see Notes 18 and 19). 3. Mix thoroughly by pipetting five to ten times, and incubate at room temperature for 5 min. 4. Place tube in magnetic rack until solution clears, 2–3 min. 5. Transfer the supernatant to a new 1.5 mL tube, without beads (see Note 20). 6. Add 50 μL (1.5× volume) of AMPure XP beads to the transferred supernatant of each sample (see Note 21) 7. Mix thoroughly by pipetting five to ten times, and let sit for 5 min at room temperature. 8. Place tube in magnetic rack until solution clears. 9. Discard the supernatant and add 500 μL of fresh 70% ethanol. Invert tubes twice while beads are still on the magnet to wash. 10. Remove ethanol by dumping or pipetting. 11. Repeat steps 9 and 10 for a total of two washes.

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Fig. 4 Traces from 40 min transposase incubation: Bioanalyzer library traces from the cells sorted in this protocol. Cells from different organs and sorts produce a range of library traces. (a) Lung Il44get GFP+ (b) Lung Il44get GFP− (c) Lymph Node Il44get GFP+ (d) Lymph Node Il44get GFP− (e) Lung IfngGreat YFP+

12. After removing the last wash, pulse spin the tube, place the tube back in the magnetic rack, and remove the last of the ethanol. 13. Leave lid open and allow samples to air dry for 5 min. Beads should be slightly damp and not cracked dry. 14. Add 25 μL of 10 mM Tris–HCl (pH 8) to each sample. Vortex then pulse spin the samples. 15. Place in magnetic rack until solution clears (see Note 22). 16. Transfer the library to a new 1.5 mL LoBind Tube. This is a safe stopping point (see Note 23). 3.7  Library Dilution and QC

1. Measure the concentration in ng/μL (see Note 24). 2. Dilute libraries to 1 ng/μL in 10 mM Tris–HCl (pH 8). 3. Run 1 μL on a Agilent Bioanalyzer High Sense DNA Chip (see Note 25). 4. Analyze traces. Refer to Fig. 4 for representative traces in the various cell populations from this protocol. While not all ­libraries look the same, they all have a maintained periodicity from 150 to 2000 bps. Observable difference may or may not reflect a correlation in overall open chromatin structure of the samples after sequencing (see Note 26).

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Fig. 5 Gene loci of the Il44get and IfngGreat CD4+ cells after helminth infection. ATAC-seq libraries were sequenced and analyzed: (a) Il4 and (b) Ifng gene loci. Data was visualized in Integrative Genome Viewer (Broad Institute). TSS transcription start site

5. After library prep, libraries were sequenced on an Illumina 2500; fragments were demultiplexed and mapped. As validation for the approach, the Il4 locus shows an open accessible chromatin structure particularly in the 5′ region of lung and lymph node Il44get GFP+ cells but not in the lung and lymph node GFP− or lung IfngGreat YFP+ cells (Fig. 5). Alternatively, the 5′ region of the Ifng locus opens and shows accessible chromatin among the IfngGreat lung YFP+ cells compared to reduced accessibility in YFP− cells and almost no accessibility in the Il44Get cells.

4  Notes 1. The authors purchase 10× PBS from Invitrogen and dilute with UltraPure Molecular Grade Water. 2. Add appropriate volume of UltraPure Molecular Grade Water minus the IGEPAL detergent volume. Then add 0.1% (v/v) Molecular biology-grade IGEPAL CA-630 (Sigma-Aldrich, cat. no. I8896), and invert tube until detergent is suspended in solution. 3. This primer is synthesized by Integrated DNA Technologies (IDT) with no additional modifications. A complete list of primers is available in Buenrostro et  al. (2013). The PCR Primer 1 (also known as NoMx) has a sequence of: A AT G ATA C G G C G A C C A C C G A G AT C TA C A C T C G TCGGCAGCGTCAGATGTG

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4. Ad Primers are synthesized by Integrated DNA Technologies (IDT) using their ultramer oligo synthesis due to the high number of bases. A complete list of primers is available in Buenrostro et al. (2013). The PCR Primer 2.1 (also known as Ad2.1_TAAGGCGA) has a sequence of: CAAGCAGAAGACGGCATACGAGATTCGCCTTAGT CTCGTGGGCTCGGAGATGT 5. Other forms of QC include running a polyacrylamide gel or using an NGS library quantification kit. We find the Bioanalyzer traces are sufficient to assess quantity and quality of libraries. 6. Lung tissue should be disrupted every 15  min with a 1  mL pipette tip (with tip slightly cut to widen the opening of the pipette) to add the digestion. 7. Target cells in this protocol were obtained by sorting on either a Sony ICyt Synergy or a BD Aria Fusion using the described strategy. 8. While this optimized method worked well for these cells, there are published ATAC-seq protocols that add the detergent directly to the enzyme master mix to avoid a separate lysis step. 9. The number of cells at this step is critical because the transposase to cell ratio will determine the distribution of generated DNA fragments. 10. Elution in 22 μL allows for 2 μL to be tested via qPCR for calculation of PCR cycles. Remaining 20  μL of sample is amplified in a standard PCR reaction. 11. At this point it is safe to freeze the eluted DNA at −20 °C or continue to PCR amplification. 12. In this qPCR step, you can use the same primer pairs in each of the samples as this is a test step. This should not be done during the PCR amplification stage. In the PCR amplification stage, you will need to be very careful with which barcodes you use depending on your pooling and sequencing strategy. 13. There is no need to run internal calculations, standards, etc. when analyzing the qPCR results. Make sure it is measuring a very basic fluorescence intensity measurement. 14. Read the number of cycles when intensity peaks, not the number of maximum intensity. This number of cycles (N) is usually between 12 and 18. 15. Ensure that samples are individually barcoded appropriately for subsequent NGS pooling and sequencing.

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16. Use individual PCR tubes for your reactions, so you can remove each sample after the appropriate number of cycles are reached. 17. This first 5 min extension at 72 °C allows extension of both ends of the primer after transposition, thereby generating amplifiable fragments. 18. Warm beads on the bench top for at least 30 min and vortex beads for at least 1 min before use. 19. Revortex beads after every eight samples as the beads settle quickly. 20. It is better to leave a few microliters behind than to transfer any beads over. The authors leave 2 μL in the tube to avoid bead carryover. 21. 1.5× AMPure XP beads in a 50 μL starting volume would be 75 μL; however, 25 μL were already added to the sample for the upper size selection, so only 50 μL more need to be added to the transferred supernatant. 22. To help beads accumulate at the top of such a small volume, put tubes in the rack halfway for ~30 s until solution clears, and then slowly slide tube fully into the magnetic rack. 23. You can store your library at −20  °C or continue to library QC. 24. The authors use the Invitrogen Qubit for accurate quantification. 25. Other fragment analyzers could be used here. 26. If there are small molecular weight fragments (130 bp or less) that look like primer or concatamer, you need to clean up the lower fragments. This smaller fragment size will disproportionally consume sequencing reads rendering ATAC-seq data unreadable. Bring total volume of library up to 50  μL.  Add 75 μL (1.5×) resuspended AMPure XP beads to the sample, and start at step 7 of Subheading 3.6. If there are higher MW fragments (2000 bp+), clean up the libraries starting at step 1 of Subheading 3.6. Elute libraries in 11.5  μL to make sure they are adequately concentrated.

Acknowledgments Thanks to the William Greenleaf lab for publishing this method and to the helpful discussions at the ATAC-seq users online Google Group forum: https://sites.google.com/site/atacseqpublic/. This research was funded in part by NIH grants RO1HL127461, RO1HL126600 (B.P.O.), and AI119004 (R.L.R).

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References 1. Buenrostro JD, Wu B, Chang HY, Greenleaf WJ (2015) ATAC-seq: a method for assaying chromatin accessibility genome-wide. Curr Protoc Mol Biol 109:21.29.21–21.29.29. https://doi. org/10.1002/0471142727.mb2129s109 2. Lara-Astiaso D, Weiner A, Lorenzo-Vivas E, Zaretsky I, Jaitin DA, David E, Keren-Shaul H, Mildner A, Winter D, Jung S, Friedman N, Amit I (2014) Immunogenetics. Chromatin state dynamics during blood formation. Science 345(6199):943–949. https://doi. org/10.1126/science.1256271 3. Shih HY, Sciume G, Mikami Y, Guo L, Sun HW, Brooks SR, Urban JF Jr, Davis FP, Kanno Y, O'Shea JJ (2016) Developmental acquisition of regulomes underlies innate lymphoid cell functionality. Cell 165(5):1120–1133. https:// doi.org/10.1016/j.cell.2016.04.029 4. Buenrostro JD, Giresi PG, Zaba LC, Chang HY, Greenleaf WJ (2013) Transposition of native chromatin for fast and sensitive epigenomic profiling of open chromatin, DNA-­ binding proteins and nucleosome position. Nat Methods 10(12):1213–1218. https://doi. org/10.1038/nmeth.2688 5. Seumois G, Chavez L, Gerasimova A, Lienhard M, Omran N, Kalinke L, Vedanayagam M, Ganesan AP, Chawla A, Djukanovic R, Ansel

KM, Peters B, Rao A, Vijayanand P (2014) Epigenomic analysis of primary human T cells reveals enhancers associated with TH2 memory cell differentiation and asthma susceptibility. Nat Immunol 15(8):777–788. https://doi. org/10.1038/ni.2937 6. Finkelman FD, Shea-Donohue T, Morris SC, Gildea L, Strait R, Madden KB, Schopf L, Urban JF Jr (2004) Interleukin-4- and interleukin-­13-mediated host protection against intestinal nematode parasites. Immunol Rev 201:139–155 7. Bao K, Carr T, Wu J, Barclay W, Jin J, Ciofani M, Reinhardt RL (2016) BATF modulates the Th2 locus control region and regulates CD4+ T cell fate during antihelminth immunity. J  Immunol 197(11):4371–4381. https://doi. org/10.4049/jimmunol.1601371 8. Mohrs M, Shinkai K, Mohrs K, Locksley RM (2001) Analysis of type 2 immunity in  vivo with a bicistronic IL-4 reporter. Immunity 15(2):303–311 9. Reinhardt RL, Liang HE, Bao K, Price AE, Mohrs M, Kelly BL, Locksley RM (2015) A novel model for IFN-gammamediated autoinflammatory syndromes. J  Immunol 194(5):2358–2368. https://doi. org/10.4049/jimmunol.1401992

Chapter 24 Identification of Functionally Relevant microRNAs in the Regulation of Allergic Inflammation Marlys S. Fassett, Heather H. Pua, Laura J. Simpson, David F. Steiner, and K. Mark Ansel Abstract Transgenic methods to manipulate CD4 T lymphocytes in vivo via forced expression of TCR transgenes and targeted “knockout” of individual genes by Cre-lox technology are fundamental to modern immunology. However, efforts to scale up functional analysis by modifying expression of larger numbers of genes in T cells ex vivo have proven surprisingly difficult. Early RNA interference experiments achieved successful small RNA transfection by using very high concentrations of short-interfering RNA (siRNA) [1], but primary T cells are generally resistant to standard electroporation, cationic liposome-, and calcium phosphate-­mediated transfection methods. Moreover, although viral vectors can successfully introduce DNA fragments of varying length, expression of these constructs in primary T cells is low efficiency and the subcloning process laborious. In this context, the relatively recent discovery of dozens of highly expressed microRNAs (miRNAs) in the immune system provides both an opportunity and a new challenge [2, 3]. How can we query the miRNAome of a cell to assign particular roles to individual miRNAs? Here, we describe an optimized technique for efficient and reproducible transfection of primary mouse CD4 T cells in vitro with synthetic miRNA mimics. Key words microRNA, T lymphocyte transfection, Th2, CD4, Electroporation, In vitro gene expression

1  Introduction Next-generation transfection systems, including the Neon™ Transfection System (Thermo Fisher Scientific) and the Amaxa® Nucleofector® (Lonza), are capable of transfecting both cell lines and primary cells with high efficiency (25–99%) and high cell viability (60–99%) [4–8]. We have found the Neon system easy to use, economical, and capable of reliably delivering small RNAs into primary mouse T cells to study miRNA function [6, 7, 9, 10]. This technique can be used to test the effect of miRNA overexpression using miRNA mimics without the need to clone into expression or targeting vectors. Electroporation with the Neon Transfection R. Lee Reinhardt (ed.), Type 2 Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1799, https://doi.org/10.1007/978-1-4939-7896-0_24, © Springer Science+Business Media, LLC, part of Springer Nature 2018

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System is amenable to testing a small set of miRNA mimics and may also be utilized for medium-throughput screens which test the function of >100 distinct miRNAs. After electroporation, CD4 T cells remain capable of proliferation, polarization, and cytokine production, allowing assessment of numerous cellular functions. Molecular consequences of altering miRNA activity in cells can be evaluated through expression studies of predicted, known, and novel mRNA targets through techniques including qPCR and RNA sequencing. In addition, Neon electroporation can be used to query the combined effects of multiple miRNAs (such as members of a miRNA family) by co-transfection [10]. This creates an opportunity to observe the potential effects of multiple miRNAs on a convergent mRNA target or pathway. The transient nature of miRNA mimic transfection in vitro provides specific advantages over stable genetic models: 1. Transfection allows temporal control in the manipulation of individual miRNA expression during T cell activation, proliferation, and differentiation. 2. Transfection facilitates rapid functional assessment of multiple miRNAs individually and in combination with one another. This is a particular challenge with genetic models because related miRNAs are often encoded in polycistrons or distributed across multiple genomic loci. 3. Transfection prevents confounding cellular changes that may be seen in stable genetic models as a result of changes in T cell differentiation or adaptations to long-term alterations in gene expression. This chapter outlines our standard protocol, which begins on day 0 with isolation, activation, and Th2 polarization of primary mouse CD4 T cells. Activated Th2-polarized cell cultures are transfected with miRNA mimics on day 1 and again on day 4 to boost miRNA expression. We include optimized instructions for operating the Neon Transfection System with primary mouse CD4 T cells. On day 5, we harvest transfected T cells for analysis by various methods, such as target gene expression and flow cytometry for cellular phenotyping.

2  Materials 2.1  Equipment (See Note 1)

1. Neon™ Transfection System (Thermo Fisher). 2. Neon™ transfection pipette. 3. Neon™ electroporation tubes (a.k.a. cuvettes). 4. Neon™ Transfection System 10 μL kit (tips). 5. 10 cm TC-treated sterile plates. 6. 96-well flat bottom TC-treated sterile plates. 7. 96-well round bottom TC-treated sterile plates.

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2.2  Cytokines and Antibodies (Table 1; See Note 2)

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1. Neutravidin (Thermo Fisher; stock 10 mg/mL in PBS). 2. Biotinylated anti-CD3 antibody (clone 2C11; stock 2.8 mg/ mL in PBS). 3. Biotinylated anti-CD28 antibody (clone 37.51; stock 2.0 mg/ mL in PBS). 4. IL-4 supernatant (10,000 U/mL stock) added as a supernatant from I3L6 cells [11]. 5. Anti-IFNγ antibody (3.5 mg/mL stock in PBS). 6. Recombinant human IL-2 (100,000 U/mL in PBS).

2.3  Solutions and Reagents

1. “Kool-Aid” media (for 1 L): 4.5 g glucose, 0.584 g l-­glutamine, 3.7 g NaHCO3, 0.116 g l-arginine HCl, 0.036 g l-asparagine, 0.006 g folic acid, 10 mL of 100× NEAA solution, and 10 mL of 100× MEM essential vitamin solution. 2. Kool-Aid-COMPLETE media: Kool-Aid media supplemented with 10% FBS, 100 μg/mL streptomycin, 100 U/mL penicillin G, 10 mM HEPES, 1 mM sodium pyruvate, 100 μM β-mercaptoethanol, and 2 mM l-glutamine. 3. siRNA buffer: (5× stock). 4. E buffer (included in Neon™ transfection 10 μL kit). 5. T buffer (included in Neon™ transfection 10 μL kit). 6. Dynabuffer: 1× PBS supplemented with 2% fetal bovine serum (FBS). 7. Dynabeads™ Mouse CD4 kit (Thermo Fisher). 8. DETACHaBEAD® (Thermo Fisher). 9. miRIDIAN miRNA mimics and chemistry-matched control miRNA mimic (Dharmacon).

3  Methods 3.1  Isolation of Mouse CD4 T Cells (Day 0)

Primary mouse CD4 T cells may be isolated from the peripheral lymph nodes and spleen by standard laboratory protocols. We favor the Dynabead Mouse CD4 kit, following the manufacturer’s instructions with slight modification. 1. Harvest the spleen and peripheral lymph nodes to obtain single cell suspensions (see Note 3). 2. Purify CD4 T cells by positive selection using the Dynabeads™ Mouse CD4 kit using anti-CD4 mAb (clone L3T4)-coated magnetic beads. We use 75 μL of Dynabeads per mouse and elute with an equal volume of the proprietary DETACHAaBEAD reagent. 3. Once cells are attached to beads, all washes should be performed to minimize agitation (see Note 4).

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Table 1 Cytokines and antibodies Reagent

Stock concentration 1× final dilution factor

1× final concentration

Biotinylated anti-CD3

2.8 mg/mL

11,200

0.25 μg/mL

Biotinylated anti-CD28

2 mg/mL

2000

1 μg/mL

Mouse IL-4

10,000 U/mL

20

500 U/mL

Anti-IFNγ mAb

3.5 mg/mL

350

10 μg/mL

Recombinant human IL-2

100,000 U/mL

5000

20 U/mL

3.2  In Vitro T Cell Activation and Th2 Polarization (Day 0)

1. Prepare tissue culture plates. Make neutravidin working solution by 2000× dilution of 10 mg/mL neutravidin stock in PBS (final 5 μg/mL). Place 10 mL neutravidin solution onto 10 cm plates, and incubate for 2 h at RT or 1 h at 37 °C, and then gently wash 2× with PBS. Keep neutravidin-coated plates warm at 37 °C in PBS briefly while preparing T cell media (Subheading 3.2, step 2). 2. Prepare one half the final culture volume of T cell polarizing media. For example, for each 10 cm plate, prepare 5 mL of Kool-Aid-COMPLETE media supplemented with T cell polarizing cytokines and blocking antibodies at 2× the desired final concentration. For Th2 culture conditions, supplemented 2× media contains biotinylated anti-CD3 (2× is 0.5 μg/mL), biotinylated anti-CD28 (2× is 2 μg/mL), IL-4 supernatant (2× is 1000 U/mL), and anti-IFNγ antibody (2× is 20 μg/mL). See Table 1 for working stock and final concentrations of all antibodies and cytokines. 3. Prepare neutravidin-coated plates with T cell media by aspirating PBS, and carefully add 2× supplemented Kool-Aid-­ COMPLETE media at half-final volume making sure to fully cover the plate surface. In a 10 cm plate, add 5 mL of 2× cytokine-­ supplemented media, and return plate to 37 °C incubator. 4. Resuspend purified CD4 T cells (from Subheading 3.1) in Kool-Aid-COMPLETE media (without cytokines or antibodies) at 2 × 106/mL. Place cells onto prepared plates such that the cell volume added is equal to the receiving 2× cytokine media, i.e., 5 mL cells in the 10 cm plate. Final cell ­concentrations will be 1 × 106/mL. If using other plate sizes, see Table 2 for appropriate cell numbers and volumes. 5. Activate cells overnight in a 37 °C incubator with 10% CO2 (see Notes 5 and 6).

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3.3  Preparation for First miRNA Mimic Transfection (Day 1)

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1. Prepare a 96-well flat bottom post-transfection culture plate (“receiving plate”) by coating with 100 μL/well of neutravidin solution for 2 h at RT or 1 h at 37 °C in PBS. Wash the neutravidin-­coated receiving plate twice with PBS, and then keep plate warm at 37 °C in PBS while preparing media (Subheading 3.3, step 2). 2. Prepare full final culture volume of T cell polarizing media. For 96-well plates, 200 μL/per well of Kool-Aid-COMPLETE media is supplemented with 1× polarizing cytokines and antibodies and will contain biotinylated anti-CD3 (1× is 0.25 μg/ mL) and biotinylated anti-CD28 (1× is 1.0 μg/mL), IL-4 supernatant (1× is 500 U/mL), and anti-IFNγ antibody (1× is 10  μg/mL) for Th2 culture conditions. Remove PBS from neutravidin-­coated plate, and add 200 μL per well of this 1× media. 3. Label the receiving plate (see Note 7), and then place in incubator to equilibrate. 4. Turn on Neon instrument and choose appropriate electroporation settings. For day 1 T cell transfection: 1550 volts, 10 ms, and 3 pulses. 5. Add 3 mL of Buffer E into the cuvette’s transfection chamber, and allow to equilibrate to RT. Carefully transfer the cuvette into Neon pipette stand without spilling any buffer (see Note 8). 6. Deposit miRNA mimics in individual Eppendorf tubes for transfection. We typically use 500 nM mimic solution in 0.5–1 μL of siRNA transfection buffer. If combining multiple miRNAs, we use a total mimic concentration of 500 nM (i.e., two miRNA mimics at 250 nM each). Set aside on ice. See Note 9.

3.4  Transfecting Th2-Polarized T Cells with miRNA Mimics (Day 1)

1. Harvest polarized Th2 cells into 15 mL conical tubes (see Note 10), take an aliquot for cell counting, and centrifuge at 400 × g-force, 5 min at RT. (Count cells during the spin.) Aspirate supernatant, and resuspend cells in 1.5 mL PBS. Transfer the needed number of each group of cells to an Eppendorf tube, and then centrifuge at 500 × g for 5 min. Aspirate supernatant, carefully removing as much as possible without disturbing pellet. 2. Resuspend cell pellet in 11 μL of T buffer per transfection of 400,000 cells. For example, 10 × 106 cells should be ­resuspended in 275 μL (see Note 11). Take care not to generate any bubbles when resuspending cells (see Note 12). 3. Remove miRNA mimic aliquots from ice, place at RT, and equilibrate for 5 min. Pipette 11 μL of cells into each Eppendorf tube pre-loaded with miRNA mimic. Pipette up and down to mix.

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Table 2 Cell numbers, media volume, and TC plate size for starting cultures Cell number

Media volume

TC plate size

1.5 × 105

200 μL

96 well

3 × 105

450 μL

48 well

5 × 10

5

750 μL

24 well

1 × 10

6

1.5 mL

12 well

4. Load a Neon pipette tip onto the Neon pipette. Fill a 50 mL conical tube with sterile PBS. 5. Slowly pipette up and down three times in PBS to wet the tip, taking care to avoid generating bubbles. Carefully aspirate the 11 μL of cells mixed with miRNA mimic into the Neon pipette tip, again avoiding bubbles. 6. Click the Neon pipette with tip into the cuvette, and then press “start” on the screen to electroporate. See Notes 13 and 14. 7. Immediately after successful electroporation, transfer transfected cells to the pre-warmed receiving plate pre-loaded with appropriate supplemented media. 8. Repeat steps 5–7 for each cell-miRNA transfection. Between transfections, wash the Neon pipette tip by pipetting up and down 3–5 times in 50 mL conical filled with PBS. The PBS does NOT need be changed between transfections. See Note 15. 9. At the end of the transfections, return plate to the incubator, and then aspirate Buffer E from the cuvette. If the cuvette has been used 90% CD4 T cells. 5. This transfection method does not work on naïve unmanipulated mouse T cells. It is necessary to activate the cells for at least 12 h prior to transfection. 6. If the cell-bead aggregate is fully disrupted during washing, there will be excessive cell loss. 7. When planning and labeling your receiving plate, be sure to account for the number of miRNA mimics multiplied by the number of target cell populations (WT, KO, etc.). Include additional wells for control mimic in each target cell population. Transfections may be carried out in technical singlets for large screens or in duplicate or triplicate to account for technical variation. 8. If you get an error message during transfection, one possible cause is a bad contact between the metal plate on the cuvette and the pipette stand. This can occur because of bad cuvette positioning or because of liquid between the contacts. If any liquid spills out of the cuvette, thoroughly dry it immediately. To minimize this risk, never pipette liquid into a cuvette already positioned in the stand. 9. When aliquoting miRNA mimics into Eppendorf tubes to prepare for transfection, you must have a separate tube for each transfection. Each combination of miRNA mimic and cells is considered one transfection.

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10. Activated T cells, particularly Th2 cells, can be surprisingly adherent to anti-CD3- and anti-CD28-coated plates. Take care to vigorously and completely blow them off the bottom of the plate. We favor using a P1000 Pipetman for this process. 11. Before resuspending the cell pellet, calculate the total number of transfections you plan to perform, and confirm that multiplying out 11 ul per 400,000 cells yields sufficient volume for that number of transfections (11 μL × # of transfections + 10% angel’s share). If you have fewer than 400,000 cells/transfection, increase the dilution. 12. If you catch a bubble in the pipette tip when drawing up cells, carefully expel the fluid and re-pipet. If you are unable to re-­ pipet successfully without bubbles a second time, expel the cells back into the Eppendorf tube, wash the tip with PBS 3–5 times, and then try to pipet the cells again. If you still are getting bubbles, check the seal between the pipette tip, and change the pipette tip if needed. A bubble within the pipette tip can cause the transfection to “arc” (often with a visible spark and popping sound) and kill the cells. 13. You will hear a click when the pipette is properly positioned within the cuvette station. Listen for this sound before pressing start, or remove and reposition the pipette. 14. The Neon instrument will beep twice if the transfection completes successfully. Wait for these beeps before transferring transfected cells to the receiving plate. See Notes 7 and 20 for additional troubleshooting tips if you do not hear the beeps. 15. In our experience, each Neon pipette tip can be reused up to 14 times. We recommend changing tips between groups to avoid contamination. 16. By day 3, T cell activation is complete, and the T cells should be “rested” in IL-2. Therefore, no neutravidin coating step is done, and there is no anti-CD3 or anti-CD28 in the media. 17. We generally do not recount cells before the day 4 transfection unless the wells look overpopulated. As a result of a low proliferation rate and increased cell death within the first day after initial transfection, the number of cells per well is usually within range. In our experience, successful transfections can be performed with as few as 50,000 cells and as many as 800,000 cells. 18. Effective miRNA overexpression in cells capable of endogenous miRNA expression may not be possible with the protocol outlined here. Limited availability of Argonaute due to high occupancy by endogenous miRNAs and/or previous saturation of the mRNA targets of highly expressed miRNAs

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may hamper transfected miRNA mimic activity in a cell. Studies using this transfection protocol can be enhanced by starting with miRNA-deficient T cells (e.g., from Dgcr8deficient animal strains) to study the gain of functional miRNA activity [6, 15]. Complementary miRNA inhibitor transfections can also be used to test loss of function [10]. 19. If performing a rescue screen in miRNA-deficient T cells from Dgcr8-deficient strains, as was pioneered in Dgcr8-deficient ES cells [16], be aware that CD4 T cells will include both miRNA-deficient cells and “escapees” that fail to delete Dgcr8 and therefore remain miRNA-sufficient. For best results, include a method to identify true knockout cells (e.g., a Cre-­ activated reporter gene) to mark the miRNA-deficient cell population prior to assessing the phenotype “rescued” by transfection with individual miRNAs [6]. 20. If you get an error message during the transfection or you do not hear the Neon instrument beep, remove the pipette from the cuvette, go back to the Eppendorf, and try to re-pipette the cells. If this does not work after several tries, you may need to jiggle the pipette spring and replace the tip. References 1. McManus MT, Haines BB, Dillon CP et al (2002) Small interfering RNA-mediated gene silencing in T lymphocytes. J Immunol 169(10):5754–5760 2. O'Connell RM, Rao DS, Baltimore D (2012) microRNA regulation of inflammatory responses. Annu Rev Immunol 30:295–312. https://doi.org/10.1146/ annurev-immunol-020711-075013 3. Baumjohann D, Ansel KM (2013) MicroRNA-­ mediated regulation of T helper cell differentiation and plasticity. Nat Rev Immunol 13(9):666–678. https://doi.org/10.1038/ nri3494 4. Chicaybam L, Sodre AL, Curzio BA, Bonamino MH (2013) An efficient low cost method for gene transfer to T lymphocytes. PLoS One 8(3):e60298. https://doi.org/10.1371/journal.pone.0060298 5. Chebel A, Rouault J-P, Urbanowicz I et al (2009) Transcriptional activation of hTERT, the human telomerase reverse transcriptase, by nuclear factor of activated T cells. J Biol Chem 284(51):35725–35734. https://doi. org/10.1074/jbc.M109.009183 6. Steiner DF, Thomas MF, Hu JK et al (2011) MicroRNA-29 regulates T-Box transcription factors and interferon-γ production in helper T

cells. Immunity 35(2):169–181. https://doi. org/10.1016/j.immuni.2011.07.009 7. Montoya MM, Ansel KM (2017) Small RNA transfection in primary human Th17 cells by next generation electroporation. J Vis Exp. https://doi.org/10.3791/55546 8. Kim JA, Cho K, Shin MS et al (2008) A novel electroporation method using a capillary and wire-type electrode. Biosens Bioelectron 23(9):1353–1360. https://doi. org/10.1016/j.bios.2007.12.009 9. Simpson LJ, Patel S, Bhakta NR et al (2014) A microRNA upregulated in asthma airway T cells promotes TH2 cytokine production. Nat Immunol 15(12):1162–1170. https://doi. org/10.1038/ni.3026 10. Pua HH, Steiner DF, Patel S et al (2016) MicroRNAs 24 and 27 suppress allergic inflammation and target a network of regulators of T Helper 2 cell-associated cytokine production. Immunity 44(4):821–832. https://doi. org/10.1016/j.immuni.2016.01.003 11. Tepper RI, Pattengale PK, Leder P (1989) Murine interleukin-4 displays potent anti-­ tumor activity in vivo. Cell 57(3):503–512 12. Schumann K, Lin S, Boyer E et al (2015) Generation of knock-in primary human T cells using Cas9 ribonucleoproteins. Proc Natl Acad

Functional Interrogation of microRNAs in CD4 T Cells Sci U S A 112(33):10437–10442. https:// doi.org/10.1073/pnas.1512503112 13. Hultquist JF, Schumann K, Woo JM et al (2016) A Cas9 ribonucleoprotein platform for functional genetic studies of HIV-­host interactions in primary human T cells. Cell Rep 17(5):1438–1452. https://doi. org/10.1016/j.celrep.2016.09.080 14. Park RJ, Wang T, Koundakjian D et al (2017) A genome-wide CRISPR screen identifies a restricted set of HIV host dependency factors.

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Nat Genet 49(2):193–203. https://doi. org/10.1038/ng.3741 15. Wang Y, Medvid R, Melton C et al (2007) DGCR8 is essential for microRNA biogenesis and silencing of embryonic stem cell self-­ renewal. Nat Genet 39(3):380–385. https:// doi.org/10.1038/ng1969 16. Wang Y, Baskerville S, Shenoy A et al (2008) Embryonic stem cell-specific microRNAs regulate the G1-S transition and promote rapid proliferation. Nat Genet 40(12):1478–1483. https://doi.org/10.1038/ng.250

Chapter 25 The Use of Biodegradable Nanoparticles for Tolerogenic Therapy of Allergic Inflammation Charles B. Smarr and Stephen D. Miller Abstract Antigen-specific tolerance is the ultimate aim of treatment of allergic diseases. Here, we describe methods for the use of biodegradable nanoparticles to safely induce tolerance for the prevention and treatment of allergic inflammation in mice. Antigen is either conjugated to the surface of carboxylated poly(lactide-co-­ glycolide) (PLG) or encapsulated within PLG nanoparticles, and the resulting antigen-associated nanoparticles are then washed prior to intravenous injection to inhibit antigen-specific allergic immune responses. Key words Tolerance, Nanoparticles, Allergy, Th2, Immunotherapy

1  Introduction The induction of antigen-specific tolerance is a long-sought goal of immunotherapy for the treatment of allergic disease. Current specific immunotherapy for the treatment of allergy involves the administration of escalating doses of soluble antigen (Ag) delivered subcutaneously or mucosally. Although this approach has been successful in the clinic, it runs the risk of inducing adverse events which necessitate a prolonged dose escalation [1]. One approach to improving the induction of antigen-specific tolerance involves the delivery of antigen in the context of a noninflammatory carrier. In murine models of inflammation, these carriers have included, among others, apoptotic cells [2–4] and nanoparticles [5–9]. Due to relative ease of manufacture and control, biodegradable antigen-­ associated nanoparticles (Ag-NP) represent the more clinically feasible implementation of this concept for immunotherapy. In a model of OVA/alum-induced Th2-driven allergic airway inflammation, both surface antigen-conjugated PLG nanoparticles (Ag-PLG) and antigen-encapsulated PLG nanoparticles [PLG(Ag)] effectively induced prophylactic tolerance to inhibit Th2 sensitization [9]. Administered post-sensitization, Ag-NP safely inhibited

R. Lee Reinhardt (ed.), Type 2 Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1799, https://doi.org/10.1007/978-1-4939-7896-0_25, © Springer Science+Business Media, LLC, part of Springer Nature 2018

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existing Th2 responses and prevented development of airway inflammation upon challenge [9]. Here, we outline methods to conjugate protein or peptide antigens to carboxylated PLG nanoparticles using 1-ethyl-3-(3′dimethylaminopropyl)-carbodiimide (ECDI) chemistry to catalyze covalent peptide bonds between amino groups on antigen and carboxyl groups on nanoparticles to generate Ag-PLG. We then describe proper preparation and administration of PLG-Ag and PLG(Ag) to the tail veins of recipient mice for the induction of antigen-specific tolerance. This protocol does not concern the production of biodegradable PLG nanoparticles, which is described in detail elsewhere [7, 10]. While previously used for induction of tolerance against OVA in an OVA/Alum model of allergic airway inflammation, these methods of tolerance induction could potentially be applied to any allergic model using a protein or peptide-­ based antigen. Thus, these methods present a tool to induce immune tolerance in an antigen-specific manner to study regulatory and inflammatory mechanisms of Th2-induced allergic inflammation.

2  Materials Use only sterile reagents throughout the protocol. 2.1  Common Reagents and Materials

1. Water: cell culture grade water. 2. PBS: Dulbecco’s phosphate-buffered saline. 3. Microcentrifuge tubes: 1.5 mL polypropylene tubes. 4. Cryovials: 2 mL high-density polyethylene cryogenic storage vials. 5. Cell strainers: 40 μm cell strainers. 6. Syringes: 1 mL tuberculin syringes. 7. Needles: 30 gauge ½ inch needles. 8. Thermal mixer with tube adapter. 9. Microcentrifuge.

2.2  Antigen-­ Conjugated PLG Nanoparticles and Antigen-­ Encapsulated PLG Nanoparticles

1. Carboxylated PLG nanoparticles: 500 nm carboxylated PLG nanoparticles. Nanoparticles can be prepared using a single emulsion technique as described in [7] or purchased from Phosphorex (see Notes 1 and 2). 2. ECDI: 1-ethyl-3-(3′-dimethylaminopropyl)-carbodiimide. 3. Antigen: Either whole protein or peptide antigens are acceptable for conjugation to carboxylated nanoparticles for the induction of tolerance. 4. Antigen-encapsulated PLG nanoparticles: 500 nm PLG nanoparticles with encapsulated antigen can be generated through a double-emulsion process as described in [10] or custom-ordered.

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3  Methods Carry out all procedures at room temperature using sterile reagents and conditions unless otherwise noted. 3.1  Antigen-­ Conjugated PLG Nanoparticles (Ag-PLG)

1. Allow reagents to warm to room temperature. 2. Weigh out 1.25 mg of lyophilized carboxylated PLG nanoparticles per dose, and transfer to a 1.5 mL microcentrifuge tube (see Note 3). 3. Resuspend nanoparticles in 200 μL of PBS (see Note 4). Add 1000  μL of PBS, and pipette thoroughly to mix. Centrifuge nanoparticles for 5 min at 3000 × g-force. 4. Repeat step 3 two additional times for a total of three washes. 5. Resuspend nanoparticles in 14.4 μL of PBS per mg of nanoparticles, and vortex for 5 s. 6. Dissolve ECDI in PBS to a concentration of 200 mg/mL immediately before use. Add 1.6 μL of ECDI solution per mg of nanoparticles to the suspension, and vortex for 5 s. 7. Add 4 μL of a 10 mg/mL Ag solution per mg of nanoparticle suspension, and vortex for 5 s (see Note 5). 8. Incubate reaction mixture for 1 h at 25 °C with shaking at 750 rpm. Vortex for 5 s every 10 min during the incubation. After 1 h, centrifuge for 5 min at 3000 × g-force. 9. Resuspend nanoparticles in 200 μL of PBS (see Note 6). Add 1000  μL of PBS, and pipette thoroughly to mix. Centrifuge nanoparticles for 5 min at 3000 × g-force. 10. Repeat step 9 two additional times for a total of three washes. 11. Resuspend nanoparticles to a final concentration of 6.25 mg/ mL in PBS. Filter nanoparticle suspension through a 40 μm cell strainer into a cryogenic vial to remove large aggregates (see Note 7). Antigen-conjugated nanoparticles are now ready for injection. 12. Vortex nanoparticles immediately before drawing for injection. Using a 1 mL syringe, draw up 240 μL of nanoparticles, remove air bubbles, and fill the void volume of a 30 gauge ½ in. needle. Inject 200 μL into the tail vein, vortexing the vial of nanoparticles and filling the syringe between each injection (see Notes 8–10).

3.2  Antigen-­ Encapsulated PLG Nanoparticles [PLG(Ag)]

1. Rehydrate antigen-encapsulated nanoparticles with cell culture grade water at a concentration of 12.5 mg/mL. Centrifuge nanoparticles for 5 min at 3000 × g-force (see Notes 3, 11, and 12).

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2. Resuspend nanoparticles in 200 μL of PBS. Add 1000 μL of PBS, and pipette thoroughly to mix. Centrifuge nanoparticles for 5 min at 3000 × g-force. 3. Repeat step 2 two additional times for a total of three washes with PBS. 4. Resuspend antigen-encapsulated nanoparticles to a final concentration of 12.5 mg/mL in PBS. Filter nanoparticle suspension through a 40 μm cell strainer into a cryogenic vial to remove large aggregates (see Note 7). Antigen-encapsulated nanoparticles are now ready for injection. 5. Vortex nanoparticles immediately before drawing for injection. Using a 1 mL syringe, draw up 240 μL of nanoparticles, remove air bubbles, and fill the void volume of a 30 gauge ½ in. needle. Inject 200 μL into the tail vein, vortexing the vial of nanoparticles and filling the syringe between each injection (see Notes 8 and 9). 3.3  Timing of Ag-NP Administration

1. Prophylactic administration: For prophylactic tolerance, nanoparticles should be administered at least 1 week prior to immunization (see Note 13). 2. Post-sensitization administration: For induction of tolerance post-sensitization, Ag-NP should be administered at least 1 week after the final sensitization, with at least 1 week allowed between doses of Ag-NP (see Note 14).

4  Notes 1. 500 nm particles have been demonstrated to be an ideal size for the induction of tolerance in murine models of inflammation [6]. In our experience, carboxylated PLG nanoparticles purchased from vendors such as Phosphorex tend to be less efficient and more variable when used to induce tolerance than comparable particles manufactured using the methods outlined in Yap et al. (2014) [7]. 2. Lyophilized nanoparticles are hygroscopic, and precautions should be taken to protect them from moisture. It is recommended to wrap nanoparticle storage containers in Parafilm and store in a desiccator under vacuum at room temperature. 3. Prepare an extra 20% of nanoparticles to account for void volume lost during injections. 4. Resuspension of pelleted nanoparticles can be difficult, requiring vigorous pipetting. Smaller volumes make this more aggressive resuspension easier. 5. If protein or peptide antigens are stored at a different concentration, simply adjust the volume of PBS used for nanoparticle

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resuspension accordingly. The reaction volume should be 20  μL per mg of nanoparticles with final concentrations of 50 mg/mL nanoparticles, 20 mg/mL ECDI, and 2 mg/mL antigen. 6. After antigen conjugation, nanoparticles will pellet in a streaky “comet” formation due to formation of particle-antigen-­ particle conjugates of varying sizes. Additional debris will coat the sides of the microcentrifuge tube. When resuspending, focus only on the pellet. 7. Nanoparticle suspensions can become stuck to the underside of the cell strainer. To maximize nanoparticle recovery, suction from the underside of the strainer using a fresh pipette tip, and add to the cryogenic vial. 8. Intravenous injection of nanoparticles is essential for efficient induction of tolerance as other administration routes such as mucosal or subcutaneous have been demonstrated to be either ineffective or pro-inflammatory [6, 7]. We have found that tail vein injection is the most consistent and efficient method of intravenous administration. 9. To dilate the tail vein and improve injection efficacy, dip tails in a beaker of warm water for a few seconds, and wipe dry immediately before injection. 10. Doing a single injection at a time is recommended to ensure even administration across multiple mice due to the quick rate at which nanoparticles settle out of suspension. 11. Thorough washing of PLG(Ag) prior to injection is critical to remove any non-encapsulated Ag remaining on the surface of lyophilized particles. Failure to wash particles may result in adverse reactions in sensitized recipient animals. 12. PLG(Ag) were successfully employed in prophylactic and postsensitization tolerance in a model of OVA-induced allergic airway inflammation [9]. In this model, tolerance was induced using a dose of 2.5 mg of PLG(OVA) delivering approximately 1 nmol of OVA. Varying encapsulation efficiencies of different antigens [8, 9] and the requirements of different models of inflammation will necessitate titration of doses in an antigenand model-dependent fashion. 13. In our experience, prophylactic tolerance with Ag-NP works best by allowing at least 1 week for development of regulatory responses to tolerogen before sensitization. With repeated sensitizations in a model of OVA-/Alum-induced allergic airway inflammation, we have had the best success with administering Ag-NP before each immunization [9]. However, optimal dosage and timing of Ag-NP administration should be determined individually for each model.

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14. In our experience in a model of OVA-/Alum-induced allergic airway inflammation, successful inhibition of inflammation was achieved with two doses of nanoparticles administered 1 and 2 weeks after the final sensitization, respectively [9]. Furthermore, superior inhibition, although not complete abrogation, of inflammation was achieved with a PLG(Ag) formulation. However, optimal dosage and timing of Ag-NP administration should be determined individually for each model.

Acknowledgments This study was supported by NIH Grant EB013198 and Juvenile Diabetes Research Society Grant 2-SRA-2014-279-Q-R. References 1. Smarr CB, Bryce PJ, Miller SD (2013) Antigenspecific tolerance in immunotherapy of Th2associated allergic diseases. Crit Rev Immunol 33(5):389–414 2. Getts DR, Turley DM, Smith CE, Harp CT, McCarthy D, Feeney EM, Getts MT, Martin AJ, Luo X, Terry RL, King NJ, Miller SD (2011) Tolerance induced by apoptotic antigen-coupled leukocytes is induced by PD-L1+ and IL-10-producing splenic macrophages and maintained by T regulatory cells. J Immunol 187(5):2405–2417. https://doi.org/ 10.4049/jimmunol.1004175 3. Prasad S, Kohm AP, McMahon JS, Luo X, Miller SD (2012) Pathogenesis of NOD diabetes is initiated by reactivity to the insulin B chain 9-23 epitope and involves functional epitope spreading. J Autoimmun 39(4):347– 353. https://doi.org/10.1016/j.jaut.2012. 04.005 4. Smarr CB, Hsu CL, Byrne AJ, Miller SD, Bryce PJ (2011) Antigen-fixed leukocytes tolerize Th2 responses in mouse models of allergy. J Immunol 187(10):5090–5098. https://doi. org/10.4049/jimmunol.1100608 5. Clemente-Casares X, Blanco J, Ambalavanan P, Yamanouchi J, Singha S, Fandos C, Tsai S, Wang J, Garabatos N, Izquierdo C, Agrawal S, Keough MB, Yong VW, James E, Moore A, Yang Y, Stratmann T, Serra P, Santamaria P (2016) Expanding antigen-specific regulatory networks to treat autoimmunity. Nature 530(7591):434–440. https://doi. org/10.1038/nature16962 6. Getts DR, Martin AJ, McCarthy DP, Terry RL, Hunter ZN, Yap WT, Ge tts MT, Pleiss M, Luo

X, King NJ, Shea LD, Miller SD (2012) Microparticles bearing encephalitogenic peptides induce T-cell tolerance and ameliorate experimental autoimmune encephalomyelitis. Nat Biotechnol 30(12):1217–1224. https:// doi.org/10.1038/nbt.2434 7. Hunter Z, McCarthy DP, Yap WT, Harp CT, Getts DR, Shea LD, Miller SD (2014) A biodegradable nanoparticle platform for the induction of antigen-specific immune tolerance for treatment of autoimmune disease. ACS Nano 8(3):2148–2160. https://doi. org/10.1021/nn405033r 8. McCarthy DP, Yap JW, Harp CT, Song WK, Chen J, Pearson RM, Miller SD, Shea LD (2017) An antigen-encapsulating nanoparticle platform for TH1/17 immune tolerance therapy. Nanomedicine 13(1):191–200. https://doi.org/10.1016/j.nano.2016. 09.007 9. Smarr CB, Yap WT, Neef TP, Pearson RM, Hunter ZN, Ifergan I, Getts DR, Bryce PJ, Shea LD, Miller SD (2016) Biodegradable antigen-associated PLG nanoparticles ­tolerize Th2-mediated allergic airway inflammation pre- and postsensitization. Proc Natl Acad Sci U S A 113(18):5059–5064. https://doi.org/10.1073/pnas. 1505782113 10. Yap WT, Song WK, Chauhan N, Scalise PN, Agarwal R, Miller SD, Shea LD (2014) Quantification of particle-conjugated or particle-encapsulated peptides on interfering reagent backgrounds. Biotechniques 57(1):39–44. https://doi.org/10.2144/ 000114190

Chapter 26 Assessing the Mouse Intestinal Microbiota in Settings of Type-2 Immune Responses Mei San Tang, Rowann Bowcutt, and P’ng Loke Abstract The microbial communities that reside within the mammalian host play important roles in the development of a robust host immune system. With the advent of sequencing technology and barcoding strategy of the bacterial 16S ribosomal RNA (rRNA) gene, microbiota studies are becoming more economical but also more important in many immunology studies. Here, we described a representative study protocol to characterize how the microbiota changes during an intestinal helminth infection, with emphasis on subtle aspects of the experimental design that are critical for data interpretation. Key words 16S rRNA, Microbiota, Helminth, Type 2 immunity

1  Introduction The microbiota is the catalog of microbes that reside on a given host. In the past few years, there is a growing interest in the field of immunology to study the functional roles that microbiota at various body surfaces play in the induction, maintenance, and education of the host mammalian immune system [1]. Throughout the course of human evolution, helminth colonization had occurred in majority of the human population, and this had remained true until quite recently [2]. As such, in order to avoid rejection, helminths have evolved mechanisms to regulate the immune system of their hosts [3], including production of various immune regulatory molecules [4] encoded into their genomes [5]. The intestinal helminths are among the most common forms of helminth colonization, and this also coincides with the locale of the majority of commensal bacteria in the human host. This sharing of the same niche points toward the possibility that intestinal worms and the microbiota interact to further modulate the host immune Mei San Tang and Rowann Bowcutt contributed equally to this work. R. Lee Reinhardt (ed.), Type 2 Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1799, https://doi.org/10.1007/978-1-4939-7896-0_26, © Springer Science+Business Media, LLC, part of Springer Nature 2018

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system in a complex, three-way interaction [6]. Indeed, recent studies in both human and mouse models have shown that intestinal worm infection can cause changes in the microbiota composition that may have beneficial effects for the host [7, 8]. As a result of reduced sequencing cost and the ability to multiplex samples using barcoding strategies for the 16S ribosomal RNA (rRNA) gene as a phylogenetic marker [9], microbiota profiling experiments are increasingly accessible, allowing more researchers to characterize the microbial communities from their model system of interest. Here, we have outlined a strategy to characterize the changes in the gut microbial communities that occur during helminth infection, both in the absence and presence of a normal type 2 immune response. In this example, we compare the microbiota of C57BL/6 mice to STAT6KO mice, which lack the transcription factor Stat6, a key regulator of type 2 immunity, over the course of an acute Trichuris muris (T. muris) infection. T. muris is a natural parasite of mice and has been used as a model for Trichuris trichiura infection in humans. T. muris has proved an important model system for understanding resistance and immunity to helminth infection [10].

2  Materials 2.1  T. muris Infection

1. C57BL/6J mice (Jackson ID: 000664). 2. STAT6KO (Jackson ID: 005977). 3. T. muris embryonated eggs (E isolate) in phosphate-buffered saline (PBS). See Fig. 1 to determine embryonated eggs. 4. Ultrapure water. 5. Magnetic stirrer. 6. Magnetic stirrer bar. 7. Dissecting microscope.

a

b

c

Fig. 1 Three possible appearances of T. muris eggs. (a) Clear visible worm structure inside the egg: embryonated. (b) Egg with granular appearance: not embryonated. (c) Empty egg: not embryonated

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8. Microscope slide. 9. P200 pipette and tips. 10. 10 mL syringe. 11. Oral gavage needle for mice. 12. 1.5 mL Eppendorf tubes for stool sample collection. 13. Click counter. 2.2  Assessing Worm Burdens

1. Petri dish. 2. Curved forceps. 3. Blunt ended scissors. 4. PBS. 5. Dissecting microscope. 6. Click counter.

2.3  16S rRNA Library Preparation

1. DNA isolation kit. 2. Tissue homogenizer. 3. NanoDrop fluorospectrometer. 4. RNase decontamination solution. 5. 70% ethanol. 6. Nuclease-free water. 7. 5PRIME HotMasterMix 2.5×. 8. Forward PCR primer construct for V4 region of the bacterial 16S rRNA gene (F515) [9]. 9. Reverse PCR barcoded primer construct (R806) for V4 region of the bacterial 16S rRNA gene (R806) [9]. 10. PCR machine. 11. 96-well PCR plate aluminum foil adhesive film for freezer storage. 12. 96-well PCR plate clear polyester adhesive film for PCR reaction. 13. PCR purification kit. 14. PicoGreen Quant-iT dsDNA kit. 15. 96-well solid black polystyrene microplate. 16. Fluorescence 96-well plate reader. 17. Qubit dsDNA BR Assay Kit. 18. Qubit fluorometer. 19. P1000, P200, and P10 filter tips. 20. 1.5 mL Eppendorf tubes. 21. (Optional) Gel extraction kit.

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3  Methods 3.1  Cohousing of Mice

1. Cohouse 6-week-old C57BL/6 (N = 5 mice) and STAT6KO (N = 5 mice) in cages containing both genotypes for 2 weeks prior to infection to normalize the gut microbiota. Mice are coprophagic, and cohousing provides some gut normalization. A less stringent strategy that is sometimes necessary would be to swap the genotypes into each other’s cages or to thoroughly mix their bedding continuously prior to the start of the experiment. The most stringent strategy that corrects for maternal cage effects is to intercross heterozygous mice, e.g., STAT6+/− mice, so that the resulting littermates will include mixture of homozygotes, heterozygotes, and wild-type mice born from the same dams.

3.2  Longitudinal Acute T. muris Infection

1. Wash T. muris eggs by centrifuging at 2000 × g for 5 min in ultrapure water. 2. Resuspend T. muris eggs in a known volume of ultrapure water. 3. Pipette 50 μL of the eggs in suspension on a microscope slide. 4. Count the number of embryonated eggs in the 50 μL aliquot using a microscope and click counter. 5. Repeat steps 3 and 4 twice with a fresh 50 μL aliquot to get an average number of embryonated eggs per 50 μL. Only eggs with the appearance shown in Fig. 1a should be counted. It does not matter if eggs with the appearance of Fig. 1b or c are also given to the mice; however, they should not be considered in the final egg count. 6. Multiply this average by 4 to get the number of eggs in 200 μL. 7. Adjust the volume of egg suspension accordingly, so you have approximately 150–200 eggs per 200 μL. 8. Tag mice in order to keep track of which mice each stool sample came from (see Note 1). 9. Infect each mice by giving 200 μL of egg solution by oral gavage. 10. Collect stool into a 1.5 mL Eppendorf from infected mice twice weekly up to day 35 post-infection. 11. Stool should be immediately frozen at −80 °C until DNA isolation step. 12. Sacrifice mice on day 35 (see Note 2).

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1. Remove the cecum and colon from the mice and place in a petri dish. At this point, the cecum and colon can be frozen at −20 °C until you have time to do the worm counts. 2. Open the cecum and colon longitudinally, and wash fecal contents in PBS. 3. Adult worms should be visible by the eye at day 35 post-­ infection. Worms appear as thin white threadlike structures (see Note 3). 4. Using curved forceps, gently pull out the worms from the mucosa. At this stage, worm burdens can be quantified if required by counting using a microscope and click counter. Alternatively, if you are just interested in the presence/absence of worms, no quantification is needed.

3.4  16S rRNA Amplification

1. Thaw stool samples from −80 °C. Isolate DNA using a DNA isolation kit of choice following the manufacturer’s protocol (see Note 4). 2. Quantitate the concentration and quality of isolated DNA samples using NanoDrop fluorospectrometer (see Notes 5 and 6). 3. Plate isolated DNA onto a 96-well PCR plate. This will allow for the use of multichannel pipette for sample transfer during the PCR amplification step and for more efficient, high-­ throughput library preparation. 4. Once DNA samples are plated completely, generate a spreadsheet containing PCR plate well number to the matching sample identifier (see Note 7). 5. Seal the plate with aluminum foil adhesive film suitable for storage in freezer. Store DNA in −20 °C, or proceed immediately to PCR amplification. 6. Designate a specific reverse barcoded primer construct to each sample, and include the information into the spreadsheet generated in step 4. 7. PCR should be performed in a clean PCR workstation. Clean workstation and all required instruments (micropipettes, pipette tip boxes) using 70% ethanol and RNase decontamination solution. Irradiate workstation and instruments for 30 min. Always use filter tips for pipetting throughout library preparation. 8. Calculate the total number of PCR reactions, and prepare a PCR mix for enough number of reactions. Each DNA sample will be PCR-amplified in triplicate. Each PCR reaction should contain:

(a) 12 μL nuclease-free water

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(b) 10 μL 5PRIME HotMasterMix 2.5×



(c) 1 μL forward primer (5 μM)—common for all samples



(d) 1  μL reverse barcoded primer (5 μM)—specific for each sample



(e) 1 μL DNA sample

9. Cycling protocol:

(a) 94 °C for 3 min



(b) 94 °C for 45 s



(c) 50 °C for 1 min



(d) 72 °C for 1.5 min



(e) Repeat steps 2–4 for 35 cycles



(f) 72 °C for 10 min



(g) 4 °C forever

10. Once PCR is completed for all samples, combine the amplification product for each sample from the triplicate PCR reactions into a single PCR plate (see Note 8). Seal the PCR plate with clear polyester adhesive film, and store in 4 °C, or proceed immediately to quantification of amplicon using PicoGreen Quant-iT kit (see Note 9). 3.5  Amplicon Quantification, Pooling, and Purification

1. Prepare a double-stranded DNA standard curve using the stock lambda DNA (100 μg/mL) provided in the Quant-iT PicoGreen kit following the table below. This is slightly different from the suggested standard curve in the Quant-iT PicoGreen kit manufacturer’s instructions. Standard concentration

Input

Nuclease-­free water

50 μg/mL

10 μL stock lambda DNA

10

25 μg/mL

5 μL lambda DNA 50 μg/mL

10 μg/mL

2 μL stock lambda DNA

5 μg/mL

2 μL lambda DNA 50 μg/mL 18

1 ng/mL

2 μL lambda DNA 10 μg/mL 18

5 18

2. Dilute the Quant-iT PicoGreen reagent 1:200 using the TE buffer provided. Each sample to be quantitated requires 198  μL of diluted PicoGreen regent. Calculate enough volume for each of the amplified samples and an additional ten reactions for the standard curve. PicoGreen reagent should be diluted fresh during each experiment and protected from light as much as possible.

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3. Prepare a 96-well solid black microplate. Designate wells for samples and wells for standard curve DNA. 4. Aliquot 198 μL of diluted PicoGreen reagent into each well of the 96-well plate. 5. Add 2 μL of amplified DNA into wells designated for samples using a multichannel pipette. Mix well. 6. Add 2 μL of standard curve DNA into wells designated for standard curve. Mix well. 7. Incubate the mixture for 5 min at room temperature, protected from light. 8. After incubation, measure the sample fluorescence using a fluorescence microplate reader and standard fluorescein wavelengths (excitation ~480 nm, emission ~520 nm). 9. Determine the concentration of the amplified DNA for each sample using the standard curve generated from step 1. 10. To decide on a quantity of DNA to pool from each sample, divide the desired DNA quantity by the concentration of the sample. Each sample should be pooled at equal DNA quantity (not equal concentration). As such, choose a DNA quantity that uses ≤70  μL volume from each of the amplified DNA sample. This is because the maximum deliverable volume from each amplified sample is slightly less than 75 μL (25 μL from each PCR reaction, a total of 75 μL from triplicate reaction, minus approximated volume loss) (see Note 10).

11. Combine all samples into a single pool, taking from each sample the volumes calculated from step 10. The amplicon pool can be kept in a 1.5 mL Eppendorf tube. If the total volume of the pooled amplicon is greater than 1.5 mL, the pool may be split into multiple 1.5 mL Eppendorf tubes. Store the pooled amplicon in 4 °C, or proceed immediately to DNA purification using a purification kit of choice for PCR product. Follow the manufacturer’s instructions (see Note 11).

12. Store purified DNA in 4 °C, or proceed immediately to the final quantitation of the purified amplicon pool with the Qubit dsDNA BR Assay Kit (see Notes 12 and 13). 13. Quantitation of the purified amplicon pool can be done by following the exact instructions provided in the Qubit dsDNA BR Assay Kit. First, prepare the Qubit working solution by diluting the Qubit dsDNA BR reagent (Dye; Component A) 1:200 in Qubit dsDNA BR buffer (Component B). 14. Prepare the two standards required for standard curve generation by adding 10 μL of each standard to 190 μL diluted Qubit dsDNA BR reagent.

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15. Prepare the sample to be quantitated by adding 2 μL of the purified amplicon pool to 198 μL of the diluted Qubit dsDNA BR reagent. 16. Quantitate the concentration of the purified amplicon pool using the Qubit fluorometer. 17. Calculate the molarity of the purified amplicon pool. DNA molarity for each amplicon pool can be calculated from the DNA concentration quantitated in step 20 using the formula: number of samples in pool × DNA concentration (ng/ μL) × 106 ÷ 650 ÷ length of amplicon. The amplicon length is 381 bp using the F515/R806 PCR primer constructs referenced in this protocol. 18. Dilute the purified amplicon pool to 50 nM. The diluted amplicon pool is now ready to be sequenced. 3.6  Considerations for Sequencing and Data Analysis

1. Batch effect: Batch effect is a well-described problem in any sequencing experiment and can arise at any step throughout the library preparation and sequencing process. Document as much technical details throughout the library preparation process as possible (e.g., the date for which each step was performed, the name of the scientist who performed that step, and where it was performed), and sequence all of the samples in a single batch. If this is not possible, design a balanced experiment such that all biological conditions of interest must be present in each of the sequencing libraries and that no biological condition is present exclusively in one sequencing library and not the others. This will avoid any biological effect being confounded by technical batch effects. 2. Sequencing assay: The decisions of sequencing depth and read length are often a consideration between multiplexing more samples to minimize sequencing cost and the ability to discover rare bacterial species. As a practical suggestion, we typically do not multiplex more than 192 samples (equivalent to samples in two 96-well plates) for sequencing on the Illumina MiSeq platform using paired-end 150 bp reads, aiming for >10,000 reads per sample. The sequencing runs can be repeated for the same set of samples, and reads from both sequencing runs merged if greater depth is desired. 3. Computational resources: There are many different computational tools available to analyze 16S rRNA sequencing data. One popular choice is the open-source software Quantitative Insights Into Microbial Ecology (QIIME). The official QIIME website has several well-documented step-by-step tutorials to help bench scientists perform basic analyses of 16S rRNA sequencing data, starting with processing steps of raw sequence reads to identifying changes in microbial diversity and identify-

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ing differential bacterial species. Processing of raw sequence reads to identify and quantitate abundance of operational taxonomic units (OTUs) will require access to a high-performance computing cluster. Once the OTU table is generated, it is feasible to perform the downstream diversity and differential analyses independent of a high-performance computing facility, although some steps that require rarefaction (random sampling of sequences) can still be computationally intensive, especially with higher sequencing depth and more samples. Installation of the QIIME software can be time- and effort-consuming, as it requires many different software dependencies. For beginners, we recommend working with your institution high-­ performance computing facility manager to have the QIIME software installed on an institution shared computing resource for the raw sequence processing step and using MacQIIME, a compiled version of QIIME, for downstream analyses on the OTU table. MacQIIME is an open-source software available for download at http://www.wernerlab.org/software/macqiime. For users without access to a high-performance computing facility or a Mac operating system, we suggest, as an alternative, the Amazon Cloud Service, where QIIME is available. 4. Learning resources: We suggest the following resources that are friendly to beginners in the bioinformatics of 16S microbiota analyses. Since QIIME is a command line-based tool, users will also have to be able to perform basic Unix operations. The following resources will also include references where users can learn basic Unix operations sufficient for their 16S microbiota analyses.

(a) Official tutorials from the QIIME website—http://qiime. org/tutorials/index.html



(b) Werner lab website—http://www.wernerlab.org/teaching/qiime/overview



(c) An article by Morgan and Huttenhower [11] providing good overview of key concepts in microbiome analysis

4  Notes 1. Mice can be tracked individually by using an ear-punch coding system to identify specific mice within the cage. It should be noted which samples came from which mouse throughout the course of the experiment. This then allows for variation that occurs between mice over the course of a longitudinal study to be estimated through multilevel modeling of OTU abundances.

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2. The natural course of T. muris infection in C57BL/6 mice terminates around day 35 post-infection due to parasite expulsion by the host’s immune response. Therefore, we suggest sacrificing mice at this time point. However, you may want to take further samples to see how long any microbiota changes persist after parasite infection is expelled. 3. On rare occasions, mice do not become infected by T. muris. Therefore, to determine if mice are infected, it would be best practice to analyze worm burdens at the end of the experiment. STAT6KO mice are susceptible to infection, and therefore, worms should be present in the cecum and large intestine of these mice at day 35 post-infection. C57BL/6 mice may still harbor a few worms at day 35, although this cannot be guaranteed as these mice are resistant to a high dose worm infection and would have nearly expelled all parasites by this time point. 4. There are several different DNA isolation kits commercially available, and this might have to be decided based on the sample type. Choose a kit specific optimized for DNA isolation from stool samples. Use the yield and quality of the isolated DNA from NanoDrop readings to guide decision-making. 5. High DNA concentration can inhibit PCR and should be diluted. In general, we suggest diluting samples with concentration >100 ng/μL by 1:2 dilution. 6. We often do not set a lower limit of DNA concentration for PCR amplification, since sequencing assay generally does not require large amount of DNA material, and it is not impossible for low concentration samples to be sufficiently amplified for sequencing purpose using this protocol. If reagent is not a limiting factor, a practical suggestion is to PCR amplify all samples (including very low concentration samples) with one round of triplicate reactions and quantitate the amplicon yield to decide if the samples should be included in the sequencing run. 7. Always make sure to be careful of plate layout and that correct samples are transferred between plates. Also be sure to include a negative control well using nuclease-free water in place of DNA sample. 8. This protocol assumes not more than 96 samples (equivalent to one 96-well plate) are being prepared for 16S rRNA sequencing. If there are more than 96 samples, samples should be pooled by PCR plates and samples from each PCR plate treated as one amplicon pool. Each amplicon pool should be quantified and purified independently in Subheading 3.4. Each of the final diluted 50 nM amplicon pool can then be combined into a single 1.5 mL Eppendorf tube for sequencing. Keeping individual amplicon pools per PCR plate helps keep

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track of the number of samples contained in each pool, and this information can be used to adjust volume of the final amplicon pool to ensure even coverage of sequencing reads. 9. For temporary storage of amplicon before quantitation with PicoGreen, the regular clear PCR plate seal can be used. However, keep in mind that these seals are not meant for longterm storage and can lead to unnecessary volume loss if left over a longer period of time. For best results, always proceed to the next steps in this protocol as soon as you can. 10. We suggest choosing a quantity between 50 and 300 ng. Choose the highest possible DNA quantity deliverable from your samples. If samples have low DNA concentrations, the PCR amplification step can be repeated a few more times to obtain enough material for the desired quantity of DNA. This can be done either for the entire set of samples or for specific samples with low DNA concentrations. 11. If using a column-based DNA purification kit, make sure not to overload the DNA-binding capacity of the purification column. 12. Optional: To ensure purity of the amplicon, run a 1.5% agarose gel with the purified amplicon, and gel purify the DNA band at ~400 bp for downstream steps using a gel extraction kit of choice. 13. There are two versions of the Qubit DNA Quantitation Assay—high sensitivity (HS) and broad range (BR). Be sure to select the correct assay while reading off the Qubit fluorometer.

Acknowledgments We thank Laurie M. Cox and the Blaser lab at the New York University School of Medicine for assistance with setting up 16S rRNA sequencing in our laboratory and for providing the reverse barcoded primer constructs. We thank David Artis for seed stock of T. muris. References 1. Belkaid Y, Hand Timothy W (2014) Role of the microbiota in immunity and inflammation. Cell 157(1):121–141. https://doi. org/10.1016/j.cell.2014.03.011 2. Girgis NM, Gundra UM, Pn L (2013) Immune regulation during helminth infections. PLoS Pathog 9(4):e1003250. https://doi.org/10.1371/journal. ppat.1003250

3. McSorley HJ, Maizels RM (2012) Helminth infections and host immune regulation. Clin Microbiol Rev 25(4):585–608. https://doi. org/10.1128/CMR.05040-11 4. Harnett W, Harnett MM (2010) Helminth-­ derived immunomodulators: can understanding the worm produce the pill? Nature Rev 10(4):278–284. https://doi.org/10.1038/ nri2730

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5. Brindley PJ, Mitreva M, Ghedin E, Lustigman S (2009) Helminth genomics: the implications for human health. PLoS Negl Trop Dis 3(10):e538. https://doi.org/10.1371/journal.pntd.0000538 6. Mutapi F The gut microbiome in the helminth infected host. Trends Parasitol 31(9):405–406. https://doi.org/10.1016/j.pt.2015.06.003 7. Lee SC, Tang MS, Lim YAL, Choy SH, Kurtz ZD, Cox LM, Gundra UM, Cho I, Bonneau R, Blaser MJ, Chua KH, Pn L (2014) Helminth colonization is associated with increased diversity of the gut microbiota. PLoS Negl Trop Dis 8(5):e2880. https://doi.org/10.1371/journal.pntd.0002880 8. Ramanan D, Bowcutt R, Lee SC, Tang MS, Kurtz ZD, Ding Y, Honda K, Gause WC, Blaser MJ, Bonneau RA, Lim YA, Loke P, Cadwell K (2016) Helminth infection pro-

motes colonization resistance via type 2 immunity. Science 352(6285):608–612. https:// doi.org/10.1126/science.aaf3229 9. Caporaso JG, Lauber CL, Walters WA, Berg-­ Lyons D, Lozupone CA, Turnbaugh PJ, Fierer N, Knight R (2011) Global patterns of 16S rRNA diversity at a depth of millions of sequences per sample. Proc Natl Acad Sci U S A 108(Suppl 1):4516–4522. https://doi. org/10.1073/pnas.1000080107 10. Grencis RK (2015) Immunity to helminths: resistance, regulation, and susceptibility to gastrointestinal nematodes. Annu Rev Immunol 33:201–225. https://doi.org/10.1146/ annurev-immunol-032713-120218 11. Morgan XC, Huttenhower C (2012) Chapter 12: human microbiome analysis. PLoS Comput Biol 8(12):e1002808. https://doi. org/10.1371/journal.pcbi.1002808

Chapter 27 The Use of CRISPR-Cas9 Technology to Reveal Important Aspects of Human Airway Biology Azzeddine Dakhama and Hong Wei Chu Abstract The CRISPR-Cas9 technology is a powerful tool that enables site-specific genome modification (gene editing) and is increasingly used in research to generate gene knockout or knock-in in a variety of cells and organisms. This chapter provides a brief overview of this technology and describes a general methodology applicable to human airway biology research. Key words CRISPR-Cas9, Gene editing, Lentivirus, Lung, Airway epithelium, Primary cells

1  Introduction Recent advances in molecular biology have led to the development of novel technologies that enable site-specific ­ modifications (­editing) at the genome level. In addition to ­facilitating studies of gene function, these technologies provide new opportunities for future applications in medicine and ­biotechnology. ­Genome-editing technologies are based on the use of highly specific nucleases such as CRIPSR-Cas9 (clustered regularly interspaced short ­palindromic repeats and CRISPRassociated protein-9 nuclease) system, which induces doublestrand breaks (DSB) at a specific location within ­targeted DNA sequences. The CRISPR-Cas9 system involves the use of a small RNA called single guide RNA (sgRNA) to guide Cas9 nuclease to a specific genomic locus. CRISPR-Cas nuclease systems are naturally found in ­bacteria and used by these microorganisms to defend t­ hemselves against invading pathogens (e.g., bacteriophages) [1–3]. The system involves two components, CRISPR and Cas nucleases. CRISPR are repetitive DNA motifs interspaced by short DNA elements with unique sequences called protospacers [4, 5]. The latter are bacteriophage DNA sequences that have been copied by bacteria during initial infection and are preserved as m ­ emory R. Lee Reinhardt (ed.), Type 2 Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1799, https://doi.org/10.1007/978-1-4939-7896-0_27, © Springer Science+Business Media, LLC, part of Springer Nature 2018

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signature of bacteriophages that infected them in the past [6, 7]. On r­ einfection, these sequences are rapidly transcribed into short RNAs, called CRISPR RNA (crRNA), which then guide Cas nuclease to locate the invading bacteriophage DNA and eliminate it by cleavage [8]. Unlike other CRISPR-Cas systems, the CRISPR-Cas9 system only relies on a single protein, Cas9, to localize and eliminate ­target DNA sequence. This property made it more suitable as a candidate technology for gene editing in mammalian cells [9, 10]. The most commonly used CRISPR-Cas9 system is derived from Streptococcus pyogenes. When guided to a target DNA sequence, this Cas9 induces double-strand DNA cleavage of three base pairs upstream of a “NGG” protospacer-associated motif (PAM) [11]. The resulting DNA break can be repaired via homology-directed repair (HDR), using a donor DNA template. In the absence of donor DNA template, the DNA break is repaired via the cellular nonhomologous end-joining (NHEJ) DNA repair pathway. This repair mechanism is error-prone and introduces insertions or ­deletions (indel) that can alter the targeted gene locus and disrupt function. Below, we describe a general methodology for the use of sgRNA-guided CRISPR/Cas9 system to disrupt gene function in primary human airway epithelial cells. The major steps include the design of sgRNA, sgRNA cloning into lentiCRISPR vector for ­co-­expression with Cas9, packaging of the expression vector into lentivirus, infection of primary airway epithelial cells, and analysis of indel mutations. For additional information, the readers are referred to the literature [10, 12].

2  Materials 2.1  Cells and Bacteria

1. Human embryonic kidney 293FT cell line (see Note 1). 2. Primary human airway epithelial cells (see Note 2). 3. Chemically competent E. coli. Stbl3 or equivalent strains are recommended for lentivector preparation.

2.2  Culture Media

1. D10 culture medium: Dulbecco’s modified Eagle medium (DMEM) containing high glucose (4.5 mg/mL), sodium pyruvate (110 mg/mL), and l-glutamine (580 mg/mL), ­ supplemented with penicillin (50 U/mL), streptomycin ­ (50 μg/mL), and 10% heat-inactivated fetal bovine serum. 2. Bronchial epithelial cell growth medium (BEGM). BEGM consists of bronchial epithelial basal medium (BEBM) ­ supplemented with SingleQuot supplements and growth ­

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f­actors (proprietary formulation), as directed by the manufacturer (Lonza Biologics Inc, Portsmouth, NH). 3. Opti-MEM I reduced serum media (Life Technologies). Use for optimal plasmid transfection into 293FT cells. 4. LB media + AMP: LB broth supplemented with ampicillin (100 μg/mL) for selection of transformed bacteria. 5. LB Agar + AMP: LB agar supplemented with ampicillin (100 μg/mL). 6. SOC medium. 2.3  Reagents and Buffers

1. All-in-one lentiCRISPR expression plasmid, available from Addgene (Cambridge, MA). 2. T4 polynucleotide kinase from New England Biolabs (NEB) (Ipswich, MA) or from a preferred supplier. 3. T4 DNA ligase and associated 10× ligation buffer from New England Biolabs or from a preferred supplier. 4. BsmBI restriction enzyme and associated 10× buffer New England Biolabs or from a preferred supplier. 5. ATP (adenosine-5′-triphosphate). 6. Polybrene (sterile, 10 mg/mL) from EMD Millipore (Billerica, MA). 7. High-fidelity DNA polymerase. 8. T7 endonuclease I and associated buffer. 9. HEPES buffer (1 M solution). 10. PBS (cell culture grade, sterile phosphate buffer solution). 11. Type I bovine collagen solution (PureCol, Advanced BioMatrix, Carlsbad, CA). 12. Nuclease-free water. All molecular reagents and reactions should be prepared with nuclease-free water. 13. Antibodies for Western blot analyses (gene target-specific).

2.4  Kits

1. Plasmid Miniprep kit (e.g., QIAprep Spin Miniprep kit from Qiagen). 2. PCR purification kit (e.g., QIAquick PCR purification kit from Qiagen). 3. Lentivirus packaging system (Lenti-Pac HIV expression ­packaging kit) from Genecopoeia (Rockville, MD) or from ­preferred supplier.

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3  Methods 3.1  Design sgRNA

1. Design sgRNA for Cas9 using available online tools to identify 20-nucleotide guide sequences located on a target gene, upstream of a “NGG” PAM (as shown below) (see Notes 3–5). Target (20 bp) PAM DNA positive strand 5′-…NNNNNNNNNNNNNNNNNNNNGG…-3′ DNA negative strand 3′-…NNNNNNNNNNNNNNNNNNNNCC…-5′

2. Determine the reverse complement sequence for sgRNA (guide sequence), either manually or using available online tools (see Note 6). 3.2  Design Oligonucleotides

1. Synthesize oligonucleotide sequences for the sgRNA and its reverse complement. For the purpose of cloning into lentiCRISPR expression plasmid (Addgene), the forward oligonucleotide sequence should be designed with a 5′-CACCG sequence (BsmBI site) added on the 5′ end of sgRNA. 2. The reverse oligonucleotide should be designed with a 5′-AAAC overhang sequence added on the 5′ end and a C nucleotide added on the 3′ end. When annealed, both oligonucleotides will have overhangs (as shown below) that can be ligated to the processed plasmid. 3. Forward oligo: 5′-CACCGNNNNNNNNNNNNNNNNNNNN-3′. 4. Reverse 3′-CNNNNNNNNNNNNNNNNNNNNCAAA-­5′.

3.3  Clone sgRNA into lentiCRISPR for Co-expression with Cas9

oligo:

LentiCRISPR is an all-in-one expression plasmid that contains the Cas9 sequence and gRNA scaffold. It also contains puromycin resistance gene for selection in mammalian cells. 1. Phosphorylate and anneal oligonucleotides (i.e., sgRNA and its reverse complement). This can be done simultaneously as a single-­step reaction. 2. In a 200 μL PCR tube, add reaction components in the following order: 12 μL of nuclease-free water, 2 μL forward oligonucleotide (from 100 μM stock), 2 μL of reverse oligonucleotide (from 100 μM stock), 2 μL of 10× ligation buffer, and 2 μL of T4 polynucleotide kinase (5 units/reaction). Place the tube in a thermal cycler, and incubate for 30 min at 37 °C (for phosphorylation), followed by 5 min at 95 °C (for denaturation), then ramping down to 25 °C at a rate of 6 °C/min (for annealing) (see Note 7).

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3. Ligate the annealed oligonucleotides to “lentiCRISPR” plasmid using the Golden Gate assembly strategy, allowing both digestion and ligation to be carried out simultaneously. (a)  Dilute the phosphorylated annealed oligonucleotides (from Step 1) 1:10 with nuclease-free water (final concentration will be 1 μM).

3.4  Transform Stable Competent E. coli



(b) Prepare a 20 μL reaction mix in 200 μL PCR tube: 1 μL plentiCRISPR plasmid (25 ng), 1 μL diluted annealed oligos (1 μM final), 2 μL 10× restriction enzyme buffer, 2 μL BsmBI (20 U), 1.5 μL T4 DNA ligase (600 U), 2 μL ATP (10 mM), and 10.5 μL nuclease-free water.



(c) Place the tube in a thermal cycler, and run the reaction with the following cycling conditions: (37 °C for 5 min, 20 °C for 5 min) × 20 cycles, followed by 1 cycle at 80 °C for 20 min to inactivate the enzymes (see Note 8).

1. Add 2 μL of the ligation product to 50 μL of ice-cold competent Stbl3 E. Coli strain. 2. Gently flick the tube four to five times to mix (but do not vortex), and incubate on ice for 30 min. 3. Heat shock at 42 °C for 30 s and return to ice for 5 min. 4. Add 50 μL of SOC medium, and plate onto LB agar plates containing 100 μg/mL ampicillin. Incubate overnight at 37 °C. 5. On the next day, pick two to three individual colonies, and check for the presence of sgRNA insert in the plasmid.

3.5  Package lentiCRISPR_sgRNA into Lentivirus, and Infect Target Cells



(a) Inoculate each individual colony into a 4 mL of LB broth containing ampicillin (100 μg/mL), and incubate for overnight culture at 37 °C with agitation (200–250 rpm).



(b) Isolate plasmid from each culture using Qiagen Miniprep plasmid kit as directed by the manufacturer (Qiagen).



(c) Analyze plasmid by sequencing using a forward primer from the U6 promoter region (e.g., LKO.1 primer: 5′-GAT ACA AGG CTG TTA GAG TAA TT-3′).



(d) Save plasmids with sequence-verified insert, and discard negative plasmids (with no inserts).

Unless large batches are prepared for virus purification and storage, this step can be timed in such way that target cells are ready for infection on the lentivirus harvest day (see Note 9).

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3.5.1  Day 0 (Morning, 8–9 AM)

Seed 293FT cells at 0.7 × 106 cells/60 mm culture dish in 4 mL of DMEM supplemented with 10% FBS and penicillin-streptomycin. The cells should be ~70% confluent by the following day (at the time of transfection).

3.5.2  Day 1 (Late Afternoon, 5–6 PM)

Transfect 293FT cells with sequence-verified lentivector expression clones (plentiCRISPR_sgRNA) and lentivirus packaging plasmids (Lenti-Pac packaging system from Genecopoeia). 1. In a sterile Eppendorf tube (Tube A), mix together 120 μL Opti-MEM, 1.5 μg lentiCRISPR_sgRNA, 1.5 μg Lenti-Pac HIV mix. 2. In a separate tube (Tube B), mix 120 μL Opti-MEM with 9 μL EndoFectin. 3. Add the diluted EndoFectin (from Tube B) dropwise to Tube A (containing the plasmids) while gently vortexing the tube. 4. Let the transfection complex form for 20 min at room temperature. 5. Add the DNA-Endofectin Lenti complex, dropwise to 293FT cells in the dish, and gently swirl the dish to distribute the complex. 6. Incubate the cells overnight at 37 °C.

3.5.3  Day 2 (Morning, 8–9 AM)

1. Replace culture media for the transfected 293FT cells. 2. Coat 6-well plate with collagen.

(a) Dilute collagen I solution to 30 μg/mL in sterile PBS.



(b) Add 1 mL of diluted collagen to each well, and incubate the plate at room temperature for 45 min.



(c) Aspirate off the solution and wash the wells twice with 1 mL of PBS.



(d) Air-dry under the hood for 15 min.

3. Seed primary human airway epithelial cells at a density of 1 × 105 cells per well in 2 mL of BEGM. 3.5.4  Day 3 (Morning, 8–9 AM)

Harvest lentivirus and infect target cells. This procedure will be repeated on days 4 and 5 (depending on the viability of transfected 293FT cells). 1. Harvest lentivirus-containing supernatant from the transfected 293FT cell culture dish, and transfer to a 15-mL conical tube. Add 4 mL of fresh medium to 293FT cells, return to the incubator for further cell culture, and harvest of freshly ­ ­packaged lentivirus.

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2. Centrifuge the 15 mL tube at 1000 × g-force for 10 min to clarify the lentivirus suspension from cell debris. Transfer the supernatant to a new tube. Add HEPES to 10 mM and ­polybrene to 8 μg/mL and mix by pipetting. 3. Infect airway epithelial cells. Remove BEGM from the 6-well culture plate, and add 2 mL of freshly harvested lentivirus ­suspension (with added polybrene and HEPES) to each well containing the target cells. 4. Centrifuge the plate at 1000 g-force for 45 min at room ­temperature. This procedure is known as spin infection and is intended to enhance the adsorption/entry of virus into the adherent cells. 5. After centrifugation, place the plate for 1 h in at 37 °C 6. Remove the virus suspension and replace with 2 mL of fresh BEGM. 3.5.5  Day 4 (Morning, 8–9 AM)

Harvest lentivirus and infect target cells again (second infection), as described above for Day 2 (Subheading 3.5.4).

3.5.6  Day 5 (Morning, 8–9 AM)

Harvest lentivirus and infect target cells again (third infection), as described above (Subheading 3.5.4) (see Notes 10 and 11).

3.6  Confirmation of Indel Mutations and Protein Expression

1. Trypsinize cells in the well and split the cell suspension into two parts. (A) Reseed one part of cells into one well of a collagen-coated 6-well plate to maintain the cells. (B) Extract DNA from the other part of cells for detection of indel mutations. 2. Perform PCR on each DNA sample using PCR primers designed to amplify about a 700-bp DNA fragment containing the sgRNA region (see Note 12). 3. Purify the PCR product on a spin column, using QIAquick PCR purification kit as directed by the manufacturer (Qiagen) or preferred method. 4. To denature and reanneal the PCR product to form DNA heteroduplexes, mix 10 μL of PCR product with 1 μL of 10× T7 endonuclease buffer in PCR tube. Incubate at 95 °C for 10 min (to denature), and then slowly cool down to 25 °C at 6 °C/ min (to reanneal). 5. Add 2 U of T7 endonuclease I to the hybridized DNA. Incubate at 37 °C for 60 min. 6. Analyze DNA fragments by electrophoresis on 2% agarose gel. 7. The T7 endonuclease cleaves mismatched DNA heteroduplexes at the indel mutation site, releasing two fragments of smaller size than the size of intact DNA homoduplexes. The smaller fragments are indicative of the presence of an indel mutation. Quantification of the relative densities of these fragments (e.g.,

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using Image J) can provide an estimate of the frequency of indel (see Note 13). 8. Confirm gene knockout at the protein level by Western blotting.

4  Notes 1. 293FT cell line is fast growing and highly suitable for lentiviral production. The cell line is maintained in D10 culture medium (see below) and should be passaged at 70–80% confluence for no more than 15 passages to maintain high performance. 2. Primary human nasal and bronchial epithelial cells can be obtained from volunteer donors, by brushing nasal cavity or bronchial airways (respectively), with a signed informed ­consent under a research protocol that is approved by the institutional review board (IRB). Both nasal and bronchial ­primary epithelial cells can also be obtained from commercial source. 3. Commonly used sgRNA design tool for CRISPR knockout, accessible at the public domain of the BROAD Institute and available at the Internet address: (http://portals.broadinstitute.org/gpp/public/analysis-tools/sgrna-design). 4. The target gene may have several transcript variants. In ­general the more variants, the less likelihood to identify a guide RNA that can target all of them at once. If the gene function to be interrupted is known to be dependent on a single variant, then targeting can be focused on that variant only. Alternatively, attempt can be made to select a guide RNA that can target most transcript variants. Also, a combination of guide RNAs can be used to target all transcript variants. However, using multiple guides will potentially increase the risk for multiple off-target DNA cleavage and mutations. 5. sgRNA can be designed on either the positive strand or the negative strand of the target DNA sequence. Since DSBs can be generated independent of the orientation of the PAM sequence, the outcome will remain the same, i.e., indel mutation with potential gene disruption. 6. Example of online tool for the design of reverse or complement sequence (http://arep.med.harvard.edu/labgc/adnan/ projects/Utilities/revcomp.html). 7. Do not use T4 PNK buffer because it does not contain ATP, which is essential for the phosphorylation. 8. RE buffer is preferably used to maintain optimal conditions for BsmBI while supporting T4 DNA ligase activity. The RE buffer usually contains 100 μg/mL BSA to stabilize the enzyme. Otherwise, BSA can be added to 100 μg/mL to the reaction

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mix. The remaining ligation product can be stored at 4 °C or at −20 °C for further use if needed. 9. This approach avoids further processing (e.g., centrifugation or freeze/thaw) that can attenuate the lentivirus activity/titer and consistently yields high transduction rates with primary human airway epithelial cells. 10. This step should be avoided if the majority of 293FT cells have detached and are dead, because the suspension will contain more debris than virus and is not suitable for infection. 11. Using the procedure outlined above, we generally obtain over 90% lentiviral transduction rate in primary human bronchial epithelial cells. However, because un-transduced cells may outgrow (at faster rate) the transduced cells over extended periods of culture, a selection step with puromycin (1 μg/mL) can help eliminate un-transduced cells within a short period of time (3–4 days). 12. It is important to use high-fidelity DNA polymerase to avoid introducing mutations during amplification that can affect the endonuclease cleavage assay. 13. Theoretically, if the sum of integrated densities of the two smaller DNA fragments equals the integrated density of the largest (undigested fragment), the indel frequency (or occurrence) is 100%. In such case, one might assume that both alleles of the target gene were mutated, resulting in target gene knockout.

Acknowledgments This work was supported by the following grants from NIH: 1U19AI125357, R01HL122321, R01AI106287, and R01HL125128. The authors wish to thank Max Seibold, Jamie Everman, and Ari Stoner (Dr. Max Seibold’s Lab, National Jewish Health, Denver) for technical advice and support. References 1. Barrangou R, Fremaux C, Deveau H, Richards M, Boyaval P, Moineau S et al (2007) CRISPR provides acquired resistance against viruses in prokaryotes. Science 315:1709–1712 2. Brouns SJ, Jore MM, Lundgren M, Westra ER, Slijkhuis RJ, Snijders AP et al (2008) Small CRISPR RNAs guide antiviral defense in prokaryotes. Science 32:960–964

3. Marraffini LA, Sontheimer EJ (2008) CRISPR interference limits horizontal gene transfer in staphylococci by targeting DNA. Science 322:1843–1845 4. Ishino Y, Shinagawa H, Makino K, Amemura M, Nakata A (1987) Nucleotide sequence of the iap gene, responsible for alkaline phosphatase isozyme conversion

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in Escherichia coli, and identification of the gene product. J Bacteriol 169:5429–5433 5. Mojica FJ, Díez-Villaseñor C, Soria E, Juez G (2000) Biological significance of a family of regularly spaced repeats in the genomes of Archaea, Bacteria and mitochondria. Mol Microbiol 36:244–246 6. Bolotin A, Quinquis B, Sorokin A, Ehrlich SD (2005) Clustered regularly interspaced short palindrome repeats (CRISPRs) have spacers of extrachromosomal origin. Microbiology 151:2551–2561 7. Pourcel C, Salvignol G, Vergnaud G (2005) CRISPR elements in Yersinia pestis acquire new repeats by preferential uptake of bacteriophage DNA, and provide additional tools for evolutionary studies. Microbiology 151:653–663

8. Marraffini LA (2015) CRISPR-Cas immunity in prokaryotes. Nature 526:55–61 9. Mali P, Yang L, Esvelt KM, Aach J, Guell M, DiCarlo JE et al (2013) RNA-guided human genome engineering via Cas9. Science 339:823–826 10. Ran FA, Hsu PD, Wright J, Agarwala V, Scott DA, Zhang F (2013) Genome engineering using the CRISPR-Cas9 system. Nat Protoc 8:2281–2308 11. Mojica FJ, Díez-Villaseñor C, García-Martínez J, Almendros C (2009) Short motif sequences determine the targets of the prokaryotic CRISPR defense system. Microbiology 155:733–740 12. Bauer DE, Canver MC, Orkin SH (2015) Generation of genomic deletions in mammalian cell lines via CRISPR/Cas9. J Vis Exp 95:e52118

Chapter 28 A Consistent Method to Identify and Isolate Mononuclear Phagocytes from Human Lung and Lymph Nodes Sophie L. Gibbings and Claudia V. Jakubzick Abstract Mononuclear phagocytes (MP) consist of macrophages, dendritic cells (DCs), and monocytes. In all organs, including the lung, there are multiple subtypes within these categories. The existence of all these cell types suggest that there is a clear division of labor and delicate balance between the MPs under steady state and inflammatory conditions. Although great strides have been made to understand MPs in the mouse lung, and human blood, little is known about the MPs that exist in the human lung and lung-­ draining lymph nodes (LNs), and even less is known about their functional roles, studies of which will require a large number of sorted cells. We have comprehensively examined cell surface markers previously used in a variety of organs to identify human pulmonary MPs. In the lung, we consistently identify five extravascular pulmonary MPs and three LN MPs. These MPs were present in over 100 lungs regardless of age or gender. Notably, the human blood CD141+ DCs, as described in the literature, were not observed in non-diseased lungs or their draining LNs. In the lung and draining LNs, expression of CD141 was only observed on HLADR+ CD11c+ CD14+ extravascular monocytes (often confused in the LN as resident DCs based on the level of HLADR expression and mouse LN data). In the human lung and LNs there are at least two DC subtypes expressing HLADR, DEC205 and CD1c, along with circulating monocytes that behave as either antigen-presenting cells or macrophages. Furthermore, we demonstrate how to distinguish between alveolar macrophages and interstitial macrophage subtypes. It still remains unclear how the human pulmonary MPs identified here align with mouse MPs. Clearly, we are now past the stage of cell surface marker characterization, and future studies will need to move toward understanding what these cell types are and how they function. Our hope is that the strategy described here can help the pulmonary community take this next step. Key words Human, Mononuclear phagocytes, Dendritic cells, Monocytes, Interstitial macrophages, Alveolar macrophages, Pulmonary, Lung

1  Introduction In the lung, alveolar macrophages are the first line of defense against invading pathogens [1, 2]. If alveolar macrophages and incoming neutrophils are incapable of containing the invading pathogens, then an adaptive immune response is initiated to assist in its clearance [3, 4]. Underneath the epithelial cells are dendritic

R. Lee Reinhardt (ed.), Type 2 Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1799, https://doi.org/10.1007/978-1-4939-7896-0_28, © Springer Science+Business Media, LLC, part of Springer Nature 2018

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cells (DCs), interstitial macrophages (IMs), and monocytes, in addition to other leukocytes and non-hematopoietic cells [5, 6]. DCs link innate and adaptive immunity by acquiring, processing, and trafficking foreign and self-antigens to the draining lymph node (LN), where they present peptides on MHC molecules and activate cognate T cells [7–10]. In addition to macrophages and DCs, monocytes can also contribute to the clearance of microbes by acquiring a macrophage-like phenotype and induce adaptive immunity by acquiring a DC-like phenotype. Lastly, we were unable to identify CD303+ or CD123+ cells in the lung or draining LNs; hence, why plasmacytoid DCs are not discussed here. For decades monocytes have been viewed as precursors to macrophages. However, we now know that monocytes continuously traffic through nonlymphoid and lymphoid tissue without becoming a bona fide macrophage or dendritic cell. In mice and humans, extravascular lung and LN monocytes are as abundant as DCs in the steady state and are even more abundant during inflammation [11, 12]. The role of monocytes in adaptive immunity is underappreciated, and although it has been shown that they can induce T cell proliferation, how they selectively activate lymphocytes is unclear [12]. In addition to monocytes, DCs, and tissue-­ specific alveolar macrophages, there are interstitial macrophages (IM). In mice we have identified three unique IM subtypes in the lung and other organs, albeit what they functionally do is still unclear [13]. In this chapter, we outline how to identify and isolate human pulmonary MPs from whole lungs en bloc with the hopes that in the near future we can functionally align these cell types with the well-characterized murine MPs. All in all, we observe consistency and reproducibility when we strictly adhere to the six cautionary steps (see Notes 1–6) for the isolation and use of human pulmonary MPs.

2  Materials 2.1  Human Lung and Lymph Nodes

1. Non-diseased human lungs and lung-draining LNs were acquired from three sources as stated in the acknowledgments (also see ref. [14]).

2.2  Bronchoalveolar Lavage (BAL) and Media

1. PBS: 1× Phosphate-buffered saline (PBS) without calcium or magnesium for perfusion and lavage. Make 1 L per lobe. 2. PBS/ETDA buffer: 1× PBS and 3 mM ethylenediaminetetraacetic acid (EDTA, from 0.5 M stock solution pH 8.0). Make 500 mL per lobe. 3. BSS-B buffer: 132 mM NaCL, 5 mM KCl, 0.5 mM NaH2PO4, 2 mM Na2HPO4, 10 mM HEPES, 1 g/L Dextrose, 1.9 mM CaCl, 1.3 mM MgSO4, pH 7.4. Make 500 mL per lobe.

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4. Scissors. 5. Large serrated forceps. 6. 2–3 pairs of hemostats. 7. 60 mL syringe. 8. 200 mL plastic beakers. 9. 500 mL plastic beakers. 10. 100 μm filter membrane. 11. 1/3 and 1 cm diameter PVC tubing: for perfusion in the pulmonary veins and inflation into the bronchus. 12. Various sized pipet tips. 13. FACS buffer: 1× PBS with 1 mM EDTA, 0.15% bovine serum albumin (BSA); keep at 4 °C. 2.3  Tissue Digestion for Lung MPs

1. Elastase buffer: 4.2 U elastase/mL in BSS-B. Approximately 150–250 mL is required for one lobe from an average adult (see Note 7). 2. 1 gallon plastic bags: strong and durable (e.g., small biohazard bag). 3. String (to tie off plastic bag). 4. Water bath at 37 °C. 5. Rolls or sheets of nylon filter membrane: 350 and 100 μm. 6. Cheesecloth. 7. Vegetable strainer. 8. Two 1 L beakers. 9. Heat inactivated fetal calf serum (FCS). 10. Large tissue culture dishes to mince tissue and 1 L plastic beakers. 11. Blender (Ninja Professional Blender). 12. Chilled centrifuge for 250 mL centrifuge tubes. 13. 50 mL conical tubes. 14. Kreb/HEPES buffer: 0.9% NaCL, PO4, KCl, HEPES. 15. Optiprep reagent density 1: 1.080 g/mL (heavy) by dilution of Optiprep reagent in Kreb/HEPES buffer in 50 mL conicals. 16. Optiprep reagent density 2: 1.040 g/mL (light) by dilution of Optiprep reagent in Kreb/HEPES buffer in 50 mL conicals. 17. Scissors. 18. Large serrated forceps. 19. 2–3 pairs of hemostats. 20. FACS buffer: 1× PBS with 1 mM EDTA, 0.15% bovine serum albumin (BSA); keep at 4 °C.

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2.4  Nonenzymatic Cell Crawl-Out Method for Lymph Node MPs

1. Scissors. 2. Forceps. 3. Culture media: RPMI 5% FCS, penicillin, streptomycin, fungizone, l-glutamine. 4. 150 × 25 mm tissue culture plate. 5. Tissue culture incubator, 37 °C, 5% CO2. 6. 100 μm filter. 7. 50 mL conical tubes and centrifuge. 8. FACS buffer: 1× PBS with 1 mM EDTA, 0.15% bovine serum albumin (BSA); keep at 4 °C. 9. Cell scraper.

2.5  Enrichment for Myeloid Cells

2.6  Staining for FACS Analysis and MP Identification

1. FACS buffer. 2. Enrichment using anti-CD11c-biotin (alternatively use CD64-­ biotin for macrophages) and anti-CD1c PE conjugated (see Table 1 for antibodies) for positive selection, either STEMCELL or Miltenyi kits can be used. 1. FACS buffer. 2. Pooled human serum. 3. FACS buffer with human serum: 1 mM EDTA, 0.15% bovine serum albumin (BSA), 20% pooled human serum; keep at 4 °C. 4. Fluorochrome conjugated antibodies (see Table 1). 5. Antibody cocktail: 3–10 μL of fluorochrome conjugated antibodies per 100 μL of FACS buffer with human serum. 6. 4’,6-Diamidine-2’-phenylindole dihydrochloride working solution: 30 μg/mL DAPI in PBS.

(DAPI)

7. Flow cytometers: analyzers (BD LSR II and Fortessa); sorter (BD ARIA fusion). 8. FlowJo software: for flow cytometric analyses.

3  Methods 3.1  Preparation of Lung Tissue for Alveolar Macrophages: Tissue Dissection, Perfusion, and Bronchoalveolar Lavage (BAL)

1. Starting with the trachea. Identify individual lobes and corresponding large bronchus. Select one lobe for BAL and tissue dissection. 2. Dissect paratracheal, subcarinal, and carinal LNs. LNs are darker, more dense tissue nodules immediately surrounding the trachea and early bronchial branches (see Note 8). 3. Identify pulmonary arteries supplying the lobe of interest. Use a 60 mL syringe attached to 1 cm PVC tubing with appropriately

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Table 1 Antibody clones used for pulmonary MP identification and isolation Antigen

Clone

Conjugate

CD1a

Ancell

FITC

CD1c

REA694

PE

CD3

UCHT1

PB

CD11b

ICRF44

PerCP

CD11c

3.9

PE-Cy7, biotin

CD14

MΦP9

V500

CD15

W6D3

PB

CD20

2H7

PB

CD26

BA5b

PE, PE-Cy7

CD36

AC106

APC

CD43

eBio84-3C1

FITC

CD45

H130

BUV395

CD56

TULY56

PB

CD64

10.1

PE

CD206

15-2

PerCP

DEC205

HD30

APC

HLA-DR

L243

APC-Cy7

sized pipet tip to follow arterial branches. Fill 60 mL syringe with PBS, and insert pipet tip into the large and small pulmonary arteries. Perfuse the lobe, and repeat several times with fresh PBS. Continue to perfuse until no more blood drains out of the pulmonary veins. The tissue should be visibly whiter. 4. Cannulate the large bronchus with 1 cm PVC tube. Secure the tubing in place with a piece of string. 5. To lavage for alveolar macrophages, fully inflate the lobe with PBS/EDTA buffer via the cannulated bronchus. Upend the tissue to drain the lavage fluid into a beaker, and gently massage the fluid out of the lobe. Drain as much fluid as possible before inflating the lung again. Lavage six times using this sequence of buffers (see Note 9):

(a) Twice with PBS/ETDA buffer



(b) Twice with 1× PBS



(c) Twice with BSS-B buffer

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6. Pass all lavage fluid through 100 μm filters to remove mucus and centrifuge at 250 × g-force for 10 min. Resuspend pellet in FACS buffer. 3.2  Preparation of Lung Single-Cell Suspension by Elastase Digestion

1. Cut excess tissue surrounding the lobe. Make sure not to pass the fissures between lobes to avoid leakage out of the lobe of interest. Transfer the lavaged lobe into a clean 1 gallon plastic bag for digestion. 2. Fully inflate the lobe with prepared elastase buffer (see Note 7). After inflation with elastase buffer, use hemostat to cross clamp the bronchus to avoid leakage of buffer. 3. Incubate in a water bath at 37 °C for 40 min. 4. Remove from water bath and transfer to a large dish. Cut away un-inflated tissue and poorly perfused patches. Remove the cannula and surrounding upper airway tissue. Cut the remaining lung tissue into large chunks (~1 × 1 × 1 inches). 5. Transfer lung pieces and fluid into a blender containing 75 mL of heat inactivated FCS and 150 mL of BSS-B buffer. Blend for two short (5 s) pulses (see Note 10). Large undigested pieces of cartilage will remain after pulse blending. These large pieces will be poured onto the cheesecloth in the following step 6. 6. The pulse-blended lobe is passed through a series of filters to remove undigested pieces and create a single-cell suspension. First, pour lung homogenate through a cheesecloth-lined vegetable strainer into a 1 L beaker. This will catch a large amount of undigested matter. Increase cell yield by washing out the blender with BSS-B buffer and pouring this over the cheesecloth. Squeeze the cheesecloth gently to drain buffer and cells. 7. Then pass the homogenate through a 350 μm filter into a second 1 L beaker and lastly through 100 μm filter to remove smaller cell clumps. Cells are passed through another 100 μm filter before centrifugation. 8. Centrifuge filtered lung cells at 250 × g-force for 10 min. 9. Resuspend the single-cell suspension in 40 mL of FACS buffer. Overlay 10 mL onto each of four prepared Optiprep density separation tubes. 10. Centrifuge at 500 × g-force for 20 min, make sure centrifuge brake is turned off, or placed on the lowest setting. 11. Collect cells within the light/heavy interface, avoiding lighter dead cells and denser red blood cells. 12. Wash once with a large volume of FACS buffer to dilute out Optiprep and then resuspend in FACS buffer for further enrichment.

Method for Human MP Isolation in Lung and LNs

3.3  Preparation of LN Single-Cell Suspension by Overnight Crawl Out

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1. Collect LNs and carefully remove and discard surrounding perinodal fat tissue. 2. In 1.5 mL eppendorf tube, use scissors and forceps to finely mince LNs in tissue culture media. 3. Transfer minced LNs into a (220 × 25 mm) TC dish and add 50 mL of fresh culture media. Only place ~2–3 LNs per dish to avoid cell overcrowding. 4. Incubate overnight (16–20 h) at 37 °C with 5% CO2. 5. The following morning, collect all single cells from the plate by pipetting off and saving all the culture media. Next gently wash adherent cells from the dish using FACS buffer and a cell scraper. Filter-collected sample through a 100 μm filter into a 50 mL conical before centrifugation. 6. Centrifuge at 250 × g-force for 5 min. Resuspend in FACS buffer. 7. Alternatively (instead of steps 1–6), LNs can be finely chopped, digested in collagenase D for 30 min at 37 °C, filtered through a 100 μm filter, collected, and resuspended in FACS buffer [7–10].

3.4  Enrichment of Myeloid Cells by CD11c+ Selection for Lung MPs and LN MPs

1. To improve MP purity, first block non-specific antibody binding by preincubating lung or LN cell suspensions with FACS buffer with human serum for at least 10 min. 2. Incubate with anti-CD11c-biotin for 15 min on ice. 3. Incubate with biotin selection cocktail for 15 min on ice. 4. Incubate with magnetic nanoparticles/beads for 15 min on ice. 5. Wash once with FACS buffer. 6. Resuspend cell pellet in 5 mL of FACS buffer and place tube into the EasySep magnet. Allow cells to bind magnet for 5 min, and then decant unbound cells, as described by STEMCELL. For Miltenyi isolation use LS columns and Miltenyi microbeads. Both STEMCELL or Miltenyi enrichments work in this protocol (see Notes 11 and 12). 7. Repeat step 6 to increase purity for STEMCELL isolation.

3.5  Staining for FACS Analysis and MP Identification

1. Resuspend 2–5 × 106 cells per sample from BAL, enriched lung, or LN in 100 μL of FACS buffer with human serum. 2. Add 100  μL of antibody cocktail, vortex, and incubate for 45 min on ice. 3. Wash once in FACS buffer and resuspend in 250 μL of FACS buffer. Place cells on ice for flow sorting or analysis. 4. Immediately prior to acquiring samples on the flow cytometer or sorter, add 50 μL DAPI working solution to 250 μL of cells. Dead cell exclusion is essential for further analysis of lung and LN MP populations (see Note 13). Use control samples to optimize FACS parameters.

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Fig. 1 CD11c enrichment for pulmonary MP identification. Left figure depicts enriched CD11c+ cells from single-­cell suspension of an entire digested lobe. Cells were first gated on live cells, which excluded dead cells (DAPI+), lineage+ cells, and subcellular debris. Then, single, CD45+ cells were gated. Top row, SSChiHLADRhiCD43+ cells were AMs (that were also CD14− not depicted here, in ref [14]). Bottom row, gates on SSCintCD45+CD43− cells, which were plotted as CD1c versus CD206 to identify extravascular cells: CD1c+CD206− and CD206+ cells. One extravascular CD1c+ MPs was identified: CD206−CD1a+ MP (designated as lung MP1). Three other extravascular MPs were CD206+ and distinguished by CD1c, CD36, and HLADR. One was CD1c+CD1aint MP (designated as lung MP2), and the other two were CD1c−CD36+HLA-DR+ and CD1c−CD36lo/-HLA-DR++ (designated as lung MP3 and MP4). Previously, we demonstrated in healthy individuals that intravascular circulating monocytes were CD206-, whereas extravascular monocytes/MPs were CD206+ [14]. Therefore, CD206 can distinguish between intra- and extravascular MPs 3.6  Gating and Analysis of Lung and LN MPs by Flow Cytometry 3.6.1  Lung MPs (Figs. 1 and 2)

1. Begin with the exclusion of dead cells and small debris using DAPI and forward scatter area (FSC-A) (see Figs. 1 and 2). 2. Use CD45, side scatter area (SSC-A), and FSC-width (W) to gate on single, hematopoietic cells. CD11c could also be used here to identify all myeloid cells. 3. Alveolar macrophages are SSChi, CD14−CD43+ cells that are uniformly CD206+ and HLA-DR+ (see ref. [14]). These tend to represent the majority of the SSChi population in the lung. SSChi CD43−HLADR− cells are mostly neutrophils (not shown), which can be confirmed by staining with CD15 or CD16. 4. From the SSCintCD43− gate, we plot CD206 vs. CD1c to separate out four different populations of lung MPs as well as intravascular blood monocytes (Fig. 1).

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Fig. 2 CD11c and CD1c enrichment for pulmonary MP sorting: five extravascular MPs in the lung designated AM and MP 1–4. Top figure depicts enriched CD11c+ cells, and bottom figure depicts enriched CD1c+ cells from single-cell suspension of an entire digested lobe. For both enrichments, cells were first gated on live cells, which excluded dead cells (DAPI+), lineage+ cells, and subcellular debris. Then, single, CD45+ cells were gated. For CD11c+ cell enrichment, SSChi/intHLADRhi cells were gated to identify and sort CD43+SSChi AMs (that were also CD14− not depicted here, in ref [14]) and two other CD1c−CD206+ pulmonary MPs. CD1c−CD206+ MPs were distinguished by CD36 and HLA-DR (designated as lung MP3 and MP4). For CD1c+ cell enrichment, SSCintHLADR+ cells were gated to exclude CD1c+ B lymphocytes. To further exclude Lin+ contaminating cells (including B cells), which were not as bright as DAPI, another gate was used to exclude any additional Lin+ cells that were not excluded in the first live cell gate. Two CD1c+ MPs were identified: CD206−CD1a+ MP (designated as lung MP1) and CD206+ CD1aint (designated as lung MP2). Previously, we demonstrated in healthy individuals that intravascular circulating monocytes were CD206-, whereas extravascular monocytes/MPs were CD206+ [14]. Therefore, CD206 can distinguish between intra- and extravascular MPs



(a)  Intravascular monocytes: CD1c− CD206− monocytes that express CD36 and lower levels of HLA-DR [14] (see Note 14).



(b) MP 1: CD1c+CD206− cells represent an HLA-DR+, CD1a+ dendritic cells.



(c) MP 2: CD1c+CD206+  (transcriptome data suggest this MP is a DC).



(d) MP 3: CD1c−CD206+CD36+ (transcriptome data suggest this is an interstitial macrophage, see Note 15).

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(e)  MP 4: CD1c−CD206+CD36−HLADR+ (transcriptome data suggest this is an interstitial macrophage, see Note 15).

3.6.2  Lymph Node MPs

(f) MP ?: CD1cintCD206+CD36+ (sorting, not shown- it is unclear whether this MP is more DC-like or Macrophage-like).

1. Begin by excluding dead cells and small debris, then doublets using a combination of DAPI and FSC-A, then FSC-W and SSC-A. 2. CD11c, SSC, and HLA-DR can be used to identify three distinct populations (Fig. 3).

Fig. 3 CD11c enrichment for lung-draining lymph node (LN) MP identification and sorting: three extravascular MPs in the lung-draining LN designated MP 1–3. Top figure depicts non-enriched cells, and bottom figure depicts enriched CD11c+ cells from the single-cell suspension of lung-draining LNs. First cells were gated on live cells, which excluded dead cells (DAPI+), lineage+ cells, and subcellular debris. Then, single, CD45+ cells were gated on. For non-enriched cells, SSCintCD11c+, HLADR−/+ cells were gated. CD11c+HLADR− cells were neutrophils (green arrow). CD11c+HLADRint were CD14+CD141+ monocytes (blue arrow), and CD11c+HLADR++ cells (red arrow) contained two DEC205+CD1c+ LN MPs. The right, top rows illustrate various stains for CD11c+ cells on the y-axis (CD1c, DEC205, CCR7, CD141, CD45, EpCAM, CD14, and CD11b) with the x-axis constant for the expression of HLA-DR [14]. For sorting, CD11c expressing cells were enriched, and three MPs were identified and sorted. Two were CD11c+HLADR++, DEC205+CD1c+ MPs that were either CD1a+CD1c+ or CD1alowCD1c+/int (designated as lymph node MP1 and MP2). The third LN MP (designated as LN MP3) was CD11c+HLADR+, CD14+CD141+ monocytes. Inserted table on the bottom left summarized all stains used to analyze the two overarching LN MPs: CD14+CD141+ and DEC205+CD1c+/int MPs [14]

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(a) CD11c+HLA-DR+/++ cells: monocytes (HLADR+) and DCs (HLADR++)

(b) CD11c+ HLA-DR− SSChi cells: Neutrophils



(c) CD11c−SSClo cells: Lymphocytes

3. Taking the CD11c+HLA-DR+/++ gate, one can repeatedly observe three MPs (Fig. 3):

(a) MP1: HLADR++DEC205+CD1c+CD1a+



(b) MP2: HLADR++DEC205+CD1c+CD1alow



(c) MP3: HLADR+CD141+CD14+ monocytes (which also express CCR2, CD36, CD206, CD11b, CD64, and CD163)

4  Notes 1. First major cautionary step during the isolation procedure of pulmonary MPs: The time to acquire, process, and isolate pulmonary MPs, followed by sorting for experiments or analyses, is very long. For this reason, many investigators may choose to save the MPs for later use by either fixing or freeze-thawing the cells. In our opinion, it is extremely important to analyze or examine the function of MPs right after isolation. From our experience, if MPs are fixed either with 1% PFA or formalin and analyzed the following day, the data quality is significantly less. This is most likely due to the following three reasons: first, fixed cells shrink thus altering their forward/side scatter (FSC/ SSC) properties; second, autofluorescence due to fixation leads to cells shifting in fluorescent channels making a­ ntibody stains less distinguishable; and lastly, fixed cells eliminate the ability to exclude dead cells, which is vital when studying human MPs. As for freeze-thaw, unlike alveolar macrophages or self-­ renewing non-hematopoietic structural cells, IMs, monocytes, and DCs do not remain viable after freezing and thawing. Therefore, we do not recommend freezing and thawing MPs. Perhaps the lack of viability is because DCs and monocytes are relatively short lived and do not self-renew or clonally expand like lymphocytes. In addition, IMs, extravascular monocytes, and DCs live within a complex extracellular matrix that cannot be replicated in vitro, and therefore, survival signals and cross talk with other cells are lacking, which most likely results in the initiation of their death when extracted from this environment. 2. Second major cautionary step during the isolation procedure of pulmonary MPs: Do not use logarithmic FSC/SSC and instead take advantage of linear FSC/SSC parameters to clearly

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exclude subcellular debris that can be DAPI negative and known to bind non-specifically to antibodies. For instance, in the figures illustrated here, CD1c+ MPs are bigger in SSC than CD1c+ B cells, thus allowing for the exclusion of not only subcellular debris but also lymphocytes. Of course it is important to distinguish live cells not only based on FSC/SSC but also with DAPI (dead cell exclusion dye) and lymphocyte and granulocyte stains for lineage dump. In our case, we exclude both dead cells and lineage cells in the same channel. Therefore, it is important to note that DAPI exclusion is higher on the log scale than lineage + (Lin+) antibodies used to exclude lymphocytes and granulocytes (see Figs. 1 and 2), and thus a second gate to exclude lineage cells should be used later in the sorting strategy. Lastly, autofluorescence, particularly for alveolar macrophages, will always be there, so try not to exclude these cells if desired for sorting or analyzing; even if they overlap with Lin+ cells (see Figs. 1 and 2), there are other ways to exclude contaminating cells from this population. 3. Third major cautionary step during the isolation procedure of pulmonary MPs relates to digestion and filtering. Liberase TM, collagenase D, and elastase all cleave away cell surface molecules used to identify pulmonary MPs. AMs are lavaged, which can then be directly sorted. Tissue MPs are digested through the alveolar epithelium to preserve the cell surface molecules on the MPs since the digestive enzymes are working directly on the epithelium rather than directly on interstitial cells. Although, we do not outline how to digest a small piece of lung tissue, small pieces of tissue can be finely minced and digested in low concentration of collagenase D [14], in a gentle shaker at 37 °C for 25 min. For LNs, due to the rapid cleaving of cell surface molecules by direct digestion, we allow LN MPs to crawl out overnight. 4. Fourth major cautionary step during the isolation procedure of pulmonary MPs: Overall, in our experience, there are some cell surface markers that are not readily cleaved by digestive enzymes, which include CD14, CD11b, HLADR, and CD11c. However, markers such as CD141, CD303, Clec9a, CD1c, and CD1a are highly susceptible to digestive cleavage (unpublished data). 5. Fifth major cautionary step during the isolation procedure of pulmonary MPs: Frequent filtering is vital to acquire and maintain cell yield and viability. The presence of fat cells in a single-cell suspension promotes clumping that also traps live cells. Particularly after centrifugation, it is impossible to disassociate live cells from clumped fat/dead cells. So to avoid clumping, or a major loss of cells, always filter before centrifugation, even after staining cells for flow and right before run-

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ning samples on the cytometer. Lastly, make sure to use 100  μm filters at first, as 70- and 40-micron filters are too small to readily allow large MPs to pass through. Studying LN MPs requires the use of relatively clean, healthy LNs. Thus, LNs not contaminated with excessive smoking or pollution particulates are preferred. To date, we are unable to recover any MPs from black LNs. 6. Sixth major cautionary step during the isolation procedure of pulmonary MPs is to be aware that intravascular and extravascular MPs exist within a piece of tissue even after extensive perfusion [13]. Therefore identifying intra- and extravascular cells in the human lung is important, and how to do this was illustrated in our previous publication [14]. 7. The volume required for digestion is best determined during the lavage by measuring the volume required to fully inflate the lobe. 4.2 U elastase/mL BSS-B buffer x volume of lobe. On average a right middle lobe from a middle-aged person requires 150–250 mL of buffer. 8. Make sure not to cut too deep into tissue, as the lobe will be inflated with lavage and enzymatic fluid for pulmonary MP isolation. 9. EDTA is included only in the first two lavages to help detach adherent macrophages from the airways epithelia. However, efficient removal of EDTA is essential to avoid subsequent inhibition of enzymatic digestion. 10. Avoid overprocessing the lung; the shear force of the blender damages cells and will result in greatly increased cell death. 11. Enrichment protocols provided by the manufacturer serve as a good starting point for titrating cell/antibody/bead ratios for optimizing enrichment efficiency based on downstream requirements. 12. Enriching LN cells using CD11c for sorting is required if optimal RNA seq analyses and functional assays are being performed. If cells are not enriched, sorting will take too long, and the viability of the LN MPs will diminish. 13. For microRNA or messenger RNA extraction from MPs, add lymphocyte and granulocyte lineage stains in addition to DAPI to insure proper exclusion during cell sorting. 14. CD206 is expressed on monocyte-derived cells and other MPs upon extravasation into tissue. CD1c+CD206− cells represent HLA-DR+, CD1a+ dendritic cells, as this was clearly shown using time-lapse video (see ref. [14]). Although we hypothesize that there are two DCs, we have yet to clearly identify the second DC in the lung, which may be the CD1c+CD206+ MP, as LN CD11c+HLADR++CD20− cells all express DEC205 and CD1c, which are divided by CD1a+ and CD1alow (see Fig. 3).

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15. The CD1c−CD206+ population further splits into two populations. One population that expresses high level of CD36 and second population that expresses lower CD36 and more HLA-DR.

Acknowledgments The authors would like to thank Drs. William Janssen and Robert Manson for collaborating and assisting in the acquisition of non-­ diseased human lungs from any of the following three sources: National Disease Research Interchange (Philadelphia, PA), the International Institute for the Advancement of Medicine (Edison, NJ), and University of Colorado Donor Alliance. This human subject research falls under federal exemption # 4. Grant support: C.V.J. NIH R01-HL115334 and R01-HL135001. References 1. Janssen WJ, Bratton DL, Jakubzick CV, Henson PM (2016) Myeloid cell turnover and clearance. Microbiol Spectr 4(6). https://doi. org/10.1128/microbiolspec. MCHD-0005-2015 2. Bharat A, Bhorade SM, Morales-Nebreda L, Mc Quattie-Pimentel AC, Soberanes S, Ridge K, DeCamp MM, Mestan KK, Perlman H, Budinger GR, Misharin AV (2015) Flow cytometry reveals similarities between lung macrophages in humans and mice. Am J Respir Cell Mol Biol 54:147–149. https://doi. org/10.1165/rcmb.2015-0147LE 3. MacLean JA, Xia W, Pinto CE, Zhao L, Liu HW, Kradin RL (1996) Sequestration of inhaled particulate antigens by lung phagocytes. A mechanism for the effective inhibition of pulmonary cell-mediated immunity. Am J Pathol 148(2):657–666 4. Yu YA, Hotten DF, Malakhau Y, Volker E, Ghio AJ, Noble PW, Kraft M, Hollingsworth JW, Gunn MD, Tighe RM (2015) Flow cytometric analysis of myeloid cells in human blood, bronchoalveolar lavage, and lung tissues. Am J Respir Cell Mol Biol 54:13–24. https://doi. org/10.1165/rcmb.2015-0146OC 5. Holt PG (2005) Pulmonary dendritic cells in local immunity to inert and pathogenic antigens in the respiratory tract. Proc Am Thorac Soc 2(2):116–120. https://doi.org/10.1513/ pats.200502-017AW

6. Sung SS, Fu SM, Rose CE Jr, Gaskin F, Ju ST, Beaty SR (2006) A major lung CD103 (alphaE)-beta7 integrin-positive epithelial dendritic cell population expressing Langerin and tight junction proteins. J Immunol 176(4):2161–2172. doi:176/4/2161 [pii] 7. Vermaelen KY, Carro-Muino I, Lambrecht BN, Pauwels RA (2001) Specific migratory dendritic cells rapidly transport antigen from the airways to the thoracic lymph nodes. J Exp Med 193(1):51–60 8. Jakubzick C, Tacke F, Llodra J, van Rooijen N, Randolph GJ (2006) Modulation of dendritic cell trafficking to and from the airways. J Immunol 176(6):3578–3584. doi:176/6/3578 [pii] 9. Jakubzick C, Helft J, Kaplan TJ, Randolph GJ (2008) Optimization of methods to study pulmonary dendritic cell migration reveals distinct capacities of DC subsets to acquire soluble versus particulate antigen. J Immunol Methods 337(2):121–131. https://doi.org/10.1016/j. jim.2008.07.005 10. Desch AN, Randolph GJ, Murphy K, Gautier EL, Kedl RM, Lahoud MH, Caminschi I, Shortman K, Henson PM, Jakubzick CV (2011) CD103+ pulmonary dendritic cells preferentially acquire and present apoptotic cell-­ associated antigen. J Exp Med

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DL, Henson PM, Janssen WJ, Jakubzick CV 208(9):1789–1797. https://doi. (2017) Three unique interstitial macrophages org/10.1084/jem.20110538 in the murine lung at steady state. Am J Respir 1 1. Jakubzick C, Bogunovic M, Bonito AJ, Cell Mol Biol 57:66–76. https://doi. Kuan EL, Merad M, Randolph GJ (2008) org/10.1165/rcmb.2016-0361OC Lymph-­ m igrating, tissue-derived dendritic cells are minor constituents within steady- 14. Desch AN, Gibbings SL, Goyal R, Kolde R, Bednarek J, Bruno T, Slansky JE, Jacobelli J, state lymph nodes. J Exp Med Mason R, Ito Y, Messier E, Randolph GJ, 205(12):2839–2850. https://doi. Prabagar M, Atif SM, Segura E, Xavier RJ, org/10.1084/jem.20081430 Bratton DL, Janssen WJ, Henson PM, 12. Jakubzick CV, Randolph GJ, Henson PM Jakubzick CV (2015) Flow cytometric analysis (2017) Monocyte differentiation and antigen-­ of mononuclear phagocytes in non-­ diseased presenting functions. Nat Rev Immunol human lung and lung-draining lymph nodes. 17:349–362. https://doi.org/10.1038/ Am J Respir Crit Care Med 193:614–626. nri.2017.28 https://doi.org/10.1164/ 13. Gibbings SL, Thomas SM, Atif SM, McCubbrey rccm.201507-1376OC AL, Desch AN, Danhorn T, Leach SM, Bratton

Chapter 29 Organoid Cultures for Assessing Intestinal Epithelial Differentiation and Function in Response to Type-2 Inflammation Bailey Zwarycz, Adam D. Gracz, and Scott T. Magness Abstract During helminth infection of the gastrointestinal tract, a complex Type-2 inflammatory response involving immunological and mucosal components is mounted to clear the infection and reestablish a physiologically normal state. This response is characterized by the secretion of key interleukins, which impact epithelial ­lineage allocation and drive tuft and goblet cell hyperplasia to lead to eventual clearance of parasitic ­organisms. While there have been advances toward understanding Type-2 inflammatory responses in the intestine, detailed cellular and molecular mechanisms of epithelial responses to general inflammation and specific inflammatory cytokines remain to be explored. Intestinal organoids represent a physiologically relevant in vitro model to study how Type-2 inflammation impacts stem cell maintenance and d ­ ifferentiation and offer a new approach for investigators to test compounds that modulate mechanisms involved in worm clearance. The methods described in this chapter include: (1) intestinal crypt and single cell isolation; (2) organoid culture and cytokine treatment, as well as methods for downstream organoid analyses; (3) gene expression analysis by qRT-PCR; (4) protein analysis by western blot, immunohistochemistry, and fl ­ orescence-activated cell sorting; and (5) organoid self-renewal by serial passaging. Key words Intestinal stem cells, Intestinal epithelial isolation, Intestinal organoids, Intestinal inflammation

1  Introduction The gastrointestinal tract is a complex organ that functions as the site of nutrient and water absorption at the interface of the lumen and the epithelial monolayer. The lumen is home to a vast number of commensal microbiota that exist in a symbiotic relationship with the host organism, and while these microbiota play a critical role in health, it is essential that they are restricted to the luminal compartment by an uncompromised epithelial monolayer. The ­ epithelial barrier can be compromised by physical injury or by ­ pathogenic microbiota that are ingested by the host. When this occurs acutely on a large scale, the host may succumb to sepsis, but at the chronic small scale, the host may develop conditions ­associated R. Lee Reinhardt (ed.), Type 2 Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1799, https://doi.org/10.1007/978-1-4939-7896-0_29, © Springer Science+Business Media, LLC, part of Springer Nature 2018

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with inflammatory bowel disease (IBD). A host infected with pathogenic microbes may not present with clinical symptoms because on an insufficient inoculum, inability of pathogenic strains to outcompete the commensal communities, or because of efficient physiologic clearance by the host immune system. Surveillance of pathogenic microorganisms by the intestinal epithelium requires complex coordination between the epithelial cells that serve as the primary barrier to luminal contents and other submucosal cell types that actuate the immune response. Epithelial tuft cells have an essential role in monitoring the luminal environment for parasitic infection and communicating this information to the underlying immune cell compartment that responds to the infection [1–5]. Tuft cells are one of six primary differentiated lineages found in the intestinal epithelium, and until recently, their function remained unknown. Decades ago, tuft cells were first described as “brush cells” based on the presence of apical tufts of stiff microvilli [5]. In homeostasis, tuft cells are considered very rare epithelial cells (~0.4–1.0%); however, following infection of the host by helminths, a type of microscopic worm, the intestinal epithelium undergoes tuft cell hyperplasia (~7.2%) in an effort to clear the worms [2]. Aside from morphological identification, a number of biomarkers are now associated with the tuft cell lineage including Dclk1, Cox1, Plcγ2, Gfi1b, Trpm5, and high levels of Sox9 [2, 3, 6, 7]. At a transcriptomic level, tuft cells demonstrate a Th2 gene expression signature suggesting that they have the capacity to respond to infection through a specific interleukin response [8]. Recent studies confirm this prediction by showing that tuft cells are involved in a positive feedback circuit initiating a Type-2 immune response to helminth infection [1–3]. In the intestine, Type-2 immune responses are commonly associated with helminth parasitic infection but also participate in IBD [9]. CD4+ lymphocytes are classically known as a mediator for Th2 immunity, but a subset of innate lymphoid cells known as ILC2 cells in the submucosa are now recognized as key mediators of Th2 responses in the intestine. ILC2 cells are characterized by expression of IL-4, IL-5, and IL-13 which in turn activate other immune cell types (basophils, mast cells, and eosinophils) that assist in clearing the infection [10]. Tuft cell function involves a sophisticated positive feedback loop between helminths, the d ­ ifferentiated epithelium, ILC2s, and the undifferentiated stem/progenitor cell compartment (Fig. 1) [1–3]. Upon parasitic i­nfection, tuft cells detect the presence of helminths or protozoa through a Trmp5dependent chemosensory pathway [2]. This causes tuft cells to secrete IL-25 that in turn acts on submucosal ILC2 cells (Fig. 1A), which secrete IL-13 (Fig. 1B). ILC2 cells are in close proximity to the stem/progenitor cell compartment, and the IL-13 secreted by ILC2 cells promotes lineage bias toward tuft and goblet cells (Fig. 1C, D). Goblet cell hyperplasia has a dual function to increase

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Fig. 1 A cellular mechanism to resolve helminth infections in the gastrointestinal tract

mucous to protect the epithelium and aid in worm clearance, while tuft cell hyperplasia serves to increase s­entinels to monitor and respond to the worm infection. Increases in tuft cell numbers continue to fuel the IL-25 positive feedback during infection ­ (Fig. 1E). Reduction in worm burden serves as a brake to tuft and goblet cell hyperplasia, and the epithelium returns to homeostasis (Fig. 1F). While significant strides have been made to understand the cellular mechanisms regulated by the tuft cell-ILC2 axis, much remains to be investigated related to the impact of other Th2 ­cytokines on ISC differentiation, the specificity of tuft cell responses to different helminth species, and additional roles that tuft cells may play in regulating the microbiome in heath and disease. Animal models have served a critical role in moving the ­Th2-­field forward; however, there is substantial merit for using culture models to address questions that are not feasible in animal models and to develop ex vivo platforms that are useful for ­screening compounds that are capable of enhancing or abrogating Th2 responses. Intestinal organoid technology has revolutionized in vitro study of gastrointestinal epithelial biology and has recently been applied to investigate Type-2 immune responses [2, 3]. Intestinal organoids are stem cell-driven structures derived from whole isolated crypts or single ISCs that can be spherical or ­budding in nature, depending on proliferation status [11]. They are n ­ on-transformed and are capable of growing indefinitely in a

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t­hree-dimensional extracellular matrix (typically Matrigel) with a defined media consisting of essential growth factors found in the ISC niche in vivo [12, 13]. Nomenclature for organoids has been further refined as “enteroids” when derived from small intestinal tissue or “colonoids” when derived from colonic tissue [14]. Organoids represent a powerful tool to study Type-2 immune responses due to their ability to (1) generate all the differentiated lineages found in the gastrointestinal tract and (2) maintain an ISC compartment that is able to respond to extrinsic signals that ­influence ISC proliferation and differentiation. While organoids are comprised exclusively of epithelial cells, the cytokine environment produced by a Th2 response can be recreated in culture media, and organoids are capable of being co-­cultured with other cell types that impact Type-2 immunity. These properties render organoid cultures highly adaptable and amenable to detailed mechanistic analysis. Organoids generated from normal mouse strains typically used in research settings can be readily evaluated for proliferation and differentiation by immunostaining, and gene expression can be ­ ­interrogated by qPCR or RNA-seq analysis. While these standard methods are useful, a number of transgenic mouse lines that express fluorescent reporter gene associated with secretory lineages involved in Type-2 immunity enable detection and isolation of live cells for analysis. Dclk1, Gfi1b, Il25, and Trpm5 represent biomarkers highly restricted to the tuft cell lineage [1, 2, 4, 6]. A Dclk1-­CreERT2 ­transgenic mouse line has been developed that enables ­identification, isolation by florescence-activated cell sorting (FACS), and e­ valuation of tuft cells when crossed to a mouse line harboring conditional fluorescent reporter allele like ROSA-flox-­ STOP-floxEGFP. Additionally, Gfi1b-EGFP, IL25-RFP, and Trpm5-EGFP transgenic mouse lines have likewise demonstrated restricted ­expression to tuft cells and do not require a separate fluorescent reporter allele [1, 2, 15]. The focus of this chapter is to provide step-by-step methods for (1) producing conditioned media that supplies the necessary growth factors for organoid culture, (2) isolation of crypts from mouse small intestine, (3) culturing crypts in ECM and c­ onditioned medial to generate organoids, (4) splitting organoids for continual maintenance in culture, and (5) analyzing organoids by immunostaining, qPCR, flow cytometry, and western blot for ­ responses to Type-2 immune responses.

2  Materials 2.1  Generating Conditioned Media

1. Cultrex R-spondin1 293T cells (Trevigen). 2. Tissue culture treated dishes 150 × 20 mm (Genesee).

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3. Dulbecco’s phosphate buffered saline (DPBS (1×), Gibco). 4. Selection media: Dulbecco’s modified eagle medium (DMEM, Gibco), 10% fetal bovine serum (Gemini), 100 U/mL penicillin/streptomycin (Invitrogen), 2 mM GlutaMax (Gibco), and 300 μg/mL zeocin (Thermo Fisher). 5. Culture media: Dulbecco’s modified eagle medium (DMEM, Gibco), 10% fetal bovine serum (FBS), 100 U/mL penicillin/ streptomycin (Invitrogen), 2 mM GlutaMax (Gibco). 6. Harvest media: advanced Dulbecco’s modified eagle medium/F12 (DMEM/F12, Gibco), 100 U/mL penicillin/ streptomycin (Invitrogen). 7. 0.22 μm bottle top filter. 8. Freezing media: DMEM, 20% FBS, 10% Dimethyl sulfoxide (DMSO). 2.2  Crypt-Enriched Intestinal Epithelial Isolation

1. Dulbecco’s phosphate buffered saline (DPBS): 1× DPBS. 2. 0.5 M EDTA, pH 8.0. 3. 3 mM EDTA. 4. 10 cm petri dishes. 5. 70% Ethanol. 6. Dissection tools: surgical scissors, dissection forceps. 7. Glass plate. 8. 100 μm cell strainer. 9. 2× ISC Medium: Advanced DMEM/F12, 200× N2 (Invitrogen), 100× B27 without Vitamin A (Gibco), 2 mM HEPES (Gibco), 4 mM Glutamax (Gibco), 200 U/mL penicillin/streptomycin (Gibco) (see Note 1).

2.3  Organoid Culture and Cytokine Treatment

1. Tissue culture plate (Genesee) (see Note 2). 2. Extracellular matrix: growth factor reduced matrigel (Corning) or Cultrex (Trevigen) (see Note 3). 3. Recombinant interleukin 4 (IL-4). 4. Recombinant interleukin 13 (IL-13). 5. 2× ISC Medium (see Note 1). 6. ENR (EGF/Noggin/R-spondin 1) medium: 50% 2× ISC media (see above), 40% Advanced DMEM/F12, 10% RSPO1-­ conditioned media (see Subheading 2.1 above for creation of this conditioned media), 50 ng/mL recombinant murine EGF, 100 ng/mL recombinant murine Noggin, and 10 μM Y-27632 (selleck chemicals) (see Notes 4–6).

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Table 1 Suggested Taqman probes for qRT-PCR analysis Gene name

Cell type

Taqman gene expression assay ID

18S

Housekeeping gene

Hs99999901_s1

Chga

Enteroendocrine cells

Mm00514341_ml

Dclk1

Tuft cells

Mm00444950_m1

Lyz2

Paneth cells

Mm00727183_s1

Muc2

Goblet cells

Mm00545872_m1

SI

Absorptive enterocytes

Mm01210305_m1

2.4  RNA Lysis and Gene Expression Analysis

1. RNAqueous Micro Kit (Ambion). 2. Dulbecco’s phosphate buffered saline. 3. 1.7 mL microcentrifuge tubes. 4. iScript cDNA synthesis kit (BioRad). 5. Taqman probes (see Table 1).

2.5  Protein Lysis for Western Blot

1. Cell recovery solution (Corning). 2. 1.7 mL microcentrifuge tubes. 3. Dulbecco’s phosphate buffered saline. 4. 2× RIPA buffer: 0.3 M NaCl, 0.1 M Tris-HcL, 0.05% Sodium Azide, 2% Triton X-100, 2% Sodium Deoxycholate (w/v), and 0.2% Sodium dodecyl sulfate in H2O. 5. Protease inhibitor cocktail (Sigma). 6. Phosphatase inhibitor cocktail (Sigma). 7. Phenylmethylsulfonyl fluoride (PMSF). 8. 2× RIPA inhibitor buffer: 1% protease inhibitor cocktail, 1% phosphatase inhibitor cocktail, and 1 μM PMSF in 2× RIPA buffer.

2.6  Immunofluorescence Analysis: Sections

1. 4% Paraformaldehyde (PFA). 2. 30% Sucrose in H20. 3. Parafilm. 4. 15 mm × 15 mm × 5 mm Cryomold. 5. Optimal cutting temperature (OCT) compound. 6. Dry ice. 7. Charged glass slides (Superfrost Plus).

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1. 4% Paraformaldehyde (PFA). 2. Dulbecco’s phosphate buffered saline (DPBS). 3. 0.5% Triton X-100 in PBS. 4. 100 mM glycine in PBS. 5. Normal goat serum (NGS). 6. Immunofluorescence (IF) Buffer: 0.1% BSA, 0.2% Triton X-100, and 0.05% Tween-20 in PBS. 7. Primary and secondary antibodies (end user determined targets). 8. Parafilm. 9. Optional: Bisbenzimide. 10. Optional: Antifade media.

2.8  Preparation of Cells for Staining and Analysis of Intracellular Markers by Flow Cytometry/FACS 2.8.1  For Crypt Dissociation Flow Cytometry/FACS

1. Hanks Balanced Salt Solution (HBSS) (Gibco). 2. 50 U/mL Dispase. 3. 20,000 U/mL Deoxyribonuclease I (DNase I) from bovine pancreas. 4. Y-27632 (Selleck Chemical). 5. Dissociation solution: 0.6 U/mL Dispase, and 120 U/mL DNase I, and 10 μM Y-27632 in HBSS. 6. Fetal bovine serum (FBS). 7. 40 μm cell strainer (Falcon). 8. Dulbecco’s phosphate buffered saline (DPBS). 9. 2× ISC Medium (see Note 1).

2.8.2  For Dissociation of Organoids for Flow Cytometry/FACS

1. TrypLE Express (Gibco). 2. Y-27632 (Selleck Chemical). 3. Water bath set to 37 °C. 4. Dulbecco’s phosphate buffered saline (DPBS (1×), Gibco). 5. 40 μm cell strainer (Falcon). 6. 2× ISC Medium (see Note 1).

2.8.3  For Intracellular Staining for Flow Cytometry/FACS

2.9  Serial Passaging for Functional Analysis

1. 1% Bovine serum albumin (BSA) in 1× DPBS. 2. 4% Paraformaldehyde (PFA) in H2O. 3. Permeabilization buffer: permeabilization buffer (10×) (eBioscience) diluted to 1× in 1% BSA in PBS. 1. TrypLE-Express (Gibco). 2. Y-27632 (Selleck Chemical). 3. 1.7 mL microfuge tubes. 4. Advanced DMEM/F12.

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5. Extracellular matrix: growth factor reduced Matrigel or Cultrex. 6. ENR (EGF/Noggin/R-spondin 1) medium (see Note 7).

3  Methods 3.1  Generating Conditioned Media

1. Plate Cultrex Rspondin1 293T cells in 15 cm tissue culture plate with 20 mL selection media. 2. Allow cells to grow to ~95% confluency, changing media every 2 days. 3. Split cells 1:25 into 25 15 cm tissue culture plates with 20 mL culture media. 4. Allow plates to grow to approximately 50% confluency. Remove culture media. Rinse plates twice with room temperature 1× DPBS. Add 20 mL harvest media to plates. 5. After ~24 h, collect media from plates. This is the “first harvest” of conditioned media. Filter through 0.22 μm filter into sterile container and store at 4 °C until second harvest. 6. Add 20 mL culture media to each plate and allow cells to grow to 80–90% confluency. Again, remove culture media. Rinse plates twice with 1× DPBS. Add 20 mL harvest media to plates. 7. After ~24 h, collect media from plates. This is the “second harvest” of conditioned media. Filter through 0.22 μm filter into sterile container and store at 4 °C (see Note 8). 8. After both collections are obtained, combine harvests in a sterile container and thoroughly mix. 9. Aliquot media into 40 mL aliquots and store frozen at −80 °C (see Note 9).

3.2  Crypt-Enriched Intestinal Epithelial Isolation

1. Prepare and label the following 50 mL conical tubes, one set (a–d) per mouse. All tubes should be prechilled on ice or at 4 °C prior to crypt isolation:

(a) Tube “P”: 10 mL DPBS



(b) Tube “E1”: 10 mL 3 mM EDTA in DPBS



(c) Tube “E2”: 10 mL 3 mM EDTA in DPBS



(d) Tube “S”: 10 mL DPBS

2. Prepare and label two 10 cm petri dishes containing 10 mL DPBS each, per mouse. 3. Euthanize mouse in accordance with institutionally approved humane practices. 4. Clean mouse abdomen around planned incision area using 70% EtOH.

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5. Open abdomen with surgical scissors and dissect desired length of intestine (see Note 10). 6. Place dissected intestine in first of two DPBS containing 10 cm petri dishes. 7. Open intestine longitudinally by cutting down the length of the lumen with sharp surgical scissors (see Note 11). 8. Rinse opened intestine by gripping tissue with forceps and gently swirling in DPBS containing petri dish to remove fecal matter. 9. Transfer intestinal tissue to tube “P” using forceps and invert gently 5–10 times to remove remaining fecal matter. 10. Transfer intestinal tissue to tube “E1” and incubate for 15 min at 4 °C, with gentle agitation (see Note 12). 11. While tissue is incubating in “E1,” prepare a glass plate or dish by cleaning with 70% EtOH followed by sterile DPBS. 12. Remove tissue from “E1” and transfer to prepared glass plate, positioned so that luminal side is facing up. 13. Using a sterile pipette tip, gently “brush” the full length of intestinal tissue, first in one direction and then in the opposite direction. This step will remove a majority of villus tissue, leaving crypts intact. Avoid brushing too forcefully or for an extended period of time in order to preserve crypts (see Fig. 2B vs. 2C). 14. Transfer brushed intestine to second DPBS containing 10 cm petri dish and rinse off remaining villi by gripping tissue with forceps and gently swirling, as in step 8. 15. Transfer tissue to lid of second DPBS containing petri dish (lid should not contain any buffer) and cut into 2–3 cm pieces using surgical scissors. 16. Transfer intestinal pieces into tube “E2” and incubate for 35 min at 4 °C, with gentle agitation, as in step 10. 17. Transfer intestinal pieces into tube “S” and shake gently for 2–5 min to remove epithelium (including crypts) (Fig. 2D) (see Note 13). 18. Add 10 mL DPBS to tube “S” and filter crypt epithelial slurry through 100 μm cell strainer into new 50 mL conical to enrich epithelial suspension for crypts (Fig. 2E). 19. Pellet crypts at 500 × g-force, 4 °C for 5 min. 20. Discard supernatant and resuspend crypt pellet in 250–500 μL 1× ISC media. 21. Prepare three 1:10 dilutions of concentrated crypt slurry and determine average number of crypts per 1 μL volume by counting on inverted light microscope (see Note 14).

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Fig. 2 Quality control steps in intestinal crypt isolation. Crypt-enriched epithelial preparations should be ­monitored by light microscopy at several critical steps in order to ensure high-quality isolation (A). First, gentle “brushing” of intestinal tissue following an initial 15 min incubation in 3 mM EDTA removes a majority of villi (B). Care should be taken to avoid over brushing or applying too much force while removing villi, as this will displace crypts as well (C). Further incubation in 3 mM EDTA for 35 min, followed by shaking of intestinal tissue in DPBS, yields a mixed fraction of crypts and villi (D). Finally, filtering the mixed crypt/villus epithelial “slurry” through a 100 μm cell strainer enriches for the crypt fraction, which can be used for crypt culture of further dissociated to single cells for analysis by flow cytometry (E)

3.3  Organoid Culture and Cytokine Treatment

1. Pre-chill all tubes and tips to be used for handling 3D extracellular matrix (ECM) reagent (see Note 15). 2. Depending on experimental needs, place a 48-well or 96-well plate(s) in cell culture incubator to pre-warm to 37 °C prior to plating ECM. 3. Prepare the total volume of ECM needed per crypt sample by transferring to a 1.7 mL conical on ice (see Notes 16 and 17). 4. Using crypt concentration previously determined in Subheading 3.2, step 21, add desired number of crypts to be plated to ECM and mix by gently pipetting to disperse crypt slurry evenly in ECM (see Notes 18–20). 5. Create “bubbles” of crypt-containing ECM by pipetting appropriate volume (10 μL for 96-well plate; 20–50 μL for 48-well plate) directly in the center of each well of the plate (Fig. 3A–D).

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Fig. 3 Best practices for placing ECM and adding/removing media from ECM-based cultures. ECM plating ­technique and media change instructions. (A) Start with the pipette tip touching center of the well base. (B) While gently ejecting the ECM, slowly raise the pipette tip from the well base. (C and D) Leaving a small amount of ECM in the pipette tip to not introduce bubbles, remove the pipette tip straight up out of the ECM patty. (E) We recommend adding media by gently pipetting down the side of the well plate to avoid direct contact with the ECM. (F) To aspirate media prior to media changes, pipet tips can be placed in contact with the base of the well, adjacent to the ECM. (G) Care should be taken to avoid making direct contact with the ECM either by pipet tip or forceful ejection of media, as this can compromise the structure of the ECM and result in loss of organoids

6. Taking care to avoid disturbing freshly plated “bubbles,” transfer well plate to 37 °C cell culture incubator and allow to polymerize for 15–20 min (see Note 21). 7. While ECM is polymerizing, prepare ENR media: 250 μL per well of 48-well plate; 100 μL per well of 96-well plate (see Note 22). 8. Following polymerization, overlay ECM “bubbles” with appropriate volume of media. To avoid damaging ECM, pipet media down the side of each well. Avoid pipetting media directly into ECM “bubble” (Fig. 3E–G). 9. Complete ENR media should be changed every 48 h throughout duration of organoid cultures. 10. Following organoid establishment, Type-2 epithelial responses (e.g.,: goblet and tuft cell differentiation) can be induced by the addition of cytokines IL-4 and IL-13. We have noted strongest induction of goblet/tuft cell hyperplasia in jejunal organoids treated with 100 ng/mL IL-13 (BioLegend), but optimal cytokine concentrations should be determined per intestinal segment studied and by manufacturer source of recombinant protein (see Note 23).

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3.4  RNA Lysis and Gene Expression Analysis

1. To lyse and isolate total RNA from organoid cultures which consist of low numbers of cells, we recommend the RNAqueous Micro Kit (Ambion). 2. Aspirate full volume of media from wells to be lysed using P200 (96-well plate) or P1000 (48-well plate) pipette tip. Take care to avoid disturbing ECM bubble (Fig. 3E–G). 3. Rinse wells once by adding and aspirating 100 μL (96-well plate) or 250 μL (48-well plate) 1× DPBS. 4. Lyse organoids in ECM bubbles by adding 200 μL (96-well plate) or 500 μL (48-well plate) lysis buffer (Ambion) directly to center of each well. Allow lysis buffer to incubate with 3D ECM/organoids for ~5–10 s at RT, and then collect lysate while scraping bottom of well with pipette tip to break up and dissolve any remaining solid ECM. 5. Transfer lysates to 1.7 mL microcentrifuge tubes and prepare RNA as per manufacturer instructions or store at −80 °C until RNA isolation. 6. Prepare cDNA and conduct qRT-PCR as per standard laboratory protocols. We recommend the iScript cDNA synthesis kit (BioRad) and Taqman probes for cDNA synthesis and gene expression analysis, respectively (see Note 24). Table 1 includes the gene names and catalog numbers of Taqman probes recommended for validation of Type-2 response in epithelial organoid cultures (see Note 25).

3.5  Protein Lysis for Western Blot

1. Due to the number of crypts required for protein analysis by western blot, we recommend using cultures in 48-well plates (250–500 crypts per well) for these assays. 2. Aspirate full volume of media from wells to be lysed using P200 (96-well plate) or P1000 (48-well plate) pipette tip. Take care to avoid disturbing ECM bubble (Fig. 3E–G). 3. Add 500 μL cell recovery solution (Corning) to each well and break up ECM patty into recovery solution by scraping bottom of well and pipetting vigorously (see Note 26). 4. Incubate organoids in cell recovery solution with end-over-end rotation for 45 min at 4 °C. 5. Pellet organoids at 5000 × g-force for 5 min at 4 °C. 6. Aspirate and discard supernatant and rinse organoid pellet twice with 500 μL of ice cold DPBS. 7. Lyse organoids in 30 μL 2× RIPA inhibitor buffer. 8. Proceed with western blot analysis as per standard protocol.

3.6  Immunofluorescence Analysis: Sections

1. Aspirate full volume of media from wells to be lysed using P200 (96-well plate) or P1000 (48-well plate) pipette tip. Take care to avoid disturbing ECM bubble (Fig. 3E–G).

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2. To fix organoids in ECM, add 100 μL (96-well plate) or 250 μL (48-well plate) freshly prepared, room temperature 4% paraformaldehyde (PFA) and incubate at room temperature for 20 min (see Note 27). 3. Aspirate and discard PFA in accordance with chemical safety standards. 4. Rinse each well with 100 μL 30% sucrose three times to remove any residual PFA. 5. Add 100 μL 30% sucrose to each well, wrap well plate in parafilm to prevent evaporation, and store samples for at least 24 h at 4 °C. 6. Take p200 tip and bend tip to use as a scraping tool. Gently scrape up organoids with bent P200 and transfer organoids and sucrose to 1.7 mL microfuge tube using P200 tip attached to pipette with tip cut to create a larger bore as to not break up organoids. 7. Centrifuge at 200 × g for 5 min at room temperature to gently pellet organoids. Remove as much 30% sucrose as possible without disturbing organoids. 8. Fill a Cryomold with OCT. 9. Add organoids and sucrose to top left region of OCT in the Cryomold, making sure not to touch the edges. To evenly distribute organoids within OCT, gently swirl the organoids and sucrose in a figure-8 shape to mix the sucrose and OCT (see Note 28). 10. Freeze organoids in OCT on dry ice and store at −80 °C. 11. To section organoids: take a series of ten 8–10 μm serial sections on separate slides, and then check for sectioned ­ organoids by light microscopy (Fig. 4). When organoid ­ ­sections are located, take serial sections until full thickness of organoid has been sectioned. Repeat until desired number of sections is procured (see Note 29). 12. Proceed with standard immunohistochemical or immunofluorescent analysis, or store slides at −80 °C until use. Examples of immunofluorescent staining for DCLK1 (Tuft cells, Abgent, Cat#AP7219b), MUC2 (Goblet cells, Santa Cruz, Cat#sc-15334, 1:500) and LYZ (Paneth cells, Diagnostic Biosystems, Cat#RP028, 1:500), are in Fig. 5. 3.7  Immunofluorescence Analysis: Whole Mount

1. Aspirate full volume of media from wells to be lysed using P200 (96-well plate) or P1000 (48-well plate) pipette tip. Take care to avoid disturbing 3D ECM bubble (Fig. 3E–G). 2. Fix organoids in ECM by adding 100 μL (96-well plate) or 250 μL (48-well plate) freshly prepared, room temperature 4% PFA and incubating at room temperature for 20 min.

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Fig. 4 Sectioned organoids can be identified by light microscopy. Following sectioning, organoids can have the appearance of dust/debris (A). When rehydrated for immunofluorescence staining, characteristic epithelial morphology is observed (B)

3. Aspirate and discard PFA in accordance with chemical safety standards. 4. Rinse fixed wells three times with 100 μL PBS. 5. Add 100 μL 0.5% Triton X-100 in PBS to permeabilize organoids and incubate at RT for 20 min. 6. Aspirate permeabilization buffer and rinse wells twice with 100 mM glycine in PBS, 15 min at RT for each wash. 7. Add 100 μL of 10% NGS in IF buffer to each well and incubate at RT for 90 min to block nonspecific antigen binding. 8. Add 100 μL of primary antibody diluted in 10% NGS in IF buffer, wrap plate in parafilm to prevent evaporation, and incubate overnight at 4 °C. 9. Aspirate primary antibody and wash each well three times in IF buffer at RT, 20 min per wash. 10. Add 100  μL of secondary antibody diluted in 10% NGS in IF buffer, incubate at RT for 2 h. 11. Aspirate secondary antibody and wash each well three times in IF buffer at RT, 20 min per wash. 12. Optional: To detect nuclei, dilute bisbenzimide 1:1000 in 1× DPBS and add to wells for 20 min at RT. 13. Wash wells three times with an excess of 1× DPBS. 14. Add 1× DPBS or antifade media to wells and image samples immediately.

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Fig. 5 Wild-type intestinal organoids contain tuft, goblet, and paneth cell populations. Canonical markers for tuft cells (DCLK1, A), goblet cells (MUC2, B), and paneth cells (LYZ2, C) can be used to assess secretory cell numbers in intestinal organoids by immunofluorescence 3.8  Analysis by Flow Cytometry

3.8.1  Dissociation of Crypts for Flow Cytometry/FACS

If analyzing primary intestinal crypts, follow methods in Subheading 3.1 to isolate crypts and then proceed to Dissociation of crypts for flow cytometry/FACS (Subheading 3.8.1). If analyzing organoid cultures, proceed to Dissociation of organoids for flow cytometry/FACS (Subheading 3.8.2). If analyzing with intracellular antibodies and fixation is necessary for analysis, proceed to Intracellular staining for flow cytometry/FACS (Subheading 3.8.3) after isolating single cells from either described method. 1. Pellet crypts at 1800 × g-force for 5 min at 4 °C. 2. Resuspend crypt pellet in 1 mL HBSS, and then add to 9 mL of dissociation solution. 3. Place tube in 37 °C water bath for 10–15 min, shaking vigorously for 30 s every 2 min. 4. After each shake, observe 10 μL aliquot of solution to assess extent of dissociation to single cells (see Note 30). 5. Once dissociation is complete, add 1 mL FBS to tube and place on ice.

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6. Filter cells through 40 μm cell strainer directly into 50 mL conical containing 5 mL ice cold sterile 1× DPBS. 7. Pellet cells at 1800 × g-force for 5 min at 4 °C. 8. Wash three times with 15 mL ice cold sterile DPBS. 9. Resuspend in 1× ISC media with 10 μM Y-27632. 10. Stain cells with antibodies at proper concentration for 1 h on ice with gentle agitation 2–3 times/h (see Notes 31 and 32). 11. Rinse twice with ice cold 1× DPBS, centrifuge at 1800 × g-force for 5 min at 4 °C. 12. Resuspend in 1× ISC media with 10 μM Y-27632 and ­perform flow cytometry/FACS. 3.8.2  Dissociation of Organoids for Flow Cytometry/FACS

1. Remove media from each well and add 100 μL (for 96-well plate well) of TrypLE Express with 10 μM Y-27632. 2. Break up enteroids and ECM with P200 tip by scraping ­bottom of well and pipetting up and down 30 times. 3. Pool all wells of cells in 5 mL of pre-warmed TrypLE Express buffer with 10 μM Y-27632. 4. Place tube in 37 °C water bath for 10–15 min. Shake tube every 1–2 min for 30 s. After each shake, observe 10 μL ­aliquot of solution to judge dissociation of organoids to single cells. 5. Filter cells through 40 μm cell strainer directly into 50 mL conical containing 5 mL ice cold sterile 1× DPBS. 6. Pellet cells at 1800 × g-force for 5 min at 4 °C. 7. Wash three times with 15 mL ice cold sterile 1× DPBS. 8. Resuspend in 1× ISC media with 10 μM Y-27632. 9. Stain cells with antibodies at proper concentration for 1 h on ice with gentle agitation 2–3 times/h (see Notes 31 and 32). 10. Rinse two times with ice cold 1× DPBS, centrifuge at 1800 × g-force for 5 min at 4 °C. 11. Resuspend in 1× ISC media with 10 μM Y-27632 and ­perform flow cytometric analysis.

3.8.3  Intracellular Staining for Flow Cytometry/FACS

1. Bring single cells to concentration of 10 million cells/mL in 1% BSA in PBS. 2. If using a surface antibody, apply now and incubate for the appropriate time and temperature for your antibody (see ­ Note 32). 3. Wash cells with 3 mL 1% BSA in PBS, spin down, and aspirate supernatant. 4. Place 100 μL of RT 4% PFA on pellet and gently pipette up and down 10× to mix. Incubate for 15 min at RT.

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5. Wash cells with 3 mL 1% BSA in PBS, spin down, and aspirate supernatant. 6. Add optimized concentration of intracellular antibody to 100 μL of permeabilization buffer and place on cell pellet, pipette up and down to mix, and incubate for 30 min at RT protected from light. 7. Wash cells twice in 3 mL permeabilization buffer, spin down, and aspirate supernatant. 8. Resuspend in 500 μL 1% BSA in PBS for analysis by flow cytometry/FACS. 3.9  Serial Passaging for Functional Analysis

1. Count number of living organoids in each well and record. 2. Remove media from each well. 3. Add 250 μL TrypLE Express with 10 μM Y-27632 to each well of 48-well plate. 4. Scrape up each ECM bubble, pipette up and down 75 times to dissociate ECM and organoids, and transfer to 1.7 mL tube containing 250 μL TrypLE Express with 10 μM Y-27632. 5. Place in 37 °C water bath for 2.5 min. 6. Pipette cells up and down 20 times. 7. Place in 37 °C water bath for 2.5 min. 8. Add 1 mL ice cold advanced DMEM/F12 and place tube on ice. 9. Centrifuge at 2000 × g-force for 5 min to pellet cells. 10. Carefully remove supernatant and add appropriate amount of ECM to each tube on ice (see Note 33). 11. Pipette up and down at least 50 times to resuspend cells within ECM and plate as directed in Subheading 3.3. 12. Allow ECM bubbles to polymerize for 15–20 min in 37 °C incubator. 13. Overlay ECM bubbles with 250 μL ENR media (see Note 34). 1 4. Allow organoids to grow in culture for 7–10 days, and then record number of living organoids present in each well. This number reflects the increase in organoid number after passaging. If necessary, repeat passaging procedure until number of required serial passages is completed (see Note 35).

4  Notes 1. Media can be made 1× by adding equal volume of advanced DMEM/F12 to 2× ISC media. 2. Other tissue culture plates may be incompatible with Matrigel/ Cultrex due to the surface charge after plastic treatment.

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3. Both commercially available matrices have been used to successfully grow intestinal organoids. To our knowledge and at the time of this publication, no studies quantifying differences in organoid performance using each matrix exist. 4. Recombinant mouse R-spondin1 (250 μg/mL, R&D Systems) can be used instead of conditioned media, but conditioned media is thought to be more biologically active. 5. Y-27632 is included for first 48 h of culture only. 6. 50 μg/mL primocin (Invivogen, 1000× of 50 mg/mL stock) can be added to media if concerned about contamination at time of crypt isolation and organoid establishment. Primocin is included for first 48 h of culture ONLY. Primocin can also be used at 100 μg/mL (500×) if contamination persists with 50 μg/mL concentration. 7. Optional: To encourage single cell growth, cultures receive 0.3 nM CHIR-99021 and 100 nM valproic acid for the first 2 days in culture only. 8. Cells can be discarded or frozen back down for storage in freezing media and subsequently used to make additional batches of conditioned media. 9. Protein activity in conditioned media can be determined by commercially available ELISA or protein activity assays, as desired. 10. Regional differences in gene expression, morphology, and performance in epithelial prep and crypt culture have been noted between duodenum, jejunum, and ileum. It is important to control for portion of intestine used when planning crypt isolation and culture experiments. 11. Fine iris scissors produce best results when opening intestines. 12. Agitation on a rocking platform is recommended. 13. Shaking force and time will affect quality and extent of crypt yield and must be determined empirically by each user. We recommend examining progress of crypt isolation from intestinal tissue by removing 10 μL aliquots from tube “S” at 1 min intervals to check for the presence of intact, well-separated crypts. If crypt yield is especially low, remnant intestinal tissue can be fixed and examined by histology for the presence of unreleased crypts. 14. To prevent drying of aliquots while counting, we recommend pipetting each 1:10 dilution into a separate region of a 10 cm petri or cell culture dish. 15. Prechilling tubes and tips is optional but recommended for users who are new to Matrigel or Cultrex. Both reagents have been used successfully by a number of groups; this protocol will refer to Matrigel and Cultrex collectively as “ECM.” End users are encouraged to test reagents empirically and decide which is best for their projects.

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16. For experiments in 48-well plates, we recommend 20–50 μL of ECM per well (depending on number of crypts plated); for experiments in 96-well plates, we recommend 10 μL of ECM per well. 17. We recommend plating crypts at a density of 5–10 crypts/μL of ECM. 18. Take care to avoid introducing bubbles in ECM; never pipet past first stop on pipette. 19. Addition of crypt slurry will dilute ECM, so using a very concentrated crypt sample will reduce the amount by which ECM is diluted. We strongly recommend avoiding dilution of ECM past 50%. 20. We recommend a crypt density of ~50–100 crypts per 10 μL droplet in 96-well plate and 250–500 crypts per 50 μL droplet in 48-well plate. Plating crypts too densely will result in poor survival of organoid cultures. 21. Polymerization may take longer than 15 min depending on dilution factor of ECM. In our experience, crypts can be left in polymerized ECM at 37 °C for up to 45 min without any notable loss in organoid-forming ability. 22. To account for pipetting error, we recommend preparing 0.5-­well volume more than needed. 23. Organoids are considered “established” when they start developing well-defined crypt-like buds (Fig. 6). The appropriate culture timepoints for organoid establishment, expansion, and cytokine treatment should be determined empirically based on the needs of the experiment. We recommend allowing organoids to establish in culture for at least 48 h prior to treatment, to allow for removal of anoikis inhibitor Y-27632.

Fig. 6 Organoids in culture form crypt buds. After 24-h inculture, crypts ball up to form spheres (A). After 3–7 days in culture, organoids develop well-defined crypt-like buds (arrow) (B)

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24. cDNA prepared from RNA lysates of 50–100 organoids per 96-well plate performs well in qRT-PCR assays when diluted in a range of 1:5–1:10. 25. We recommend using Muc2 and Dclk1 as controls for goblet and tuft cell hyperplasia, respectively, and Lyz2, Chga, and SI to validate that paneth, enteroendocrine, and absorptive enterocyte lineages remain unaffected by cytokine treatment. 26. Cell recovery solution is critical to eliminate as much ECM as possible prior to crypt lysis. Residual ECM will affect total protein assays and may affect western blot results, depending on proteins of interest. 27. Avoid adding cold PFA to plates, as this can compromise integrity of ECM and result in loss of organoids. 28. Evenly mixing fixed organoids into OCT is important for downstream sectioning; if organoids are not evenly distributed within the Cryomold, it will be more difficult to acquire ­high-­quality sections. 29. Organoid sections are not visible by naked eye unless organoids are very large at time of processing. Examining serial sections by microscopy is essential for obtaining sectioned tissue. 30. Many factors can cause differences in the length of time it takes for crypt dissociation, including mouse age/genotype, intestinal region, or treatment. We recommend constantly checking each sample for single cell dissociation. 31. If staining for live/dead discrimination, vital dyes should be added immediately before running samples on flow cytometer/ FACS instrument. 32. Antibodies and antibody concentrations must be empirically determined by end user. 33. The amount of ECM to cells is dependent on experimental conditions and size of plate; we recommend 20–25 μL for each well of a 48-well plate and 8–10 μL for 96-well plate. 34. To encourage single cell growth, cultures receive 0.3 nM CHIR-99021 and 100 nM Valproic Acid for the first 2 days in culture only. 35. Organoids will eventually become very crowded within each well, so we suggest keeping organoid concentration similar across treatments and passages by increasing the passaging ratio (1:2, 1:4, etc.) as cultures become crowded. It is ­important to record this passaging ratio to accurately reflect number of organoids resulting from serial passaging even if all organoids are not plated/counted.

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Chapter 30 Utilization of Air–Liquid Interface Cultures as an In Vitro Model to Assess Primary Airway Epithelial Cell Responses to the Type 2 Cytokine Interleukin-13 Jamie L. Everman, Cydney Rios, and Max A. Seibold Abstract The airway epithelium lines the respiratory tract and provides the primary protective barrier against inhalational insults including toxic environmental substances and microorganisms. The airway epithelium also plays a critical role in regulating airway immune responses. The airway epithelial response to the type 2 cytokine, interleukin-13 (IL-13), is critical to airway inflammation, mucus production, and airway hyperresponsiveness present in asthma. Relevant primary cell models of the human airway epithelium are needed to investigate the biology of IL-13-mediated airway epithelial effects. Here, we describe the generation of a differentiated mucociliary human airway epithelium using an in vitro air–liquid interface (ALI) culture model system. We also describe methods to stimulate this culture model with IL-13 and harvest cells and biomolecules to interrogate cellular and molecular aspects of the airway epithelial IL-13 response. Key words Airway epithelial cells, Air–liquid interface, IL-13, Gene expression, Asthma

1  Introduction Chronic airway diseases, including asthma, have a multifactorial etiology involving multiple genetic and environmental factors. This complexity results in a heterogeneous asthmatic population with respect to pathobiological mechanisms (i.e., disease endotypes) driving disease development and persistence. The most common asthma endotype is driven by the production of type 2 cytokines (Interleukin-4, Interleukin-5, and Interleukin-13) from airway infiltrating immune cells [1, 2]. This type 2-high asthma endotype is present in ~50% of asthmatics and is characterized by increased airway eosinophilia, mucus production, and airway hyperresponsiveness (AHR) [3, 4]. Although these cytokines are all important in the disease process, IL-13 has been demonstrated to be particularly important in triggering asthma symptoms using in vitro epithelial and animal models [5–8]. IL-13 mediates its effects through the induction of cytokine signaling in a range of R. Lee Reinhardt (ed.), Type 2 Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1799, https://doi.org/10.1007/978-1-4939-7896-0_30, © Springer Science+Business Media, LLC, part of Springer Nature 2018

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cell types including T cells, B cells, eosinophils, airway smooth muscle cells, fibroblasts, and airway epithelial cells [5]. While the response of each of these cell types is relevant for asthma disease, IL-13 stimulation of the mouse airway epithelium alone has been shown to be sufficient for mediating cardinal type 2-high disease features, including AHR and mucus metaplasia [6]. IL-13 mediates its effects on the airway epithelium by signaling through the heterodimeric IL-13Rα1 and IL-4Rα receptor and activation of STAT6 driven transcription, setting off a cascade of gene regulation that drives mucus metaplasia and the production of inflammatory mediators [6]. Despite intense study, the full biology of the airway epithelial response to IL-13, including induced genes, transcription factors activated, and pathways induced and repressed, is incompletely known, especially in the human airway epithelium. Moreover, much work remains to understand the effects of chronic IL-13 stimulation and how this stimulus interacts with or is affected by other important disease driving environmental and genetic factors. Additionally, studies are needed to investigate effects of inhibiting various aspects of the IL-13 response in airway epithelium for potential therapy. In summary, reliable models of the human airway epithelium are greatly needed to investigate IL-13 and its role in airway epithelial dysfunction. The collection of airway epithelial cells through epithelial brushings, performed in the context of bronchoscopies, provides a rich source of airway cells for research. Moreover, recent studies showing the ease of nasal airway epithelium collection by brushings, and high overlap of the nasal and bronchial airway epithelium on a cellular, molecular, and functional level, have made collection of primary human airway epithelial cells even more feasible [9]. The characteristics of these isolated cells can be directly studied through evaluation of cell type percentages, RNA/protein expression, and other cellular assays, all of which can be related to clinical traits. While evaluation of these ex vivo cells is valuable, more mechanistic, controlled studies of airway epithelial function require an organized epithelial layer that is viable for an extended period of time and is capable of being experimentally manipulated. To accomplish this, basal airway epithelial cells can be recovered from the brushings described above and expanded in culture to generate millions of airway epithelial cells for further in vitro studies [10– 12]. Although many questions relevant to the airway can be answered by studying basal airway epithelial cells, other biology and disease-relevant questions can only be answered by the generation of a complete mucociliary epithelium. The characteristics of this epithelium include the formation of tight junctions and apical/basolateral polarization, the generation of mucus and other airway secretions, and ciliary motion. This mucociliary epithelium can be generated in culture from basal airway epithelial cells when differentiated on a semipermeable transwell insert. The differentiation process is stimulated by a cocktail of growth media

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supplements and exposure of the culture to air on the apical surface of the epithelial layer. Following approximately 21 days of culture, this process generates a fully differentiated epithelium composed of basal, goblet, club, and ciliated airway epithelial cell types. Characterization of these differentiated cultures indicates that they closely mimic responses seen within the in vivo airway, including changes in transcription [13], apical mucus production [14], cilia development and beat frequency patterns [15, 16], and responses to toxic environmental and pathogenic stimuli [17–20]. This chapter details the air–liquid interface methodology we use to generate differentiated mucociliary airway epithelial cultures from primary human basal airway epithelial cells. Additionally, we describe the methods we use for stimulation of the cultures with IL-13 and for the subsequent characterization of the cellular and molecular responses within the airway epithelium. Although this chapter is written with IL-13 investigation in mind, the generation of the cultures and many of the steps in stimulation and characterization are relevant and applicable to other studies aimed at further exploring additional environmental and immunological stimuli using this in vitro air–liquid interface model system.

2  Materials 2.1  Seeding and Expansion of Basal Airway Epithelial Cells on Transwell Membrane Inserts 2.0.1  Preparation of Airway Epithelial Cell Air–Liquid Interface (ALI) Expansion Medium

1. DMEM/BEBM/F6 base medium: 250 mL Dulbecco’s Modified Eagle’s Medium (DMEM) containing low glucose (1 g/L) and sodium pyruvate, 250 mL Bronchial Epithelial Basal Medium (BEBM—Lonza), 2 mL Bronchial Epithelial Cell Growth Medium (BEGM) SingleQuot bovine pituitary extract (Lonza), 500 μL BEGM SingleQuot hydrocortisone (Lonza), 500  μL BEGM SingleQuot epinephrine (Lonza), 500 μL BEGM SingleQuot gentamicin/amphotericin B (Lonza), 400 μL BEGM SingleQuot insulin (Lonza), and 250 μL BEGM SingleQuot transferrin (Lonza), filter sterilize using a 500 mL filter unit (0.2 μm), and store at 4 °C (see Table 1). 2. Bovine serum albumin (BSA) stock solution: Prepare 5% BSA in molecular grade water, sterilize with a 0.2 μm filter, and store aliquots at −20 °C. 3. Ethanolamine (EA) stock solution: Prepare 0.5% EA in sterile PBS, and store aliquots at −20 °C. 4. Retinoic acid (RA) stock solution: Prepare a 0.3 μg/μL RA in DMSO, and store aliquots at −20 °C. 5. Human epithelial growth factor (hEGF) stock solution: Prepare a 10 ng/μL hEGF in sterile PBS, and store aliquots at −20 °C. 6. MgCl2 stock solution: Prepare 0.3 M MgCl2 in molecular grade water, sterilize with a 0.2 μm filter, and store solutions at 4 °C.

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Table 1 DMEM/BEBM/F6 base medium recipe Reagent

Volume

DMEM (low glucose with sodium pyruvate)

250 mL

BEBM basal medium

250 mL

BEGM SingleQuot—bovine pituitary extract

2 mL

BEGM SingleQuot—hydrocortisone

500 μL

BEGM SingleQuot—epinephrine

500 μL

BEGM SingleQuot—gentamicin/amphotericin B

500 μL

BEGM SingleQuot—insulin

400 μL

BEGM SingleQuot—transferrin

250 μL

Filter sterilize and store at 4 °C

500 mL

7. MgSO4 stock solution: Prepare 0.4 M MgSO4 in molecular grade water, sterilize with a 0.2 μm filter, and store solutions at 4 °C. 8. CaCl2 stock solution: Prepare 1 M CaCl2 in molecular grade water, sterilize with a 0.2 μm filter, and store solutions at 4 °C. 9. Y-27632 dihydrochloride stock solution: 10 mM Y-27632 (ApexBio Technology) in sterile molecular grade water, and store solutions at −20 °C. 10. Complete ALI expansion medium: 20 mL DMEM/BEBM/ F6 base medium, 200 μL bovine serum albumin (BSA) stock solution, 20 μL ethanolamine (EA) stock solution, 20 μL human epithelial growth factor (hEGF) stock solution, 20 μL MgCl2 stock solution, 20 μL MgSO4 stock solution, 20 μL CaCl2 stock solution, 2 μL retinoic acid (RA) stock solution, and filter sterilize using a 0.2 μm filter (see Table 2). 11. Complete ALI expansion medium + Y-27632: 20 mL DMEM/BEBM/F6 base medium, 200 μL bovine serum albumin (BSA) stock solution, 20 μL ethanolamine (EA) stock solution, 20 μL human epithelial growth factor (hEGF) stock solution, 20 μL MgCl2 stock solution, 20 μL MgSO4 stock solution, 20 μL CaCl2 stock solution, 2 μL retinoic acid (RA) stock solution, filter sterilize using a 500 mL filter unit (0.2 μm), and add 20 μL Y-27632 dihydrochloride stock solution to pre-filtered media (see Table 2). 2.1.1  Collagen Coating Transwell Inserts

1. Bovine collagen solution: 3 mg/mL type I collagen (Advanced BioMatrix). 2. 6.5 mm transwell, 0.4 μm pore, polyester membrane insert (Corning) (see Note 1).

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Table 2 Complete ALI expansion medium recipe

Reagent

For 20 mL of media

DMEM/BEBM/F6 base medium

20 mL

Bovine serum albumin (BSA) stock solution

200 μL

Ethanolamine (EA) stock solution

20 μL

Human epithelial growth factor (hEGF) stock solution

20 μL

0.3 M MgCl2 stock solution

20 μL

0.4 M MgSO4 stock solution

20 μL

1 M CaCl2 stock solution

20 μL

Retinoic acid (RA) stock solution

2 μL

Filter sterilize Y-27632 dihydrochloride (10 mM stock solution) (for initial cell seeding only)

20 μL

3. 10× sterile phosphate buffered saline without calcium/magnesium (10× PBS). 4. 1× sterile phosphate buffered saline without calcium/magnesium (1× PBS). 5. 24-well tissue culture treated cell culture plate. 2.1.2  Seeding Airway Epithelial Cells to Transwell Inserts

1. Primary basal airway epithelial cells. 2. Hank’s balanced salt solution (HBSS) with calcium/magnesium (Corning). 3. 2× DNase solution: 0.5 mg/mL deoxyribonuclease I from bovine pancreas dissolved in Hank’s balanced salt solution (HBSS) with calcium/magnesium, 0.2 μm filter sterilized. 4. Hemocytometer. 5. Trypan blue solution: 0.4% trypan blue in PBS. 6. Humidified tissue culture incubator at 37 °C with 5% CO2.

2.1.3  Expansion of Basal Cells on Transwell Inserts

1. Complete ALI expansion medium without Y-27632 dihydrochloride stock solution. 2. Inverted bright-field microscope.

2.2  Establishing the Air–Liquid Interface and Differentiation of Airway Epithelial Cell Monolayers

1. PneumaCult-ALI medium kit (StemCell Technologies): PneumaCult-­ALI basal medium, PneumaCult-ALI 10× supplement, and PneumaCult-ALI maintenance supplement (100×). 2. Heparin solution (StemCell Technologies): 0.2% heparin sodium salt solution in PBS.

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3. Hydrocortisone stock solution (96 μg/mL) (StemCell Technologies). 4. Penicillin (10,000 IU)/streptomycin (10,000 μg/mL) stock solution. 5. Complete PneumaCult-ALI differentiation medium: 450 mL PneumaCult-ALI basal medium, 50 mL PneumaCult-ALI 10× supplement, 5 mL PneumaCult-ALI maintenance supplement (100×), 1 mL heparin solution, 2.5 mL hydrocortisone stock solution, 2.5 mL penicillin/streptomycin stock solution (see Table 3). 6. Inverted bright-field microscope. 2.3  Stimulation of Differentiated ALI Cultures with IL-13 Cytokine

1. 1× phosphate buffered saline without magnesium/calcium (1× PBS). 2. 1000× recombinant human (rh) IL-13 cytokine stock: 10 ng/ μL rh IL-13 in 1× PBS with 0.1% bovine serum albumin. 3. 1000× bovine serum albumin stock: 1.5 mg/mL in 1× PBS. 1. 1× phosphate buffered saline without magnesium/calcium (1× PBS).

2.4  Harvesting ALI Cultures for Cellular and Molecular Characterization

2. PBS/DTT wash solution: 10 mM dithiothreitol (DTT) in 1× PBS, warmed to 37 °C.

2.4.1  Harvesting Samples from ALI Cultures

3. Accutase solution: accutase (Fisher Scientific), 5 mM EDTA, 5 mM EGTA; warmed to 37 °C. 4. Protein lysis buffer: RIPA lysis buffer (Sigma-Aldrich) with 1× complete mini protease inhibitors (Sigma-Aldrich). 5. RNA/DNA Lysis buffer: RLT plus lysis buffer (Qiagen) plus 40 mM DTT. 6. ALI fixation buffer: 3.2% paraformaldehyde, 3% sucrose in 1× PBS; sterilize with a 0.2 μm filter, and store solutions at 4° C. 7. Cytospin buffer: 3.2% paraformaldehyde, 1.5% sucrose in 1× PBS; sterilize with a 0.2 μm filter, and store solutions at 4° C. Table 3 Complete PneumaCult-ALI differentiation medium recipe Reagent

For 500 mL of media

PneumaCult-ALI basal medium

450 mL

PneumaCult-ALI 10× supplement

50 mL

PneumaCult-ALI maintenance supplement (100×) 5 mL Heparin solution (0.2%)

1 mL

Hydrocortisone stock solution (96 μg/mL)

2.5 mL

Penicillin/streptomycin stock solution

2.5 mL

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3  Methods 3.1  Seeding and Expansion of Basal Airway Epithelial Cells on Transwell Membrane Inserts 3.4.1  Preparation of Airway Epithelial Cell Air–Liquid Interface (ALI) Expansion Medium 3.1.1  Collagen Coating Transwell Inserts

1. Prepare DMEM/BEBM/F6 base medium as indicated in Table 1. 2. Prepare 1 mL of complete ALI expansion media supplemented with Y-27632 dihydrochloride per insert to be seeded, scaled appropriately, as indicated in Table 2, and warm in a 37 °C water bath (see Note 2).

1. Using sterile forceps, add the desired number of transwell inserts to a sterile 24-well cell culture plate. 2. To prepare bovine collagen suspension, mix 0.5 mL sterile 10× PBS with 4 mL of bovine collagen solution by vortexing for 10 s. Add 150 μL of bovine collagen suspension to the apical chamber of each transwell insert and allow inserts to incubate for 1 h at room temperature. 3. Following incubation, remove the apical suspension using a micropipette. Wash both chambers of the insert by gently adding 800 μL of 1× PBS to the basolateral chamber and 200 μL of 1× PBS to the apical chamber. Remove and discard PBS wash from both chambers (see Note 3). 4. Once wash has been completed, cell suspension should be immediately seeded to the apical chamber of the insert as described in Subheading 3.1.3, step 4 below (see Note 4).

3.1.2  Seeding Airway Epithelial Cells to Transwell Inserts

1. Bring primary airway epithelial cell suspension volume to 5 mL using HBSS. Add 5 mL of 2× DNase solution and incubate suspension in a 37 °C water bath for 5 min. 2. Add 10 mL of room temperature HBSS to the suspension and pellet cells at 250 × g-force for 5 min at 4 °C; discard supernatant. 3. Suspend cell pellet in 1 mL of pre-warmed complete ALI expansion media supplemented with Y-27632. Prepare a 1:10 dilution of cell suspension in 4% trypan blue solution and determine cell concentration using a hemocytometer. 4. To seed 6.5 mm transwell inserts (growth area of 0.33 cm2), prepare a master mix such that each insert will be seeded with 2.0 × 104 cells in a total volume of 200 μL of complete ALI expansion medium + Y-27632; mix well (see Note 5). 5. Carefully add 200 μL of cell suspension master mix to the apical chamber of each transwell insert (see Note 6). Following the seeding of cells to each apical chamber, add 500 μL of

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complete ALI expansion medium + Y-27632 to the basolateral chambers of each insert. 6. Incubate seeded transwell inserts in a humidified tissue culture incubator at 37 °C with 5% CO2. 3.1.3  Expansion of Basal Cells Seeded to Transwell Insert Membranes

1. After 24 h of incubation post-seeding, carefully remove apical media from each insert and make observations as to the attachment and state of the cells on the membrane; remove basolateral media from each well (see Note 7). 2. Replace with fresh, pre-warmed complete ALI expansion medium (without Y-27632 supplement) to each insert, adding 200 μL to each apical chamber and 500 μL to each basolateral chamber; return cultures to tissue culture incubator (see Note 8). 3. Epithelial monolayers on the transwell membrane should be observed daily for confluence under 5× magnification, and fresh pre-warmed complete ALI expansion medium (without Y-27632 supplement) should be replaced in both chambers every 48 h.

3.2  Establishing the Air–Liquid Interface and Differentiation of Airway Epithelial Cell Monolayers 3.2.1  Establishing the Air–Liquid Interface of Epithelial Monolayers

3.3  Stimulation of Differentiated ALI Cultures with IL-13 Cytokine

1. Prepare complete PneumaCult-ALI differentiation medium as indicated in Table 3 (see Note 9). 2. Once airway epithelial cells reach 100% confluence on the transwell membrane, remove ALI expansion media from both apical and basolateral chambers of each transwell insert. Add 500 μL of pre-warmed complete PneumaCult-ALI differentiation medium to the basolateral chamber only, leaving the apical chamber free of medium and exposed to air; return cultures to tissue culture incubator (see Note 10). 3. Replace complete PneumaCult-ALI differentiation medium in the basolateral chamber of each insert every 48 h until cultures are fully differentiated (21+ days post-airlift), observing the cultures under 5× or 10× magnification at each media change and noting changes in the secretion of mucus and development of cilia over the course of differentiation. 1. Prior to stimulations, wash apical membranes of ALI inserts by adding 200 μL of pre-warmed 1× PBS to each apical membrane, and incubate cultures in a tissue culture incubator for 5 min. Aspirate and discard PBS wash and existing basolateral media. 2. For cytokine stimulations, prepare a 10 ng/mL solution of IL-13 by diluting the 1000× recombinant human (rh) IL-13 cytokine stock in complete PneumaCult-ALI differentiation medium. For paired control stimulations, dilute the 1000× bovine serum albumin stock to a 1× solution in complete

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PneumaCult-­ALI differentiation medium. Warm solutions in a 37 °C water bath (see Note 11). 3. Using the pre-warmed media suspensions of either IL-13 (experimental) or BSA (control), initiate stimulations by adding 20 μL to the apical chamber and 500 μL to the basolateral chamber of each respective culture; return cultures to tissue culture incubator. 4. Discard and replace fresh IL-13- or BSA-containing media in the apical and basolateral chambers every 24 h for the duration of the experimental stimulation. Observe cultures under 5× or 10× magnification at each restimulation to note any phenotypic changes in the cultures between control and IL-13 stimulated conditions (see Note 12). 3.4  Harvesting Differentiated ALI Cultures for Functional Characterization

A wide variety of information can be obtained from ALI cultured airway epithelial cells, depending on the type of samples harvested from cultured cells, as described in Table 4. Ultimately, the samples collected are dependent on the experimental questions being asked, and the number of ALI inserts should be scaled accordingly depending on the timepoints, number of cells, and replicates necessary for each experiment. For harvesting sample(s) from ALI cultures, a single sample type may be collected from one whole insert, or multiple samples may be harvested from a single insert after cells have undergone dissociation from the insert membrane, as described below.

Table 4 Applications for samples harvested from cultured ALI epithelial cells Sample type

Application

# Cells to harvest

RNA

qPCR gene expression analysis Next-generation RNA sequencing analysis

≥5 × 104 cells

DNA

Next-generation DNA sequencing Verification of gene editing

≥2 × 104 cells

Protein lysate

SDS-PAGE/western blot Protein-protein interactions Shotgun proteomic analysis

≥1.0 × 105 cells

Fixed ALI inserts

Immunohistochemistry (histology)

Whole insert

Fixed single cells

Immunofluorescence staining (cytospins) Flow cytometry analysis

Varies

Single-cell suspensions

FACS cell sorting Single-cell RNA sequencing

Varies

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3.4.1  Harvesting a Single Sample Type from Whole Intact ALI Cultures

1. For RNA or DNA lysates: Add 200 μL of RNA/DNA lysis buffer directly to the apical chamber of the transwell insert. Allow sample to incubate for 1 min at room temperature. Collect lysate and bring to a minimum of 500 μL total volume in RNA/DNA lysis buffer; samples can be isolated immediately or stored at −80 °C. 2. For immunohistochemical staining: Wash apical chamber with warm 1× PBS for 5 min at 37 °C; discard apical wash and basolateral media. Add ice cold ALI fixation buffer to the apical (200 μL) and basolateral (500 μL) chambers; incubate on ice for 20 min. Remove fixation buffer and wash both chambers two times with ice cold 1× PBS. Store samples at 4 °C or process for histology sectioning.

3.4.2  Harvesting Multiple Sample Types from ALI Cultures Using Cellular Dissociation from ALI Membrane

1. Wash cells by adding 200 μL of PBS/DTT wash solution to the apical chamber of each insert to be harvested. Incubate in tissue culture incubator for 5 min; aspirate and discard apical and basolateral media. 2. Add 1× PBS to the apical (200 μL) and basolateral (500 μL) chambers, aspirate and discard. 3. To dissociate cells from the membrane, add 200 μL of accutase solution to the apical chamber of the insert and incubate in tissue culture incubator for 30 min (see Note 13). 4. Following the incubation, harvest cells in the apical chamber and collect in a 15 mL conical tube. Wash apical chamber with accutase solution to remove and collect all cells off of the membrane, and pool all collections in the same tube. Check all inserts under 5× magnification to verify that all cells have been dislodged and harvested from each insert (see Note 14). 5. Centrifuge cells at 250 × g-force for 5 min at 4 °C, resuspend cell pellet in 1 mL of PBS/DTT wash, and repeat centrifugation step; discard supernatant. 6. Suspend pellet in 1 mL of 1× PBS and centrifuge cells at 250 × g-force for 5 min at 4 °C; discard supernatant. 7. Suspend pellet in 1 mL of 1× PBS and determine cell count. 8. The appropriate number of cells can be harvested as listed in Table 4 (minimum guidelines) or for additional experiments at the user’s discretion.

4  Notes 1. A variety of transwell insert sizes and membrane composition options are available for use. Choice of pore size and membrane type is dependent upon the cell type being used and the downstream experiments to be conducted. For airway epithe-

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lial cells, the protocol here recommends the use of transwells that have 0.4 μm pores and are composed of polyester membrane composition. These inserts are optically clear, allowing the user to visualize the culture using a microscope for the duration of the experiment. 2. For optimal results, preparation of fresh complete ALI expansion media is recommended prior to the seeding of basal cells to transwell inserts. 3. The collagen coating on the membrane in the apical chamber is thin and fragile. While an aspirator or vacuum unit can be used to remove the PBS wash from the basolateral chamber, it is advised to use a micropipette to gently remove these washes from the apical chamber. Disruption of the collagen coating could have an adverse effect on the attachment of the epithelial cells on the membrane. 4. Various alternate protocols that detail seeding cells to air–liquid interface suggest allowing collagen coat to dry prior to cell seeding. However, for optimal results with the protocol described here, we highly recommended to not let more than 5 min pass between the final wash and plating of cells to collagen-­coated inserts. 5. Cell seeding densities of transwell inserts can range from 6.6 × 104 cells/cm2 to 3.9 × 105 cells/cm2 depending on transwell growth area size, cell type, passage number, and culture conditions. It is advised to determine the optimal cell seeding density based on user-specific cell lines and basal cell expansion conditions prior to air–liquid interface experiments. 6. Care must be taken in the seeding of the cell suspension as to not touch the collagen-coated surface in the apical chamber of the transwell insert. Scratching of the collagen deposited on the membrane may result in sub-optimal cell attachment. 7. Although some cellular debris or clumps of cells may be seen, healthy cultures should have 20–80% confluency (depending on the seeding density), and cells should be attached to the membrane. Should the majority of the cells being rounded or floating, cultures should be monitored closely as high levels of cell death may result in extended time to form a complete monolayer and potentially poor or inconsistent air–liquid interface differentiation. 8. This media change 24 h post-seeding is to remove the Y-27632 dihydrochloride supplement from the culture. The presence of this molecule retains airway epithelial cells in a stem-like state enabling them to replicate in the basal cell state more efficiently [10], and removal is necessary for cellular differentiation.

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9. Complete PneumaCult-ALI differentiation medium can be made and stored at 4 °C for up to 2 weeks. Depending on experimental culture needs, culture medium recipe can be scaled up or down according the user. 10. Cell monolayers should resemble a tight cobblestone-like pattern and must be 100% confluent prior to establishing the air-­ lift interface. It is important to ensure that monolayers are complete along the edges of the insert and no small holes or gaps exist throughout the culture. Inserts air-lifted prior to full confluence may result in failed cultures or incomplete or absent differentiation of the epithelial monolayer. 11. The IL-13 cytokine stock is suspended in a 0.1% BSA/PBS solution for stability and long-term storage of the cytokine. For paired controls in these stimulations, mock stimulated samples should be incubated with the same amount of BSA in complete media alone to account for any cellular response to the resuspension buffer itself. 12. Duration of IL-13 stimulation is dependent on the objective and experimental question being analyzed. While stimulations ranging from 24 h to 10 days have been validated using this model, stimulations may need to be optimized for different cell types or modifications to the culture model described in this chapter. 13. To encourage the cell suspension to disassociate from the membrane, remove plate from the incubator at 10 or 15 min intervals and pipet suspension up and down 3–5 times. Return plate to incubator for the duration of the incubation time. 14. Air–liquid interface cultures can differentiate into cultures with a wide range of characteristics, including mucus production levels and cell numbers which can affect the dissociation rate of the cells from the membrane. If cells are difficult to dislodge from the membrane after 30 min incubation, they may be gently agitated with a pipet tip to remove from the membrane and harvested. It is important to note that if this technique is utilized, some cell death may occur, and it is very important to ensure the cells are in a single-cell suspension prior to collection of particular samples for downstream analysis. References 1. Fahy JV (2015) Type 2 inflammation in asthma—present in most, absent in many. Nat Rev Immunol 15(1):57–65. https://doi. org/10.1038/nri3786

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Index A Adoptive cell transfer���������������������������������136, 142, 146–147 Air-liquid Interface��������������������������������������������������419–430 Airway hyperresponsiveness (AHR)��������������������� 5, 419, 420 Allergens house dust mite (der p)����������������������������������� 7, 154, 166 peanut (ara h)���������������������������������������������������������39–46 Allergic conjunctivitis (AC)������������������������������������������49–55 Allergic eye disease (AED)�������������������������������������������49–55 Allergic immunity/inflammation�������������� 2, 60, 93, 341–350, 353–358 Allergy������������������������ 39–46, 49–51, 153, 161, 211, 247, 353 Anaphylaxis������������������������������������� 40, 45, 71, 72, 74, 77–79 Antigen-presenting cells������������������������������������������� 165, 208 Anti-inflammatory�������������������������������������������������������������27 Assay for Transposase-Accessible Chromatin with high throughput sequencing (ATAC-seq)������327–339 Asthma�������������� 1, 59, 153, 165, 166, 211, 225, 247, 304, 419

B B cells��������� 103, 105, 110, 135, 212, 226–231, 233, 234, 238, 247–263, 389, 392, 420

C Chromatin immunoprecipitation (ChIP)��������������������� 266, 268, 271–273, 303–325 Chromatin immunoprecipitation followed by sequencing (ChIP-Seq)����������������������303–325 CRISPR-Cas9���������������������������������������������������������371–379 Cytokine reporter mice���������������������������������������������211–222 Cytokines stem cell factor (SCF)���������������������������������������������������83 See also Interleukins

D Dendritic cells (DCs)����������59, 166, 185–186, 196, 199–201, 206–208, 226, 229, 231, 237–239, 248, 262, 382, 389, 393

E Electroporation���������������������������������� 195, 341, 342, 345–348

Endothelial cells lymphatic endothelial cells��������������������������������������������60 Endotypes�������������������������������������������������������������������������419 Eosinophil��������� 1, 4, 5, 8, 50, 55, 60, 260, 265–273, 398, 419 Epigenetics���������������������������������������������������������������303–325 Epithelial cells airway epithelium��������������������������������������� 237, 420, 421 alveolar epithelial cells (AECs) type 1����������������������������������������������������������������������60 type 2����������������������������������������������������������������������60 bronchiolar epithelial cells (BECs)������������������� 59, 65–67 intestinal epithelial isolation������������������������������� 401, 404 intestinal epithelium, 398

F Fibrosis�������������������������������������������������������������������������������50 Filariasis�����������������������������������������������������������������������11–26 Flow cytometry�������� 66, 68, 75, 79, 81, 82, 85, 87, 88, 94, 97, 102–105, 114, 116–117, 121–132, 136, 166, 184, 188, 190–194, 196, 200, 201, 206, 207, 238, 248–251, 253, 258–260, 329, 332, 342, 388, 400, 403, 404, 406, 411–413, 427 FluoroSpot���������������������������������������������������������������155–161 Follicular dendritic cell (FDCs)��������� 226–229, 231, 233, 262 Food allergy������������������������������������������������������������������ 40, 71 Fungi�������������������������������������������������������������������� 2, 168, 384

G Gene editing������������������������������������������������������������� 372, 427 Gene regulation�������������������������������������������������������� 303, 420 Germinal center (GC)212, 214, 226, 227, 232, 247, 253, 254, 256, 257, 261, 262

H Helminth Hymenolepis diminuta�����������������������������������28, 29, 34, 37 Litomosoides sigmodontis������������������������������������������������11 Nippostrongylus brasiliensis���������������������������������� 212, 328 Helminthic therapy������������������������������������������������ 28, 29, 34 Histone acetylation H3K27ac������������������������������������������������������������ 304, 306 Hybridoma�������������������������137, 183, 184, 187, 200–202, 239

R. Lee Reinhardt (ed.), Type 2 Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1799, https://doi.org/10.1007/978-1-4939-7896-0, © Springer Science+Business Media, LLC, part of Springer Nature 2018

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Type 2 Immunity: Methods and Protocols 434  Index

  

I

P

Immunoglobulin IgE������������������������������������������������������������������������������248 immunoglobulin E reporter mouse verigem, 248 IgG1, 41, 43, 212, 226 Immunohistochemistry������������������6, 212, 248, 409, 427, 428 Immunotherapy�������������������������������������������������� 46, 153, 353 Inflammation����������1–3, 27, 28, 60, 81–91, 93–106, 109, 136, 165–181, 211, 353–358, 382, 397–416 Innate lymphoid cells group 1 innate lymphoid cell (ILC1)��������������������������110 group 2 innate lymphoid cell (ILC2)��� 1, 60, 93, 110, 398 group 3 innate lymphoid cell (ILC3)��������������������������110 Interleukin IL-3������������������������������������������������������������������������ 72, 83 IL-4��������������������������������������������������� 1, 94, 249, 401, 419 IL-5������������������������������������������������������������������ 1, 94, 419 IL-13�������������������������������������������������1, 94, 401, 419–430 Intestinal organoids������������������������������������������ 399, 411, 414 Intestinal stem cells����������������������������������������������������������399 In vitro gene expression����������������������������������������������������342

Parasite�������������� 11, 12, 18, 20–21, 25, 26, 252, 260, 360, 368 Peanut allergy���������������������������������������������������������������39–46 Peptide MHC complex������������������������������������135, 165–167, 170 Peritoneum�������������������������������������20, 72, 82–86, 90, 98, 142 Plasma cells (PC)�����������������������247, 253, 256, 257, 259, 261 Precision cut lung slices (PCLS)������������������������������237–246 Protease������������������������1–8, 71, 166, 268, 270, 271, 305, 306, 311, 402, 424 Pulmonary������������ 98, 102, 237, 382–385, 388, 389, 391–393

L Lentivirus��������������������������������������������������372, 373, 375–379 Localization������������������������������������������������������ 237–246, 372 Lung��������� 1–8, 11, 59–68, 82, 94, 97–99, 101–103, 106, 109, 136, 139, 142–144, 148, 149, 165, 166, 212–219, 221, 222, 237–246, 327–339, 381–394 Lymphatics������������������������������������������������������11, 12, 60, 238

M Macrophages alveolar macrophages������59, 381, 382, 384, 388, 391, 392 interstitial macrophages (IMs)������������������������������������382 mononuclear phagocytes�������������������������������������381–394 Mast cell bone marrow mast cell���������������������������72–73, 75, 82, 87 degranulation���������������������������������������������������� 55, 72–76 MHC tetramer�������������������������� 121, 166, 167, 176, 178–180 Microbiota�������������������������������������������������109, 359–369, 397 MicroRNA������������������������������������������������������� 341–350, 393 Microscopy confocal microscopy���������������������������� 238–240, 243–244 live cell imaging��������������������������� 239, 240, 243–245, 248 multi-photon/two-photon microscopy�������������� 226–230, 232, 240 Monocytes���������������������������������������������5, 382, 388–391, 393 Mouse model����������������������� 29, 32, 34, 35, 37, 39–46, 49–55, 136, 184, 360

N Nanoparticles��������������������������������������������������� 353–358, 387 Next generation sequencing (NGS) library preparation���338

R Rhinosinusitis�����������������������������������������������������������������������1 RNA-Seq single cell RNA-Seq������������ 276, 281, 285, 296, 299–301 low-input RNA-Seq, 76, 96 Smart-Seq 2, 276, 285

S Skin��������������������������12, 20, 22, 71, 72, 75, 81–84, 86, 88–90, 109, 136, 161 16S rRNA��������������������������������� 360, 361, 363–364, 366, 368

T Tagmentation��������������������� 304, 306, 313, 314, 317, 323, 325 T cell epitope������������������������������������������������������������153–162 T-cell fusion��������������������������������������������� 184–185, 197–199 T-cell receptor TCR-Vγ/δ������������������������������������������������������������������136 TCR-Vα/β, 205 T cell receptor transgenic mouse������������������������������ 183, 205 T cells CD4+ T cell������������������������� 166, 206, 213, 304, 327–339 natural killer T cell (NKT)������������������������� 121–132, 304 γδ T cells������������������������������������������������������������146–147 T follicular helper cell (Tfh)����������������������� 212, 225–234 T-helper 1 cell (Th1)���������������������������123, 136, 304, 347 T-helper 2 cell (Th2)���������������1, 54, 59–60, 94, 123, 136, 165–166, 211, 212, 304, 342, 344–347, 349, 353, 354, 398–400 Thymocyte development���������������������������������� 122, 123, 209 Timothy grass (TG)�������������������������� 154–156, 158, 159, 162 T lymphocyte transfection������������������������������������������������341 Tn5 transposase��������������������������������� 304, 328–330, 332, 333 Tolerance���������������������� 40, 141–142, 183, 353, 354, 356, 357 Transcription factor���������������93, 94, 109–112, 117, 122–124, 126, 129, 132, 266, 304, 321, 327, 360, 420 Transcriptomics����������������������������������������������������������������398 Type-2 immunity�����������11–26, 135–149, 211–222, 360, 400 Tyramide signal amplification (TSA)�������������������������������212

V Vascular endothelial cells (VECs)�������������������������� 60, 65–67