Lipid-Activated Nuclear Receptors: Methods and Protocols [1st ed.] 978-1-4939-9129-7, 978-1-4939-9130-3

Table of contents :
Front Matter ....Pages i-xi
Quantification of Oxysterol Nuclear Receptor Ligands by LC/MS/MS (Lilia Magomedova, Carolyn L. Cummins)....Pages 1-14
A Stable Luciferase Reporter System to Characterize LXR Regulation by Oxysterols and Novel Ligands (Samantha A. Hutchinson, James L. Thorne)....Pages 15-32
Isolation and Analysis of Human Monocytes and Adipose Tissue Macrophages (Jean-Baptiste Julla, Raphaelle Ballaire, Tina Ejlalmanesh, Jean-François Gautier, Nicolas Venteclef, Fawaz Alzaid)....Pages 33-48
Glucan-Encapsulated siRNA Particles (GeRPs) for Specific Gene Silencing in Adipose Tissue Macrophages (Emelie Barreby, André Sulen, Myriam Aouadi)....Pages 49-57
Isolation and Purification of Tissue Resident Macrophages for the Analysis of Nuclear Receptor Activity (Laura Alonso-Herranz, Jesús Porcuna, Mercedes Ricote)....Pages 59-73
Bone Marrow-Derived Macrophage Immortalization of LXR Nuclear Receptor-Deficient Cells (Ana Ramón-Vázquez, Juan Vladimir de la Rosa, Carlos Tabraue, Antonio Castrillo)....Pages 75-85
Dual Cross-Linking Chromatin Immunoprecipitation Protocol for Next-Generation Sequencing (ChIPseq) in Macrophages (David A. Rollins, Inez Rogatsky)....Pages 87-98
Analysis of LXR Nuclear Receptor Cistrome Through ChIP-Seq Data Bioinformatics (Juan Vladimir de la Rosa, Ana Ramón-Vázquez, Carlos Tabraue, Antonio Castrillo)....Pages 99-109
Quantifying Cellular Cholesterol Efflux (Sabrina Robichaud, Mireille Ouimet)....Pages 111-133
Methods for Assessing the Effects of LXR Agonists on Macrophage Bacterial Infection (Estibaliz Glaría, Jonathan Matalonga, Annabel F. Valledor)....Pages 135-141
Measuring Apoptotic Cell Engulfment (Efferocytosis) Efficiency (Matthew C. Gage)....Pages 143-152
Methods to Study Monocyte and Macrophage Trafficking in Atherosclerosis Progression and Resolution (Ada Weinstock, Edward A. Fisher)....Pages 153-165
Preparation of Frozen Liver Tissues for Integrated Omics Analysis (Ning Liang, Rongrong Fan, Saioa Goñi, Eckardt Treuter)....Pages 167-178
Identification of Nuclear Receptor Targets by Chromatin Immunoprecipitation in Fatty Liver (Natalia Becares, Inés Pineda-Torra)....Pages 179-188
The LPS/D-Galactosamine-Induced Fulminant Hepatitis Model to Assess the Role of Ligand-Activated Nuclear Receptors on the NLRP3 Inflammasome Pathway In Vivo (Yasmine Sebti, Lise Ferri, Mathilde Zecchin, Justine Beauchamp, Denis Mogilenko, Bart Staels et al.)....Pages 189-207
Analyzing T-Cell Plasma Membrane Lipids by Flow Cytometry (Kirsty E. Waddington, Inés Pineda-Torra, Elizabeth C. Jury)....Pages 209-216
Back Matter ....Pages 217-220

Citation preview

Methods in Molecular Biology 1951

Matthew C. Gage Inés Pineda-Torra Editors

Lipid-Activated Nuclear Receptors Methods and Protocols

METHODS

IN

MOLECULAR BIOLOGY

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

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

Lipid-Activated Nuclear Receptors Methods and Protocols

Edited by

Matthew C. Gage and Inés Pineda-Torra Division of Medicine, Centre for Cardiometabolic Medicine, University College London, London, UK

Editors Matthew C. Gage Division of Medicine Centre for Cardiometabolic Medicine University College London London, UK

Ine´s Pineda-Torra Division of Medicine Centre for Cardiometabolic Medicine University College London London, UK

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-4939-9129-7 ISBN 978-1-4939-9130-3 (eBook) https://doi.org/10.1007/978-1-4939-9130-3 Library of Congress Control Number: 2019930086 © Springer Science+Business Media, LLC, part of Springer Nature 2019 This work is subject to copyright. All rights are reserved by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, express or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. This Humana Press imprint is published by the registered company Springer Science+Business Media, LLC, part of Springer Nature. The registered company address is: 233 Spring Street, New York, NY 10013, U.S.A.

Preface Nuclear receptors comprise a family of ligand-activated transcription factors with crucial roles in the maintenance of cholesterol and lipid homeostasis (including biosynthesis, catabolism, and uptake by tissues and cells) while having multiple actions on innate and adaptive immunity. Indeed, the transcriptional regulation of ligand-receptor complexes constitutes an integral part of the signal transduction exerted by the lipophilic ligands of these receptors. This is exemplified by the fact that nuclear receptor aberrant activation or function has been implicated in a wide range of pathologies including autoimmune, metabolic, and vascular diseases. Nuclear receptor biology is therefore a very active field of research which employs a wide range of methodologies to explore and define their function, cross talk with other lipid- and hormone-activated nuclear receptors, and the significance of their contributions to various aspects of these different diseases. This book covers a wide range of methodologies and detailed protocols to study the actions some of these receptors have in tissues and immune cell types that can be considered classic metabolic “powerhouses” (liver, adipose tissue, and monocytes/macrophages). To these, a chapter on the identification of lipids in T cells has been added, reflecting the newly appreciated importance of metabolism in the immune actions of these cells. Although most of the chapters appear to be focused on the biology of the oxysterol receptor or liver X receptor (LXR), the majority of the methods described can be easily applied to multiple nuclear receptors, and in some cases, the authors have provided notes on how they can be adapted to other tissues or cell types. This edition in the Methods in Molecular Biology series comprehensively describes established and cutting-edge techniques, from biochemistry through molecular biology to bioinformatic analyses, which will allow the reader to confidently perform experiments to analyze the function of LXR or other nuclear receptor in their field of interest. In most cases, the detailed experimental protocols are introduced by an overview of the receptor or the cell type being discussed. As editors, we have enormously benefitted from the multidisciplinary and international network of researchers in the nuclear receptor community, and it truly has been a pleasure to go through this experience with such dedicated and eminent scientists. We are very grateful for their efforts to bring their tried and tested protocols to the rest of the research community. Finally, we sincerely hope the readers will enjoy this edition and successfully implement these methods in their labs. London, UK

Matthew C. Gage Ine´s Pineda-Torra

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

v ix

1 Quantification of Oxysterol Nuclear Receptor Ligands by LC/MS/MS . . . . . . . Lilia Magomedova and Carolyn L. Cummins 2 A Stable Luciferase Reporter System to Characterize LXR Regulation by Oxysterols and Novel Ligands . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Samantha A. Hutchinson and James L. Thorne 3 Isolation and Analysis of Human Monocytes and Adipose Tissue Macrophages . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jean-Baptiste Julla, Raphaelle Ballaire, Tina Ejlalmanesh, Jean-Franc¸ois Gautier, Nicolas Venteclef, and Fawaz Alzaid 4 Glucan-Encapsulated siRNA Particles (GeRPs) for Specific Gene Silencing in Adipose Tissue Macrophages . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Emelie Barreby, Andre´ Sulen, and Myriam Aouadi 5 Isolation and Purification of Tissue Resident Macrophages for the Analysis of Nuclear Receptor Activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Laura Alonso-Herranz, Jesu´s Porcuna, and Mercedes Ricote 6 Bone Marrow-Derived Macrophage Immortalization of LXR Nuclear Receptor-Deficient Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ana Ramon-Va´zquez, Juan Vladimir de la Rosa, Carlos Tabraue, and Antonio Castrillo 7 Dual Cross-Linking Chromatin Immunoprecipitation Protocol for Next-Generation Sequencing (ChIPseq) in Macrophages . . . . . . . . . . . . . . . . . David A. Rollins and Inez Rogatsky 8 Analysis of LXR Nuclear Receptor Cistrome Through ChIP-Seq Data Bioinformatics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Juan Vladimir de la Rosa, Ana Ramon-Va´zquez, Carlos Tabraue, and Antonio Castrillo 9 Quantifying Cellular Cholesterol Efflux. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sabrina Robichaud and Mireille Ouimet 10 Methods for Assessing the Effects of LXR Agonists on Macrophage Bacterial Infection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Estibaliz Gları´a, Jonathan Matalonga, and Annabel F. Valledor 11 Measuring Apoptotic Cell Engulfment (Efferocytosis) Efficiency . . . . . . . . . . . . . Matthew C. Gage 12 Methods to Study Monocyte and Macrophage Trafficking in Atherosclerosis Progression and Resolution . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ada Weinstock and Edward A. Fisher 13 Preparation of Frozen Liver Tissues for Integrated Omics Analysis . . . . . . . . . . . . ˜ i, and Eckardt Treuter Ning Liang, Rongrong Fan, Saioa Gon

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Contents

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Identification of Nuclear Receptor Targets by Chromatin Immunoprecipitation in Fatty Liver . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 179 Natalia Becares and Ine´s Pineda-Torra 15 The LPS/D-Galactosamine-Induced Fulminant Hepatitis Model to Assess the Role of Ligand-Activated Nuclear Receptors on the NLRP3 Inflammasome Pathway In Vivo . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 189 Yasmine Sebti, Lise Ferri, Mathilde Zecchin, Justine Beauchamp, Denis Mogilenko, Bart Staels, He´le`ne Duez, and Benoit Pourcet 16 Analyzing T-Cell Plasma Membrane Lipids by Flow Cytometry . . . . . . . . . . . . . . 209 Kirsty E. Waddington, Ine´s Pineda-Torra, and Elizabeth C. Jury Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Contributors LAURA ALONSO-HERRANZ  Myocardial Pathophysiology Area, Centro Nacional de Investigaciones Cardiovasculares Carlos III (CNIC), Madrid, Spain FAWAZ ALZAID  Centre de Recherche des Cordeliers, INSERM, Sorbonne Universite´, USPC, Universite´ Paris Descartes, Universite´ Paris Diderot, Paris, France MYRIAM AOUADI  Department of Medicine, Integrated Cardio Metabolic Center (ICMC), Karolinska Institutet, Stockholm, Sweden RAPHAELLE BALLAIRE  Centre de Recherche des Cordeliers, INSERM, Sorbonne Universite´, USPC, Universite´ Paris Descartes, Universite´ Paris Diderot, Paris, France; Inovarion, Paris, France EMELIE BARREBY  Department of Medicine, Integrated Cardio Metabolic Center (ICMC), Karolinska Institutet, Stockholm, Sweden JUSTINE BEAUCHAMP  European Genomic Institute for Diabetes (E.G.I.D.), Lille, France; UNIV LILLE, Lille, France; INSERM UMR 1011, Lille, France; CHU Lille, Lille, France; Institut Pasteur de Lille, Lille, France NATALIA BECARES  Division of Medicine, Centre of Cardiometabolic Medicine, University College of London, London, UK ANTONIO CASTRILLO  Instituto de Investigaciones Biome´dicas “Alberto Sols”, Consejo Superior de Investigaciones Cientı´ficas (CSIC), Centro Mixto CSIC-Universidad Autonoma de Madrid, Madrid, Spain; Unidad de Biomedicina IIBM-ULPGC (Unidad Asociada al CSIC), Universidad de Las Palmas de Gran Canaria, Las Palmas, Spain; Grupo de Investigacion Medio Ambiente y Salud (GIMAS), Instituto Universitario de Investigaciones Biome´dicas y Sanitarias (IUIBS) de la ULPGC, Las Palmas, Spain CAROLYN L. CUMMINS  Department of Pharmaceutical Sciences, University of Toronto, Toronto, ON, Canada JUAN VLADIMIR DE LA ROSA  Instituto de Investigaciones Biome´dicas “Alberto Sols”, Consejo Superior de Investigaciones Cientı´ficas (CSIC), Centro Mixto CSIC-Universidad Autonoma de Madrid, Madrid, Spain; Unidad de Biomedicina IIBM-ULPGC (Unidad Asociada al CSIC), Universidad de Las Palmas de Gran Canaria, Las Palmas, Spain; Grupo de Investigacion Medio Ambiente y Salud (GIMAS), Instituto Universitario de Investigaciones Biome´dicas y Sanitarias (IUIBS) de la ULPGC, Las Palmas, Spain HE´LE`NE DUEZ  European Genomic Institute for Diabetes (E.G.I.D.), Lille, France; UNIV LILLE, Lille, France; INSERM UMR 1011, Lille, France; CHU Lille, Lille, France; Institut Pasteur de Lille, Lille, France TINA EJLALMANESH  Centre de Recherche des Cordeliers, INSERM, Sorbonne Universite´, USPC, Universite´ Paris Descartes, Universite´ Paris Diderot, Paris, France RONGRONG FAN  Department of Biosciences and Nutrition, Karolinska Institutet, Huddinge, Sweden LISE FERRI  European Genomic Institute for Diabetes (E.G.I.D.), Lille, France; UNIV LILLE, Lille, France; INSERM UMR 1011, Lille, France; CHU Lille, Lille, France; Institut Pasteur de Lille, Lille, France

ix

x

Contributors

EDWARD A. FISHER  Departments of Medicine (Cardiology) and Cell Biology, and the Marc and Ruti Bell Program in Vascular Biology, New York University School of Medicine, New York, NY, USA MATTHEW C. GAGE  Division of Medicine, Centre for Cardiometabolic Medicine, University College of London, London, UK JEAN-FRANC¸OIS GAUTIER  Centre de Recherche des Cordeliers, INSERM, Sorbonne Universite´, USPC, Universite´ Paris Descartes, Universite´ Paris Diderot, Paris, France; Assistance Publique-Hoˆpitaux de Paris, Lariboisie`re Hospital, Department of Diabetes, Clinical Investigation Centre (CIC-9504), Paris, France ESTIBALIZ GLARI´A  Nuclear Receptor Group, Department of Cell Biology, Physiology and Immunology, School of Biology, University of Barcelona, Barcelona, Spain; Institute of Biomedicine of the University of Barcelona (IBUB), Barcelona, Spain SAIOA GON˜I  Department of Biosciences and Nutrition, Karolinska Institutet, Huddinge, Sweden SAMANTHA A. HUTCHINSON  School of Food Science and Nutrition, University of Leeds, Leeds, UK JEAN-BAPTISTE JULLA  Centre de Recherche des Cordeliers, INSERM, Sorbonne Universite´, USPC, Universite´ Paris Descartes, Universite´ Paris Diderot, Paris, France; Assistance Publique-Hoˆpitaux de Paris, Lariboisie`re Hospital, Department of Diabetes, Clinical Investigation Centre (CIC-9504), Paris, France ELIZABETH C. JURY  Division of Medicine, Centre for Rheumatology, University College of London, London, UK NING LIANG  Department of Biosciences and Nutrition, Karolinska Institutet, Huddinge, Sweden LILIA MAGOMEDOVA  Department of Pharmaceutical Sciences, University of Toronto, Toronto, ON, Canada JONATHAN MATALONGA  Nuclear Receptor Group, Department of Cell Biology, Physiology and Immunology, School of Biology, University of Barcelona, Barcelona, Spain DENIS MOGILENKO  European Genomic Institute for Diabetes (E.G.I.D.), Lille, France; UNIV LILLE, Lille, France; INSERM UMR 1011, Lille, France; CHU Lille, Lille, France; Institut Pasteur de Lille, Lille, France MIREILLE OUIMET  University of Ottawa Heart Institute, Ottawa, ON, Canada INE´S PINEDA-TORRA  Division of Medicine, Centre for Cardiometabolic Medicine, University College of London, London, UK JESU´S PORCUNA  Myocardial Pathophysiology Area, Centro Nacional de Investigaciones Cardiovasculares Carlos III (CNIC), Madrid, Spain BENOIT POURCET  European Genomic Institute for Diabetes (E.G.I.D.), Lille, France; UNIV LILLE, Lille, France; INSERM UMR 1011, Lille, France; CHU Lille, Lille, France; Institut Pasteur de Lille, Lille, France ANA RAMO´N-VA´ZQUEZ  Instituto de Investigaciones Biome´dicas “Alberto Sols”, Consejo Superior de Investigaciones Cientı´ficas (CSIC), Centro Mixto CSIC-Universidad Autonoma de Madrid, Madrid, Spain; Unidad de Biomedicina IIBM-ULPGC (Unidad Asociada al CSIC), Universidad de Las Palmas de Gran Canaria, Las Palmas, Spain; Grupo de Investigacion Medio Ambiente y Salud (GIMAS), Instituto Universitario de Investigaciones Biome´dicas y Sanitarias (IUIBS) de la ULPGC, Las Palmas, Spain MERCEDES RICOTE  Myocardial Pathophysiology Area, Centro Nacional de Investigaciones Cardiovasculares Carlos III (CNIC), Madrid, Spain SABRINA ROBICHAUD  University of Ottawa Heart Institute, Ottawa, ON, Canada

Contributors

xi

INEZ ROGATSKY  Graduate Program in Immunology and Microbial Pathogenesis, Weill Cornell Graduate School of Medical Sciences, New York, NY, USA; The David Rosensweig Genomics Center, Hospital for Special Surgery Research Institute, New York, NY, USA DAVID A. ROLLINS  Graduate Program in Immunology and Microbial Pathogenesis, Weill Cornell Graduate School of Medical Sciences, New York, NY, USA; The David Rosensweig Genomics Center, Hospital for Special Surgery Research Institute, New York, NY, USA; Mayo Clinic School of Medicine M.D. Program, Rochester, MN, USA YASMINE SEBTI  European Genomic Institute for Diabetes (E.G.I.D.), Lille, France; UNIV LILLE, Lille, France; INSERM UMR 1011, Lille, France; CHU Lille, Lille, France; Institut Pasteur de Lille, Lille, France BART STAELS  European Genomic Institute for Diabetes (E.G.I.D.), Lille, France; UNIV LILLE, Lille, France; INSERM UMR 1011, Lille, France; CHU Lille, Lille, France; Institut Pasteur de Lille, Lille, France ANDRE´ SULEN  Department of Medicine, Integrated Cardio Metabolic Center (ICMC), Karolinska Institutet, Stockholm, Sweden CARLOS TABRAUE  Instituto de Investigaciones Biome´dicas “Alberto Sols”, Consejo Superior de Investigaciones Cientı´ficas (CSIC), Centro Mixto CSIC-Universidad Autonoma de Madrid, Madrid, Spain; Unidad de Biomedicina IIBM-ULPGC (Unidad Asociada al CSIC), Universidad de Las Palmas de Gran Canaria, Las Palmas, Spain; Grupo de Investigacion Medio Ambiente y Salud (GIMAS), Instituto Universitario de Investigaciones Biome´dicas y Sanitarias (IUIBS) de la ULPGC, Las Palmas, Spain JAMES L. THORNE  School of Food Science and Nutrition, University of Leeds, Leeds, UK ECKARDT TREUTER  Department of Biosciences and Nutrition, Karolinska Institutet, Huddinge, Sweden ANNABEL F. VALLEDOR  Nuclear Receptor Group, Department of Cell Biology, Physiology and Immunology, School of Biology, University of Barcelona, Barcelona, Spain; Institute of Biomedicine of the University of Barcelona (IBUB), Barcelona, Spain NICOLAS VENTECLEF  Centre de Recherche des Cordeliers, INSERM, Sorbonne Universite´, USPC, Universite´ Paris Descartes, Universite´ Paris Diderot, Paris, France KIRSTY E. WADDINGTON  Division of Medicine, Centre for Cardiometabolic Medicine, University College of London, London, UK; Division of Medicine, Centre for Rheumatology, University College of London, London, UK ADA WEINSTOCK  Departments of Medicine (Cardiology) and Cell Biology, and the Marc and Ruti Bell Program in Vascular Biology, New York University School of Medicine, New York, NY, USA MATHILDE ZECCHIN  European Genomic Institute for Diabetes (E.G.I.D.), Lille, France; UNIV LILLE, Lille, France; INSERM UMR 1011, Lille, France; CHU Lille, Lille, France; Institut Pasteur de Lille, Lille, France

Chapter 1 Quantification of Oxysterol Nuclear Receptor Ligands by LC/MS/MS Lilia Magomedova and Carolyn L. Cummins Abstract Oxidative derivatives of cholesterol such as 22(R)-hydroxycholesterol, 24(S)-hydroxycholesterol, 25-hydroxycholesterol, and (25S),26-hydroxycholesterol are endogenous ligands for the liver X receptors (LXRα and LXRβ). The LXRs are nuclear hormone receptors known as “intracellular cholesterol sensors” because of their ability to bind to and be activated by oxysterols at circulating concentrations. Oxysterols are expressed in a tissue-specific manner and are generally at least 104 to 106-fold less abundant than cholesterol. Thus, the extraction and measurement of oxysterols from plasma and tissues are facilitated by the removal of bulk sterols by solid phase extraction prior to quantitative analysis by mass spectrometry. In this chapter we describe step by step methods for extracting and quantitating oxysterols from biological samples using electrospray ionization LC/MS/MS. Key words Nuclear receptors, Liver X receptor, Oxysterols, LC/MS/MS, Deuterated internal standards, 27-Hydroxycholesterol

1

Introduction Oxysterols are oxidized derivatives of cholesterol. They serve important roles in the regulation of cholesterol and immune homeostasis in part by serving as signaling molecules for nuclear receptor transcription factors. The liver X receptors, LXRα and LXRβ, are activated by the oxysterols 22(R)-hydroxycholesterol, 24(S)-hydroxycholesterol, 25-hydroxycholesterol, (25R),26hydroxycholesterol, and 24,25-epoxycholesterol [1]. Changes to oxysterol levels occur in response to various physiologic conditions such as atherogenic diet feeding [2] and activation of macrophages with TLR4 ligand [3]. Activation of LXRs by oxysterols can initiate a program of gene transcription to help decrease intracellular cholesterol in a feed-forward loop [4]. Apart from LXRs, other nuclear receptors have also been found to be regulated by specific oxysterols. For example,

Matthew C. Gage and Ine´s Pineda-Torra (eds.), Lipid-Activated Nuclear Receptors: Methods and Protocols, Methods in Molecular Biology, vol. 1951, https://doi.org/10.1007/978-1-4939-9130-3_1, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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Lilia Magomedova and Carolyn L. Cummins

27-hydroxycholesterol (equivalent to (25R),26-hydroxy-cholesterol) was shown to act as an endogenous selective estrogen receptor modulator [5]. The brain-derived oxysterol, 24(S)hydroxycholesterol, was also shown to be an inverse agonist of RORα and RORγ [6]. Oxysterols are present in plasma and tissues both in their free form and as fatty acid esters. Cytochrome P450 enzymes mediate the bulk of the enzymatic conversions of cholesterol to oxysterols (i.e., 22-, 24-, 25-, and 26- hydroxycholesterol species). However, limiting the exposure of samples to oxygen is recommended to prevent the nonenzymatic oxidation of cholesterol to specific oxysterols during sample handling. Major auto-oxidation products of cholesterol include 5,6-epoxy-, 7α-hydroxy-, 7β-hydroxy-, and 7-keto- cholesterol [7]. Steps to minimize the formation and/or measure the extent of non-specific oxidation are taken by the inclusion of antioxidants, incubation of samples in the absence of O2, and the spiking in of deuterium-labeled surrogate/internal standards at the start of the procedure [7]. Fatty acid esters represent as much as 40–90% of the total oxysterols present in a given tissue, although this is highly dependent on the tissue and oxysterol in question [7]. Removal of the fatty acid ester from the oxysterol is achieved by saponification of the plasma or tissue extracts prior to LC/MS/MS analysis. In this protocol we review procedures for extracting and quantitating oxysterols from plasma and tissues. The procedure to extract oxysterols from plasma is based on methods described by McDonald et al. [8, 9] and Lund and Diczfalusy [10] with minor modifications. Extraction of oxysterols from tissues was performed using the modified Folch method [10, 11]. After extraction, saponification of the oxysterol fatty acid esters is performed, and then oxysterols are separated from bulk cholesterol and fatty acids (or triglycerides and phospholipids if not saponified) using solid phase extraction. Levels of oxysterols are then quantified by LC/MS/MS using deuterated internal standards to control for extraction efficiency and matrix effects.

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Materials Prepare all solutions using HPLC-grade solvents and analytical grade reagents. Diligently follow all local waste disposal regulations when disposing of waste materials and solvents.

2.1 Blood and Tissue Sample Collection

1. Collect blood samples into EDTA-containing collection tubes. Centrifuge at 500
g for 20 min at 4  C. Transfer plasma into a new Eppendorf tube and freeze at 80  C until ready to use (within 3 months). 2. Collect tissue (~100 mg) and snap freeze in liquid nitrogen. Store at 80  C until ready to use.

LC/MS/MS Analysis of Oxysterols

2.2

Lipid Extraction

3

1. Butylated hydroxytoluene (BHT). Make 25 mg/mL in ethanol, and store at room temperature. 2. Dichloromethane/methanol (CH2Cl2/MeOH, 1:1, v/v). 3. Phosphate-buffered saline (PBS). 4. Dichloromethane. 5. Chloroform/methanol (CHCl3/MeOH, 2:1, v/v). 6. 50 mM NaCl. 7. Glass culture tubes with PTFE-lined caps (16
125 mm, see Note 1). 8. Table-top centrifuge capable of accommodating glass culture tubes (see Note 2). 9. Sample concentrator supplied with N2 gas (see Note 3).

2.3

Saponification

1. Ethanol. 2. 10 N KOH and 3.5 M KOH in water. 3. 0.35 M ethanolic KOH (prepare fresh from 1 part 3.5 M KOH in 9 parts EtOH). 4. 10 mL disposable borosilicate glass pipettes. 5. Portable Pipet-Aid. 6. Pasteur pipettes (900 borosilicate). 7. Concentrated phosphoric acid (85% v/v).

2.4 Solid Phase Extraction (SPE)

1. Prepacked 100 mg Isolute Silica solid phase extraction (SPE) columns (see Note 4). 2. Sample processing vacuum manifold (see Note 5). 3. Toluene. 4. Hexane. 5. 0.5% (v/v) isopropanol in hexane. 6. 30% (v/v) isopropanol in hexane. 7. 14 mL borosilicate glass test tubes (16
100 mm). 8. 8 mL amber glass vials (17
60 mm). 9. PTFE-lined caps for 8 mL amber vials.

2.5 Oxysterol and Internal Standards

1. Purified oxysterol standard for the preparation of an external standard curve. 2. Matching deuterated oxysterol standard for each analyte of interest (see Note 6). 3. Stock solutions: dilute each standard in a glass vial to 10 mM using MeOH, and store at 80  C after flushing with N2 or Ar gas. 4. Prepare the internal standard stock solution: pool stock solutions to a final concentration of 10 μM each.

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Lilia Magomedova and Carolyn L. Cummins

LC/MS/MS

1. 2 mL amber glass sample vials. 2. Vial inserts, 250 μL glass with polymer feet. 3. Guard Cartridge Eclipse XDB-C18 column (4.6
12.5 mm, 5 μm). 4. HPLC Column: TSK-gel ODS-120 T column (4.6
250 mm, 5 μm). 5. 5 M ammonium acetate (HPLC grade). 6. HPLC with degasser, binary pump, temperature-controlled column compartment, and autosampler (see Note 7). 7. Triple quadrupole mass spectrometer equipped with ESI source (see Note 7). 8. Mobile phase: 7% HPLC-grade water and 93% HPLC-grade methanol containing 5 mM ammonium acetate.

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Methods Carry out all procedures at room temperature unless otherwise specified (see Note 8). It is important to minimize the use of plastics when performing extractions with organic solvents to limit the leaching of plasticizers into the sample (see Note 9).

3.1 Plasma Oxysterol Extraction

Perform the following steps in a fume hood to minimize inhalation of volatile organic solvents. 1. Equilibrate frozen plasma samples to room temperature for 10 min prior to extraction. 2. Transfer 100 μL of plasma into a 16
100 mm borosilicate glass test tube containing 10 μL of internal standard mix. 3. Add 2 mL of dichloromethane/methanol (CH2Cl2/MeOH, 1:1, v/v) solution containing 50 μg/mL of BHT (see Note 10). 4. Vortex for 15 s, leave at room temperature for 10 min, and centrifuge at 2465
g at 25  C for 5 min. 5. Carefully decant the supernatant into a 16
125 mm glass screw cap tube (see Note 11). 6. Add 2 mL of dichloromethane/methanol (CH2Cl2/MeOH, 1:1, v/v) solution to the sample pellet and vortex for 30 s to dislodge the pellet. 7. Centrifuge at 2465
g at 25  C for 5 min and pool the supernatant with the initial extract. 8. To saponify the extract, add 200 μL of 10 N KOH and vortex for 10 s (see Note 12). 9. Flush with N2 for 10 s and incubate for 1.5 h at 35  C (see Note 13).

LC/MS/MS Analysis of Oxysterols

5

10. To induce phase separation and extract the lipids, add 2 mL of PBS and vortex for 15 s. 11. Centrifuge at 2465
g at 25  C for 5 min. 12. Using a Pasteur pipette, transfer the organic layer (bottom) to a new 16
100 mm glass test tube and set aside (see Note 14). 13. Re-extract the remaining sample by adding 2 mL of dichloromethane, vortex for 15 s, and centrifuge at 2465
g at 25  C for 5 min. Combine the lower organic layer with the initial sample (see Note 15). 14. Dry the extracted lipids under constant stream of N2 using a sample concentrator. 15. Dissolve dried lipid extract in 1 mL toluene, vortex for 15 s, and proceed to Subheading 3.3 SPE extraction. 3.2 Tissue Oxysterol Extraction

Keep samples frozen prior to extraction. Perform the following steps in a fume hood to minimize inhalation of volatile organic solvents. 1. Record the weight of the frozen tissue prior to extraction (see Note 16). 2. To a 14 mL glass test tube, add 10 μL of internal standard mix and 4 mL of chloroform/methanol (CHCl3/MeOH, 2:1, v/v) solution containing 50 μg/mL BHT (see Notes 8–10). 3. Homogenize ~100 mg of tissue in the glass tube containing the organic solvent for 30 s and let it sit at room temperature for 10 min. 4. Add 1 mL of 50 mM NaCl and vortex for 15 s; centrifuge at 1500
g at 25  C for 30 min. 5. Carefully remove organic phase (bottom) from under the “tissue debris” and place into a new 16
125 mm glass screw cap tube (see Note 17). 6. Dry the extracted lipids under constant stream of N2 using a sample concentrator. 7. Dissolve dried lipids in 1 mL of ethanol. Vortex for 30 s to fully resuspend the sample. 8. Add 3.5 mL of 0.35 M ethanolic KOH; vortex for 10 s (see Notes 12 and 18). 9. Flush with N2 gas for 10 s and incubate for 1.5 h at 35  C (see Note 13). 10. Neutralize with 10.5 μL of concentrated phosphoric acid solution. 11. Extract the lipids by adding 2.1 mL of 0.9% NaCl and 6.3 mL CHCl3. Vortex for 30 s and centrifuge at 2465
g for 5 min at 25  C.

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Lilia Magomedova and Carolyn L. Cummins

12. Transfer the organic layer (bottom) to a new glass test tube and dry under a constant stream of N2 (see Notes 17 and 19). 13. Dissolve lipid extract in 1 mL toluene, vortex for 15 s, and proceed to Subheading 3.3 SPE extraction. 3.3 Solid Phase Extraction (SPE)

Do not let the SPE column dry out between solvent additions. Stop the vacuum when the majority of the liquid has been drawn into the SPE column and only a small meniscus remains. Reinitiate the vacuum once the next solvent has been added to the reservoir. 1. Assemble the 100 mg Isolute silica cartridges on the sample vacuum manifold (see Notes 4 and 5). 2. Condition the column with 2 mL hexane. Discard the eluate (see Note 20). 3. Apply the sample in 1 mL toluene. Discard the eluate (see Note 21). 4. Wash the column with 4 mL hexane. Discard the eluate. 5. Elute cholesterol with 8 mL 0.5% 2-propanol in hexane. Discard the eluate (see Notes 21 and 22). 6. Elute oxysterols with 5 mL 30% 2-propanol in hexane into 8 mL brown glass vials (see Note 23). 7. Dry the oxysterol fraction under constant stream of N2 using a sample concentrator (see Notes 24 and 25).

3.4

LC/MS/MS

Oxysterols are resolved using reverse phase HPLC and detected using a triple quadrupole mass spectrometer. The gas temperature and collision energy parameters will need to be optimized for each analyte based on the mass spectrometer that is available for use. Parameters used to couple the HPLC to an Agilent 6410 QQQ are provided below. Those used to couple to an SCIEX 4000 QTrap have been reported previously by McDonald et al. [8]. 1. Dissolve dried lipid extracts in 150 μL of 93% HPLC-grade methanol/7% HPLC-grade water, vortex for 30 s, and transfer into labeled Eppendorf tubes. 2. Centrifuge samples at 21,000
g to pellet any insoluble material (see Note 26). 3. Pipette samples into amber mass spec vials with borosilicate glass inserts and place into the autosampler (see Note 27). 4. Operate the mass spectrometer in multiple reaction monitoring (MRM) mode using electrospray ionization (ESI). 5. Use the following ESI source parameters: Gas temp 175  C, gas flow 10 L/min, nebulizer 35 psi, and capillary voltage 6000 V (see Table 1 and Note 28).

LC/MS/MS Analysis of Oxysterols

7

Table 1 ESI acquisition parameters for the analysis of oxysterols using Agilent 6410 QQQ

Compound

MW Precursor Product Fragment or Collision Retention (g/Mol) ion ion voltage (V) energy time (min)

20S-Hydroxycholesterol

402.7

402

385

80

1

12.8

22R-Hydroxycholesterol

402.7

420

385

70

7

10.9

22S-Hydroxycholesterol

402.7

420

385

70

7

11.9

24R-Hydroxycholesterol

402.7

420

385

70

7

13.8

24S-Hydroxycholesterol

402.7

420

385

70

7

14.2

25-Hydroxycholesterol

402.7

420

367

70

3

16.7

25(R),26-Hydroxycholesterola

402.7

420

385

70

7

20.8

25(S),26-Hydroxycholesterol

402.7

420

385

70

7

21.6

400.6

418

383

155

3

20.6

24(R/S)-Hydroxycholesterol-d7 409.7

427

392

110

3

13.6, 14.0

25-Hydroxycholesterol-d6

408.7

426

373

80

7

16.5

25(R/S), 26-Hydroxycholesterol-d4

406.7

424

389

80

3

20.5, 21.3

24(R/S),25-Epoxycholesterol

b

a

Frequently referred to as 27-hydroxycholesterol in the literature Isomers are not chromatographically separated on this column

b

6. Inject 25 μL of sample and separate over 30 min on a TSK-gel ODS-120T column (4.6
250 mm, 5 μm) with C18 guard column with the column compartment set at 16  C. Run an isocratic gradient of 7% water in methanol containing 5 mM ammonium acetate at 1 mL/min (see Note 29). 7. The chromatogram expected after injection of purified oxysterol standards is shown in Fig. 1a (see Notes 30 and 31). Oxysterol profiles of the 420 -> 385 transition for mouse brain, liver, and adrenal gland extracts are shown in Fig. 1b. The high level of 24S-hydroxycholesterol was evident in the brain extract. In contrast, the adrenal gland was the only tissue tested in which 22R-hydroxycholesterol was observed (see Fig, 1). 3.5

Quantification

Oxysterol quantification is done based on the principle of isotope dilution, where a deuterated internal standard or mix of standards of known amounts is spiked into the sample matrix at the beginning of the extraction procedure. The deuterated analogues are expected to behave similarly to their endogenous counterparts and should, in theory, be susceptible to degradation, auto-oxidation, and ionization to the same extent as the unlabeled oxysterols (see Note 32). Addition of the internal standard at the start of the extraction will

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Lilia Magomedova and Carolyn L. Cummins

A

10 μM Standards

x104 +ESI MRM Frag=70.0V CID@** (420.0000 -> 367.0000) 3 16.5

B

5 x10 +ESI MRM Frag=70.0V CID@** (420.0000 -> 385.0000) 14.2 1.2

Brain Extract

1.0

2

0.8 1

0.6

0

0.4

x104 +ESI MRM Frag=70.0V CID@** (420.0000 -> 385.0000) 3 14.1 10.9 2 11.9 1

0.2 20.6 21.3

1.0

Liver Extract

20.7

0.8

0 x104 +ESI MRM Frag=155.0V [email protected] (418.0000 -> 383.0000) 3

0.6 0.4 20.6

2 1

0.2 3

0 x104 +ESI MRM Frag=80.0V [email protected] (402.0000 -> 385.0000) 3 2

0 4 x10 +ESI MRM Frag=70.0V CID@** (420.0000 -> 385.0000) 1.2

x10 +ESI MRM Frag=70.0V CID@** (420.0000 -> 385.0000) Adrenal Extract 3.0 10.9 2.5 20.6 2.0 1.5

12.8

1.0 1

0.5

0 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29

Counts vs. Acquisition Time (min)

7

8

9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25

Counts vs. Acquisition Time (min)

Fig. 1 LC/MS/MS chromatograms of pure standards (1 μL injection, 10 pmol on column) (a) and mouse tissue samples (25 μL injection) (b) extracted from the brain (~100 mg), liver (~100 mg), or pooled adrenal glands (~30 mg). Note that the y-axis is variable in panel B highlighting the relative abundance of different oxysterols in each tissue. Note that the deuterated internal standards were included prior to extraction and saponification but are not shown in this figure

also account for the efficiency of extraction or any losses incurred during sample transfer (see Note 33). 1. Prepare a 6-point external calibration curve in MeOH with concentrations that span an order of magnitude on either side of the expected analyte concentration (see Note 34). Each calibration standard will contain the exact same amount of deuterated internal standard mixture that was added to the unknowns (10 μL). Increasing concentrations of pure unlabeled oxysterols are added to generate a curve that has a constant amount of deuterated internal standard and variable amounts of unlabeled oxysterols. This curve will allow you to determine the limit of detection for each oxysterol on your specific instrument. 2. Prepare a second standard curve using the same matrix as the samples to be analyzed (i.e., plasma). It is critical to include a blank matrix sample that has been spiked with the deuterated internal standards. This will be used to subtract the endogenous background levels of oxysterols for the calibration curve analysis (see Note 35).

LC/MS/MS Analysis of Oxysterols

9

3. Perform the identical extraction of the calibration standards as was done for the unknowns (i.e., include saponification and SPE extraction). 4. Run samples and calibration standards together in the same worklist (see Note 36). 5. Extract the MRM transitions for each of the oxysterols and internal standards (see Note 37). 6. Integrate the area under the curve for the analyte of interest and corresponding deuterated internal standard and calculate the peak area ratio of analyte/internal standard. 7. Generate a regression line for the calibration standards by plotting the ratio of peak area of analyte to the peak area of the corresponding internal standard on the y-axis and the oxysterol concentration on the x-axis. Determine the equation that describes the line of best fit using linear regression (see Note 38). 8. Compare the slopes of the calibration curves prepared in MeOH vs. matrix to determine whether the relative response of the analyte is accounted for by the internal standard (see Note 39). 9. Use the appropriate calibration curve (and the line of best fit) to interpolate the concentration of oxysterols in the samples of interest based on the peak area ratio of analyte/internal standard (see Notes 40 and 41).

4

Notes 1. These tubes can be difficult to source. We use Wheaton 358607 tubes for our experiments. 2. We use a 5810 Eppendorf centrifuge with swinging bucket rotor but any similar piece of equipment that can accommodate test tubes can be used. 3. We use a Techne Sample Concentrator with temperature control. Any similar piece of equipment capable of sample concentration under N2 will work well. 4. The size of the SPE column to use is determined by your application. We find that 100–300 mg is generally suitable for this application which uses between 1 mL and 3 mL for conditioning. The Isolute® Si column is available from Biotage. 5. SPE manifolds such as Supelco Visiprep, Agilent Vac Elut, or others can be used. 6. Common vendors include Avanti Polar Lipids, Sigma-Aldrich, and C/D/N isotopes.

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Lilia Magomedova and Carolyn L. Cummins

7. For example, Agilent 1200 HPLC system coupled to an Agilent 6410 triple quadrupole mass spectrometer. 8. Performing the liquid-liquid extraction at room temperature allows for more rapid phase separation between solvents and increases reproducibility. 9. Plasticizers can leach from plastic tubes and pipet tips when exposed to organic solvents. It is highly recommended to perform as many steps as possible using glass test tubes and glass pipets. We have previously reported the detection of highly ionizable plasticizers derived from microcentrifuge tubes with MRM pairs that could interfere with oxysterol analysis [12]. 10. Add the BHT to the solvent mixture fresh each time a new extraction is initiated. Dispense the organic solvent mixture using a glass pipet. Take care not to immerse the pipet so that the graduated markings do not leach off into the solution as this will happen with repeated exposure to this solvent system. 11. During this step, take care not to dislodge the pellet. If pellet becomes dislodged, centrifuge again for another 10 min. 12. If only free oxysterol levels need to be determined, omit this step. 13. Flush with Ar or N2 for 10 s to prevent non-specific oxidation with air during the incubation. This incubation can be performed in a water bath, sand bath, or other heating blocks that provide uniform heating. 14. To minimize sample loss, use a dedicated Pasteur pipet for each sample. Prepare a holding test tube to keep individual sample pipets separated. 15. Perform the transfer step using the same Pasteur pipet. 16. This will be used in the final step to normalize the oxysterol levels to tissue weight (i.e., reporting as ng oxysterol/mg tissue). 17. When inserting the Pasteur pipet, be sure to depress the rubber bulb while passing through the aqueous layer to prevent liquid (or tissue debris) from entering it. When working with larger tissues, a small debris disk will be formed which can be gently pushed aside with the pipette tip. Release the bulb when the pipet has touched the bottom of the tube to aspirate the organic layer. 18. Prepare fresh from a 3.5 M stock solution. 19. Samples can be warmed to 37  C while under N2 to speed up the drying process. 20. A 14 mL test tube will be sufficient to collect the washes.

LC/MS/MS Analysis of Oxysterols

11

21. Apply vacuum to allow a flow through rate of 1 drop per second. 22. If desired, this fraction can be kept for analysis of less polar metabolites. 23. If not using internal standards for oxysterols, elute with 6 mL. This will remove less cholesterol but prevent loss of oxysterols. 24. If desired, remove any remaining analytes from the column with 2 mL methanol. 25. Samples can be dried under N2 with gentle heating (37  C) and stored at 80  C until resuspended just in time for analysis by LC/MS/MS. 26. Alternatively, samples can be filtered using spin filters. 27. Ensure that no air bubble has been trapped at the base of the glass insert by briefly vortexing the sample prior to loading into the autosampler. 28. The gas temperature for the Agilent 6410 QQQ is normally set to 350  C for ESI mode. However, for oxysterols, we have found that the most common adduct observed is the ammonium adduct [MþNH4]+ which is much better preserved in the source when the gas temperature is low which is why it is set at  175 C for this assay. The loss of one [MþNH4+NH3H2O]+ or two [MþNH4+NH32H2O]+ water molecules is the most common ion loss for oxysterols. Interestingly, 20(S)-hydroxycholesterol is different in that the molecular ion is consistent with [MþNH4+H2O]+. 29. To prevent carryover of more lipophilic sterols into the next run, you can add a post-column cleanup step with 100% methanol and/or extend the run time of the method to allow all late eluting compounds to emerge. While the SPE procedure will have removed the majority of the neutral sterols from the sample, there will be some remaining that may elute at a later time point and could interfere will subsequent runs. 30. If separation of isobaric compounds is not essential for your analysis, other standard C18 columns (i.e., Agilent XDB C18) can be used but will show co-elution of R/S isomers (for 24and 26-hydroxycholesterol) and overlap between peaks for 24and 25-hydroxycholesterol. 31. All oxysterols listed in Table 1 are detectable at 100 fmol on column with these mass spectrometry conditions. 32. Ideally, the deuterated analogue of the oxysterol of interest should be used as an internal standard, if it is commercially available. If the specific oxysterol is not available, a related heavy-labeled analogue of similar structure and retention time should be used. For example, 24(R/S)-hydroxycholesterol-d7

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Lilia Magomedova and Carolyn L. Cummins

or 25-hydroxycholesterol-d6 can be used as a surrogate for 20S-, 22R/S-, and 24 R/S-hydroxycholesterol; and 25 (R/S),26-hydroxycholesterol-d4 could be used as a surrogate for 24,25-epoxycholesterol. 33. Oxysterol extraction efficiency may vary based on the sample matrix and should be assessed for each tissue prior to sample preparation. Ideally, we aim to have >80% extraction efficiency for each analyte. Lower extraction efficiencies might be acceptable if analyte abundance is not limiting and the internal standard exhibits a similar level of extraction such that it appropriately controls for the overall extraction efficiency. 34. For example, if the expected plasma concentration for (25R),26-hydroxycholesterol is 100 ng/mL, prepare standards ranging between 10 and 1000 ng/mL (i.e., 10, 25, 100, 250, 500, 1000 ng/mL). More standards can always be included at the low and high end of the curve to estimate the dynamic range of the instrument. Two additional controls to prepare are (1) 0 ng/mL concentration (containing only the internal standard spiked in) and (2) a true blank (with no internal standard spiked in). 35. To prepare the calibration standards, use control matrix from the same species as the samples to be analyzed. In addition, ensure that all of the calibration standards are made from the same batch of matrix (if required, pool multiple blank samples prior to re-aliquoting to prepare the calibration curve). 36. When setting up the autosampler worklist, a full calibration curve is run before and after each set of samples to monitor for any drift in detector sensitivity. Be sure to include several injections of blanks (i.e., mobile phase 93% MeOH) at the beginning of the run (to prime the system) and after the highest concentration in the calibration curve to check for carryover. If the worklist is long and will be running overnight, add injections of calibration standards every 10 to 15 samples bracketed by 2 blank injections. Although we recommend preparing a full standard curve for each batch of samples, once the full standard curve has been established, it is also acceptable to include two quality controls (a high and low concentration) with new batches of samples (in place of a full standard curve). If the calibration curve prepared in MeOH shows a different slope from that prepared in matrix, the matrix calibration curve should be used to correct for unanticipated matrix effects that are not fully accounted for by the internal standard. 37. The software used for this step will vary depending on the mass spectrometer manufacturer (i.e., in this protocol Agilent’s MassHunter software was used).

LC/MS/MS Analysis of Oxysterols

13

38. Do not forget to account for the amount of endogenous oxysterol present in the blank matrix. This can be done by subtracting the peak area ratio obtained for the blank from all other calibration standards or by including the peak area ratio of the blank at X ¼ 0 and including this point in the linear regression when drawing the line of best fit. 39. Performing the calibration curve in MeOH and sample matrix allows you to determine whether the extraction efficiency is consistent across a wide range of concentrations as the slope of the curve should be similar. Once this is established, future standards can be prepared in MeOH for simplicity. 40. If identical volumes of plasma were extracted between samples and standards and the same injection volume was used, the unknown oxysterol concentrations can be read directly from the standard curve. 41. For tissue levels, convert the concentration of oxysterol measured for the sample into the corresponding amount (i.e., interpolated concentration
150 μL); convert this amount from nmol to ng using the molecular weight of the analyte (i.e., nmol
MW (ng/nmol)); and normalize to the weight of the piece of tissue extracted to obtain a final value in ng oxysterol/mg tissue.

Acknowledgments The authors would like to thank Dr. Jeffrey McDonald for his input on this protocol, the Natural Sciences and Engineering Research Council (RGPIN 03666-14) for supporting research activities in the Cummins lab, and the Canadian Foundation for Innovation for supporting the purchase of the analytical equipment used in this protocol. References 1. Janowski BA, Grogan MJ, Jones SA, Wisely GB, Kliewer SA, Corey EJ, Mangelsdorf DJ (1999) Structural requirements of ligands for the oxysterol liver X receptors LXRalpha and LXRbeta. Proc Natl Acad Sci U S A 96 (1):266–271 2. Zhang Z, Li D, Blanchard DE, Lear SR, Erickson SK, Spencer TA (2001) Key regulatory oxysterols in liver: analysis as delta4-3-ketone derivatives by HPLC and response to physiological perturbations. J Lipid Res 42 (4):649–658 3. Bauman DR, Bitmansour AD, McDonald JG, Thompson BM, Liang G, Russell DW (2009)

25-Hydroxycholesterol secreted by macrophages in response to toll-like receptor activation suppresses immunoglobulin a production. Proc Natl Acad Sci U S A 106 (39):16764–16769. https://doi.org/10. 1073/pnas.0909142106 4. Lund EG, Menke JG, Sparrow CP (2003) Liver X receptor agonists as potential therapeutic agents for dyslipidemia and atherosclerosis. Arterioscler Thromb Vasc Biol 23 (7):1169–1177. https://doi.org/10.1161/ 01.ATV.0000056743.42348.59 5. Umetani M, Domoto H, Gormley AK, Yuhanna IS, Cummins CL, Javitt NB, Korach

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KS, Shaul PW, Mangelsdorf DJ (2007) 27-Hydroxycholesterol is an endogenous SERM that inhibits the cardiovascular effects of estrogen. Nat Med 13(10):1185–1192. https://doi.org/10.1038/nm1641 6. Wang Y, Kumar N, Crumbley C, Griffin PR, Burris TP (2010) A second class of nuclear receptors for oxysterols: regulation of RORalpha and RORgamma activity by 24S-hydroxycholesterol (cerebrosterol). Biochim Biophys Acta 1801(8):917–923. https://doi.org/10.1016/j.bbalip.2010.02. 012 7. Schroepfer GJ Jr (2000) Oxysterols: modulators of cholesterol metabolism and other processes. Physiol Rev 80(1):361–554. https:// doi.org/10.1152/physrev.2000.80.1.361 8. McDonald JG, Smith DD, Stiles AR, Russell DW (2012) A comprehensive method for extraction and quantitative analysis of sterols and secosteroids from human plasma. J Lipid Res 53(7):1399–1409. https://doi.org/10. 1194/jlr.D022285

9. McDonald JG, Thompson BM, McCrum EC, Russell DW (2007) Extraction and analysis of sterols in biological matrices by high performance liquid chromatography electrospray ionization mass spectrometry. Methods Enzymol 432:145–170. https://doi.org/10.1016/ S0076-6879(07)32006-5 10. Lund EG, Diczfalusy U (2003) Quantitation of receptor ligands by mass spectrometry. Methods Enzymol 364:24–37 11. Folch J, Lees M, Sloane Stanley GH (1957) A simple method for the isolation and purification of total lipides from animal tissues. J Biol Chem 226(1):497–509 12. McDonald JG, Cummins CL, Barkley RM, Thompson BM, Lincoln HA (2008) Identification and quantitation of sorbitol-based nuclear clarifying agents extracted from common laboratory and consumer plasticware made of polypropylene. Anal Chem 80 (14):5532–5541. https://doi.org/10.1021/ ac8005632

Chapter 2 A Stable Luciferase Reporter System to Characterize LXR Regulation by Oxysterols and Novel Ligands Samantha A. Hutchinson and James L. Thorne Abstract Nuclear receptors (NRs) are ligand-activated transcription factors. Class 2 NRs, such as the liver X receptor (LXR)α and (LXR)β, are typically retained in the nucleus bound to the DNA in both the presence and absence of ligand. Upon binding ligands including hydroxylated cholesterol, LXR releases corepressor proteins in exchange for coactivators resulting in target gene transcription. Activity of the LXRs therefore depends on a combination of the local ligand concentration(s) and cofactor expression, which itself is a function of cell and tissue type, mutation load, and epigenetic regulation. Cross talk with other transcription factors or signaling pathways can also alter LXR activity. The role that LXR plays in both normal physiology and disease progression is becoming increasingly apparent, and a better understanding of how and when LXR is activated or repressed is pressing biological and clinical questions. The complexity of LXR regulation makes identifying novel ligands and determining LXR activity in new cell types challenging. Generating cell lines that contain a stably integrated luciferase reporter gene with an upstream LXR-dependent promoter provides a quick, cheap, robust, efficient, and high-throughput solution to identify novel ligands and assess ligand activity in new cell types. Transplant of these stable cell culture cell lines as xenografts allows reporter activation to be assessed in vivo. Here we describe the generation of stable LXR reporter cell lines, how to confirm transgene insertion and select single cell clones, as well a method to assess transgene activity in vitro. Key words Nuclear receptors, Liver X receptor, Oxysterols, Luciferase reporter assay

1

Introduction The nuclear receptor (NR) superfamily is an unusual class of transcription factors (TF) as they are regulated not just by cofactors and posttranslational modifications as is the case for most other TFs but also by ligand binding. NRs sense and integrate exogenous, endocrine, or metabolic signals and respond by influencing key cellular pathways such as proliferation, apoptosis, differentiation, or senescence [1]. These regulatory features make NRs exceptional candidates for novel drug design in a range of diseases.

Matthew C. Gage and Ine´s Pineda-Torra (eds.), Lipid-Activated Nuclear Receptors: Methods and Protocols, Methods in Molecular Biology, vol. 1951, https://doi.org/10.1007/978-1-4939-9130-3_2, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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Samantha A. Hutchinson and James L. Thorne

There are two liver X receptor (LXR) isoforms (α and β), and their principal role is to sense changes in cellular requirements for fat and lipid metabolism and enact appropriate transcriptional changes. LXR activation has been implicated in progression or amelioration of several diseases, including cardiovascular disease [2], cancer [3], inflammatory disease [4], and autoimmune pathologies [5]. The LXRs have an overlapping ligand-binding repertoire (typically oxidized cholesterol compounds termed “oxysterols”) but show differences in tissue localization and regulation by endogenous cofactors [6]. Sensitivity of nuclear receptors to their ligands is regulated by cofactors, and disruption of cofactor balance is a common mechanism for disease progression [7, 8]. Importantly, there are selective modulators of LXR which result in different transcriptional outcomes depending on cellular and chromatin context. This ambiguity in ligand function results in divergent roles for LXR in disease. In cancer, for example, oxysterol endogenous LXR ligands have tumor suppressor roles in prostate cancer and melanoma [9, 10] but are pro-metastatic in breast cancer [11, 12]. The context in which LXR activation occurs is therefore an increasingly important clinical research question. Oxysterols are a class of lipids derived through cholesterol hydroxylation by members of the cytochrome P450 family and were identified as the endogenous LXR ligands by Janowski and colleagues in 1996 [13]. An array of exogenous LXR ligands have since been found in nature [14] or developed using pharmacological approaches [15]. In vitro single oxysterols can modulate LXR activity at low micromolar concentrations [16]. When combined the concentration of LXR-activating oxysterols in human serum is in the high nano- to low micromolar range; these measurements in tissues are limited [17–20]. Synthetic ligands such as T0901317, GW3965, and GSK2033 function at nanomolar concentrations with LXR transcriptional activity being altered by as much as 80-fold relative to basal conditions [16, 21, 22]. Selective LXR modulators such as riccardin c or the phytosterols appear to exert their effects over a similar concentration range as the oxysterols [14, 21, 23]. Through exposing cell lines, stably transduced (viral integration) or transfected (non-viral integration) with a reporter gene under the control of an LXR-responsive promoter, to a serial dilution of ligand, LXR activity can be determined. Different ligands and/or different cell types can be compared side by side allowing robust characterization of ligand efficacy and how permissive the cell type is to LXR activation. In this chapter we set out the protocols needed to create a stable reporter cell line utilizing commercially available inactivated lentiviral particles, measure transcriptional activation with the luciferase reporter system, and explain how to validate and select clones. Lentiviral particles which contain the transgene of interest, an antibiotic selection cassette, and

LXR Reporter Activation by Oxysterols

17

replication-incompetent insertion sequences provide a relatively quick and robust route to generating stable reporter cell lines. There are multiple advantages to generating stable reporters: experiments can be run over a longer time period and transplanted for in vivo evaluation of LXR activity. Importantly for delineating the complex LXR regulatory mechanisms, stable reporters can be combined with other types of experiments such as transient knockdown or overexpression of cofactors or regulatory miRNAs. Generating stable reporter lines is more time-consuming and initially expensive (although if hundreds of assays are to be run then stables should again be considered), so the choice of stable or transient experiments should be made. If transient is deemed more appropriate, then the reader is directed to the methods described by Gage et al. [24]. Luciferase reporter assays are well-established tools for evaluating promoter activity. The production of luciferase and its capacity to convert D-luciferin into oxyluciferin are directly proportional to the activation of its promoter. Therefore, a standard plate reader with bioluminescence detection can be used to accurately measure promoter activity using commercially available reagents. We find this assay is a reproducible and cost-effective way to assess LXR activity in response to panels of proposed LXR ligands, to evaluate LXR activity in new cell types, and to determine how LXR activity is altered following experimental perturbation to its regulatory cofactors. The method is transferrable to other NRs (the choice of promoter sequence is the only change required) and works in a broad range of common cell types.

2

Materials Read all protocols and notes before purchasing reagents and ligands. The experiment should be carefully designed and the suitability of stable over transient experiments confirmed before starting. Store commercially bought reagents as directed in product data sheets and all other reagents at room temperature or as otherwise stated. Generation of stably transduced cell lines may be subject to GMO legislation in your region; ensure appropriate guidelines are followed and approvals acquired before commencing work.

2.1 Puromycin Titration Curve

1. Aspiration polystyrene pipette, sterile. 2. Polystyrene serological pipettes, sterile: 25 mL, 10 mL, and 5 mL. 3. Sterile phosphate buffered saline (PBS) 1
: 2.6 mM KCl, 1.47 mM KH2PO4, 137.93 mM NaCl, and 8.06 mM Na2HPO4, pH 7.0–7.3.

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Samantha A. Hutchinson and James L. Thorne

4. Puromycin 25 mg, stored at 20  C. 5. Fetal bovine serum (FBS), stored at 20  C (see Note 1). 6. DMEM-10: Dulbecco modified eagle medium (DMEM) supplemented with 10% FBS, stored at 4  C (see Note 2). 7. Trypsin 1
, sterile. 8. Trypan blue. 9. Inverted microscope. 10. Hemocytometer/cell counter. 11. Single channel pipettes and sterile filter tips. 12. P200 multichannel pipette and sterile filter tips. 13. 15 mL and 50 mL sterile tubes. 14. 96-well culture plates, sterile. 15. Centrifuge capable of spinning at 225
g, for 50 mL tubes. 16. Sterile trough, for use with multichannel pipette. 2.2 Transduction with Lentiviral Particles

1. PBS 1
, sterile. 2. Trypsin 1
, sterile. 3. Puromycin 25 mg, stored at 20  C. 4. Lentiviral particles, stored at 80  C. 5. SureENTRY transduction reagent, stored at 20  C. 6. Distilled water. 7. DMEM-10: DMEM supplemented with 10% FBS, stored at 4  C. 8. DMEM-10P: DMEM supplemented with 10% FBS and 1 μg/ mL puromycin, stored at 4  C. 9. DMEM-10 N: DMEM supplemented with 10% FBS and 0.1 mM NEAA, stored at 4  C. 10. DMEM-10 Np/s: DMEM supplemented with 10% FBS, 0.1 mM NEAA, and 100 U/mL penicillin and 100 μg/mL streptomycin, stored at 4  C. 11. Inverted microscope. 12. Hemocytometer/cell counter. 13. Trypan blue. 14. Centrifuge capable of spinning at 225
g, for 50 mL tubes. 15. 15 mL and 50 mL sterile tubes. 16. Polystyrene aspiration pipettes, sterile. 17. Polystyrene serological pipettes, sterile: 25 mL, 10 mL, and 5 mL. 18. 6-well cell culture plate, sterile.

LXR Reporter Activation by Oxysterols

19

19. 48-well cell culture plates, sterile. 20. Tissue culture flasks (T25 and T75), sterile. 21. Single channel pipettes and sterile filter tips. 22. P200 multichannel pipette and sterile filter tips. 2.3

Luciferase Assay

1. PBS 1
, sterile. 2. Trypsin 1
, sterile. 3. Luciferase reporter assay components including luciferase substrate, luciferase buffer, and lysis buffer (see Note 3). 4. Distilled water. 5. DMEM-10: DMEM supplemented with 10% FBS, stored at 4  C. 6. DMEM-10P: DMEM supplemented with 10% FBS and 1 μg/ mL puromycin, stored at 4  C. 7. Dimethyl sulfoxide (DMSO), grade: Extra pure. 8. Ethanol (ETOH), grade: ACS, 100% ethanol absolute. 9. Ethanol flushed with nitrogen, grade: ACS, 100% ethanol absolute. 10. Stock solution of LXR antagonist GSK2033 at 10 mM in ethanol, grade: ACS, 100% ethanol absolute (see Note 4). 11. Stock solution of LXR agonist T0901317 at 10 mM in DMSO, grade: Extra pure (see Note 4). 12. Stock solution of LXR endogenous ligand 22-OHC at 10 mM in nitrogen-flushed ethanol (see Note 4). 13. 15 mL and 50 mL tubes, sterile. 14. Centrifuge capable of spinning at 225
g, for 15 mL and 50 mL tubes. 15. Inverted microscope. 16. Hemocytometer/cell counter. 17. Trypan blue. 18. Gilson single channel pipettes and sterile filter tips. 19. Gilson P200 multichannel pipette and sterile filter tips. 20. Plate rocker. 21. Luminescence reader (Tecan spark 10 M). 22. 96-well white culture plates, sterile.

2.4

Clone Selection

1. PBS 1
, sterile. 2. Trypsin 1
, sterile. 3. DMEM-10: DMEM supplemented with 10% FBS, stored at 4  C.

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Samantha A. Hutchinson and James L. Thorne

4. 15 mL and 50 mL tubes, sterile. 5. Centrifuge capable of spinning at 225
g, for 15 mL and 50 mL tubes. 6. Inverted microscope. 7. Hemocytometer/cell counter. 8. Trypan blue. 9. 96-well cell culture plates, sterile. 10. Single channel pipettes and sterile filter tips. 11. P200 multichannel pipette and sterile filter tips. 12. Tissue culture flasks (T25 and T75), sterile. 2.5 Storage/Freezing Down Cells

1. Cell culture freezing media: 10% DMSO in 90% FBS (this will vary for different cell lines). 2. Centrifuge capable of spinning at 225
g, for 15 mL and 50 mL tubes. 3. Cryotubes for liquid nitrogen storage. 4. Liquid nitrogen and storage tank.

2.6 Copy Number Analysis: DNA Extraction, Quantification, and qPCR

1. Chloroform (fume hood use only). 2. Ethylenediaminetetraacetic acid (EDTA), 0.5 M. 3. DNA buffer: 1 M Tris–HCL, pH 8.0, 0.5 M EDTA in dH2O. 4. Ethanol 100% and 70%. 5. Isoamyl alcohol (3-methyl-1-butanol). 6. Phenol (stored at 4  C, fume hood use only). 7. PBS 1
, sterile. 8. 100% isopropanol (2-propanol). 9. Proteinase K 20 mg/mL, stored at 20  C. 10. Sodium acetate (NaOAc), 3 M, pH 5.2. 11. Sodium dodecyl sulfate (SDS), 20%. 12. Tris–HCL, 1 M, pH 8.0. 13. Tris acetate/disodium EDTA buffer (TAE buffer), 1
: 40 mM Tris pH 7.6, 20 mM acetic acid, and 1 mM EDTA. 14. Trypsin 1
, sterile, stored at 4  C. 15. Distilled water. 16. Nuclease-free water. 17. qPCR master mix kit. 18. Shaker (e.g., Thermomixer Eppendorf AG 22331, Hamburg). 19. Centrifuge capable of spinning at 225
g, for 15 mL tubes.

LXR Reporter Activation by Oxysterols

21

Table 1 Genomic DNA qPCR gene primer sequences Gene

Strand direction Primer sequence

β-Actin

Forward Reverse

Luciferase ORF Forward Reverse

CAACTCCATCATGAAGTGTGAC CCACACGGAGTACTTGCGCTC GAGATACGCCCTGGTTCCTG GCATACGACGATTCTGTGATTTG

20. Centrifuge capable of spinning at 225
g, 956
g, and 20,817
g for 1.5 mL tubes. 21. 1.5 mL Eppendorf tubes, sterile. 22. Polypropylene 15 mL tubes, sterile. 23. Nucleic acid Nanodrop).

quantification

instrument

(e.g.,

Tecan,

24. qPCR 96-well plates. 25. Optical adhesive films for 96-well qPCR plates. 26. qPCR machine. 27. Primers or TaqMan assays for housekeeping gene and firefly luciferase (see Table 1).

3

Methods Carry out all procedures at room temperature unless otherwise specified. Methods described were optimized for use with the breast cancer cell line MDA.MB.231, and similar conditions were applicable in MDA.MB.468, MCF7, and HepG2 cells. Where cell line-specific optimization of steps may be required, the reader is directed to the relevant notes.

3.1 Puromycin Titration Curve

Key step: It is important to find the lowest puromycin concentration that kills the entire cell culture in the absence of the selection cassette. Puromycin at this concentration, but no higher, should then be applied to transduced cells to select positive clones. 1. Culture MDA.MB.231 cells as described (see Note 5). 2. When cells are 80% confluent, wash with 5 mL sterile PBS 1
using a serological pipette. 3. Discard PBS 1
. 4. Add 3 mL trypsin 1
for a T75 flask, and incubate until cells detach (typically about 3 min). 5. Add 7 mL DMEM-10 to the cells to inactivate the trypsin.

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6. Resuspend cells as a single cell suspension. 7. Using a hemocytometer with trypan blue, or a cell counter with live/dead gating, count cells, and resuspend at 1.5
105 live cells/mL in DMEM-10. 8. Seed the cells at 3
104 cells per well (200 μL from a 1.5
105 cell suspension) into a sterile 96-well tissue culture plate (see Note 6). 9. Place 96-well plates at 37  C in incubator (5% CO2) overnight, and allow cells to attach. 10. The next morning set up the puromycin titration in 50 mL tubes. Set up two tubes containing 50 mL of DMEM-10 and 5 tubes containing 25 mL DMEM-10. Label these as follows: 8 μg/mL and 0 μg/mL (50 mL DMEM-10), and 4 μg/mL, 2 μg/mL, 1 μg/mL, 0.5 μg/mL and 0.25 μg/mL (each with 25 mL DMEM-10). 11. Add 16 μL puromycin (stock concentration 25 mg/mL) to the 8 μg/mL tube, and homogenize well with a Stripette. 12. Perform a twofold serial dilution starting with placing 25 mL from the 8 μg/mL tube to the 4 μg/mL tube. Mix and repeat to generate 2 μg/mL, 1 μg/mL, and 0.5 μg/mL finishing with 0.25 μg/mL. The last tube should contain 50 mL DMEM10 only. 13. Remove cells to be treated from the incubator, visually confirm attachment and that confluency is suitable (see Note 7). 14. Remove the DMEM-10 from the entire 96-well plate, wash with PBS 1
, and add 200 μL of each puromycin-containing media concentration (including the puromycin negative growth media) to eight replicate wells per puromycin concentration. 15. Evaluate cell death daily and apply fresh DMEM-10P at day 3. 16. After 5 days of DMEM-10P, evaluate the cell cultures to determine the lowest puromycin concentration that resulted in 100% cell death. 17. The chosen concentration should be confirmed with fresh serial dilutions of puromycin in separate cell passages before continuing with the transductions and clone selection. 3.2 Cell Transduction with Signal Particles

Perform cell transduction procedure under sterile conditions. When handling lentiviral particles, pay attention to local regulations for handling GMO materials (see Note 8). 1. Begin protocol during exponential phase of cell culture (20 min. 18. Resuspend the DNA dried pellet in 20 μL of sterile H2O, and shake at 3.3
g overnight at 37  C. 19. Quantify DNA and store at 4  C. 3.8 Copy Number qPCR

1. Run qPCR using primers targeted to an endogenous gene with known copy number in your cell lines (many cell lines have non-diploid karyotype) and targeted to the luciferase open reading frame (ORF) with unknown copy number. See Table 1 for primer sequences. 2. Determine copy number by comparing the Ct of the endogenous target (typically two copies) with the Ct of the luciferase ORF. In both the non-transduced cells and the transduced reporter line, the endogenous target should be equal. In the non-transduced cells, the luciferase ORF should not be detected, and in the stable reporter line, the copy number can be calculated by comparing the luciferase ORF with Ct for the endogenous control.

4

Notes 1. Fetal bovine serum (FBS) contains lipids, growth factors, hormones, steroids, and other components that may activate the reporter. Charcoal stripping of FBS removes these factors but may alter growth kinetics of cell lines (e.g., loss of estradiol may impair ER-positive breast cancer cell lines). In the experiments described here, we used non-charcoal stripped FBS to allow evaluation of LXR antagonists and repression of basal reporter activity. The choice of FBS is assay-dependent. 2. There is controversial evidence that glucose stimulates LXR activity [16]. Both this and cell line requirements should be considered when planning these experiments. As such, any potential contribution by glucose to LXR activation should be evaluated in your specific cell line system.

LXR Reporter Activation by Oxysterols

29

3. To analyze luciferase activity, a range of commercial kits are available which are convenient and cost-effective. Typically, they include (a) a cell lysis buffer, (b) a substrate, and (c) a buffer to dissolve the substrate. It may be possible to optimize and reduce buffer volumes to increase the number of assays by as much as 50% that can be performed per kit without loss of sensitivity of detection. This should be assessed when first testing the stably transduced cell cultures. 4. GSK2033 is used at a final concentration of 1 μM from a 10 mM stock diluted in ETOH. T0901317 is used at a final concentration of 1 μM from a 10 mM stock diluted in DMSO. Other agonists may need to be diluted or dissolved in other solvents. For example, oxysterols are highly sensitive to autooxidation and so should be dissolved in nitrogen-flushed ethanol to impair loss of activity. Frequent purchasing of oxysterols in small batches also helps reduce problems with loss of signal caused by nonenzymatic oxidization over time. 5. Culture of MDA.MB.231, MDA.MB.468, and MCF7 cell lines: Passage cells during exponential phase of cell culture ( 5 mL). 7. Add 5 mL MACS™ buffer to column, and immediately elute labelled cells by pushing down on the plunger. 8. Centrifuge cells at 300  g for 10 min at 4  C, aspirate supernatant, and continue to downstream applications (see Note 7). 3.3 Cell Culture and Differentiation

1. Following PBMC isolation and counting, centrifuge cells for 5 min at 350  g at 4  C.

3.3.1 PBMC and Monocyte Cell Culture

2. Discard supernatant, and resuspend cells in PBMC culture medium at a concentration of 2  106 cells/mL (106 cells/ mL for selected or enriched monocytes) (see Note 8). 3. Seed cells in low-attachment plates for immediate use can be maintained in culture for 72 h to 96 h. 4. For differentiation of monocytes to macrophages, see Subheading 3.3.2.

3.3.2 Differentiation to Macrophages (Human Monocyte-Derived Macrophages; HMDM)

1. For differentiating monocytes into macrophages, cells should be seeded into poly-L-lysine (0.1 mg/mL)-coated plates to facilitate cell adherence. 2. Following adherence of cells (2–3 h at 37  C), replace medium with HMDM differentiation medium. 3. Cells differentiate into macrophages in approximately 10 days (differentiation is visually confirmed; cells are larger and exhibit bright protruding membranes with a central pronounced nucleus). Throughout differentiation, change medium every other day (see Note 9).

3.4 Flow Cytometry for Quantitative Analysis of Monocyte Phenotypes

Monocyte phenotyping can be carried out on ~400 μL whole blood. For basic phenotyping of monocytes, a cytometer equipped with 4–5 parameters is sufficient (see Table 1 for suggested panel setup and Fig. 4 for gating strategy). The in-depth application of flow cytometry is beyond the scope of this chapter, and thus for a detailed insight into the setting up of a cytometer, compensation settings, and antibody titration, please refer to Maecker and Trotter [14]. This protocol can be applied to both whole blood and PBMCs (Subheading 3.1), in which large granular cells will be removed by the density gradient, and thus gating will be more straightforward (see Note 10). For carrying out the protocol from

Monocyte and Macrophage Isolation and Analysis

Morphological gating around monocytes

SSC-A

FSC-A Q2

Q4 CD14

Q3

Live haematopoietic cells

Monocytes (Lineage negative)

Viability

Q1

CD14

CD16

FSC-A

Doublet exclusion

SSC-H

FSC-H

SSC-A

Doublet exclusion

43

Lineage

CD45

Q1: Non-classical monocytes Q2: Intermediate monocytes Q3: Classical monocytes Q4: non-monocytes

Fig. 4 Gating strategy to be applied for basic analysis of circulating monocytes in humans. Q1–Q4 quadrants, FSC forward scatter, SSC side scatter, -A area signal, -H height signal, CD cluster of differentiation, CD45 hematopoietic cell marker, CD14 monocyte marker, CD16 monocyte activation marker

PBMCs, following counting starts at step 3. When working with fresh human blood, work under sterile conditions, in a cell culture cabinet dedicated to human material; work rapidly and on ice, unless otherwise stated. 1. After homogenizing whole-blood tubes by repeated pipetting, transfer 400 μL of blood to a FACS tube, and add 4 mL of red blood cell lysis buffer (create a tube of pooled cells from all samples to have a final volume of 400 μL; this tube will be used for unstained cells) (see Note 11). 2. Cap tube and seal with Parafilm, agitate samples horizontally at room temperature for 10 min, and then centrifuge at 350  g for 5 min at 4  C. 3. Discard supernatant, pulse vortex cell pellet, and resuspend in FcR blocking reagent (diluted 1:100 in FACS buffer); incubate on the bench at room temperature for 10 min. 4. Add 2 mL of FACS buffer and centrifuge at 350  g for 5 min at 4  C. 5. Prepare antibody cocktail in FACS buffer, including viability dye, and maintain on ice. Calculate a minimum staining volume of 50 μL per sample. From this point, protect antibodies and samples from light.

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6. Discard supernatant following the centrifugation (step 4), pulse vortex cell pellet and resuspend cells in antibody cocktail, incubate in the dark at 4  C for 30–45 min, and resuspend unstained cells in 50 μL FACS buffer. 7. Add 2 mL of FACS buffer and centrifuge at 350  g for 5 min at 4  C. 8. Resuspend cells in 400 μL FACS buffer, and filter through a 40 μm filter into a new FACS tubes. 9. Cap tubes and keep at 4  C in the dark until acquisition within 4 h (although not recommended, acquisition can be carried out later if cells are fixed) (see Note 12). 10. Acquisition may also be carried out on a cell sorter, which will allow sorting cells for downstream culture applications or for RNA/protein analysis. When sorting for RNA and protein analysis, sorting cells directly into lysis buffer is recommended. 3.5 Adipose Tissue Macrophage Isolation

Adipose tissue biopsies are often kept in PBS following excision; once the biopsy is obtained, the tissue must be weighed (as results are most commonly reported as a function of tissue weight, i.e., cells/g) and treated immediately for downstream applications. In this digestion protocol, care is taken to be able to isolate both the stromal-vascular fraction (containing immune cells, endothelial cells, etc.) and the adipocyte fraction (see Note 13). At all times that tissue is fresh, work under sterile conditions, in a cell culture cabinet dedicated to human material. 1. Warm collagenase digestion solution to 37  C in a water bath. 2. Place adipose tissue in a small dish, and cut into small pieces a few millimeters thick. 3. Transfer adipose tissue to a 15 mL centrifuge tube, and fill tube up to the 8 mL mark with warmed collagenase digestion solution; this solution is in excess for a biopsy of under 5 g (scale up if needed). 4. Cap tube and seal with Parafilm, shake vigorously by hand for a few seconds, and then transfer to a shaking water bath or a shaking chamber maintained at 37  C. Allow to digest for 25–30 min. 5. Remove tube from shaker, shake vigorously by hand for a few seconds, and pass the lysate through a 200 μm cell strainer into a 50 mL Falcon tube. 6. Allow to stand at room temperature, adipocytes will float to the top within 5–10 min, and lower phase will become more transparent. Remove adipocytes with a plastic Pasteur pipette to continue macrophage isolation (if adipocytes will be kept for analysis, see Note 14).

Monocyte and Macrophage Isolation and Analysis

45

7. Adjust volume of the lower phase to 25 mL with PBS þ 2% (w/v) BSA; cap and centrifuge at 350  g for 5 min at room temperature. 8. Discard supernatant, and resuspend pellet in red blood cell lysis buffer adjusting volume to 10 mL; incubate at room temperature for 3 min. 9. Pass the suspension through a 70 μm cell strainer into a new 50 mL centrifuge tube; centrifuge at 350  g for 5 min at 4  C. 10. Discard supernatant and resuspend pellet in 5 mL of PBS; this is the stromal-vascular fraction of adipose tissue; count cells (see Note 15). 11. To analyze macrophage polarization, apply flow cytometric staining protocol (Subheading 3.4 from step 3 onward), with the use of a 6-parameter cytometer to differentiate M1 and M2 macrophages. The panel described in Table 2 and Fig. 5 will

FSC-H

SSC-A

Doublet exclusion

SSC-H

Doublet exclusion

Morphological gating

FSC-A

FSC-A

SSC-A Haematopoietic cells

Macrophages

Lineage

FSC-A

CD11c

Viability

Viability

CD68

Live lineage negative cells

Q1

Q2

Q4 CD206

Q3

CD45

Q1: M1 macrophages Q2: M1-2 macrophages Q3: M2 macrophages Q4: M0 macrophages

Fig. 5 Gating strategy to be applied for basic analysis of adipose tissue macrophages in humans. Q1–Q4 quadrants, FSC forward scatter, SSC side scatter, -A area signal, -H height signal, CD cluster of differentiation, CD45 hematopoietic cell marker, CD68 macrophage marker, CD11c M1 macrophage marker, CD206 M2 macrophage marker

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allow quantification of total immune cells, macrophages, and polarized macrophages from adipose tissue.

4

Notes 1. For most applications, venous blood can be collected into EDTA or heparin-coated tubes; however, do check specifications of any secondary analysis kits for anticoagulant compatibility. 2. The use of an automated cell counter is recommended, such as a slide reader or benchtop cytometer (e.g., Countess II, Invitrogen; MACSQuant, Miltenyi Biotec). With such instruments consistent gating can be used across all samples to count cells of a desired size. A hemocytometer can also be used. Viability dyes should be included when counting. 3. The use of CD14 microbeads or classical monocyte isolation kit: Both reagents/kits are antibody based; while CD14 microbeads will allow isolation of all monocytes, the CD14 receptor is targeted with an antibody and thus will no longer be functional. The classical monocyte isolation kit is based on negative selection; therefore classical monocytes remain untargeted; thus all classical monocyte receptors are actionable in downstream applications. However the classical monocyte isolation kit will not allow isolation of intermediate nor nonclassical monocytes, due to their co-expression of CD16, a negative lineage marker targeted by the classical monocyte isolation kit. Both approaches in mind, CD14 microbeads are better suited to RNA or protein analysis of the total monocyte population without downstream experiments targeting monocyte function. The classical monocyte isolation kit is more suited to in vitro approaches that target monocyte activation or functional properties. 4. (Optional) Wash cells by adding 2–5 mL of MACS buffer per 1 mL of whole blood, and centrifuge at 445  g for 10 min at room temperature in a swinging bucket rotor without brake. Aspirate supernatant carefully. Do not disturb the cell pellet. Leave a residual volume of supernatant (approximately 1–2 mm in height) to avoid cell loss. Resuspend cell pellet by adding MACS buffer to a total volume of 1 mL. This wash step will allow removing any residual unbound antibodies and will allow reducing the probability of contaminating cells being sorted. 5. Here the protocol for LS columns is applied; if using MS columns, adapt 3 mL volumes to 500 μL and 5 mL volume to 1 mL. The use of MS columns is suitable when starting biological material is limiting.

Monocyte and Macrophage Isolation and Analysis

47

6. When using CD14 microbeads, the positive labelled fraction that is retained in the magnetic field is the monocytecontaining fraction. When using the classical monocyte isolation kit, the negative fraction that flows through the column while in the separator’s magnetic field is the fraction containing classical monocytes. 7. If RNA analysis is to be carried out, repeat centrifugation wash step using only PBS in order to remove any residual BSA which may interfere with RNA isolation. 8. Throughout the culture procedure, aimed to achieve confluence of monocytes, enriched samples will have a higher proportion of monocytes, and thus the total number of cells must be tittered down when samples have been enriched by the use of a gradient. 9. Add 1.5 the habitual volume of medium to allow cultivation over the weekend. 10. Following Ficoll-Paque gradient, morphological gating (first window, Fig. 4) will not represent the topmost population of cells. These cells are eliminated in the gradient due to their high granularity/poly-lobed nuclei. 11. Unstained cells are important when setting detection voltages of the morphological gate; the placement of an appropriate voltage of the photomultipliers for the forward and side scatter detectors is essential to having the representative morphological plot from blood (first window, Fig. 4). 12. Fixation is carried out by resuspending cells in ethanol- or formalin-based fixative and then washing cells with FACS buffer centrifugation before acquisition. Cell fixation is not recommended as this may distort cell morphology and invalidate the viability dye used. 13. When working with adipose tissue, do not work on ice until the adipocyte fraction has been removed. This is because adipocytes tend to solidify at cold temperatures (as any lipid-rich milieu). 14. If adipocytes are kept for analysis, they must be washed in successive rounds of PBS. This is carried out by adding 10 mL of PBS to adipocytes, capping and gently shaking the tube and then allowing adipocytes to float to the top. Repeat this washing step two more times, recover adipocytes with a plastic Pasteur pipette, snap freeze, or continue to downstream analysis. 15. Further purification of macrophages can be carried out through positive or negative selection strategies described above (Subheading 3.2).

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References 1. Kraakman MJ, Murphy AJ, Jandeleit-Dahm K, Kammoun HL (2014) Macrophage polarization in obesity and type 2 diabetes: weighing down our understanding of macrophage function? Front Immunol 5:470 2. Bensinger SJ, Tontonoz P (2008) Integration of metabolism and inflammation by lipidactivated nuclear receptors. Nature 454 (7203):470–477 3. Allen JN, Dey A, Nissly R, Fraser J, Yu S, Balandaram G, Peters JM, Hankey-Giblin PA (2017) Isolation, characterization, and purification of macrophages from tissues affected by obesity-related inflammation. J Vis Exp (122). https://doi.org/10.3791/55445 4. ImmGen C (2016) Open-source ImmGen: mononuclear phagocytes. Nat Immunol 17 (7):741 5. El-Sahrigy SA, Mohamed NA, Talkhan HA, Rahman AM (2015) Comparison between magnetic activated cell sorted monocytes and monocyte adherence techniques for in vitro generation of immature dendritic cells: an Egyptian trial. Cent Eur J Immunol 40 (1):18–24 6. Davies LC, Taylor PR (2015) Tissue-resident macrophages: then and now. Immunology 144 (4):541–548 7. Bertani FR, Mozetic P, Fioramonti M, Iuliani M, Ribelli G, Pantano F, Santini D, Tonini G, Trombetta M, Businaro L, Selci S, Rainer A (2017) Classification of M1/M2polarized human macrophages by label-free hyperspectral reflectance confocal microscopy and multivariate analysis. Sci Rep 7(1):8965

8. Guilliams M, Ginhoux F, Jakubzick C, Naik SH, Onai N, Schraml BU, Segura E, Tussiwand R, Yona S (2014) Dendritic cells, monocytes and macrophages: a unified nomenclature based on ontogeny. Nat Rev Immunol 14(8):571–578 9. Cignarella A, Tedesco S, Cappellari R, Fadini GP (2018) The continuum of monocyte phenotypes: experimental evidence and prognostic utility in assessing cardiovascular risk. J Leukoc Biol https://doi.org/10.1002/JLB. 5RU1217-477RR. [Epub ahead of print] 10. Ginhoux F, Guilliams M (2016) Tissueresident macrophage ontogeny and homeostasis. Immunity 44(3):439–449 11. Boutens L, Stienstra R (2016) Adipose tissue macrophages: going off track during obesity. Diabetologia 59(5):879–894 12. de Jong AJ, Pollastro S, Kwekkeboom JC, Andersen SN, Dorjee AL, Bakker AM, Alzaid F, Soprani A, Nelissen R, Mullers JB, Venteclef N, de Vries N, Kloppenburg M, Toes REM, Ioan-Facsinay A (2018) Functional and phenotypical analysis of IL-6-secreting CD4 (þ) T cells in human adipose tissue. Eur J Immunol 48(3):471–481 13. Ouchi N, Parker JL, Lugus JJ, Walsh K (2011) Adipokines in inflammation and metabolic disease. Nat Rev Immunol 11(2):85–97 14. Maecker HT, Trotter J (2006) Flow cytometry controls, instrument setup, and the determination of positivity. Cytometry A 69 (9):1037–1042

Chapter 4 Glucan-Encapsulated siRNA Particles (GeRPs) for Specific Gene Silencing in Adipose Tissue Macrophages Emelie Barreby, Andre´ Sulen, and Myriam Aouadi Abstract Macrophages are cells of the immune system that have been suggested as important regulators of wholebody metabolism in mammals. In obesity, adipose tissue macrophages (ATMs) are thought to play both a detrimental and a beneficial role in the regulation of insulin sensitivity. Here, we describe a protocol to prepare and administer glucan-encapsulated RNAi particles (GeRPs), for specific delivery of siRNA and subsequent gene silencing in ATMs in obese mice. Using the GeRP technology to silence genes provides a unique method to study the function of factors expressed by ATMs in the regulation of metabolism. Key words RNAi, Delivery, Insulin sensitivity, Macrophages, Adipose tissue, Gene silencing, Obesity, Diabetes, Inflammation, Innate immunity

1

Introduction Adipose tissue (AT) is an endocrine organ involved in the regulation of whole-body metabolism. In obesity, AT dysfunction, including a reduced ability to store excess nutrients in forms of fat, is associated with metabolic diseases such as insulin resistance (IR) and type 2 diabetes (T2D) [1]. The AT is a heterogeneous tissue composed of fat-storing cells, adipocytes, and the stromalvascular fraction containing progenitors, endothelial cells, and immune cells. Macrophages are the main immune cell type in the AT, and their abundance increases with obesity and IR in animals and humans [2, 3]. ATMs have been shown to produce factors that are both detrimental and beneficial for the regulation of insulin sensitivity [4, 5]. The in vivo relevance of these ATM-expressed factors and their influence on whole-body metabolism are often investigated by the use of transgenic animal models, either complete knockout models or myeloid-specific knockouts using the Cre-Lox recombinase system. However, these approaches are not specific to ATMs, and results cannot be delineated from

Matthew C. Gage and Ine´s Pineda-Torra (eds.), Lipid-Activated Nuclear Receptors: Methods and Protocols, Methods in Molecular Biology, vol. 1951, https://doi.org/10.1007/978-1-4939-9130-3_4, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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macrophages in other tissues. A solution to this non-specificity is the use of glucan-encapsulated siRNA particles (GeRPs) that can silence gene expression in phagocytic cells in a tissue-specific manner [6–9]. Tissue specificity of GeRP-mediated gene silencing depends on the route of administration; intraperitoneal and intravenous administration results in specific gene silencing in ATMs and liver macrophages, respectively [8, 9]. This technology has been employed to elucidate several interesting aspects of ATMs in the regulation of metabolism during obesity. Gene silencing of ATM-derived inflammatory cytokines, tumor necrosis factor alpha (Tnfa) and osteopontin (Opn/Spp1), improves glucose tolerance during obesity by decreasing inflammation [8]. Conversely, silencing of lipoprotein lipase (LPL) worsens glucose tolerance by reducing the ability of macrophages to store excess lipids in the AT [10]. Here we provide a detailed protocol for the process of preparing GeRPs for ATM-specific gene silencing (Fig. 1). This includes the generation of glucan shells (GS) from Baker’s yeast (Saccharomyces cerevisiae), labeling of the GS with fluorescein isothiocyanate (FITC) for fluorescent detection in tissues and cells, loading the particles with siRNA and finally in vivo delivery.

Fig. 1 Generation of fluorescent GeRPs. GeRPs are generated from Baker’s yeast by a series of alkaline and solvent extractions to remove the cell wall and other cell components, yielding porous 2–4 μm hollow β1, 3-D-glucan particles (scheme of particles, top; procedure, bottom; microscopy of particles). Empty β1, 3-D-glucan particles are then loaded with complexes of Dy547-labeled siRNA (red) and Endo-Porter

Gene Silencing in Adipose Tissue Macrophages

2

51

Materials

2.1 Preparation of Glucan Shells

1. SAF-Mannan yeast (S. cerevisiae). 2. 0.5 M sodium hydroxide solution: Add 20 g of NaOH to 1 L of deionized water. Prepare fresh. 3. Distilled water. 4. Isopropanol. 5. Acetone. 6. Erlenmeyer flask (2 L). 7. Magnetic stirring bar. 8. Hot plate with magnetic stirrer. 9. Thermometer. 10. High-speed centrifuge. 11. 250 mL centrifuge bottles. 12. 1.5 mL Eppendorf tubes. 13. Spatula. 14. Brinkman Polytron homogenizer or equivalent. 15. Fume hood. 16. 200 μM mesh filter. 17. Electric coffee grinder. 18. Hemocytometer.

2.2 Fluorescent Labeling of Glucan Shells

1. Glucan shells. 2. Sodium carbonate buffer pH 9.2: Prepare a 0.1 M solution of sodium carbonate and a 0.1 M solution of sodium bicarbonate. To obtain 500 mL of buffer, mix 50 mL of 0.1 M sodium carbonate with 450 mL of 0.1 M sodium bicarbonate. Store at RT. 3. Fluorescein isothiocyanate (FITC) solution: 1 mg/mL FITC in absolute ethanol. 4. Absolute ethanol. 5. Filtered water. 6. Sterile water. 7. Analytical balance. 8. Erlenmeyer flask (1 L). 9. 50 mL centrifuge tubes. 10. Probe sonicator. 11. Vortex mixer. 12. Lyophilizer.

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2.3 Loading of Glucan Shells with siRNA and Delivery to Adipose Tissue Macrophages In Vivo

1. FITC-labeled GS. 2. 1 mM siRNA: dilute siRNA stock to 1 mM using 1 siRNA buffer (Dharmacon). 3. 5 mM Endo-Porter (EP; Gene Tools): Prepare a 5 solution of the 1 mM EP stock by placing the uncapped tube in a speed vacuum on high heat for approximately 3 h until completely dry. Resuspend the pellet in 200 μL RNase-free water, and heat again at 45  C for 15 min to solubilize the dried Endo-porter (see Note 1). 4. 30 mM sodium acetate buffer: Prepare 250 mL buffer, and adjust pH to 4.8 using glacial acetic acid. Sterile filter. 5. Filters (22 μm pore size). 6. PBS. 7. 1 mL syringe. 8. Vortex mixer. 9. Probe sonicator.

3

Methods Perform the preparation of glucan shells in a fume hood.

3.1 Preparation of Glucan Shells

1. Resuspend 100 g SAF-Mannan yeast in 1 L 0.5 M NaOH solution in a 2 L Erlenmeyer flask with a stir bar. Heat the solution to 80–85  C for 1 h while stirring to maintain suspension. Make sure to maintain the temperature range by inserting a thermometer in the flask. Caution: hot caustic. 2. Leave the solution to cool to 40–50  C. Then transfer the suspension into 6  250 mL centrifuge bottles. Centrifuge using a high-speed centrifuge at 15,000  g for 10 min (see Note 2). 3. Decant off the supernatant, and resuspend the pellet in deionized water (1 L in total) using a homogenizer at 5000 rpm (see Note 3). 4. Once the pellet has dissolved, transfer the solution back to a clean 2 L flask. 5. Add 20 g of NaOH to make a 0.5 M solution. 6. Heat to 80–85  C for 1 h while stirring to maintain suspension. Make sure to maintain temperature range by inserting a thermometer in the flask. 7. Leave the solution to cool to 40–50  C.

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8. Transfer the suspension into the 250 mL centrifuge bottles. Centrifuge at 15,000  g for 10 min. Decant off the supernatant. 9. Wash the pellets first in sterile water by adding 200 mL deionized water to each centrifuge tube. Use the homogenizer to resuspend the pellets. 10. Centrifuge at 22,000  g for 15 min and decant the supernatant. 11. Repeat the wash step with water until the supernatant is clear (see Note 4). 12. Wash the pellets with isopropanol by adding 200 mL isopropanol to each centrifuge tube. Use the homogenizer to resuspend the pellets. 13. Centrifuge at 15,000  g for 15 min, and decant the supernatant (see Note 5). 14. Continue to wash the pellets with isopropanol until the supernatant is clear (see Note 4). 15. Wash the pellets in acetone by adding 200 mL acetone to each of the six tubes. Use the homogenizer to resuspend the pellets. 16. Centrifuge at 15,000  g for 20 min and decant the supernatant. Repeat the wash with acetone twice. 17. Centrifuge the suspension at 15,000  g for 30 min to collect the glucan shells. Decant the supernatant. 18. Break and mix the packed pellet using a spatula, and allow the acetone to evaporate off under a fume hood overnight. 19. Grind the resulting pellet in a coffee grinder into a fine powder. 20. Filter the powder through a 200 μm mesh filter, and transfer into tubes for storage at 20  C. 21. The yield can be determined with the use of a hemocytometer (see Note 6). 22. Measurement of hydrodynamic volume: to measure the volume that can be absorbed by the GS, weigh 10 mg of glucan shells in a 1.5 mL Eppendorf tube. Record the weight of the tube containing the GS, and then resuspend the GS in 400 μL water by vortexing. Allow to incubate at RT for 1 h; vortex every 10 min. Thereafter, pellet the GS by centrifugation at 9000  g for 10 min, remove excess water, and weigh the tube again. The hydrodynamic volume is the difference in weight before and after the addition of water divided by the weight of the dry GS (mg water/mg GP) and should be around 20–30 μL per mg of GS.

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3.2 Fluorescent Labeling of Glucan Shells

Glucan shells can be labeled with different fluorophores; here we describe the process of how to label them with FITC (see Note 7). 1. Wash glucan shells with sodium carbonate buffer by adding 100 mL of sodium carbonate buffer to 1 g of GS in an Erlenmeyer flask. 2. Centrifuge at 3000  g for 10 min. 3. Remove the supernatant, and resuspend the particles in fresh 100 mL of sodium carbonate buffer. 4. Add 10 mL of FITC solution to the buffered glucan shell suspension. 5. Stir at room temperature for ~16 h protected from light. 6. Transfer the solution to 50 mL centrifuge tubes. 7. Centrifuge at 4700  g for 5 min. 8. Wash with 40 mL filtered water per tube. Resuspend the particles by thoroughly vortexing the tubes, and then sonicate until solution is homogenous. 9. Centrifuge at 4700  g for 5 min and remove the supernatant. 10. Repeat the wash until the supernatant is clear (see Note 8). 11. Wash the particles with 40 mL absolute ethanol. 12. Rehydrate the particles by resuspending the pellet in 40 mL sterile water inside a tissue culture hood. 13. Centrifuge at 4700  g for 10 min and remove the supernatant. 14. Add 10 mL sterile water in the hood. 15. Vortex thoroughly and sonicate until solution is homogenous. 16. Freeze the particles in liquid nitrogen and dry using a lyophilizer. 17. Store at

3.3 Loading of Glucan Shells with siRNA and Delivery to Adipose Tissue Macrophages In Vivo

20  C protected from light.

1. Incubate acetate buffer, siRNA, and Endo-porter at RT for 15 min according to the volumes described in Table 1 (see Notes 9–12). 2. Add the siRNA/EP solution from step 1 to the FITC-labeled glucan shells according to the ratio in Table 1. 3. Carefully vortex to mix and incubate at RT for 1 h in the dark. 4. Add PBS (see volume in Table 1), and sonicate for 5  10 s at 30% amplitude. 5. Administer the glucan-encapsulated RNAi particles by daily intraperitoneal (ip) injections for 5–10 days with a volume of 200 μL. Vortex the suspension before each injection (see Notes

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Table 1 Reagents for loading glucan shells with siRNA. The following amounts are for the preparation of GeRPs for 5  200 μL injections For 1 dose

Volume (μL)

1 mM siRNA

3 nmol

3

5 EP

50 nmol

10

Glucan shells

1 mg

–

30 mM acetate buffer, pH 4.8

–

13

PBS

–

974

Fig. 2 Intraperitoneally delivered FITC-labeled GeRPs localize in epididymal adipose tissue without targeting macrophages in other tissues including the heart, liver, kidney, lung, or spleen. After administration of GeRPs, tissues were collected, fixed, paraffin embedded, and sectioned before H&E staining. The fluorescence of the FITC-GeRPs was evaluated on unstained sections. All images were acquired with a Zeiss Observer Z1 microscope. White arrows indicate location of GeRPs

13 and 14). Injecting obese mice ip will result in an accumulation of GeRPs specifically in the adipose tissue (Fig. 2). 6. Mice are sacrificed, and tissues harvested 24 to 48 h after the last injection.

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Notes 1. EP is an amphipathic peptide composed of leucine and histidine residues used to bind the siRNA to be loaded into glucan shells (Fig. 1). 2. We use a Beckman Avanti JXN-30 with a 250 mL flask rotor. 3. When using the homogenizer, the solution can foam, so make sure to start with 1 L of water for the washing steps so that you know how much water is added to the pellets. 4. The supernatant usually becomes clear after three to five washes. 5. Optional stopping point: The protocol can be stopped after the first wash with isopropanol, and bottles containing the glucan shell pellet can be left at 4  C overnight. 6. With a starting amount of 100 g of yeast, the final product should be around 6–8 g GS with 1  109 GS per mg. 7. The same protocol can be used to label with other fluorophores, e.g., rhodamine. 8. The supernatant usually becomes clear after five to eight washes. 9. Depending on the efficiency of the sequence, the amount of siRNA used per mg of GS can be varied from 1 to 5 nM. The efficiency is determined by performing a dose response. 10. The volume of acetate buffer used should be equal to the volume of siRNA and EP combined and needs to be adjusted if the concentration of siRNA is altered. 11. It is possible to load the GS with multiple siRNAs as long as the final concentration does not exceed the maximum amount recommended (5 nM per mg of GS). 12. Prepare GeRPs for one extra mouse for every five mice since small amounts will be lost each time when loading the syringes for injection. 13. The duration of the treatment depends on the target. Gene silencing is generally observed between 5 and 10 daily intraperitoneal injections with 200 μL GeRPs. 14. GeRPs can be stored at 4  C for up to 1 week if needed. For longer period of times, aliquot the volume necessary for one daily dose, flash freeze, and store at 20  C. GeRPs cannot be stored at 20  C without flash freezing first; otherwise they form aggregates that are difficult to break down.

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Acknowledgment We would like to thank Tarja Schro¨der for the tissue histology and Kjell Hultenby for the electron microscopy. This work was supported by research grants from AstraZeneca through the ICMC (M.A.), the Swedish Research Council (M.A.: 2015-03582), and the Novo Nordisk Foundation, including the Tripartite Immunometabolism Consortium (M.A. and TrIC; NNF15CC0018486) and the Strategic Research Program in Diabetes at Karolinska Institutet (M.A.). References 1. Kahn SE, Hull RL, Utzschneider KM (2006) Mechanisms linking obesity to insulin resistance and type 2 diabetes. Nature 444 (7121):840–846. https://doi.org/10.1038/ nature05482 2. Lumeng CN, Bodzin JL, Saltiel AR (2007) Obesity induces a phenotypic switch in adipose tissue macrophage polarization. J Clin Invest 117(1):175–184. https://doi.org/10.1172/ JCI29881 3. Wu D, Molofsky AB, Liang HE, RicardoGonzalez RR, Jouihan HA, Bando JK, Chawla A, Locksley RM (2011) Eosinophils sustain adipose alternatively activated macrophages associated with glucose homeostasis. Science 332(6026):243–247. https://doi. org/10.1126/science.1201475 4. Weisberg SP, McCann D, Desai M, Rosenbaum M, Leibel RL, Ferrante AW Jr (2003) Obesity is associated with macrophage accumulation in adipose tissue. J Clin Invest 112(12):1796–1808. https://doi.org/10. 1172/JCI19246 5. Xu H, Barnes GT, Yang Q, Tan G, Yang D, Chou CJ, Sole J, Nichols A, Ross JS, Tartaglia LA, Chen H (2003) Chronic inflammation in fat plays a crucial role in the development of obesity-related insulin resistance. J Clin Invest 112(12):1821–1830. https://doi.org/10. 1172/JCI19451 6. Aouadi M, Tesz GJ, Nicoloro SM, Wang M, Chouinard M, Soto E, Ostroff GR, Czech MP (2009) Orally delivered siRNA targeting

macrophage map 4k4 suppresses systemic inflammation. Nature 458(7242):1180–1184. https://doi.org/10.1038/nature07774 7. Tesz GJ, Aouadi M, Prot M, Nicoloro SM, Boutet E, Amano SU, Goller A, Wang M, Guo CA, Salomon WE, Virbasius JV, Baum RA, O’Connor MJ Jr, Soto E, Ostroff GR, Czech MP (2011) Glucan particles for selective delivery of siRNA to phagocytic cells in mice. Biochem J 436(2):351–362. https://doi.org/ 10.1042/BJ20110352 8. Aouadi M, Tencerova M, Vangala P, Yawe JC, Nicoloro SM, Amano SU, Cohen JL, Czech MP (2013) Gene silencing in adipose tissue macrophages regulates whole-body metabolism in obese mice. Proc Natl Acad Sci U S A 110(20):8278–8283. https://doi.org/10. 1073/pnas.1300492110 9. Tencerova M, Aouadi M, Vangala P, Nicoloro SM, Yawe JC, Cohen JL, Shen Y, GarciaMenendez L, Pedersen DJ, Gallagher-DorvalK, Perugini RA, Gupta OT, Czech MP (2015) Activated Kupffer cells inhibit insulin sensitivity in obese mice. FASEB J 29(7):2959–2969. https://doi.org/10.1096/fj.15-270496 10. Aouadi M, Vangala P, Yawe JC, Tencerova M, Nicoloro SM, Cohen JL, Shen Y, Czech MP (2014) Lipid storage by adipose tissue macrophages regulates systemic glucose tolerance. Am J Physiol Endocrinol Metab 307(4): E374–E383. https://doi.org/10.1152/ ajpendo.00187.2014

Chapter 5 Isolation and Purification of Tissue Resident Macrophages for the Analysis of Nuclear Receptor Activity Laura Alonso-Herranz, Jesu´s Porcuna, and Mercedes Ricote Abstract Tissue resident macrophages (TRMs) are multifunctional immune cells present in all tissues, contributing to the correct development, homeostasis, and protection against pathogens and injury. TRMs are morphologically and functionally heterogeneous, as a result of both the diversity of tissue environments in which they reside and their complex origin. Furthermore, some specific TRM populations are controlled by nuclear receptors. A widely used method for studying the role of nuclear receptors in immune cells is flow cytometry. Although flow cytometry is extensively used in tissues such as the peripheral blood, lymph nodes, peritoneal cavity, and bone marrow, there is a need for protocols for the study TRMs in solid tissues. In this chapter, we describe a comprehensive protocol for obtaining single-cell suspensions of resident macrophages from the pleural cavity, heart, lung, spleen, and kidney, and we present detailed gating strategies for the study of nuclear receptor activity in different TRM subsets within these tissues. Key words Tissue macrophages, Heart, Pleural cavity, Kidney, Spleen, Lung, Nuclear receptors, Flow cytometry

1

Introduction ´ lie Metchnikoff in 1892 The term macrophage was first coined by E to identify cells involved in the phagocytosis process during inflammation. Macrophages are immune cells that play roles in many specific processes in order to maintain tissue homeostasis [1]. Tissue resident macrophages (TRM) are multifunctional, specialized, and heterogeneous cells found in most mammalian tissues. They act as sentinel cells or as an immune barrier against pathogens, protecting against homeostatic imbalance and tissue damage. However, TRMs have many other functions required not only in adulthood but also during development; moreover, many of these functions vary from one tissue to another [2, 3]. Macrophages were for many years

Laura Alonso-Herranz and Jesu´s Porcuna contributed equally to this work. Matthew C. Gage and Ine´s Pineda-Torra (eds.), Lipid-Activated Nuclear Receptors: Methods and Protocols, Methods in Molecular Biology, vol. 1951, https://doi.org/10.1007/978-1-4939-9130-3_5, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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thought to always originate from adult bone marrow; however, lately it has become evident that most adult TRMs originate during embryonic development and not from circulating monocytes [2, 4–6]. The contribution of circulating monocytes to TRM populations is restricted to a few specific tissues, including the gut, the skin, and the heart, or during inflammation and injury [4, 7, 8]. In adulthood, most TRMs are replaced by local selfrenewal independently of bone marrow-derived precursors [4, 9, 10]. Some TRM subtypes are regulated by specific types of nuclear receptors (NRs) [11–14]. NRs are ligand-dependent TFs that regulate diverse aspects of development and homeostasis [15–17]. Important NRs involved in TRM regulation include LXRα in marginal zone splenic macrophages [11], PPARγ in alveolar macrophages [13], and RAR in peritoneal macrophages [12]. The complex transcriptional control of TMR subsets creates a need for powerful tools allowing scientists to distinguish among the many different populations. Flow cytometry is a powerful and established tool in the study of the immune system (e.g., immunophenotyping of peripheral blood cells, analysis of apoptosis, and detection of cytokines) [18, 19]. Flow cytometry is based on the light scattering properties of the cells under investigation. Light scattering at different angles can distinguish differences in size and internal complexity; in addition, light emitted from fluorescently labeled antibodies can identify a great variety of cell surface and cytoplasmic antigens [20]. These properties make flow cytometry a powerful tool for the rapid and detailed analysis of complex populations. State-ofthe-art flow cytometers are able to analyze up to millions of cells, recognizing and differentiating, depending on the system, more than ten fluorochromes, and thus allowing the precise identification of several populations in a single sample. Flow cytometry techniques are improving continually. Moreover, the combination of time-of-flight flow cytometry (CyTOF) detectors and sophisticated algorithms (tSNE) is helping researchers to identify new subsets within already known TRM populations, ensuring an unbiased study of the cells. In this chapter, we describe how to harvest, digest (if needed), and prepare single-cell suspensions of purified TRM samples from the pleural cavity, heart, lung, spleen, and kidney. These protocols generate high yields of leukocytes, and specifically macrophages, from the indicated tissues. Furthermore, the staining antibodies we propose here are optimized to give excellent fluorescence resolution, allowing the identification of diverse TRM subsets within each tissue. The protocol described here is intended for the study of the contribution of NRs to TRM development, identity, and function. The protocol can be used to characterize TRMs from wild-type and

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NR knockout mice in order to analyze the role of the NR under study. This protocol can also be used to obtain pure TRM populations by cell sorting for further analysis, such as genome-wide studies (RNA-Seq, ATAC-Seq, ChIP-Seq, GRO-seq), qPCR, and ligand-based activation of NRs in cell culture.

2

Materials 1. Adult (8–12 weeks old) C57BL/6 mice. 2. Collagenase type IV from Clostridium histolyticum. 3. Hybridization oven with rocker. 4. 70% ethanol in distilled water. 5. Sterile surgical tools: tweezers and scissors. 6. 1 PBS, cold (around 4  C). 7. 1, 2, and 10 mL syringes. 8. 18, 21, and 25 G needles. 9. 1.5, 15, and 50 mL polypropylene conical test tubes. 10. CO2 chamber. 11. 1.5 mL test tubes. 12. Red blood cell lysis buffer: 4.13 g NH4Cl, 0.5 g KHCO3, 100 μL EDTA, 500 mL distilled H2O. 13. Portable electric pipette controller. 14. Sterile 10, 200, and 1000 plastic tips. 15. Compensation beads. 16. Pipettes. 17. 10 mL glass pipettes. 18. Vortex. 19. Precision balance. 20. Neubauer chamber. 21. Trypan blue. 22. Centrifuge. 23. Bright-field microscope. 24. Aluminum foil. 25. Tape. 26. FACS buffer: 1 PBS, 1% inactive fetal bovine serum (FBS), and 5 mM EDTA. 27. Heart and kidney digestion buffer: 1 PBS with 100 μg/mL collagenase IV. 28. Lung digestion buffer: 1 PBS with 10 μg/mL collagenase IV. 29. Cytometer tubes.

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Table 1 Proposed panel of antibodies for the gating strategies described in Figs. 1–5. For each antibody, we indicate the clone we use and the concentration. Note that the concentration needs to be adjusted according to the fluorescence intensity of the fluorochrome selected for each clone Antibody

Clone

Concentration (μg/mL)

Pleural macrophages

CD45-APC-Cy7 B220-PerCPCy5.5 CD11b-AF647 CD115-PE F4/80-PECy7 MHCII-BV605

30-F11 RA3-6B2 M1/70 AFS98 BM8 M5/114.15.2

2 2 2 2 2 0.67

Cardiac macrophages

CD45-PerCPCy5.5 CD11b-PECy7 F4/80-PE Ly6C-APC MHCII-BV605 CCR2-APC

30-F11 M1/70 BM8 AL-21 M5/114.15.2 475,301

2 2 6 4 0.67 5

Alveolar macrophages

CD45-PerCPCy5.5 CD11b-PECy7 CD11c-FITC SiglecF-PE

30-F11 M1/70 N418 E50-2440

2 2 5 4

Splenic macrophages

CD45-APC-Cy7 CD11b-AF647 F4/80-PECy7 MHCII-BV605

30-F11 M1/70 BM8 M5/114.15.2

2 2 2 0.67

Kidney macrophages

CD45-APC-Cy7 CD11b-AF647 F4/80-PECy7 Tim 4-PE

30-F12 M1/71 BM9 RMT4-54

2 2 2 2

30. Round-bottom 96-well plate. 31. Antibodies for flow cytometry (see Table 1). 32. 70 and 100 μm nylon cell strainers. 33. 70 μm conical filters. 34. Flow cytometer/cell sorter.

3

Methods

3.1 Setting Up Tools and Reagents in the Lab

1. Set up the hybridization oven at 37  C. 2. Set up the centrifuge at 4  C. 3. Thaw collagenase type IV from

20  C to 4  C.

4. Remove the red blood cell lysis buffer from the fridge, and place it at room temperature.

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3.2 Mouse Sacrifice and Tissue Harvesting

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1. Euthanize mice according to the relevant regulatory laws for animal experimentation. 2. Place the mouse on its back. Clean the chest and abdomen by soaking all the fur with 70% ethanol. 3. Using tweezers and scissors, open the animal (see Note 1). First, make a small incision in the skin with the scissors. With the scissors near-closed, introduce them under the skin, and then open them up. Remove the scissors, and repeat this process until you have completely separated the skin from the peritoneum and ribs. After this, gently pull the fur back to expose the inner skin lining the peritoneal cavity and chest (see Note 2). 4. Cap a 2 mL syringe with a 25 G needle, and insert the needle between the fifth and sixth rib of the mouse (see Note 3). Introduce 2 mL of cold 1 PBS into the pleural cavity. 5. Using a 2 mL syringe capped with a 20 G needle, recover as much pleural lavage as possible. We suggest injecting the needle through the diaphragm. It is extremely important not to damage the lungs. 6. Transfer the pleural lavage into a 15 mL test tube, and fill it to 10 mL with cold 1 PBS. 7. Use the scissors to open the chest under the sternum, and cut the ribs to expose the heart. Cut the right atrium, and, using a 10 mL syringe capped with a 25 G needle, gently perfuse the animal through the apex with 20 mL of cold 1 PBS (see Note 4). 8. Repeat step 7, but this time perfuse through the right ventricle. 9. Dissect the atria from the ventricles and discard. 10. Transfer the ventricles to 1.5 mL tubes containing 1 mL cold 1 PBS. 11. Collect the lungs. Transfer the multilobular part to 1.5 mL tubes with cold 1 PBS. 12. Cut the splenic vessels and remove the spleen with the help of tweezers. 13. Transfer the spleen to a 1.5 mL tube with cold 1 PBS. 14. Next, cut the renal vessels, and harvest the kidneys in cold 1 PBS in 1.5 mL tubes. 15. Weigh the solid tissues on a precision balance (see Note 5).

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3.3 Preparation of Single-Cell Suspension from the Pleural Cavity (For Pleural Resident Macrophages)

1. Transfer the pleural lavage to a 15 mL test tube, and fill up to 10 mL with cold 1 PBS. 2. Centrifuge for 5 min at 500  g and discard the supernatant. 3. If blood is visible, resuspend the pellet with a vortex in 300 μL of red blood cell lysis buffer for 3 min. Then wash with 30 mL of 1 PBS (see Notes 6 and 7). 4. Centrifuge for 5 min at 500  g and discard the supernatant. Keep samples on ice until ready to proceed to Subheading 3.8.

3.4 Preparation of Single-Cell Suspension from Cardiac Tissue (For Cardiac Resident Macrophages)

1. Place the heart in one 1.5 mL tube with 1 mL of digestion buffer (see Note 8). 2. Finely chop the heart into small pieces with small sterile scissors (see Notes 9–11). 3. To digest the heart, place the sample in the hybridization oven, and incubate at 37  C for 45 min with gentle shaking (30 rpm). Hold the samples in place by sticking them to the oven surface with tape (see Note 11). 4. Mechanically homogenize the digested tissue with up and down motions through a 1 mL syringe capped with an 18 G needle. Work on ice from this point on (see Note 12). 5. To remove the tissue stroma, transfer the digested sample to a 50 mL tube through a 100 μm nylon cell strainer. Gently press the digested tissue against the cell strainer, and wash it with about 30 mL of 1 PBS (see Note 13). 6. Centrifuge for 10 min at 500  g at 4  C and discard the supernatant. 7. To eliminate erythrocytes, resuspend the pellet by vortexing in 600 μL of red blood cell lysis buffer. Incubate for 3 min at room temperature (see Note 14). 8. Wash with 10 mL FACS buffer, centrifuge for 5 min at 500  g at 4  C, and discard the supernatant. Keep samples on ice until ready to proceed to Subheading 3.8.

3.5 Preparation of Single-Cell Suspension from the Lung (For Alveolar Macrophages)

1. Place the lung sample in one 1.5 mL tube with 1 mL of lung digestion buffer (see Note 8). 2. Finely chop the lung sample into small pieces with small sterile scissors (see Notes 9–11). 3. To digest the lung tissue, place the sample in the hybridization oven, and incubate at 37  C for 30 min with gentle shaking (30 rpm). Hold the samples in place by sticking them to the oven surface with tape (see Note 11). 4. Mechanically homogenize the digested tissue with up and down motions through a 1 mL syringe capped with an 18 G needle. Work on ice from this point on (see Note 12).

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5. To remove the tissue stroma, transfer the digested sample to a 50 mL tube through a 70 μm nylon cell strainer. Gently press the digested tissue against the cell strainer, and wash it with about 30 mL of 1 PBS (see Note 13). 6. Centrifuge for 5 min at 500  g at 4  C and discard the supernatant. 7. To eliminate erythrocytes, resuspend the pellet by vortexing in 500 μL of red blood cell lysis buffer. Incubate for 3 min at room temperature (see Note 14). 8. Wash with 10 mL FAC buffer, centrifuge for 5 min at 500  g at 4  C, and discard the supernatant. Keep samples on ice until ready to proceed to Subheading 3.8. 3.6 Preparation of Single-Cell Suspension from the Spleen (For Spleen Resident Macrophages)

1. Place the spleen on a pre-wetted (with 1 PBS) 100 μm nylon filter over an open 50 mL tube. Squeeze the spleen against the filter with the plunger of a 2 mL syringe. 2. Pass 50 mL of 1 PBS through the filter. 3. Centrifuge for 5 min at 500  g at 4  C and discard the supernatant. 4. Resuspend the pellet by vortexing in 500 μL of red blood cell lysis buffer for 5 min at room temperature. Then wash with 30 mL of 1 PBS. 5. Centrifuge for 5 min at 500  g and discard the supernatant. Keep samples on ice until ready to proceed to Subheading 3.8.

3.7 Preparation of Single-Cell Suspension from the Kidney (For Kidney Resident Macrophages)

1. Cut the kidney into two halves with sterile scissors. 2. Select the half to be processed, and weigh it in the precision balance (see Note 15). 3. Finely mince the tissue in 500 μL digestion buffer in a 1.5 mL tube. Ensure that no visible tissue pieces remain (see Notes 9–11). 4. To digest the half kidney, place it in the hybridization oven, and incubate at 37  C for 45 min with gentle shaking (30 rpm). Hold the samples in place by sticking them to the oven surface with tape (see Note 16). 5. Transfer the digested tissue to a 50 mL test tube through a 100 μm nylon cell strainer. Ensure you recover all the sample by washing the digestion tube with up to 30 mL of 1 PBS. 6. Centrifuge for 5 min at 500  g and discard the supernatant 7. Resuspend the pellet by vortexing in 500 μL of red blood cell lysis buffer for 3 min at room temperature. Wash with 30 mL of 1 PBS. 8. Centrifuge for 5 min at 500  g and discard the supernatant. Keep samples on ice until ready to proceed to Subheading 3.8.

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3.8 Staining of TRMs for Flow Cytometry 3.8.1 Cell Counting

The following steps are the same for all five tissues.

1. Mix 10 μL of the cell suspension with 10 μL Trypan blue. 2. Count the viable cells in a Neubauer chamber under a brightfield microscope fitted with a 10  objective. 3. Centrifuge the cell suspension for 5 min at 500  g and discard the supernatant.

3.8.2 Staining of Pleural, Cardiac, Alveolar, Spleen, and Kidney Resident Macrophages for Flow Cytometry

You will need to use the entire cell suspension for the pleural cavity and heart. For spleen resident macrophages, use 5  106 cells; for alveolar macrophages, use 1/7 of the sample; and for kidney resident macrophages, use 2/5 of the sample. 1. Block the cells by adding 100 μL of anti-CD16/CD32 Ab (1:100) in FACS buffer per sample. 2. Incubate for 10 to 15 min at 4  C. 3. For staining, transfer the samples to a round-bottom 96-well plate or cytometer tubes (see Notes 17 and 18). 4. Wash with FACS buffer: 200 μL for 96-well plate or 1 mL for cytometer tubes. 5. Centrifuge for 5 min at 500  g at 4  C, and discard the supernatant. 6. Use compensation beads to establish the cytometer settings according to your staining panel. Put one drop of compensation beads in empty wells of the p96-well plate or in fresh cytometer tubes (one for each fluorochrome used). Add one tested antibody to one well or tube containing compensation beads; the antibody should have the same concentration you will use in the staining panel. From this point, the compensation beads and experimental samples must follow the same steps of incubation, washing, and centrifugation. 7. Add 50 μL (96-well plate) or 100 μL (cytometer tube) of antibody mix per sample. Incubate for 30 min at 4  C with gentle shacking (30 rpm). Protect samples from light with aluminum foil (see Notes 19 and 20). 8. Propose antibody panels (see Table 1). 9. Transfer samples stained in 96-well plates to cytometer tubes. Wash with 1 mL FACS buffer, and centrifuge for 5 min at 500  g at 4  C, and discard the supernatant. 10. Resuspend in 200–300 μL FACS buffer. 11. Filter the samples through 70 μm conical filters, and transfer them to fresh cytometer tubes. 12. Place the samples on ice and go to the cytometer.

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1. Use a small fraction of one stained sample to adjust the voltage settings. Remember to do this for each tissue analyzed. 2. Run the compensation beads to create the compensation matrix. Remember to do this for every antibody panel. 3. Run the negative and fluorescence minus one (FMO) controls to create the layout of the gating strategy for every tissue. 4. Run your stained samples, and acquire the data using the specifications of your particular cytometer. 5. Export the data for later analysis with the software of your choice.

3.8.4 Flow Cytometry Analysis of Pleural Resident Macrophages

Within the single viable cell population, we gate leukocytes (CD45+ cells). We then exclude B cells by gating only B220 cells (see Note 21). Macrophages are gated within the CD11b+ population. We can then differentiate two macrophage populations based on their levels of F4/80 and MHCII expression: large pleural macrophages (LPMs) are F4/80highMHCII /low cells, and small pleural macrophages (SPMs) are defined as F4/80-/lowMHCIIhigh cells (see Fig. 1).

Fig. 1 Example gating strategy for pleural resident macrophages

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Fig. 2 Example gating strategy for cardiac resident macrophages

3.8.5 Flow Cytometry Analysis of Cardiac Resident Macrophages

Within the single viable cell population, we gate leukocytes (CD45+ cells). We then select macrophages (CD11b+F4/80+ cells) and differentiate two cardiac macrophage populations based on their level of MHCII expression: MHCIIhighLy6Clow macrophages and MHCIIlowLy6Clow macrophages (see Fig. 2). We can also identify monocytes as Ly6Chi cells.

3.8.6 Flow Cytometry Analysis of Alveolar Macrophages

Within the single viable cell population, we gate leukocytes (CD45+ cells). We then select Ly6G /CD11b+ cells to exclude neutrophils. Alveolar macrophages are distinguished as CD11chighSiglecFhigh cells (see Fig. 3).

3.8.7 Flow Cytometry Analysis of Spleen Resident Macrophages

Within the single viable cell population, we gate leukocytes (CD45+ cells). We then select myeloid cells as CD11b+ cells. Two macrophage subsets can be differentiated according to the expression of F4/80 and Tim-4: Tim4+ macrophages and red pulp macrophages that are F4/80+ (see Fig. 4).

3.8.8 Flow Cytometry Analysis of Kidney Resident Macrophages

Within the single viable cell population, we gate leukocytes (CD45+ cells). We can then differentiate two myeloid subsets according to the expression levels of CD11b and F4/80: CD11blowF480high (kidney resident macrophages) and CD11bhighF480low (see Fig. 5).

Isolation of Tissue Resident Macrophages

Fig. 3 Example gating strategy for alveolar macrophages

Fig. 4 Example gating strategy for spleen resident macrophages

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Fig. 5 Example gating strategy for kidney resident macrophages

4

Notes 1. We recommend use blunt-ended scissors. 2. If you also intend to do a peritoneal lavage, be careful not to break the peritoneum. 3. It is extremely important to introduce only the needle tip; otherwise, the lungs can be damaged, and the pleural lavage will be contaminated with red blood cells and circulating monocytes. A sign that the pleural inflation is proceeding well is movement of the liver toward the intestine. 4. It is important to perfuse the mouse well in order to eliminate erythrocytes and circulating cells as much as possible. This avoids having a mix of TRMs and circulating myeloid cells during the flow cytometry analysis. 5. Weighing the tissues allows calculation of TRM abundance (TRMs/mg of tissue). 6. Sometimes, blood may be visible due to the injection or inflation. We recommend lysing red blood cells to ensure correct labeling in subsequent steps. 7. If the lungs are damaged during lavage, we strongly recommend discarding the sample because of the risk of contamination by circulating leukocytes. 8. We strongly recommend that you fine-tune the conditions for your own lab, varying digestion times by 10 min among tests,

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trying different collagenase concentrations, and even using digestion buffers different from those proposed here, for example, Hanks’ balanced salt solution with Ca2+. Be careful with collagenase incubation times; exposures longer than 1 h will dramatically decrease the yield of viable cells. 9. Tissue pieces must be small enough to pass through an 18 G needle. 10. If you have more than one sample, keep the already minced tissues on ice while mincing the rest. 11. Thorough mincing with shaking is very important to ensure proper cell extraction. 12. The combination of enzyme digestion and mechanical tissue disruption improves TRM yield during digestion and isolation. 13. We recommend the use of the soft end of a 2 mL syringe plunger to squeeze the sample residues. 14. Although the mouse is perfused through the heart, erythrocyte lysis is required afterward to eliminate residual red blood cells. Note that before the lysis, the pellet is still red, and after the lysis it should be completely white. 15. Maintain the other half in 1 PBS on ice; you many need it in case something goes wrong. 16. We strongly recommend shaking the tissue by hand every 5 min to increase the cell yield. 17. When several stainings are required, we recommend dividing the sample among different wells or tubes. Nevertheless, avoid this practice when studying rare populations. 18. When testing a new antibody, it is advisable to use fractions of the cell sample for a negative control (50,000 cells) and a fluorescence minus one (FMO) control (50,000 cells). We recommend that these fractions are taken from the sample with the highest cell number. In the negative control, cells are not labeled with antibody, which allows the negative and the positive populations to be discerned. In the FMO control, the sample is stained with all the antibodies except the one being measured, revealing false-positive signals due to the effect of the other fluorochromes. In both controls, the cells must be processed with the same protocol as the experimental samples. 19. You can adjust the incubation time to your needs (minimum 15 min, maximum 2 h). 20. It is very important to protect antibodies from light in order to prevent bleaching of conjugated fluorochromes. 21. Macrophages are highly autofluorescent; as a result, the population of F4/80+ cells in the B220 gate seems to be partly B220+.

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Acknowledgment We thank Simon Bartlett for English editing. This work was supported by grants from the Spanish Ministry of Economy and Competitiveness (SAF2015-64287R, SAF2017-90604-REDT), Fundacio´ Marato´ TV3 (121931), and the Community of Madrid (B2017/BMD-3684) to M.R. L.A-H. is funded by a fellowship from Obra Social “La Caixa”. The CNIC is supported by the MEIC and the Pro CNIC Foundation and is a Severo Ochoa Center of Excellence awardee (MEIC award SEV-2015-0505). References 1. Varol C, Mildner A, Jung S (2015) Macrophages: development and tissue specialization. Annu Rev Immunol 33:643–675 2. Davies LC, Jenkins SJ, Allen JE, Taylor PR (2013) Tissue-resident macrophages. Nat Immunol 14:986–995 3. Gautier EL, Shay T, Miller J, Greter M, Jakubzick C, Ivanov S, Helft J, Chow A, Elpek KG, Gordonov S, Mazloom AR, Ma’ayan A, Chua WJ, Hansen TH, Turley SJ, Merad M, Randolph GJ (2012) Geneexpression profiles and transcriptional regulatory pathways that underlie the identity and diversity of mouse tissue macrophages. Nat Immunol 13:1118–1128 4. Ginhoux F, Guilliams M (2016) Tissueresident macrophage ontogeny and homeostasis. Immunity 44:439–449 5. Gomez Perdiguero E, Klapproth K, Schulz C, Busch K, Azzoni E, Crozet L, Garner H, Trouillet C, de Bruijn MF, Geissmann F, Rodewald HR (2015) Tissue-resident macrophages originate from yolk-sac-derived erythromyeloid progenitors. Nature 518:547–551 6. Ginhoux F, Greter M, Leboeuf M, Nandi S, See P, Gokhan S, Mehler MF, Conway SJ, Ng LG, Stanley ER, Samokhvalov IM, Merad M (2010) Fate mapping analysis reveals that adult microglia derive from primitive macrophages. Science 330:841–845 7. Epelman S, Lavine KJ, Beaudin AE, Sojka DK, Carrero JA, Calderon B, Brija T, Gautier EL, Ivanov S, Satpathy AT, Schilling JD, Schwendener R, Sergin I, Razani B, Forsberg EC, Yokoyama WM, Unanue ER, Colonna M, Randolph GJ, Mann DL (2014) Embryonic and adult-derived resident cardiac macrophages are maintained through distinct mechanisms at steady state and during inflammation. Immunity 40:91–104 8. Lavine KJ, Epelman S, Uchida K, Weber KJ, Nichols CG, Schilling JD, Ornitz DM,

Randolph GJ, Mann DL (2014) Distinct macrophage lineages contribute to disparate patterns of cardiac recovery and remodeling in the neonatal and adult heart. Proc Natl Acad Sci U S A 111:16029–16034 9. Guilliams M, Scott CL (2017) Does niche competition determine the origin of tissueresident macrophages? Nat Rev Immunol 17:451–460 10. Hashimoto D, Chow A, Noizat C, Teo P, Beasley MB, Leboeuf M, Becker CD, See P, Price J, Lucas D, Greter M, Mortha A, Boyer SW, Forsberg EC, Tanaka M, van Rooijen N, Garcı´a-Sastre A, Stanley ER, Ginhoux F, Frenette PS, Merad M (2013) Tissue resident macrophages self-maintain locally throughout adult life with minimal contribution from circulating monocytes. Immunity 38:792–804. https:// doi.org/10.1016/j.immuni.2013.04.004 11. N AG, Guillen JA, Gallardo G, Diaz M, de la Rosa JV, Hernandez IH, Casanova-Acebes M, Lopez F, Tabraue C, Beceiro S, Hong C, Lara PC, Andujar M, Arai S, Miyazaki T, Li S, Corbi AL, Tontonoz P, Hidalgo A, Castrillo A (2013) The nuclear receptor LXRalpha controls the functional specialization of splenic macrophages. Nat Immunol 14:831–839 12. Okabe Y, Medzhitov R (2014) Tissue-specific signals control reversible program of localization and functional polarization of macrophages. Cell 157:832–844 13. Schneider C, Nobs SP, Kurrer M, Rehrauer H, Thiele C, Kopf M (2014) Induction of the nuclear receptor PPAR-gamma by the cytokine GM-CSF is critical for the differentiation of fetal monocytes into alveolar macrophages. Nat Immunol 15:1026–1037 14. Lavin Y, Winter D, Blecher-Gonen R, David E, Keren-Shaul H, Merad M, Jung S, Amit I (2014) Tissue-resident macrophage enhancer landscapes are shaped by the local microenvironment. Cell 159:1312–1326

Isolation of Tissue Resident Macrophages 15. Roszer T, Menendez-Gutierrez MP, Cedenilla M, Ricote M (2013) Retinoid X receptors in macrophage biology. Trends Endocrinol Metab 24:460–468 16. Valledor AF, Ricote M (2004) Nuclear receptor signaling in macrophages. Biochem Pharmacol 67:201–212 17. Rosenfeld MG, VV L, Glass CK (2006) Sensors and signals: a coactivator/corepressor/epigenetic code for integrating signal-dependent programs of transcriptional response. Genes Dev 20:1405–1428 18. Herzenberg LA, Tung J, Moore WA, Herzenberg LA, Parks DR (2006) Interpreting flow

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cytometry data: a guide for the perplexed. Nat Immunol 7:681 19. Walter W, Alonso-Herranz L, Trappetti V, Crespo I, Ibberson M, Cedenilla M, Karaszewska A, Nunez V, Xenarios I, Arroyo AG, Sanchez-Cabo F, Ricote M (2018) Deciphering the dynamic transcriptional and posttranscriptional networks of macrophages in the healthy heart and after myocardial injury. Cell Rep 23:622–636 20. Adan A, Alizada G, Kiraz Y, Baran Y, Nalbant A (2017) Flow cytometry: basic principles and applications. Crit Rev Biotechnol 37:163–176

Chapter 6 Bone Marrow-Derived Macrophage Immortalization of LXR Nuclear Receptor-Deficient Cells Ana Ramo´n-Va´zquez, Juan Vladimir de la Rosa, Carlos Tabraue, and Antonio Castrillo Abstract Macrophages are professional phagocytic cells that play key roles in innate and adaptive immunity, metabolism, and tissue homeostasis. Lipid metabolism is tightly controlled at the transcriptional level, and one of the key players of this regulation in macrophages and other cell types is the LXR subfamily of nuclear receptors (LXRα and LXRβ). The use of LXR double knockout (LXR-DKO) macrophages in vitro has yielded extensive benefits in metabolism research, but this technique is hindered by primary macrophage cell expansion capability, which diminishes along terminal cell differentiation process. Here we detail a method to immortalize LXR double knockout bone marrow-derived macrophage cells at an early stage of differentiation, using a retroviral delivery of a combination of murine v-myc and v-raf oncogenes. This methodology enables the generation of autonomous self-renewing macrophages bearing an LXR-DKO genetic background, as a valuable tool for research in lipid metabolism and other LXR receptor-mediated effects. Key words Bone marrow-derived macrophages, Immortalization, J2 retrovirus, LXR nuclear receptors, GM-CSF

1

Introduction Macrophages are professional phagocytic cells that take part in pivotal immune processes such as pathogen clearance and cytokine secretion, but they also perform other crucial functions like regulation of metabolism and maintenance of tissue homeostasis. Some of these metabolic-related functions, exemplified by fatty acid synthesis and cholesterol metabolism, are performed through transcriptional regulation exerted by the liver X receptors: LXRα and LXRβ. These are members of the nuclear receptor superfamily that share around 77% of sequence similarity, both in their DNA and ligand binding domains (DBD and LBD) [1]. Despite their highly similar

Matthew C. Gage and Ine´s Pineda-Torra (eds.), Lipid-Activated Nuclear Receptors: Methods and Protocols, Methods in Molecular Biology, vol. 1951, https://doi.org/10.1007/978-1-4939-9130-3_6, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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sequence, the expression of LXR proteins differs greatly: while LXRα is expressed in tissues with a metabolic function (liver, adipose tissue, intestine, kidney, and tissue macrophages), LXRβ is found ubiquitously. Their implication in cholesterol and fatty acid metabolism has been demonstrated by the fact that LXR nuclear receptors control the transcription of several genes whose protein products are involved in different aspects of metabolic processes [2–4]. Additionally, LXRs participate in other transcriptional aspects of macrophage biology, such as control of inflammatory pathways [5], apoptotic cell clearance [6], limitation of adaptive immune responses [7], macrophage cell survival [8, 9], and development of macrophage population in spleen [10], among other functions. An important drawback associated to LXR double knockout genetic background is that, upon aging, male mice experience a severe and progressive lipid accumulation in their caput epididymides, causing epithelium disruption, abnormal tubule morphology, and occlusion, leading ultimately to production of fragile spermatozoa and premature infertility [11]. In addition, within the framework of the principles of the 3Rs to reduce the number of mice in experimentation emerges the need to find alternative methods to comply with ethical procedures in science. Therefore, using this technique to immortalize murine bone marrow-derived macrophages will prospectively prevent the sacrifice of a larger number of rodents. Several retroviral-based immortalization methods have been largely applied to establish autonomous self-renewing macrophage and dendritic cell lines from diverse tissue origins [12–14]. The method described here takes advantage of the integration of two murine oncogenes (v-raf and v-myc) in a single retroviral vector, engineered in 1985 (pHWJ-2) [15], achieving a cooperative, and therefore more efficient, cell transformation by both oncogenes. A helper virus-free system has also been established: a packaging cell line derived from NIH/3T3 fibroblastic cells stably transformed with a Moloney murine leukemia virus strain (leuk M-MuLV) [15] constituting the J2 retrovirus-producing cells. Primary murine monocytic cells immortalized with J2 retrovirus have been extensively analyzed for homogeneity and identity after the immortalization process, having demonstrated to preserve conventional surface macrophage markers and characteristic cellspecific functions [12, 13]. As a consequence, immortalized macrophages can be used to model cell responses under homeostatic and inflammatory settings, as to allow the study of LXR-dependent functions with an impact on macrophage biology. This tailoring process for cell line generation represents an advantage over other commercially available macrophage cell lines like RAW 264.7, which most clones lack or display a practically undetectable LXRα gene expression, hampering the possibility to conduct experiments

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in this cell line related to this LXR nuclear receptor unless previous gene transduction is performed [8]. As a final remark, due to their intrinsically active proliferative ability, cell cycle and signaling pathways related to cell division processes cannot be assayed in these immortalized cells, i.e., tumor growth studies, constituting the only relevant caveat worth pointing out [16, 17]. In this chapter we describe the method used to immortalize most effectively bone marrow-derived macrophages (BMDMs), bearing an LXR-DKO genetic background, at an early differentiation stage, based on the retroviral transduction of two murine oncogenes. Also described are the retrovirus-producing cell maintenance and retroviral suspension preparation.

2 2.1

Materials J2 Culture

1. J2-producing cells [15] (see Note 1). 2. Complete medium: DMEM (Dulbecco’s Modified Eagle Medium) high glucose, containing L-glutamine, 10% FBS, 1% sterile-filtered penicillin, and streptomycin (10 mg/mL). 3. Treated cell culture dishes (100  20 mm). 4. Syringe filter units of 0.45 μm pore size. 5. Sterile conical tubes (50 mL). 6. Needleless sterile syringe (10 mL). 7. Bleach or other disinfectant. 8. Optical microscope with 10 and 20 magnifications.

2.2

BMDM Isolation

1. 70% ethanol. 2. Paper towels. 3. Sterile tweezers and scissors (at least 2 pairs of each). 4. Sterile conical tubes (50 mL). 5. PBS 1. 6. Treated cell culture dishes (100  20 mm). 7. Sterile syringes (10 mL). 8. 25 G needles. 9. Cell strainers (70 μm pore size). 10. Red blood cell lysis buffer 1 (RBC buffer): 0.15 M NH4Cl (8.29 g/L), 0.01 M KHCO3 (1 g/L), 0.01 M EDTA (0.038 g/L), pH 7.4. 11. GM-CSF- or M-CSF-conditioned medium (see Note 2). 12. Hexadimethrine bromide. 13. Dimethyl sulfoxide, DMSO, pure.

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14. Cryogenic vials. 15. Cell freezing medium: 90% FBS and 10% DMSO. 16. 0.5% trypsin-EDTA in PBS. 17. Centrifuge. 18. Cell scrapers. 19. Sterile polystyrene pipettes (10 mL, 5 mL). 20. Pipette controller. 21. Single-channel manual pipette (1 mL, 200 μL, 10 μL). 22. Optical microscope with 10 and 20 magnifications. 23. Cell incubator with programmable control of CO2 and temperature. 2.3 BMDM Immortalization

1. Immortalization medium 1: 85% J2 supernatant (see Subheading 3.1), 5% FBS, 10% GM-CSF- or M-CSF-conditioned medium (or complete medium, supplemented with the recombinant murine GM-CSF or M-CSF cytokine (see Note 2)), in a total volume of 30 mL. Add hexadimethrine bromide last and only before use, to a final concentration of 10 μg/mL, and mix well. Prepare immortalization medium freshly before use, and store it at 4  C when not in use. 2. Immortalization medium 2: 20% J2 supernatant (see Subheading 3.1), 15% GM-CSF- or M-CSF-conditioned medium and fill to 30 mL with complete DMEM medium. Add hexadimethrine bromide last and only before use, to a final concentration of 10 μg/mL, and mix well. Prepare immortalization medium freshly before use, and store it at 4  C when not in use. 3. Treated cell culture dishes (100  20 mm). 4. Sterile conical tubes (50 mL). 5. PBS 1. 6. Sterile polystyrene pipettes (10 mL, 5 mL). 7. Pipette controller. 8. Optical microscope with 10 and 20 magnifications.

3

Methods

3.1 Culture and Maintenance of J2 Retrovirus-Producing Cells

1. Thaw cryogenic vial containing J2-producing cells (see Note 1) at room temperature, and proceed immediately to step 2. 2. Transfer the total volume of the cryogenic vial to a tube containing 9 mL of complete DMEM medium. Spin the tube 5 min at 1000  g to pellet cells.

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3. Resuspend cell pellet in 1 mL of complete DMEM medium, and add 9 mL more. Plate the total volume in a 100  20 mm treated culture dish. 4. Maintain cells until confluence in an incubator under standard cell culture conditions at 37  C and 5% CO2. Check cell dishes under the microscope every other day to monitor growth rate (see Note 3). 5. As soon as cells reach confluence, maintain cells for additional 48 h, and then proceed to collect supernatant. 6. Remove culture medium from dishes with a needleless syringe, attach to the tip a filtering unit of 0.45 μm pore size, and filter the medium onto a sterile tube, to remove cells. At this point dishes with cells can be discarded (see Notes 4 and 5). 7. Pour a considerable amount of bleach on contaminated material for at least 1 h before discarding it. Please follow your local and state regulations for biological waste disposal. Viral supernatant can be stored at 20  C for 3–4 months or used freshly. If used freshly, schedule the viral supernatant preparation on the same day that bone marrow-derived cells are isolated, and store it at 4  C when not in use. 3.2 Isolation of Bone Marrow-Derived Cells

1. Euthanize one LXR-DKO mouse according to the relevant regulatory laws for animal experimentation (see Note 6). 2. Place mouse on a dissection board, previously moistened with 70% ethanol. 3. Clean with 70% ethanol the skin around the abdomen and hind legs of the mouse. 4. Using sterile tweezers and scissors, lift and hold the abdomen skin, and cut in a transversal direction. 5. Pull the skin down, by hand, firmly toward the feet to expose the hind leg muscles. Cut the remaining skin around the feet if necessary, to free each limb. 6. Dissect the legs, using a different pair of scissors and holding the foot with clean tweezers. Cut and detach the muscles around femurs and tibiae. Be very careful not to cut the bones. 7. Remove the femur from the hip joint twisting and pulling gently the leg until the femoral head is released (see Note 7). 8. Clean the remaining muscle from the bone rubbing it with a paper towel soaked in 70% ethanol. 9. Severe the feet and cut the knee joint to separate femur and tibia. Wash bones with 70% ethanol, and collect them in a 15 mL tube with PBS 1 (see Note 8).

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10. Inside the laminar flow hood, place the bones on a cell culture dish, and cut the bone epiphyses using sterile scissors. 11. Flush out the bone marrow with a syringe attached to a 25 G needle, inject 10 mL of PBS 1 on one end of the bone, and collect it from the other end on a 50 mL sterile tube, holding the bone in upright position with sterile tweezers. Repeat the washing procedure until the bone cavity becomes white. 12. Transfer the volume of PBS with bone marrow cells to a different tube, filtering through a 70 μm cell strainer, to remove cell clumps and bone fragments. 13. Spin the tube 5 min at 1000  g to pellet cells. Resuspend cell pellet in 1 mL of RBC 1 buffer, and let it stand for 2 min at room temperature. Next, dilute at least ten times the volume of cells with PBS 1. 14. Transfer the volume to a different sterile tube, filtering through a 70 μm cell strainer to remove lysed erythrocytes. 15. Spin bone marrow progenitors 5 min at 1000  g. 3.3 Immortalization of Bone MarrowDerived Macrophages

1. Resuspend bone marrow progenitor cell pellet in 1 mL of immortalization medium 1, and pipette gently. Add immortalization medium 1 (see Subheading 2.3) to a total volume of 30 mL. 2. Distribute 7.5 mL in each 100  20 mm treated culture dish (4 dishes in total). 3. Incubate overnight in a cell incubator. 4. On the next day, collect supernatant with detached cells in a tube. 5. Add 6 mL of immortalization medium 2 to each dish. 6. Spin tube with supernatants 5 min at 1000  g. 7. Resuspend cell pellet in 1 mL of immortalization medium 2 (see Subheading 2.3). Add immortalization medium 2 to a total volume of 6 mL. 8. Distribute 1.5 mL in each 100  20 mm treated culture dish. 9. Incubate dishes for 7 days under standard cell culture conditions, monitor under the microscope every other day. 10. Remove culture medium from dishes. Wash dishes gently with 6 mL of PBS 1 per dish, twice. 11. Add 6 mL of complete DMEM medium per dish, and scrape adherent cells. Collect the total volume of medium in a tube and spin 5 min at 1000  g.

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12. Resuspend cell pellet in 1 mL of complete DMEM medium supplemented with 10% GM-CSF or M-CSF, and add 14 mL more. 13. Distribute 7.5 mL into two 100  20 mm treated culture dishes. 14. Monitor cell growth rate on the following days, and replace culture medium when cell debris is observed, preferably before medium color changes. When cell density in the dish is over 40%, change medium approximately every other day. When cells start to proliferate autonomously, forming circular cell clusters, GM-CSF, M-CSF cytokine, or conditioned medium may no longer be added to the culture medium (see Note 9). Maintain cells in 100  20 mm culture dishes, proliferation rate varies among immortalized clones, therefore, cell passages should be empirically determined, typically 1:3 or 1:4 every other day. Cell dilution resulting in less than a 15% final cell density may cause growth arrest.

4

Notes 1. The originally described recombinant retrovirus carrying v-myc and v-raf oncogenes [15] are not commercialized as such, but several already immortalized cell lines following this method can be purchased to produce this type of retrovirus: AMJ2-C11 [18] or PMJ2-R [19], obtained from alveolar and peritoneal macrophages, respectively. 2. GM-CSF- or M-CSF-conditioned medium can be obtained from the cell culture supernatant of BHK-HM5 [20] or L929 (subclone 929 of parental L strain) cell lines [21], respectively, or, alternatively, medium supplemented with 10 ng/mL of the commercial recombinant murine GM-CSF or M-CSF cytokines. 3. J2 cells under the microscope should have a fibroblast-like appearance, i.e., multipolar or polygonal, and be elongated in shape and should attach well to the dish (Fig. 1). Before confluence is reached, cell culture medium can be replaced with fresh complete DMEM medium, and cells can be detached (see Note 4) in order to expand the cell population or kept frozen (see Note 5) as a J2 cell stock. 4. To detach J2 cells, add 5 mL of 0.5% trypsin-EDTA to one 100  20 mm culture dish, and place in the incubator for at least 2 min. Pipette cells very gently and transfer to a sterile tube. Add at least 2 the volume of trypsin used, to neutralize the enzyme activity.

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Fig. 1 Cultured J2 retrovirus-producing cells display a typical spindle-shaped and fibroblastic appearance. Optical microscope, magnification 10

5. To store a J2 cell stock at 20  C, resuspend pelleted cells in 1 mL freezing medium, use cryogenic vials. Freeze immediately at 80  C after cell resuspension. 6. Generally, one 8–14-week-old LXR-DKO mouse should yield enough bone marrow progenitors to perform macrophage immortalization following this protocol. 7. The femur can also be released cutting the hip, above the hip joint, but it involves a risk of cutting the bone, compromising the bone marrow sterility. Twist and pull gently until the femoral head, with the appearance of a small round white ball, is released from the hip. Twisting the leg to dislocate the hip joint is a slower and more conservative procedure. 8. At this point, femurs and tibiae can be stored at 4  C in a 15 mL tube filled with 7 mL of PBS 1 or sterile cell culture media for less than 24 h. 9. GM-CSF and M-CSF cytokines are growth factors that stimulate macrophage cell proliferation. Surviving cells after day 7 that autonomously proliferate independently of this stimulus have been retrovirally immortalized and no longer require cytokine stimulation. At this point cells become round, refringent under the microscope, and loosely adherent to the dish, forming circular cell clusters (Fig. 2).

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Fig. 2 Growth stages and cell appearance of immmortalized bone marrow-derived macrophages. (a, b) Cells on early stages of immortalization acquire a round shape and can form cell clumps. (c–e) After 7–10 days in culture, cells continue to increase in number and become typically arranged in circular aggregates. (f–h) At late stages, immmortalized cells divide autonomously at a constant rate, progressively filling the culture dish surface in loosely adherent cell clusters

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macrophages. Nat Immunol 14(8):831–839. https://doi.org/10.1038/ni.2622 11. Frenoux JM, Vernet P, Volle DH, Britan A, Saez F, Kocer A, Henry-Berger J, Mangelsdorf DJ, Lobaccaro JM, Drevet JR (2004) Nuclear oxysterol receptors, LXRs, are involved in the maintenance of mouse caput epididymidis structure and functions. J Mol Endocrinol 33 (2):361–375. https://doi.org/10.1677/jme. 1.01515 12. Blasi E, Mathieson BJ, Varesio L, Cleveland JL, Borchert PA, Rapp UR (1985) Selective immortalization of murine macrophages from fresh bone marrow by a raf/myc recombinant murine retrovirus. Nature 318 (6047):667–670. https://doi.org/10.1038/ 318667a0 13. Lutz MB, Granucci F, Winzler C, Marconi G, Paglia P, Foti M, Assmann CU, Cairns L, Rescigno M, Ricciardi-Castagnoli P (1994) Retroviral immortalization of phagocytic and dendritic cell clones as a tool to investigate functional heterogeneity. J Immunol Methods 174(1-2):269–279. https://doi.org/10. 1016/0022-1759(94)90031-0 14. Girolomoni G, Lutz MB, Pastore S, Assmann CU, Cavani A, Ricciardi-Castagnoli P (1995) Establishment of a cell line with features of early dendritic cell precursors from fetal mouse skin. Eur J Immunol 25 (8):2163–2169. https://doi.org/10.1002/ eji.1830250807 15. Rapp UR, Cleveland JL, Fredrickson TN, Holmes KL, Morse HC, Jansen HW, Patschinsky T, Bister K (1985) Rapid induction of hemopoietic neoplasms in newborn mice by a raf(mil)/myc recombinant murine retrovirus. J Virol 55(1):23–33 16. Flaveny CA, Griffett K, El-Gendy Bel D, Kazantzis M, Sengupta M, Amelio AL, Chatterjee A, Walker J, Solt LA, Kamenecka TM, Burris TP (2015) Broad Anti-tumor Activity of a Small Molecule that Selectively Targets the Warburg Effect and Lipogenesis. Cancer Cell 28(1):42–56. https://doi.org/ 10.1016/j.ccell.2015.05.007 17. Tavazoie MF, Pollack I, Tanqueco R, Ostendorf BN, Reis BS, Gonsalves FC, Kurth I, Andreu-Agullo C, Derbyshire ML, Posada J, Takeda S, Tafreshian KN, Rowinsky E, Szarek M, Waltzman RJ, McMillan EA, Zhao C, Mita M, Mita A, Chmielowski B, Postow MA, Ribas A, Mucida D, Tavazoie SF (2018) LXR/ApoE Activation Restricts Innate Immune Suppression in. Cancer Cell 172 (4):825–840.e818. https://doi.org/10. 1016/j.cell.2017.12.026

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Chapter 7 Dual Cross-Linking Chromatin Immunoprecipitation Protocol for Next-Generation Sequencing (ChIPseq) in Macrophages David A. Rollins and Inez Rogatsky Abstract Macrophages arise from distinct progenitor cell populations throughout development and are one of the most diverse cell types, capable of performing discrete functions, undergoing distinct modes of activation, and infiltrating or residing in numerous niches in the body. In adapting to their environments, macrophages display high levels of plasticity which is associated with profound epigenomic and transcriptional changes. Understanding these changes has been greatly facilitated by the next-generation sequencing (NGS)-based approaches such as RNAseq and chromatin immunoprecipitation (ChIP)seq. Despite the recent advances, obtaining quality ChIPseq data in macrophages for endogenous factors and especially coregulators recruited to DNA indirectly has proved to be extremely challenging. Here, we describe a dual crosslinking protocol for ChIPseq in macrophages that we developed for difficult-to-ChIP transcription factors, coregulators, and their posttranslational modifications. Further, we provide guidance on crucial optimization steps throughout this protocol. Although our experience has been predominantly in murine and human macrophages, we believe our protocols can be modified and optimized to study signal-induced epigenomic changes in any cell type of choice. Key words Macrophages, Transcriptional coregulators, Dual cross-linking, Chromatin immunoprecipitation, Next-generation sequencing

1

Introduction Macrophages (MΦ) are innate immune cells that arise in development from the myeloid progenitors in the yolk sac and fetal liver or from monocytic precursors in the bone marrow and are found in virtually every organ—as tissue residents (e.g., liver Kupffer cells, alveolar MΦ, or microglia in the brain)—or as infiltrates that arrive through the bloodstream. As critical mediators of homeostasis, development, and disease, MΦ are capable of responding to diverse environmental stimuli and adopting variegated activation states [1, 2]. For example, MΦ sense pathogens or cellular damage via

Matthew C. Gage and Ine´s Pineda-Torra (eds.), Lipid-Activated Nuclear Receptors: Methods and Protocols, Methods in Molecular Biology, vol. 1951, https://doi.org/10.1007/978-1-4939-9130-3_7, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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recognition receptors, such as the Toll-like receptors, and acquire inflammatory or phagocytic phenotypes. Tissueresident MΦ perform a variety of organ-specific functions: microglia carry out synaptic pruning of neurons during development, alveolar MΦ clear dust and foreign particles in the lung tissue, and Kupffer cells in the liver store hemosiderin so that it is available for the production of hemoglobin [1, 3]. In addition to their roles in normal physiology, MΦ contributions in autoimmune conditions such as rheumatoid arthritis or lupus, metabolic syndrome and type 2 diabetes, cardiovascular disease, and atherosclerosis continue to be recognized [1, 4], further expanding their repertoire. MΦ activation involves profound changes in their transcriptomic makeup which in turn relies on assembly and DNA binding of numerous signal-specific transcription factor (TF) complexes and chromatin modifiers which together dynamically affect the chromatin landscape of a cell. In our interrogation of these changes, the development and optimization of genomic techniques such as chromatin immunoprecipitation followed by nextgeneration sequencing (ChIPseq) has been instrumental, leading to enormous advancements in understanding how histone modifications, TFs, and components of basal transcriptional machinery interact with chromatin genome-wide. Critical advantages over older methods such as ChIP followed by qPCR or microarray (ChIPchip) include increased spatial resolution that enables more precise mapping of binding motifs and enhancer elements, an unbiased nature, a better signal-to-noise ratio, and an impressive linear range of signal quantitation [5]. Sequencing costs have commonly been cited as obstacles to ChIPseq; however the price of NGS has plummeted over recent years, making this approach an increasingly more feasible research tool [6]. As ChIPseq has become more available and affordable, some of the more significant difficulties that researches face are technical and have to do with specific cell types, lack of ChIP quality antibodies, or the architecture of TF complexes of interest. For example, TFs or coregulators that do not bind DNA directly are less detectable by ChIPseq [7–9]. This problem is further exacerbated when chromatin coregulators of interest are low-abundance—a common feature for coregulators whose limiting protein levels are a part of regulatory mechanism that ensures precise coregulator recruitment and signal-specific redistribution [10]. Although overexpression and epitope-tagging overcomes some of these obstacles, validating conclusions drawn in such artificial systems would require much additional experimentation. One obstacle to efficient detection of chromatin regulator interactions with DNA by ChIP has been the small size of formaldehyde molecule, the cross-linking reagent most commonly

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employed, and its higher reactivity toward protein–DNA rather than protein-protein complexes. As such, formaldehyde more efficiently links macromolecules that are less than 2 A˚ apart and interactions that occur directly on DNA as opposed to more distant protein–protein interactions. One method to increase ChIP efficiency for longer-distance interactions is to introduce a secondary cross-linker, such as N-hydroxysuccinimide esters DSG (7.7 A˚) or ˚ ), which has proven efficacious for p65 and STAT3 DSP (12 A ChIPs [11]. We employed a dual cross-linking procedure in our ChIPseq workflow for TFs and coregulators in macrophages of multiple species [12, 13]. Below, we present our ChIPseq protocols and provide a guide to troubleshooting the most common difficulties in ChIPseq experiments. We discuss multiple strategies used to optimize ChIPseq signals, including dual cross-linking, the use of magnetic beads, modified washes, and post-ChIP quality control analyses. The purpose of this chapter is to aid the investigators in optimizing the wet lab steps in ChIPseq; we will not be discussing in detail computational aspects of the NGS data analysis as there are numerous resources available that specifically address the bioinformatic workflow and troubleshooting when dealing with the ChIPseq data [14, 15].

2

Materials 1. Dynabeads Buffer: PBS with 1% BSA, filtered and stored at 4
C. 2. Disuccinimidyl glutarate (DSG) 0.5 M stocks in nuclease-free DMSO stored at 20
C (stable for 3–4 months). 3. Sterile PBS; two bottles—one stored at RT and the other at 4
C. 4. Nuclease-free, molecular biology grade water. 5. 16% methanol-free formaldehyde in closed vials, stored at RT protected from light. 6. 10 Fixing Buffer: 500 mM HEPES-KOH, pH 7.5, 1 M NaCl, 10 mM EDTA, pH 8.0, 5 mM EGTA, pH 8.0. Filtered and stored at 4
C. 7. 2.5 M glycine in nuclease-free water, sterile and filtered. Stored at RT. 8. 5 lysis buffer: 250 mM HEPES-KOH, pH 7.5, 700 mM NaCl, 5 mM EDTA, pH 8.0, 50% glycerol, 5% NP-40, 1.25% Triton X-100. Filtered and stored at 4
C as 10–20 mL aliquots.

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9. 10 wash buffer: 100 mM Tris–HCl, pH 8.0, 2 M NaCl, 10 mM EDTA, pH 8.0, 5 mM EGTA, pH 8.0. Filtered and stored at 4
C. 10. 1 shearing buffer: 0.1% sodium dodecyl sulfate (SDS), 10 mM EDTA, pH 8.0, 50 mM Tris, pH 8.0, filtered and stored at 4
C in 1.5 mL aliquots. 11. 1 Pico Shearing Buffer: 10 mM Tris–Cl pH 8.0, 100 mM NaCl, 1 mM EDTA pH 8.0, 0.5 mM EGTA pH 8.0, 0.1% Na-Deoxycholate, 0.5% N-lauroylsarcosine. Filtered and stored at 4
C in 1.5 mL aliquots. 12. 1 Elution Buffer: 1.9 mL TE (10 mM Tris, pH 8.0 and 1 mM EDTA, pH 8.0) þ 0.1 mL 10% SDS solution. Filtered and stored at RT. 13. 1 Modified RIPA Buffer: 50 mM HEPES, pH 7.5, 100 mM LiCl, 1 mM EDTA, pH 8.0, 1% NP-40, 0.7% Na-deoxycholate. Filtered and stored at 4
C. 14. Protease and phosphatase inhibitors. 15. Dynabeads Protein A or G, stored at 4
C. 16. Recombinant, PCR-grade 20 mg/mL Proteinase K. 17. DNase- and protease-free 10 mg/mL RNase A. 18. PCR Purification Kit. 19. 1.5 mL tube magnetic stand. 20. 1–10 μg antibody of your choice. 21. Cells: 10  106 MΦ per condition. 22. Sterile cell scrapers with 1.1800 blade. 23. Sterile 1.5 and 15 mL tubes. 24. Eppendorf DNA LoBind 1.5 mL tubes. 25. Sonication tubes: 1.5 mL TPX tubes for Bioruptor, 1.5 mL microtubes for Bioruptor Pico, or microTUBE with afa 130 μL tubes for the Covaris S220 sonicator. 26. Sonicators: Bioruptor, Bioruptor Pico (Diagenode), or Covaris S220. 27. Ice bucket. 28. Magnetic stirrer and hot plate. 29. Centrifuge/microcentrifuge. 30. Nutating mixer. 31. 55
C and 65
C water baths. 32. Tissue culture dishes, 150  20 mm.

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Methods

3.1 ChIP Day 0: Prepare Beads for Immunoprecipitation

See Notes 1–4 before proceeding. 1. Calculate the amount of Dynabeads magnetic beads slurry you will need based on the number of immunoprecipitations þ1, at 40 μL Dynabeads per reaction (e.g., for 4 reactions, prepare 5  40 ¼ 200 μL Dynabeads). 2. Pipet the calculated amount of Dynabeads into a 1.5 mL Eppendorf tube, and place in a magnetic stand on ice to aspirate buffer. 3. Wash the Dynabeads three times with 1 mL of Dynabeads buffer on ice, using the magnetic stand and aspirating the buffer. Dynabeads washes are performed by sandwiching 1.5 mL tubes between two pre-chilled Eppendorf racks and going in an up and down motion at least two to three times to resuspend the Dynabeads. 4. Resuspend the Dynabeads in 500 μL Dynabeads buffer, and add 1–10 μg of your antibody of choice on ice. 5. Nutate overnight at 4
C on a nutating mixer.

3.2 Dual CrossLinking

See Note 5 before proceeding. 1. Prepare 0.2 mM DSG in 37
C PBS, 15 mL per condition, at least 30 min before cells are ready as follows: (a) Heat PBS to 37
C on a magnetic stirrer—hot plate in a sterile 250 mL beaker with continuous stirring. (b) Thaw out 0.5 M DSG stocks and, using a P200 or P1000 pipet, slowly add 1:250 to the 37
C PBS to create a 2 mM DSG solution. 2. Take your 10  106 cells (in 150  20 mm dishes) out of the incubator and remove media. Wash cells gently with 5 mL RT or 37
C PBS, remove PBS and add 15 mL 0.2 mM DSG-PBS to each plate. Incubate for 30 min at RT. 3. Prepare 1 fixing buffer, 20 mL per condition, from 10 fixing buffer with sterile water at RT. Add 1:16 of 16% methanol-free formaldehyde to the 1 fixing buffer to create a 1% solution. 4. When step 2 is completed, remove DSG solution, wash gently with 5 mL RT PBS, remove PBS and add 20 mL 1% formaldehyde in Fixing Buffer from step 3 to plates. Incubate for 10 min at RT. 5. Quench cross-linking by adding 1.2 mL of 2.5 M glycine (20) to each plate. Incubate for 5 min at RT.

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6. Aspirate buffer and wash once with 5 mL PBS at RT. Aspirate PBS and add 5 mL of 4
C PBS. 7. Scrape the cells from plates on ice into 15 mL tubes and pellet cells at 600  g for 10 min. 8. Aspirate all PBS. Cell pellets can be flash frozen in liquid nitrogen and stored at 80
C (stopping point) or used directly in ChIP. 3.3 ChIP Day 1: Sonication and IP

Before starting this day, make sure sonicators are set up and have been degassed or cooled down to 4
C. See Note 6 before proceeding. 1. Prepare 1 lysis buffer from 5 lysis buffer, add protease and phosphatase inhibitors, and sterile water on ice. Lyse cell pellets in 3.5 mL of 1 lysis buffer. 2. Using a 5 mL pipette, resuspend the pellet by pipetting up and down five times, avoiding air bubbles. 3. Close the tubes tightly and place them on a nutator for 10 min at 4
C and pellet cell nuclei by centrifugation at 600  g for 10 min at 4
C. Aspirate lysis buffer on ice. 4. Prepare 1 wash buffer from 10 wash buffer, protease, and phosphatase inhibitors, and sterile water on ice. Resuspend nuclear pellets in 3.5 mL of 1 wash buffer. 5. Repeat steps 2 and 3. Make sure to remove as much wash buffer as possible without disturbing nuclear pellet. 6. Resuspend cells in 1 shearing buffer, and incubate for 15 min on ice before sonication. The type and amount of buffer depends on the type of sonicator to be used: (a) For Covaris S220 sonicator, resuspend pellet in 130 μL of 1 Covaris shearing buffer with inhibitors, and transfer to 130 μL Covaris AFA fiber tubes. Avoid bubbles! (b) Bioruptor sonicator: resuspend pellet in 300 μL of 1 Covaris shearing buffer with inhibitors, and transfer to 1.5 mL TPX tubes, avoiding bubbles. (c) Bioruptor Pico: resuspend pellet in 300 μL of 1 Pico shearing buffer, and transfer to 1.5 mL Pico microtubes, avoiding bubbles. 7. Sonication will be performed in a water-bath sonicator and is one of the most variable steps in ChIP protocols. Sonication conditions we have optimized for MΦ are provided below for your reference. (a) Covaris S220 sonicator with the conditions of 200 cycles/ burst, duty factor of 10, and 120 s have been used for murine bone marrow-derived MΦ.

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(b) Diagenode Bioruptor sonicator on the high setting, cycles of 30 s on and 30 s off, and a cycle number of 26 have been used for PMA-differentiated THP1 MΦ. Do not do more than ten cycles at a time, waiting 5–10 min in between. (c) Diagenode Bioruptor Pico sonicator with cycles of 30 s on and 30 s off has been used in MCSF-differentiated, CD14þ human MΦ from buffy coats with a cycle number of 10 and in BV2 murine microglia with a cycle number of 12. Do not do more than 10 cycles at a time, waiting 5–10 min in between. 8. Following sonication, transfer nuclear lysates to 1.5 mL Eppendorf tubes, and bring up the volume to 450 μL with the 1 shearing buffer. 9. Add 1:10 volume (50 μL) of 10% Triton X-100 to a final concentration of 1% to sequester the SDS. 10. Clear the lysates by centrifuging tubes at 14,000  g in a pre-cooled to 4
C microcentrifuge for 20 min. 11. Transfer the supernatant to a new DNA LoBind 1.5 mL tube on ice. Save 20 μL of the supernatant for input in a separate 1.5 mL tube on ice, and store at 4 or 20
C. 12. Once Dynabeads from day 0 have nutated overnight, use a magnetic stand to pellet and aspirate buffer. 13. Wash Dynabeads three times with 1 mL of Dynabeads buffer, using the magnetic stand to pellet and aspirating the buffer. Bring the volume up to that calculated for 40 μL per IP using Dynabeads buffer. 14. Aliquot 40 μL antibody-conjugated Dynabeads per IP. Incubate on a nutating mixer at 4
C overnight. 3.4 ChIP Days 2–3: Washes and DNA Isolation

See Notes 7 and 8 before proceeding. 1. Wash Dynabeads antibody-bound complexes eight times with 1 modified RIPA buffer containing protease inhibitors and once with TE þ 50 mM NaCl (no inhibitors). Dynabeads washes are performed by sandwiching 1.5 mL tubes between two pre-chilled Eppendorf racks and going in an up and down motion at least 2–3 times to resuspend the Dynabeads followed by a 5-min incubation on a nutating mixer at 4
C. Each time, use the magnetic stand on ice to collect the beads and a pipette to aspirate the buffer. Make sure to aspirate any remaining liquid in the cap of the tube after the last wash. 2. Prepare elution buffer with 1 μg/μL DNase-free RNase A (1 μL of 10 mg/mL stock per 100 μL elution buffer) and 200 μg/mL Proteinase K (1 μL of 20 mg/mL stock per 100 μL elution buffer). Aliquot 100 μL elution buffer with

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Proteinase K into each IP, vortex briefly, and incubate in a 55
C water bath for 1.5 h. 3. Remove the tubes from the water bath, vortex, and place into a 65
C water bath for 6 h (up to overnight) to reverse crosslinks. 4. Purify eluted DNA using the QIAquick PCR Purification Kit according to manufacturer’s directions. (we use 600 PB buffer at the start and 14 μL of nuclease-free, molecular biology grade water to aliquot the DNA into 1.5 mL DNA LoBind tubes at the end). 5. Transfer 2 μL of eluted DNA into a 1.5 mL tube for qPCR analysis (can bring the final volume up to 250 μL with nucleasefree water). 6. Transfer 2 μL of eluted DNA into a 1.5 mL tube for quality control analysis (see Subheading 3.5 below). Store the remaining 10 μL of eluted DNA at 20
C. 3.5 Quality Control Before Preparing ChIP Libraries and Sequencing

See Notes 9 and 10 before proceeding. 1. Determine concentration of eluted and input DNA using a Qubit fluorometer and fragment size distribution using Agilent Bioanalyzer. These steps are often performed by the core facility that handles sequencing. Ideally, this protocol yields DNA concentrations of at least 0.3–1.5 ng/μL with at least 7–15% (or higher) of DNA in a desired 130–230 nt range. These parameters should be close to each other across samples in a group (IP or input). 2. Perform qPCR with your samples, normalizing to input (2 μL of input sample can be diluted in up to 250 μL of nuclease-free water) and choosing positive and negative controls for enrichment using validated primers to specific genomic sites. Note that samples with qPCR fold enrichment of SRR6051682.sam

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3.6 Creating a Tag Directory

The next step requires a Tag Directory created from the SAM file. At this point, the reads are split into separated files based on their chromosomes. It will help to speed up the analysis of large datasets without requiring too much memory: 1. Run the following command to create a Tag Directory: makeTagDirectory Input-ChIP-Seq/ SRR6051682.sam -format samThe first argument is required to specify the output directory, and the second argument recognizes the input format but it is optional. 2. The resulting Tag Directory will contain several *.tags.tsv files; these files will be used for further downstream analysis by several tools contained within the HOMER package.

3.7 TagCounts Normalization

1. Normalize the number of reads between experiments in order to compare data from separate experiments. To do this, HOMER has a tool to resample an experiment to get equal number of reads to make experiments more comparable: getRandomReads.pl Input-ChIP-Seq/ 10000000 > output. tags.tsv In the example above, the first parameter identifies the Tag Directory object to be resampled, the second parameter indicates the total number of tags that will be sampled from the original Tag Directory, and the last parameter is the output file. 2. Create a new Tag Directory with the sampled tags using the t option: makeTagDirectory Resampled-Input-ChIP-Seq/ -t output.tags. tsv

3.8 Making Genome Browser Files

The visualization of ChIP-seq read densities is a common task that allows the comparison between different signal profiles as potential binding sites for transcription factors. HOMER presents a group of tools that helps to create various formatted files that can be used in different genome browsers (Fig. 2). 1. Here, we use UCSC to visualize bedGraph generated files (see Note 9) as follows: makeUCSCfile Resampled-Input-ChIP-Seq/ -o autoThe -o auto parameter automatically generates an output file placed in the same Tag Directory called Resampled-Input-ChIP-Seq.ucsc.bedGraph. gz. The output file can be named differently by specifying -o outputfilename. 2. To visualize bedgraph files in the UCSC Genome Browser, select Mouse GRCm38/mm10 in the genome section, then

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Fig. 2 Snapshot from UCSC genome browser showing binding events within the ABCA1 locus. Lanes represent control DKO, 3FLAG-LXRα and 3FLAG-LXRβ immortal bone marrow macrophages

click on the “add custom tracks” button, select the bedgraph file created, and click on Submit button. 3.9

Peak Calling

The main goal of ChIP-seq experiments is to identify transcription factor binding regions at the whole-genome level. The principle is to identify regions in the genome where more sequencing reads are found compared to what would be expected by chance. There are different methods to identify enrichment regions, such as the peak calling tool integrated into the HOMER package to identify peaks corresponding to 3FLAG-LXRα or 3FLAG-LXRβ, compared to a double knockout background for LXR as a negative control (see Note 10). 1. Find peaks for a transcription factor using the following command: findPeaks Resampled-3FLAG_LXRa-ChIP-Seq/ -style factor -o auto -i Resampled-DKO-ChIP-Seq/ In the above example, the first argument is the Tag Directory for the target sample. The second option -style specifies the type of ChIP-seq (in this case, we use factor). The argument -o specifies where the resulting peak file will be saved. If -o is not specified or -o auto is specified, the peak file will be written to /Resampled3FLAG_LXRa-ChIP-Seq /peaks.txt. The argument “-o” determines the place where the output file will be placed; in this example the output file will be stored in the target sample Tag Directory, and the name of the file will be peaks.txt.

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3.10 Annotating Peak Dataset

After peak calling is set, a genomic feature (genes, promoters, introns, exons, intergenic regions, etc.) to those peaks found should be assigned for further analysis (see Note 11). HOMER integrates a useful tool that helps with peak annotation. 1. Annotate peaks using the next command: annotatePeaks.pl peaks.txt mm10 > outputfile.txt The two first arguments, peak.txt and mm10, are required and indicate the peak file and the reference genome used to annotate. The last argument indicates the output file that is a data table that can be opened with Microsoft Excel or a similar program.

3.11

Motif Analysis

The enrichment regions found in the peak calling step (Subheading 3.9) are correlated with genomic sequences with a high affinity for specific transcription factors or with factors that can bind in the neighborhood. To search for enriched motifs on those regions, HOMER has a very powerful tool that runs a differential motif discovery algorithm, using background sequence regions as a control. This tool performs de novo motif analysis as well as checks the enrichment of known motifs. 1. Run the following command to perform an analysis of the peak file for discovery motifs: findMotifsGenome.pl peaks.txt mm10 3FLAG-LXRa_MotifOutput/ -size 200The first parameter specifies the peak file of interest, the second parameter indicates the reference genome, and the third parameter is the output directory where the resulting files will be saved; the -size parameter is mandatory and indicates the size from the peak center where the motif analysis will take place (see Note 12). 2. HOMER will create an output directory with a variety of output files including the motif enrichment analysis result displayed as HTML files (knownResults.html and homerResults. html).

3.12 Creating Heatmaps

An ultimate task of ChIP-seq experiments is to interpret the biological meaning of the genome-wide information about a given transcription factor occupancy or chromatin modification. For this purpose, we use seqMINER, an integrated user-friendly platform developed in Java (see Note 5), that allows graphical representations of the data as heatmaps (Fig. 3) to illustrate specific properties of the cistrome landscape in particular states of cell biology. Although seqMINER supports multiple file formats, we use BED format generated with HOMER from the Tag Directory:

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Fig. 3 Heatmap generated with seqMINER, showing binding peak intensity of LXR in control DKO, 3FLAG-LXRα and 3FLAG-LXRβ in immortalized bone marrow macrophages. Each panel in the figure represents 2 kb regions from the center of the binding event

1. Use a HOMER tool to change the file format as follows: tagDir2bed.pl Resampled-3FLAG_LXRa-ChIP-Seq/ 3FLAG_LXRa-ChIP-Seq.bed

>

2. Repeat for all the files that need to be reformatted. Once all the BED files are created, the process to create a heatmap is divided into three steps. 3. Load all necessary datasets (the peak and BED files) in the respective fields using the “browse” button. 4. Select the load aligned reads and load into the program by clicking the load file(s) >> button. 5. Once the datasets are loaded, change the peak extension in the Options section located in the Tool bar menu. 6. Select desired datasets and extract the data corresponding to the peak extensions from the BED files by clicking in the Extract data bottom. 7. Dataset clustering can be done by selecting a specific method from the dropdown menu and clicking the “Clustering” button. 8. A new window with the heatmap result will emerge.

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Notes 1. In this case, the sequencing data from the SRA repository was obtained through the command line in the Linux work environment. For this step it is important to know exactly where the SRA file is stored. The ftp (file transfer protocol) address where a file is hosted presents the following structure ftp://ftptrace.ncbi.nih.gov/sra/sra-instant/reads/ByRun/sra/SRR/ SRR605/SRR6051682/SRR6051682.sra, where SRR6051682 represents the SRA run accession that contains the sequencing data for a particular sequencing experiment. 2. When all the SRA files needed are the focus of a specific study, the URL structure is slightly different, ftp://ftp-trace.ncbi.nih. gov/sra/sra-instant/reads/ByStudy/sra/SRP/SRP118/ SRP118142/, where SRP represents the study accession that contains the project metadata describing a sequencing study or project. 3. If a wget -r command is used to retrieve the files, the option -r will recursively download the files, and you may also need to add a / to the end of the URL to make sure wget knows that it is a directory, like this wget -r ftp://ftp-trace.ncbi.nih.gov/sra/srainstant/reads/ByStudy/sra/SRP/SRP118/SRP118142/ 4. If the path of the Bowtie2 (*.bt2) files is stored in the BOWTIE2_INDEXES variable, you only need to specify the prefix for the Bowtie2 (*.bt2) index files in the Bowtie2 command; if not, then you need to specify the full path name of the index files. 5. Java Runtime Environment must be installed. 6. In our example we have establish eight processors/threads. This will vary depending on your work station. 7. mm10 is the common prefix for the *.bt2 files that were downloaded in Subheading 3.3. 8. The default output and the recommended storing read alignments is a SAM file. HOMER does not handle directly BAM format and should be converted into SAM format for downstream analysis. 9. Because bedgraph files can be very large, it is very useful to use bigWig files instead. HOMER can produce these files by running the specific tool for this task (bedGraphToBigWig). The only drawback is that a webserver is necessary to host the resulting bigWig files. In this case, instead of uploading the whole file to UCSC, the browser looks for the data file on the webserver where it is hosted and takes only the indispensable parts.

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10. It is highly recommended to use an Input, IgG, or a knockout sequencing run as negative control. A separate Tag Directory for the negative control experiment should be created. 11. An important prerequisite for peak annotation is that the mouse genome features must be configured to use with HOMER. See HOMER configuring section to do that. 12. The parameter “-size” is one of the most important in the findMotif script. For Transcription Factor peaks, most of the motifs are found 100 bp upstream or downstream from the peak center, making it better to use a fixed-size “-size 200 bp” rather than depend on the peak size “-size given.” References 1. Hong C, Tontonoz P (2014) Liver X receptors in lipid metabolism: opportunities for drug discovery. Nat Rev Drug Discov 13:433–444. https://doi.org/10.1038/nrd4280 2. Joseph SB, Castrillo A, Laffitte BA, Mangelsdorf DJ, Tontonoz P (2003) Reciprocal regulation of inflammation and lipid metabolism by liver X receptors. Nat Med 9:213–219. https://doi.org/10.1038/nm820 3. Ito A, Hong C, Rong X, Zhu X, Tarling EJ, Hedde PN, Gratton E, Parks J, Tontonoz P (2015) LXRs link metabolism to inflammation through Abca1-dependent regulation of membrane composition and TLR signaling. Elife 4: e08009. https://doi.org/10.7554/eLife.08009 4. Sequence Read Archive Submissions Staff (2011) Downloading SRA data using command line utilities. In: SRA Knowledge Base. National Center for Biotechnology Information (US), Bethesda (MD) https://www.ncbi. nlm.nih.gov/books/NBK158899/ 5. Andrews S. (2010). FastQC: a quality control tool for high throughput sequence data. http://www.bioinformatics.babraham.ac.uk/ projects/fastqc

6. Li H, Handsaker B, Wysoker A et al (2009) The sequence alignment/map format and SAMtools. Bioinformatics 25:2078–2079. https://doi.org/10.1093/bioinformatics/ btp352 7. Langmead B, Salzberg SL (2012) Fast gappedread alignment with Bowtie 2. Nat Methods 9:357–359. https://doi.org/10.1038/nmeth. 1923 8. Heinz S, Benner C, Spann N et al (2010) Simple combinations of lineage-determining transcription factors prime cis-regulatory elements required for macrophage and B cell identities. Mol Cell 38:576–589. https://doi.org/10. 1016/j.molcel.2010.05.004 9. Ye T, Krebs AR, Choukrallah MA, Keime C, Plewniak F, Davidson I, Tora L (2011) seqMINER: an integrated ChIP-seq data interpretation platform. Nucleic Acids Res 39(6):e35. https://doi.org/10.1093/nar/gkq1287 10. Karolchik D, Hinrichs AS, Kent WJ (2009) The UCSC Genome Browser. Curr Protoc Bioinformatics. Chapter 1. Unit1.4. https:// doi.org/10.1002/0471250953.bi0104s28

Chapter 9 Quantifying Cellular Cholesterol Efflux Sabrina Robichaud and Mireille Ouimet Abstract Measuring cholesterol efflux involves the tracking of cholesterol movement out of cells. Cholesterol efflux is an essential mechanism to maintain cellular cholesterol homeostasis, and this process is largely regulated via the LXR transcription factors and their regulated genes, the ATP-binding cassette (ABC) cholesterol transporters ABCA1 and ABCG1. Typically, efflux assays are performed utilizing radiolabeled cholesterol tracers to label intracellular cholesterol pools, and these assays may be tailored to quantify the efflux of exogenously delivered cholesterol or alternatively the efflux of newly synthesized (endogenous) cholesterol, in different cell types (macrophages, hepatocytes). Cholesterol efflux may also be customized to quantify cholesterol flux out of the cell to various exogenous cholesterol acceptors, such as apolipoprotein A-I, highdensity lipoprotein, or methyl-beta-cyclodextrin, depending on the purpose of the experiment. Here, we provide comprehensive protocols to quantify the net flux of cholesterol out of cells and recommendations on how this assay may be tailored as a function of the experimental question at hand. Key words Efflux, Endogenous cholesterol, Exogenous cholesterol, Phospholipid, ApoA-I, HDL, Methyl-beta-cyclodextrin, ABCA1, ABCG1, Cholesterol mass assay, Fluorescent cholesterol efflux

1

Introduction Measuring cholesterol efflux has had a major impact in the field of lipoprotein metabolism. From characterization of the mechanisms of apolipoprotein A-I (apoA-I) lipidation for high-density lipoprotein (HDL) formation to understanding the reverse cholesterol transport (RCT) pathway, quantification of cholesterol efflux has been a tremendous experimental tool. Cholesterol efflux, the movement of cholesterol out of cells, is a fundamental mechanism critical to maintaining cellular and whole-body cholesterol homeostasis. Efflux of cholesterol is largely regulated by the nuclear liver X receptors (LXR) that induce the expression of a host of genes involved in cholesterol efflux, such as the ATP-binding cassette (ABC) efflux transporters A1/G1, and the sterol regulatory element-binding proteins (SREBPs) transcription factors that regulate lipid homeostasis [1, 2].

Matthew C. Gage and Ine´s Pineda-Torra (eds.), Lipid-Activated Nuclear Receptors: Methods and Protocols, Methods in Molecular Biology, vol. 1951, https://doi.org/10.1007/978-1-4939-9130-3_9, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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LXRα and LXRβ bind to LXR response elements (LXREs) in target genes to activate gene expression, in conjunction with the retinoid X receptor (RXR) [3]. Under sterol-rich conditions, natural derivatives of cholesterol—oxysterols—are produced and bind to and activate LXRs [4]. LXR activation results in increased expression of cholesterol trafficking genes, such as Niemann-Pick type C (NPC) 1 and 2; several genes in the RCT pathway, such as macrophage ABCA1, ABCG1, and apoE; and genes regulating hepatic bile synthesis. Additionally, LXR can suppress the uptake of low-density lipoprotein (LDL) through induction of Idol (inducible degrader of the LDLR) that targets the LDL receptor (LDLR) for degradation [5]. Activated LXRs also reduce intestinal cholesterol absorption and promote hepatic cholesterol excretion by increasing the expression of ABCG5/G8 in the intestine and liver. Because of this concerted action of LXRs on regulating the first and last steps of RCT, namely, the rate-limiting efflux of cholesterol out of peripheral cells and its excretion into the feces, LXRs have reached the status of master regulators of this anti-atherogenic pathway and have been extensively studied in the context of cardiovascular disease. In turn, under sterol-depleted conditions, SREBP2 is activated to promote cholesterol biosynthesis via its transcriptional control over hydroxymethyl-glutaryl-coenzyme A [HMG-CoA] reductase, the rate-limiting enzyme in cholesterol synthesis, and LDL uptake via regulation of LDLR expression [2]. SREBP2 also exerts control over the expression of ABCA1, ABCG1, and NPC1 via miR-33, an intronic microRNA within the SREBP2 gene co-transcribed with SREBP2 during states of cholesterol depletion to limit cholesterol export [6]. More recently, miR-33 was shown to control cholesterol availability upstream of the ABCA1/G1 efflux transporters, by suppressing numerous genes of the autophagy pathway that contributes to lipid droplet (LD) catabolism [7]. Finally, in addition to being potent LXR agonists, oxysterols also regulate the SREBP pathway, and therefore there is an important cross talk between the two regulatory pathways. INSIGs serve as oxysterol sensors [8], and SCAP detects cellular cholesterol levels [9]; when oxysterols are bound to INSIG, the INSIG-SREBP-SCAP complex remains ER-bound, and thus oxysterols negatively modulate cholesterol uptake and biosynthesis. Following receptor-mediated endocytosis, lipoproteinassociated cholesterol esters (CE) and triglycerides (TG) are hydrolyzed by the lysosome-resident lysosomal acid lipase (LAL). Newly synthesized cholesterol as well as lipoprotein-derived cholesterol can be incorporated into LDs via ER-resident acyl-CoA/cholesterol acyltransferase (ACAT) that catalyzes the esterification of excess cholesterol for storage in cytosolic LDs. LD cholesterol undergoes constitutive cycles of esterification and hydrolysis that regulate cholesterol availability for cell membranes and efflux.

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Unesterified cholesterol released from LDs via lipolysis can be effluxed to a cholesterol acceptor if one is present, resulting in net CE hydrolysis, or otherwise it is re-esterified by ACAT [10, 11]. Two mechanisms contribute to LD-associated CE hydrolysis: (1) extralysosomal, cytoplasmic lipolysis that is mediated by neutral CE hydrolases [10, 12] and (2) autophagy-mediated LAL-dependent LD lipolysis, which involves autophagic sequestration and delivery of LDs in the lysosomal lumen for degradation [13, 14]. Interestingly, while autophagy doesn’t appear to mediate LD catabolism in “unloaded” cells, lipid loading specifically triggers this process to promote the clearance of excess cellular lipid [13, 15]. In macrophage foam cells, for example, autophagy is responsible for ~50% of LD-associated CE hydrolysis, while the remaining is attributable to neutral lipolysis [13]. The first step of RCT is cholesterol efflux out of peripheral cells, and the ABCA1/G1 cholesterol pumps are critical to this pathway [16]. Using the in vivo RCT assay to track the appearance of a 3 [H]-tracer from 3[H]-cholesterol-loaded macrophages into the plasma, liver, and feces of mice, ABCA1/G1 were shown to coordinate macrophage cholesterol removal for its clearance from the body [17]. Mechanistically, ABCA1 was shown to transfer phospholipids and cholesterol to HDL apolipoproteins, primarily apoAI that comprises ~80% of HDL protein [18], that are associated with very little or no lipid [19]. While ABCA1-mediated apoA-I lipidation occurs exclusively at the plasma membrane [20], ABCA1 traffics between the cell surface and late endocytic vesicles [21] and preferentially mobilizes cholesterol deposited in late endosomes/ lysosomes to stimulate cholesterol efflux [22], and internalization and shuttling of ABCA1 is functionally important for efflux of cholesterol out of endosomal compartments [23]. In turn, efflux to HDL involves passive diffusion of cholesterol as well as active cholesterol transfer, and there are three main transporters (ABCA1, ABCG1, and SR-BI) that mediate lipid transfer to HDL particles [24–27]. Macrophage ABCG1 expression has been shown to specifically stimulate net cholesterol efflux to HDL, but not to lipidpoor apoA-I [28], and a critical role for ABCG1 in preventing macrophage lipid accumulation in vivo has been established. Finally, synthetic cholesterol acceptors such as methyl-ß-cyclodextrin (mß-CD) can be efficient at extracting cellular cholesterol and hold potential to stimulate RCT in vivo [29, 30]. This chapter provides protocols for designing cholesterol efflux assays using radiolabeled cholesterol to quantify efflux of cholesterol out of primary murine macrophages. When thinking about designing an efflux experiment, two key elements must first be determined: (1) How will the cells be labeled? and (2) which efflux acceptor will be used? To address these questions, please see Notes 1 and 2. In addition, not all cell types express abundant levels of

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ABCA1/G1, i.e., HEK293 cells, and therefore might not represent a good model for studying cholesterol efflux. Below, we describe cholesterol efflux assays using primary murine macrophages with express scavenger receptors and ABCA1/G1 cholesterol pumps. These cells play a critical role in scavenging modified LDL that accumulates in the arterial wall and mediate the first step of the anti-atherogenic RCT pathway. See Fig. 1 for an overview of cellular cholesterol metabolism.

Fig. 1 Overview of cellular cholesterol metabolism. Lipoprotein-derived cholesterol esters internalized via receptor-mediated endocytosis (e.g., LDL via LDLR) are hydrolyzed in lysosomes. The ensuing free cholesterol can then be shuttled to membranes for efflux from the cell (if a cholesterol acceptor is present) or alternatively be stored in lipid droplets (if in excess or in the absence of a cholesterol acceptor) via ACAT-dependent esterification. Elevated cholesterol and its derivatives, oxysterols, activate nuclear receptors PPAR, LXR, and RXR to promote cholesterol efflux. Free cholesterol can pumped out of cells by ABCA1, ABCG1, and SR-BI to Apo-A1, nascent HDL, and HDL, respectively. On the other hand, low levels of cellular cholesterol/oxysterols activate the SREBP2 transcription factor to promote cholesterol biosynthesis and limit cholesterol efflux. Illustration created using BioRender

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Materials

2.1 Murine Bone Marrow-Derived Macrophages

1. Macrophage differentiation media: Dulbecco’s modification of Eagle’s medium (DMEM) with 4.5 g/L glucose, L-glutamine, and sodium pyruvate supplemented with 20% L929conditioned media (see Note 3), 10% heat-inactivated FBS, and 1% penicillin/streptomycin (P/S). 2. Dissection kit with dissecting and operating scissors and forceps. 3. 70% ethanol. 4. 1
PBS. 5. Sterile PCR tubes. 6. 1.5 mL microcentrifuge tube with lock lids. 7. 18 G needle. 8. 15 cm tissue culture dishes. 9. 5 mM EDTA in 1
PBS, ice-cold. 10. Cell scrapers. 11. Microcentrifuge set at 4  C. 12. Centrifuge.

2.2 Murine Peritoneal Macrophages

1. Macrophage media: Dulbecco’s modification of Eagle’s medium (DMEM) with 4.5 g/L glucose, L-glutamine, and sodium pyruvate supplemented with 10% heat-inactivated FBS, 1% P/S. 2. 70% ethanol. 3. Dissection kit with dissecting and operating scissors and forceps. 4. 18–20 G needle attached to a 5–10 mL syringe. 5. 1
PBS. 6. Desired tissue culture plate format for assay (i.e. 12-well, 24-well). 7. Centrifuge. 8. Red blood cell lysis buffer.

2.3 Equilibration Media

1. Dulbecco’s modification of Eagle’s medium (DMEM) with 4.5 g/L glucose, L-glutamine, and sodium pyruvate. 2. Fatty acid-free bovine serum albumin. 3. 50 mL syringe. 4. 0.22 μm syringe filter. 5. Vortex.

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2.4 KBr Density Solution

1. Potassium bromide. 2. Glass Pyrex dish (9 in. by 9 in.). 3. 2 M NaOH. 4. EDTA. 5. NaN3. 6. Oven set to 80  C.

2.5 Radiolabeled Lipids

1. Radiolabeled cholesterol: [3H]cholesterol or [14C]cholesterol. 2. Radiolabeled cholesterol ester: cholesteryl oleate [cholesteryl1,2-3H(N)]. 3. Radiolabeled mevalonate (cholesterol precursor): Mevalonolactone RS-[5-3H(N)]. 4. Radiolabeled acetate (cholesterol precursor): [3H]acetic acid or [14C]acetic acid. 5. Radiolabeled choline chloride (phospholipid precursor): choline chloride [methyl-3H].

2.6 Methyl– β-Cyclodextrin Cholesterol

1. Cholesterol. 2. Methyl-β-cyclodextrin. 3. DMEM media. 4. Glass tube. 5. 0.22 μm syringe filter. 6. 20 mL syringe. 7. Heating sonicating water bath at 37  C. 8. 37  C incubator. 9. Tube rotator.

2.7 Human PlasmaDerived Lipoprotein

1. Plasma. 2. Centrifuge set at 4  C. 3. EDTA. 4. NaN3. 5. PMSF. 6. BHT. 7. Ultracentrifuge. 8. Fixed-angle rotor (70Ti). 9. 39 mL Quick-Seal Polypropylene Tubes. 10. Tube cutter.

2.8 Lipoprotein Dialysis

1. Three glass beakers filled with 3 L of sterile 1X PBS. 2. 10,000 MWCO dialysis cassette.

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3. 18 G needle. 4. Lowry reagents. 5. Endotoxin-free lipoprotein. 2.9

Acetylated LDL

1. Low-density lipoprotein. 2. Sodium acetate. 3. Acetic anhydride. 4. NaCl. 5. 0.22 μm syringe filter. 6. Lowry Reagents.

2.10

Oxidized LDL

1. A glass beaker filled with 3 L of sterile 1
PBS at 37  C. 2. CuSO4. 3. Dialyzed lipoprotein. 4. 18 G needle attached to a 30 mL syringe. 5. BHT. 6. EDTA. 7. 0.22 μm syringe filter. 8. Lowry reagent.

2.11

Aggregated LDL

1. Endotoxin-free LDL. 2. Vortex. 3. Plain media.

3

Methods

3.1 Murine Bone Marrow-Derived Macrophages (BMDMs) Preparation (See Fig. 2)

Murine BMDMs can be differentiated from hematopoietic progenitors within the bone marrow of mouse femurs. From one mouse, approximately two confluent 15 cm dishes of BMDMs can be obtained. For this: 1. Harvest mouse femurs and tibias by removing the leg above the hip joint and below the knee joint (femur) and just below the knee joint to where the red bone marrow ends in the bone (tibia). 2. Thoroughly clean bones from any remaining tissue, and cut off both ends of femurs and tibias to expose the bone marrow. 3. Disinfect the bones by quickly dipping it in 70% ethanol before rinsing in DMEM and inserting in a PCR tube with a small hole at the tip. Use one tube for each leg (one femur and one tibia per tube).

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Fig. 2 Preparation of bone marrow-derived macrophages (BMDMs). Flowchart of the isolation of bone marrow cells and their differentiation into BMDMs for cholesterol efflux assays. Illustration created using BioRender

4. Place the PCR tube in a 1.5 mL microcentrifuge tube with a lock lid containing 100–150 μL of macrophage differentiation media. 5. Centrifuge at 8000
g for 2 min at 4  C. 6. Collect the bone marrow at the bottom of the 1.5 microcentrifuge tube by pipetting thoroughly up and down to resuspend the pellet.

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7. Transfer the cells to a 15 cm dish containing 20 mL of macrophage differentiation media. 8. Add an additional 5–10 mL of differentiation media between day 3 and 5 of differentiation. 9. Cells are ready 5 to 8 days after initial plating. 10. To lift cells, wash thoroughly with ice-cold 1
PBS twice to remove any residual FBS that will quench EDTA. 11. Add 10 mL of ice-cold 5 mM EDTA in 1X PBS (dissociation solution) to each dish, and incubate for 20 min at 4  C. 12. Gently lift away cells from the dish using a cell lifter. 13. Verify under the microscope that there are no residual cells on the plate. If cells remain, add 5–10 mL of dissociation solution, and scrape again until there are no cells left. 14. Rinse dish with DMEM containing 10% FBS, and add this volume containing residual cells to the lifted cells, to inactivate the EDTA from the dissociation solution. 15. Centrifuge the cells at 2000
g for 8 min at 4  C. 16. Resuspend the cells in growth media. 17. Plate cells in desired culture plates. 18. Cells will take 3–5 h to adhere completely to the cell culture plate. 3.2 Murine Peritoneal Macrophages Preparation

Peritoneal macrophages can be harvested by peritoneal lavage 3 to 5 days (optimally 4 days) following intraperitoneal (i.p.) injection of thioglycolate that elicits an inflammatory reaction recruiting monocytes to the peritoneum, which subsequently differentiate into macrophages in an attempt to resolve the inflammation. Typically, the yield for 1 mouse is ~20–30 million cells. 1. Inject 1 mL of 3% thioglycolate medium i.p. per mouse using an insulin syringe. 2. Wait 4 days and harvest peritoneal cells. 3. To harvest peritoneal cells, the peritoneal cavity can be washed by injecting 5–10 mL of 1
PBS with a 18–20 G needle attached to a 5–10 mL syringe. Keep collecting the 1
PBS until the PBS comes out clear and not a white cloudy solution. 4. Centrifuge the harvested peritoneal cells at 2000
g for 8 min. 5. Lyse red blood cells by resuspending the pellet in red blood cell lysis buffer and incubating at room temperature for 3–5 min. 6. Resuspend the cells in DMEM media supplemented with 10% FBS, 1% P/S. 7. Plate cells in desired culture plates (24-well, 48-well, or 96-well plates). For 24-well plates, plate peritoneal macrophages at

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500,000 cells per well in 500 μL of DMEM media supplemented with 10% FBS, 1% penicillin/streptomycin (P/S). 8. Cells will take 3–5 h to adhere properly to the cell culture plate. 3.3 Equilibration Media Preparation

2 mg/mL bovine serum albumin (BSA) medium is used to equilibrate cells subsequent to cholesterol loading where applicable, and it is also the media used to dilute cholesterol acceptors for cholesterol effluxes. 1. Weigh fatty acid-free BSA to have a final concentration of 2 mg/mL. For a 500 mL bottle of media, use 1 g of fatty acid-free BSA. 2. Transfer 30 mL of DMEM media from a fresh bottle into a 50 mL conical tube. 3. Add 1 g of fatty acid-free BSA, and vortex until the BSA is completely dissolved. 4. Filter-sterilize the concentrated BSA solution into the fresh DMEM bottle from which 30 mL was removed to dissolve the BSA, using a sterile 0.22 μm filter attached onto a sterile 50 mL syringe. 5. Supplement media with 1% P/S.

3.4 KBr Density Solutions Preparation

1. Dry KBr by placing a layer of KBr into a glass Pyrex 9
9 dish, and leave overnight in 80  C oven. 2. Remove from oven and transfer into 50 mL tubes. Store on desiccating crystals, and minimize the opening of the container. 3. To make a 1.006 g/mL density solution, start with: NaCl

11.40 g

NaOH

500 μL of 2 M solution

EDTA

0.1 g

NaN3

0.3 g

ddH2O

to 1 L

4. Add an extra 3 mL of ddH2O and store at 4  C. 5. Using a refractometer to verify the density of solutions, add KBr to increase the density or ddH2O to decrease the density as needed to achieve desired densities. 6. All solutions should be stored at 4  C. The ddH2O blank should be 0 on the refractometer.

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The following procedure describes the preparation of 10 mL of a 5 mM mß-CD/cholesterol solution with a mß-CD/cholesterol molar ratio of 8:1. Final concentration of cholesterol is 242 μg/mL [31]. This solution may be used to load cells with cholesterol. 1. Dissolve 2.41 mg of cholesterol and 66.90 mg of methyl-betacyclodextrin into 10 mL of DMEM media, in a glass tube. 2. Sonicate the solution at 37  C for 3 min, in a sonicating water bath. 3. Incubate the solution overnight on a tube rotator at 37  C. 4. Filter-sterilize the solution and store at 4  C away from light.

3.6 Human PlasmaDerived Lipoproteins Preparation

Human LDL and HDL lipoproteins may be isolated by sequential density ultracentrifugation or alternatively purchased. All of the following steps, adapted from Chapman et al. ([32, 33]), are carried out in a sterile environment where possible, using sterile and endotoxin-free glassware, reagents, plasticware, and stir bars (see Notes 4–9). Plasma preparation: 1. Spin blood at 1500
g for 15 min at 4  C to pellet red blood cells. 2. Transfer the plasma to a new tube, and add 1 mM EDTA, 0.02% NaN3, 0.5 mM PMSF, and 20 μM BHT. Do not add the latter if you plan on subsequently oxidizing the lipoproteins. Isolation of lipoproteins from human plasma using sequential density ultracentrifugation: Centrifugation steps can be done using a fixed angle 70Ti rotor and 39 mL Quick-Seal Polypropylene Tubes. Tubes need to be filled to their maximum volumes at all times. If required, fill tubes completely by supplementing to their maximum volume using the appropriate density buffer. For this: 1. Transfer plasma to quick-seal tubes. 2. Centrifuge plasma for 20 min at 20,000 rpm or 41,000
g at 4  C. 3. Remove the top layer containing chylomicrons, and keep the bottom fraction. 4. Centrifuge the bottom fraction 40,000 rpm or 165,000
g at 4  C.

for

18

h

at

5. Collect the top layer and keep the bottom. The top layer contains the VLDL. 6. Calculate the proper amount of KBr needed to have a density of 1.019 g/mL according to Table 1. Assume that plasma has a starting density of 1.006 g/mL. 7. Adjust the density of the bottom fraction of step 6 to 1.019 g/ mL with dry KBr by slowly stirring the solution for 30 min at

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Table 1 Quantity of KBr to be added to 100 mL of plasma to change the density from d1 to d2. Units are shown in grams

4  C to completely dissolve the KBr. Transfer to Quick-Seal tubes, fill, and seal. 8. Centrifuge for 18 h at 40,000 rpm or 165,000
g at 4  C. 9. Collect the top fraction containing intermediate-density lipoprotein (IDL) using a tube cutter. Aim to collect the smallest volume possible. Keep the bottom fraction. 10. Adjust the density of the bottom fraction of step 9 to 1.063 g/mL with dry KBr by slowly stirring the solution for 30 min at 4  C to completely dissolve the KBr. Transfer to Quick-Seal tubes, fill, and seal. 11. Centrifuge for 20 h at 40,000 rpm or 165,000
g at 4  C.

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12. Collect the top fraction containing LDL using a tube cutter. Keep bottom fraction. 13. Adjust the density of the bottom fraction of step 12 to 1.125 g/mL with dry KBr by slowly stirring the solution for 30 min at 4  C to completely dissolve the KBr. Transfer to Quick-Seal tubes, fill, and seal. 14. Centrifuge for 20 h at 40,000 rpm or 165,000
g at 4  C. 15. Collect the top fraction containing HDL2 using a tube cutter. Keep bottom fraction. 16. Adjust the density of step 15 to 1.21 g/mL with dry KBr by slowly stirring the solution for 30 min at 4  C to completely dissolve the KBr. Transfer to Quick-Seal tubes, fill, and seal. 17. Centrifuge for 40 h at 40,000 rpm or 165,000
g at 4  C. 18. Collect the top fraction containing HDL3 using a tube cutter. Keep bottom fraction. 19. Adjust the density of the bottom fraction of step 18 to 1.25 g/ mL with dry KBr by slowly stirring the solution for 30 min at 4  C to completely dissolve the KBr. Transfer to Quick-Seal tubes, fill, and seal. 20. Centrifuge for 40 h at 40,000 rpm or 165,000
g at 4  C. 21. Collect the top fraction containing VHDL using a tube cutter. The bottom layer now consists of lipoprotein-deficient serum (LPDS). 3.7 Lipoprotein Dialysis

1. Immerse a dialysis cassette (10,000 MWCO) into a pre-sterilized 4 L glass beaker filled with 3 L of cold, sterile, and endotoxin-free 1
PBS for at least 1–2 min. 2. Remove the hydrated cassette, and add lipoproteins of interest to the cassette using a syringe with an 18 G needle. 3. After adding the lipoprotein solution, remove all the air from the cassette, and mark the port used. 4. Dialyze the LDL against 3 L of 1
PBS for 4 h at 4  C while gently stirring. Keep the beaker covered at all times to keep the solution sterile. 5. Transfer the dialysis cassette into a new 4 L of cold, sterile, and endotoxin-free 1
PBS. 6. Dialyze overnight at 4  C. 7. Using a new port on the dialysis cassette, inject a little air, then aspirate the lipoprotein solution using a 18 G needle attached to a 30 mL syringe, and transfer into a 50 mL conical tube. 8. Measure the final lipoprotein concentration by Lowry assay. Volumes of 1 μL, 2.5 μL, and 5 μL of sample should be enough for quantification. Keep at least 100 μg of lipoprotein for endotoxin testing.

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3.8 Acetylated LDL Preparation

LDL can be acetylated by repetitive additions of acetic anhydride [34] or alternatively purchased. For LDL acetylation: 1. Add equal volumes of LDL and saturated sodium acetate into a small glass beaker, with a stir bar. 2. Slowly stir the mixture, at the lowest speed to avoid aggregation, in the cold room at 4  C. Every 5 min, add 5 μL of acetic anhydride until you reach the total volume of acetic anhydride required to completely acetylate LDL lysine residues (see Note 10). 3. Dialyze the acetylated LDL against 4 L of 1
0.15 M NaCl three times (4 h minimum or overnight at 4  C). 4. Filter-sterilize the dialysis product with a 0.22 μm filter. 5. Measure the concentration with Lowry assay and perform endotoxin test.

3.9 Oxidized LDL (oxLDL)

Moderately oxLDL may be obtained via oxidation of LDL (protocol adapted from [35]). Extensively oxidized LDL can be obtained by incubation with CuSO4 for 24 h as previously described [36] or alternatively purchased, and minimally oxidized LDL can be obtained by incubation with CuSO4 for 2 h. 1. Add 3 L of warm 1
PBS to a 4 L glass beaker. Warm up the PBS the night before. 2. Add 15 μL of 1 M CuSO4 to the 1
PBS. 3. Take the dialysis cassette, and add it to the fresh beaker with 1
PBS and CuSO4. 4. Dialyze the LDL against 5 μM of CuSO4 for exactly 6 h at 37  C. 5. After 6 h, remove the cassette from the beaker, and extract the dialysis product from the cassette using a 30 mL syringe attached to a 18 G needle. 6. Inject some air first into the dialysis cassette before aspirating the oxLDL. The solution should now be white, not yellow. 7. Transfer the oxLDL into a 50 mL tube. 8. To stop the oxidation, add 50 μM BHT and 0.2 μM EDTA to the oxLDL. Gently mix the sample with a pipette. 9. Filter-sterilize the oxLDL and store at 4  C away from light. 10. Keep an aliquot of around 300 μL for measuring the concentration via a Lowry assay and endotoxin test.

3.10 Aggregated LDL (agLDL)

Protocol adapted from Otero-Vinas et al. [37]. 1. In a 15 mL conical tube, add the correct volume of sterile endotoxin-free LDL to obtain a final concentration of 1 mg/ mL of LDL in plain DMEM (i.e., 1 mL of 5 mg/mL LDL solution into 4 mL of plain DMEM). 2. Vortex at maximum speed for 5 min. The agLDL solution should be cloudy.

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1. In a glass tube that can be capped, dissolve the required amount of radiolabeled cholesterol needed to load the cells (for a final concentration of 0.5 μCi/mL to 5 μCi/mL) into a small volume (~500 μL of plain DMEM media) (see Note 11). 2. Add the required amount of lipoprotein to the mixture, enough to load all the cells of the plate (or the amount of wells needed) at a final concentration of 50 μg/mL. 3. Incubate at 37  C overnight (or for at least 30 min before adding to cells) to allow for the radiolabeled cholesterol to incorporate into the lipoprotein. 4. Following the preincubation of lipoprotein and cholesterol, add the amount of complete DMEM media (10% FBS or LPDS with 1% P/S) needed for the experiment. 5. Filter-sterilize and add to cells of the culture plate.

3.12 Cholesteryl Ester Incorporation

1. Take 5 mL of LPDS with 1% P/S, and add the required amount of lipoprotein needed. 2. Dissolve the radiolabeled cholesteryl ester in DMSO, in a volume equivalent to 1/100 of the final volume (see Note 12). Using an insulin syringe, aspirate the cholesteryl ester (100 μL), and rapidly inject into the LPDS/lipoprotein solution. If done successfully, no precipitate should form. If one does form, this step must be repeated. 3. Incubate the mixture overnight at 37  C. 4. Filter-sterilize before use.

3.13 Quantifying Efflux of Exogenously Delivered Cholesterol

The following protocols are designed to be used with adherent cells such as macrophages (see Notes 13–18).

3.13.1 LDL-Derived Cholesterol

To maximize LDL endocytosis, cells may be preincubated for 24 h in LPDS-containing media 24 h prior to incubation with [3H] cholesterol-LDL to upregulate LDLR. 1. Remove media from the wells. 2. Add the correct volume of DMEM containing 10% LPDS (see step 21 of Subheading 3.6). Incubate at 37  C for 24 h prior to loading. 3. Proceed to step 1 of Subheading 3.13.2 below, with the exception that LDL loading in step 2 in that section can be done in 10% LPDS, 1% P/S DMEM.

3.13.2 Modified LDL-Derived Cholesterol

1. Aspirate media from the wells. 2. Add the correct volume of radiolabeled lipoproteins, for example, 0.5 mL/well of [3H]cholesterol-acLDL, [3H]cholesterol-

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oxLDL, or [3H]cholesterol-agLDL, for a 24-well plate. Typically, we use 50 μg/mL of lipoproteins preincubated with [3H]cholesterol for a final concentration of 5 μCi/mL (see Note 19). 3. Incubate at 37  C for 30 h in 10% FBS media containing 1% P/S (see Note 20). 4. Carefully aspirate the media, and wash twice with sterile, endotoxin-free 1X PBS to wash away excess lipoproteins. 5. Incubate cells in equilibration media (2 mg/mL BSA, 1% P/S) overnight (18 h) allowing for equilibration of the newly incorporated cholesterol into the cells. 6. To begin efflux, aspirate the equilibration media, and add cholesterol acceptors such as human recombinant apo-A1 (50 μg/mL) prepared as [38] or purchased, or HDL (50 μg/ mL) to serum-free media (fresh 2 mg/mL fatty acid-free BSA DMEM), to half of the wells. The other half of the wells will contain serum-free media alone. 7. Incubate for 4–24 h. 8. To stop efflux, transfer the well supernatants to microcentrifuge tubes. 9. Briefly centrifuge to remove nonadherent cells, at 2,500
g for 8 min. 10. Dissolve the remaining cells in the wells by adding 0.5 mL of 0.5M NaOH/well. 11. Incubate for at least 2 h at room temperature on a rocker to lyse the cells (see Note 21). 12. Measure radioactivity in 200 μL of the supernatants and in 200 μL the dissolved cells by scintillation counting, adding ~3 mL of scintillation cocktail to each vial (see Note 22). 13. Cell lysates and media can be frozen at 20  C for future use. 3.14 Quantifying Efflux of Newly Synthesized Cholesterol

To quantify the efflux of newly synthesized cholesterol, a precursor in the cholesterol biosynthesis pathway such as mevalonate or acetate may be used [30]. 1. Label cells for 48 h with [3H]mevalonate (10 μCi/mL) or [3H] acetate (20 μCi/mL) DMEM with 1% FBS P/S. 2. Perform efflux as in Subheading 3.13.2, using human recombinant apoA-I. 3. Immunoprecipitate apoA-I in supernatant aliquots with a polyclonal antihuman apoA-1 rabbit antiserum and Protein G Sepharose. 4. Collect immunoprecipitates by centrifugation according to Protein G Sepharose manufacturer’s specifications.

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5. Wash immunoprecipitates four times with 1X PBS. 6. Resuspend in a final volume of 0.5 mL for scintillation counting. 7. Subtract the radioactivity of the apoA-I-free media to that of the apoA-I-containing media, and normalize to total cellular protein levels. 8. Determine total cellular protein levels by Markwell Lowry assay [39]. 3.15 HDL Genesis/ ApoA-I Lipidation

To determine the lipidation of apoA-I with cholesterol or phospholipids, as a measure of HDL biogenesis: 1. Label cells with [3H]mevalonate (10 μCi/mL) or [3H]acetate (20 μCi/mL) for 48 h in DMEM with 1% FBS P/S, to quantify the lipidation of apoA-I with cholesterol. Alternatively, cells may be labeled with [3H]choline (5 μCi/mL) to quantify phospholipid lipidation of apoA-I. 2. Incubate cells with serum-free medium for 24 h. 3. Stop efflux as in Subheading 3.13.2. 4. Immunoprecipitate the secreted apoA-I directly from the cell supernatants and subject to scintillation counting as detailed in [40].

4

Notes 1. How the cells will be labeled depends on the cellular cholesterol pool of interest. If this is endogenous cholesterol, then a precursor in the cholesterol biosynthesis pathway such as mevalonate may be used. Alternatively, when tracking the fate of lipoprotein-derived cholesterol, native LDL or modified LDL in which radiolabeled cholesterol or CE has been incorporated can be used. If LD biology and lipolysis are the focus, then cells must be loaded with an excess of cholesterol to stimulate LD biogenesis. In this scenario, modified LDL should be employed to generate foam cells enriched in cytosolic LDs. It is important to note however that not all cells express scavenger receptors (such as scavenger receptor class A [SRA], cluster-determinant 36 [CD36]) that mediate the recognition and internalization of modified LDL such as aggregated LDL (agLDL), oxidized LDL (oxLDL), or acetylated LDL (acLDL), a commonly used model modified LDL. Such cells, hepatocytes, for example, may alternatively be loaded with cholesterol-cyclodextrin complexes or ß-VLDL remnants, or efflux of newly synthesized cholesterol (mevalonate precursor) or phospholipids (choline chloride precursor) may be studied in the context of apoA-I lipidation/HDL biogenesis.

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2. For the question of which efflux acceptor to be used, this depends on which cholesterol pool the researcher wants to deplete. For example, at 37  C uncomplexed mß-CD is delivered to late endosomes and lysosomes via micropinocytosis, where it efficiently promotes cholesterol removal from these cell compartments [41]. However, if used at 4  C, mß-CD will selectively deplete the cell membrane of its cholesterol [30]. In turn, apoA-I efflux is primarily ABCA1-dependent, and ABCA1 promotes efflux of endo-lysosomal cholesterol, while HDL efflux is primarily ABCG1-dependent. It is also important to note that cholesterol efflux is bi-directional and follows a gradient concentration, from the highest concentration to the lowest, and therefore efflux assays should be complemented by cholesterol mass assays to ascertain the net flux of cholesterol out of cells [42]. Other factors to take into consideration are: (a) The duration of the efflux assay—If efflux of cholesterol arising from neutral (constitutive) or acid lipolysis (induced under certain conditions and requires longer incubations with apoA-I) is of interest. (b) Dissociation of the cholesterol reesterification arm of the CE cycle—An ACAT inhibitor (ACATi) may be included during the incubation of cells with cholesterol acceptors to prevent reesterification of hydrolyzed CE during efflux. (c) Basal versus maximal efflux capacity—Cells should be treated with an LXR agonist such as T0901317 (synthetic LXR ligand) or oxysterols (natural LXR ligands) prior to efflux to upregulate ABCA1 and ABCG1 and maximally stimulate efflux. This may be useful if one is anticipating a reduction in efflux caused by their treatment of interest. 3. L929-conditioned media is collected from L929 cells that secrete macrophage colony-stimulating factor (M-CSF), a lineage-specific growth factor that promotes the proliferation and differentiation of myeloid progenitors into macrophages. L929-conditioned media can be obtained by collecting the culture media of L929 cells after 3–7 days of culture [43]. Cells should be around 90% confluent at the time of the last collection. Media is then pooled and filter-sterilized using 0.2 μm filter. L929-conditioned media can be stored at 20  C until use. 4. Endotoxin testing: lipoprotein preparations should be tested for endotoxin contamination. All required reagents for endotoxin testing may be purchased from, e.g., Associates of Cape Cod Incorporated: Pyrotell Gel clot Formulation, Single Test Vial (STV) (Cat# SKGS2505), and Control Standard Endotoxin (CSE), Escherichia coli O113:H10, 0.5 μg/vial (Cat# SKE00055). When preparing lipoproteins, the risk of

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endotoxin contamination can be reduced by baking glassware overnight and by cleaning plasticware with an endotoxin removal detergent prior to use. 5. When isolating lipoproteins, it is important to not stir the lipoproteins vigorously to avoid aggregation. 6. During lipoprotein isolation, it is possible to skip the IDL isolation step and proceed directly to LDL isolation from the VLDL fraction. 7. To collect all HDL particles together, adjust the density of the bottom layer of the LDL fractionation spin with dry KBr to 1.21 g/mL directly, which corresponds to the HDL3 particles. 8. It is possible to concentrate the LDL fraction further using ultracentrifugation. For this, add the LDL fraction to a new mL Quick-Seal® Polypropylene Tube (Cat# 362248), and centrifuge using the TLA110 rotor from Beckman Coulter Life Sciences at 60,000 rpm or 200,000
g for 18 h at 4  C. 9. Lipoproteins should be stored at 4  C protected from light for less than 1 month; otherwise they will become oxidized. In the case of oxLDL, various degrees of LDL oxidation will produce a substrate for distinct scavenger receptors. For example, similar to acLDL, minimally oxidized LDL binds to SRA and CD14 but not CD36, whereas mildly oxLDL is a ligand for CD36 [35]. 10. The correct volume of acetic anhydride to be used to acetylate LDL can be calculated using the following equation: Mass of LDL added
1:5 ¼ volume of acetic anhydride required in μL:

11. For [3H]cholesterol or [14C]cholesterol, it typically comes solubilized in EtOH. Ensure that the volume of radiolabeled cholesterol-EtOH is not greater than 30 μL; otherwise it may cause the lipoprotein to precipitate during incubation. If needed, dry down cholesterol with a stream or nitrogen, and resuspend in 30 μL of EtOH by thoroughly vortexing, and then add the plain media and vortex to homogenize. 12. Cholesteryl ester typically comes in toluene. Dry down the required amount of cholesteryl ester with nitrogen, then add 3 mL of CH3Cl, and dry down completely again. Add 20 μL of CH3Cl, resuspend the cholesteryl ester, and then add ~50 μL DMSO. Vortex vigorously, and let it sit until all the liquid has collected in the bottom. 13. Efflux is expressed as a percentage of 3[H]-cholesterol in medium/(3[H]-cholesterol in mediumþ3[H]-cholesterol in cells)
100%. Specific effluxes to apoA-I or HDL are calculated by subtracting effluxes of the wells without apoA-I or

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HDL from those containing apoA-I or HDL, respectively. Alternatively, for HDL genesis/apoA-I lipidation assays, cell lysates may be used to quantify protein content in order to express efflux as CPM/cell protein. 14. The equilibration time can be altered to suit experimental needs. For instance, if immortalized (dividing) J774 macrophages are used rather than primary (nondividing) murine macrophages, it might be useful to shorten the equilibration and efflux duration to avoid cell over-confluency. 15. Effluxes may be performed in 12-well plates (best for apoA-I lipidation assays in hepatic cells), 24-well plates, or 48-well plates (both good for macrophage effluxes, but 24-well plates might be best if cell lysates will be used for both total counts and for TLCs). 16. It is possible to collect some media over the course of the efflux assay, for example, if it’s unknown whether the maximal efflux difference between treatments occurs at 4 h, 8 h, or 24 h. For this, for a 24-well plate (0.5 mL/well), 50 μL might be removed at 4 h and another 50 μL at 8 h, and the efflux stopped at 24 h. For best accuracy, we recommend counting 40 μL of the 50 μL for the 4 h and 8 h time points and 200 μL of the 24 h time point. Each time point must be taken into consideration for the total amount of radioactivity in calculating cholesterol effluxes for each time point. 17. Although this chapter primarily describes assays to quantify radiolabeled cholesterol, fluorescent cholesterol analogs may alternatively be used to perform cellular cholesterol efflux assays, such as dehydroergosterol (DHE) [44], natural fluorescent sterol from yeast, or BODIPY-cholesterol [45]. 18. The amount of radioactivity used might depend on the needs of the experiments, for example, the length of the time of the efflux assay, the type of cell used, etc. A range of 0.5 μCi/ mL–5 μCi/mL is appropriate for cholesterol efflux assays. 19. Lipids, for example, total cellular cholesterol (free and esterified) if the cells contain radiolabeled cholesterol or a precursor of it, can be quantified from the NaOH cell lysates by thinlayer chromatography. For this, total lipids are extracted [46] and separated by thin-layer chromatography (TLC) on silica gel plates using a nonpolar solvent system (hexane/diethyl ether/acetic acid, 70:30:1, v/v) for separation of cholesterol and CE. The bands corresponding to cholesterol and cholesteryl esters are excised and counted for radioactivity. 20. During modified LDL loading, we recommend using DMEM media containing 10% FBS, 1% P/S. In our experience, this yields comparable loading to using media containing LPDS, given that in contrast to LDLR the expression of scavenger

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receptors that mediate the uptake of modified LDL is not affected by the serum lipoprotein content and, more importantly, cell viability is superior in the presence of FBS- as compared to LPDS-containing media. However, if directly comparing LDL versus modified LDL loading, one would need to use comparable loading media (in this case LPDScontaining media), and if the LPDS media preincubation step is performed for LDL-loaded cells, then it must be done for modified LDL-loaded cells alongside. 21. To store cell lysates for longer, transfer them into microcentrifuge tubes, or wrap the plate in saran wrap along with moist paper towels (good for a few hours at room temperature or overnight at 4  C) to prevent evaporation. 22. It is good practice to determine cellular mass of cholesterol, in parallel to cholesterol efflux [42]. For this, the Biovision Cholesterol Quantitation Kit or other commercially available kits can be employed, following the manufacturer’s instructions. Variations in total cellular cholesterol esters before and after efflux can be expressed as % cholesterol ester hydrolysis, as a previously described [13]. References 1. Chinetti G, Lestavel S, Bocher V, Remaley AT, Neve B, Torra IP, Teissier E, Minnich A, Jaye M, Duverger N, Brewer HB, Fruchart JC, Clavey V, Staels B (2001) PPAR-alpha and PPAR-gamma activators induce cholesterol removal from human macrophage foam cells through stimulation of the ABCA1 pathway. Nat Med 7(1):53–58. https://doi.org/ 10.1038/83348 2. Horton JD, Goldstein JL, Brown MS (2002) SREBPs: activators of the complete program of cholesterol and fatty acid synthesis in the liver. J Clin Invest 109(9):1125–1131. https://doi. org/10.1172/jci15593 3. Hong C, Tontonoz P (2008) Coordination of inflammation and metabolism by PPAR and LXR nuclear receptors. Curr Opin Genet Dev 18(5):461–467. https://doi.org/10.1016/j. gde.2008.07.016 4. Shibata N, Glass CK (2010) Macrophages, oxysterols and atherosclerosis. Circ J 74 (10):2045–2051 5. Zelcer N, Hong C, Boyadjian R, Tontonoz P (2009) LXR regulates cholesterol uptake through idol-dependent ubiquitination of the LDL receptor. Science 325(5936):100–104. https://doi.org/10.1126/science.1168974 6. Moore KJ, Rayner KJ, Suarez Y, FernandezHernando C (2010) microRNAs and

cholesterol metabolism. Trends Endocrinol Metab 21(12):699–706. https://doi.org/10. 1016/j.tem.2010.08.008 7. Ouimet M, Ediriweera H, Afonso MS, Ramkhelawon B, Singaravelu R, Liao X, Bandler RC, Rahman K, Fisher EA, Rayner KJ, Pezacki JP, Tabas I, Moore KJ (2017) microRNA-33 regulates macrophage autophagy in atherosclerosis. Arterioscler Thromb Vasc Biol 37(6):1058–1067. https://doi.org/ 10.1161/ATVBAHA.116.308916 8. Radhakrishnan A, Ikeda Y, Kwon HJ, Brown MS, Goldstein JL (2007) Sterol-regulated transport of SREBPs from endoplasmic reticulum to Golgi: oxysterols block transport by binding to Insig. Proc Natl Acad Sci U S A 104(16):6511–6518. https://doi.org/10. 1073/pnas.0700899104 9. Radhakrishnan A, Sun LP, Kwon HJ, Brown MS, Goldstein JL (2004) Direct binding of cholesterol to the purified membrane region of SCAP: mechanism for a sterol-sensing domain. Mol Cell 15(2):259–268. https:// doi.org/10.1016/j.molcel.2004.06.019 10. Brown MS, Goldstein JL, Krieger M, Ho YK, Anderson RG (1979) Reversible accumulation of cholesteryl esters in macrophages incubated with acetylated lipoproteins. J Cell Biol 82 (3):597–613

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Chapter 10 Methods for Assessing the Effects of LXR Agonists on Macrophage Bacterial Infection Estibaliz Gları´a, Jonathan Matalonga, and Annabel F. Valledor Abstract Macrophages are phagocytic cells that actively engulf and kill microorganisms within a specialized phagolysosomal system. Several pathogenic bacteria, however, actively co-opt host mechanisms and escape from microbial digestion to establish intracellular replication within macrophages. This chapter highlights detailed protocols to measure the effects of the LXR pathway on bacterial infection of murine bone marrow-derived macrophages. Key words LXR, Macrophage, Bacteria, Infection

1

Introduction Macrophages play essential roles in the immune response against pathogens. Upon recognition of pathogen-associated molecular patterns and/or opsonins, they internalize microorganisms, including bacterial cells, through phagocytosis. After engulfment, macrophages kill and digest the internalized material within the phagolysosomal system [1]. Although a wide range of microorganisms are successfully eliminated by phagocytes, several pathogenic bacteria have developed strategies, including the capability to actively invade host cells and escape from microbial digestion within phagolysosomes, to survive within the host. Paradoxically, despite harboring an arsenal of microbicidal tools, macrophages represent a cellular compartment in which many pathogens establish for intracellular replication and subsequent dissemination [2]. As an example, Salmonella enterica serovar Typhimurium (S. Typhimurium) actively promotes its own uptake by macropinocytosis through the use of a type III secretion system that allows the bacterium to inject effectors that target the host cell cytoskeleton [3]. Later on, the bacterium is able to modify the phagosome transforming it into a Salmonella-containing vacuole that supports

Matthew C. Gage and Ine´s Pineda-Torra (eds.), Lipid-Activated Nuclear Receptors: Methods and Protocols, Methods in Molecular Biology, vol. 1951, https://doi.org/10.1007/978-1-4939-9130-3_10, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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bacterial cell survival and replication. Infection and intracellular survival in macrophages is required for full virulence of S. Typhimurium in vivo [4]. Nuclear receptors are a family of ligand-activated transcription factors that control many aspects of physiology. Within this family, liver X receptors (LXRs) are activated by specific oxidized forms of cholesterol (oxysterols) and intermediaries of cholesterol biosynthesis to subsequently regulate the expression of genes involved in lipid and glucose homeostasis and in immune responses [5, 6]. Two LXR isoforms have been described (LXRα and β), and each of them forms heterodimers with retinoid X receptors (RXRs) to positively modulate target gene expression. Recent work from our group has identified a molecular mechanism by which LXR agonists interfere with the capability of S. Typhimurium to infect murine macrophages [7]. This mechanism involves transcriptional activation of the NADase CD38, which translates in reduced intracellular NAD+ levels and interferes with pathogen-induced changes in the F-actin cytoskeleton, limiting the capability of non-opsonized Salmonella to infect macrophages. In this chapter we describe in detail protocols based on the use of flow cytometry and confocal microscopy to measure the effects of the LXR pathway on infection of murine bone marrow-derived macrophages by an invasive S. Typhimurium strain.

2

Materials All the materials must be sterile and endotoxin-free. 1. LXR agonists: T0901317, 25-hydroxycholesterol.

GW3965,

and

2. RXR agonist: LG100268. 3. Dimethyl sulfoxide (DMSO). 4. High-glucose DMEM with L-glutamine and without sodium pyruvate. 5. Fetal bovine serum (FBS). 6. Phosphate-buffered magnesium.

saline

(PBS)

without

calcium/

7. Bacterial liquid growth medium 2xYT: 1.6% Bacto tryptone, 1% Bacto yeast extract, 0.5% NaCl, pH 7. 8. Bacterial liquid growth medium Super Optimal Broth (SOB): 2% Bacto tryptone, 0.5% Bacto yeast extract, 10 mM NaCl, 2.5 mM KCl, 10 mM MgCl2, 10 mM MgSO4, pH 7. 9. Ampicillin.

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10. Transformation buffer 1: 0.1 M RbCl, 50 mM MnCl2·4H2O, 30 mM KAc, 10 mM CaCl2·2H2O, 10% glycerol, pH 5.8. 11. Transformation buffer 2: 0.2 M MOPS, 10 mM RbCl, 75 mM CaCl2·2H2O, 12% glycerol, pH 6.8. 12. 100 mL and 1 L Erlenmeyer flasks. 13. Tissue culture plates (6-well and 24-well). 14. Cell scrapper. 15. 50 mL polypropylene tubes. 16. 1.5 mL polypropylene microtubes. 17. Centrifuge. 18. Hemacytometer. 19. Liquid bath. 20. Incubator at 37
C. 21. Humidified incubator at 37
C, 5% CO2. 22. Horizontal shaker. 23. Petri dishes containing solid LB agar. 24. Bunsen burner. 25. L-Shaped glass spreader. 26. 75% ethanol. 27. Spectrophotometer. 28. Paraformaldehyde (PFA) 1% and 4% in PBS, prepared fresh. 29. Wheat germ agglutinin (WGA) conjugated to Alexa Fluor 488. 30. DAPI (1 μg/mL). 31. Mounting medium. 32. Microscopy slides and coverslips. 33. Milli-Q H2O. 34. Flow cytometer (equipped with an excitation red laser (561 nm) and fluorescence detector at 610/20 nm). 35. Confocal microscope (equipped with lasers exciting at 405 nm, 488 nm, and 561 nm wavelengths).

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Methods All the steps before sample processing must be carried out under sterile conditions.

3.1 Macrophage Plating and Culture

1. Murine bone marrow-derived macrophages should be generated as described [8]. 2. Plate macrophages using the following indications:

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3.1.1 For Flow Cytometry

1. Plate 1.5  106 macrophages per well in 6-well plates with 2 mL of DMEM-10% FBS, and allow them to attach for 2 h in a humidified incubator at 37
C, 5% CO2.

3.1.2 For Confocal Microscopy

1. Plate 2  105 macrophages per well in 24-well plates containing UV-sterilized coverslips. Cover the cells with 1 mL of DMEM-10% FBS, and allow them to attach for 2 h in a humidified incubator at 37
C, 5% CO2.

3.2 Treatment with LXR/RXR Agonists

1. Prepare a stock solution of LXR (or RXR) agonist (1–10 mM in DMSO). Store stock at 80
C. 2. Once the cells have attached to the plates, add LXR/RXR agonists at 1 μM each for the desired period of time (see Note 1). Culture cells in a humidified incubator at 37
C, 5% CO2. For control samples, incubate the cells with DMSO (vehicle) at the same dilution than the one in the samples treated with ligands.

3.3 Obtention of Bacterial Cells Expressing Red Fluorescent Protein (RFP) 3.3.1 Prepare Competent Bacterial Cells for Heat Shock Transformation

This protocol shows steps used for the generation of fluorescent S. Typhimurium strain SL1344. To avoid contamination by other microorganisms, perform all the steps next to a Bunsen burner or inside a biological safety cabinet with laminar flow.

1. Inoculate a bacterial colony in a 100 mL Erlenmeyer flask containing 30 mL of SOB. 2. Grow bacteria overnight at 37
C with horizontal shaking at 250 rpm. 3. Subculture the cells in a 1 L Erlenmeyer flask using a 1:50 dilution in 100 mL of SOB. 4. Grow bacteria at 37
C with horizontal shaking at 250 rpm until the optical density of the culture at 600 nm (OD600) reaches 0.45 (mid-log phase). 5. Distribute the volume to four 50 mL Falcon tubes and keep them on ice for 15 min. 6. Centrifuge for 15 min at 3000  g, 4
C to pellet the bacterial cells. 7. Remove supernatants, and resuspend bacterial cell pellets in 10 mL of transformation buffer 1. 8. Centrifuge for 15 min at 3000  g, 4
C. 9. Remove supernatants and resuspend in 4 mL of transformation buffer 2. 10. Prepare 500 μL aliquots and freeze at 80
C.

LXRs and Macrophage Bacterial Infection 3.3.2 Transform S. Typhimurium Strain SL1344 with pBR.RFP.1 Plasmid Encoding Red Fluorescent Protein (RFP) [9] (See Note 2) by Heat Shock

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1. Mix 100 μL of competent bacteria with 1–10 ng of supercoiled plasmid DNA. For the negative control, use H2O instead of plasmid DNA. 2. Keep on ice for 30 min. 3. Perform the heat shock at 42
C for 1.5 min (in a liquid bath). 4. Put the cells on ice for 5 min. 5. Add 2xYT medium up to 1 mL, and grow the transformed bacterial cells in a 1.5 mL microtube for 1 h at 37
C with horizontal shaking at 250 rpm. 6. Plate 50 μL bacterial growth onto LB agar plates supplemented with 100 μg/mL ampicillin, and allow the colonies to grow for 15 h at 37
C.

3.4 Bacterial Cell Growth

1. Pick a bacterial colony with a sterile pipette tip, and transfer it to a 50 mL tube containing 10 mL 2xYT medium supplemented with 100 μg/mL ampicillin. To obtain a saturated culture, let the bacteria grow for at least 16 h at 37
C with horizontal shaking at 250 rpm. 2. Once at saturation (OD600 ¼ 2), dilute the bacterial cell culture 1:100 by transferring 100 μL to a new 50 mL tube with 10 mL 2xYT, and culture it at 37
C, 250 rpm for 2–3 h (at this time bacterial growth would be in log phase, and bacterial cells would express optimal levels of effectors for invasion). 3. Measure the OD600 of the bacterial cell culture using a spectrophotometer, and estimate the bacterial cell concentration using a standard curve (see Note 3).

3.5 Macrophage Infection

1. Add bacteria to the macrophage culture at the desired multiplicity of infection (MOI) (see Note 4). Let the infection occur for 30 min in an incubator at 37
C, 5% CO2. Also include two types of controls: (a) negative control (noninfected cells) and (b) control for attachment without engulfment (macrophages are incubated with the bacteria for 30 min at 4
C). 2. Place the plates on ice to stop infection and bacterial replication. Remove culture media, and wash the cells three times with 2 mL of ice-cold PBS to remove non-internalized bacteria.

3.6 Process the Cells for the Analysis of Infection 3.6.1 Analysis by Flow Cytometry

1. Scrape cells in 1 mL PBS and transfer them to 1.5 mL microtubes. 2. Centrifuge for 5 min at 200  g, 4
C to pellet the cells. 3. Remove supernatant, and fix the cells in 150 μL 1% PFA for 30 min at room temperature. 4. To measure infection, analyze RFP fluorescence in macrophages by flow cytometry using a 561 nm laser for excitation and a 610/20 nm filter for the detection of emission. Check

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bacterial cell fluorescence by comparing it to a negative control (bacterial cells not transformed with an RFP encoding plasmid). 3.6.2 Analysis by Confocal Microscopy

1. Fix cells on coverslips (inside the cell culture plates) with 300 μL 4% PFA for 30 min at room temperature. 2. Wash cells with 1 mL PBS. 3. Stain cell membranes with 2.5 μg/mL fluorescent WGA for 30 min at room temperature. 4. Stain nuclei with 1 μg/mL DAPI for 5 min at room temperature. 5. Wash cells twice for 5 min with 1 mL PBS. 6. Wash cells with 1 mL mQ H2O. 7. Plate coverslips on microscopy slides with a drop of mounting medium (follow recommendations for the specific mounting medium). 8. Store samples in the dark at 4
C until subsequent analysis by confocal microscopy (see Note 5). 9. For each fluorochrome, collect serial 1 μm z-axis optical images from whole cells using a 63 objective. Use the following lasers: 405 nm (DAPI, nuclei), 488 nm (WGA-Alexa Fluor 488, membranes), and 561 nm (RFP, bacteria).

3.7 Estimation of MOI After the Infection

1. Make dilutions of bacterial culture in 1.5 mL microtubes with 2xYT (see Note 6). 2. Dispense 100 μL of each dilution on LB agar plates (in duplicates), and incubate overnight at 37
C. 3. Count colonies to calculate the exact MOI used in the experiment.

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Notes 1. LXRs form heterodimers with RXRs to activate transcription of their target genes. Agonists for both nuclear receptors can be used (1 μM each) to obtain synergistic effects on induction of gene expression. 2. Transformation with a plasmid which, in addition to fluorescence, confers antibiotic resistance enables the selection of bacteria of interest and avoids contamination by other bacteria. When using antibiotic-resistant colonies, antibiotics should be added to the solid LB agar plates and to the bacterial growth medium.

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3. To obtain a standard curve, make serial dilutions of a saturated bacterial cell culture ranging from the original concentration to 1:100 dilution (e.g., 1:2, 1:4, 1:10, 1:100). Measure OD600 and dispense 100 μL of each dilution on solid LB agar plates. Incubate the plates with bacteria overnight at 37
C. Then count colonies to calculate the bacterial cell concentration, and generate the standard curve by plotting cell concentration vs absorbance. 4. At MOI 5–10, invasive Salmonella will infect 10–40% macrophages within 30 min, and a reduction of infection by LXR agonists can be detected. 5. For optimal results, analysis of infection by confocal microscopy should be performed no longer than 2 weeks after sample preparation. 6. Despite the fact that an estimation of the bacterial MOI is performed before the infection based on the OD600 of the bacterial growth, it is recommended to calculate the exact MOI used in each experiment by plating bacterial cell dilutions on LB agar plates. Usually, dilutions 106 to 107 work best for subsequent counting of viable bacterial colonies on plates.

Acknowledgments This work was supported by a grant from the Spanish Ministry of Economy and Competitivity (SAF2017-89510-R) to A.F. Valledor. Estibaliz Gları´a is supported by a fellowship from the University of Barcelona (Ajuts de Personal Investigador predoctoral en Formacio´, APIF). References 1. Flannagan RS, Cosı´o G, Grinstein S (2009) Antimicrobial mechanisms of phagocytes and bacterial evasion strategies. Nat Rev Microbiol 7:355–366 2. Price JV, Vance RE (2014) The Macrophage Paradox. Immunity 41:685–693 3. Guiney DG, Lesnick M (2005) Targeting of the actin cytoskeleton during infection by Salmonella strains. Clin Immunol 114:248–255 4. Haraga A, Ohlson MB, Miller SI (2008) Salmonellae interplay with host cells. Nat Rev Microbiol 6:53–66 5. Hong C, Tontonoz P (2014) Liver X receptors in lipid metabolism: opportunities for drug discovery. Nat Rev Drug Discov 13:433–444

6. Pascual-Garcı´a M, Valledor AF (2012) Biological roles of liver x receptors in immune cells. Arch Immunol Ther Exp 60:235–249 7. Matalonga J, Glaria E, Bresque M et al (2017) The Nuclear Receptor LXR Limits Bacterial Infection of Host Macrophages through a Mechanism that Impacts Cellular NAD Metabolism. Cell Rep 18:1241–1255 8. Valledor AF, Comalada M, Xaus J et al (2000) The differential time-course of extracellularregulated kinase activity correlates with the macrophage response toward proliferation or activation. J Biol Chem 275:7403–7409 9. Birmingham CL, Smith AC, Bakowski MA et al (2006) Autophagy controls Salmonella infection in response to damage to the Salmonellacontaining vacuole. J Biol Chem 281:11374–11383

Chapter 11 Measuring Apoptotic Cell Engulfment (Efferocytosis) Efficiency Matthew C. Gage Abstract Efferocytosis is the process of recognizing and removing dead and dying cells, performed by a variety of phagocytic cells including macrophages. It has recently been shown that liver X receptor (LXR) signaling in macrophages regulates the expression of important efferocytosis receptors, bridging and signaling molecules. Here we describe a sensitive yet robust efferocytosis assay, optimized to measure bone marrowderived macrophage (BMDM) apoptotic cell engulfment capability. This assay can be applied to genetically or pharmacologically altered BMDMs. Key words Nuclear receptors, Efferocytosis, Apoptosis, Macrophages, BMDM, Jurkats, Engulfment

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Introduction Efferocytosis is the term given to the process of recognizing, engulfing, and processing dead and dying cells [1] which is performed by a variety of phagocytes including macrophages [2]. Macrophages are a heterogeneous population of phagocytotic leukocytes [3] which have high efferocytotic capability [4], shown to engulf dead and dying cells in healthy tissues with high cell turnover such as the bone marrow, spleen, and thymus [5]. Macrophages also perform their efferocytic duties in atherosclerosis [6]— the leading cause of cardiovascular disease [7]—where macrophage efferocytosis capacity in atherosclerotic plaques has been linked to the progression of necrotic cores through, among others, the tyrosine kinase receptor Mer (MerTK) [6]. Macrophages express nuclear receptors, several of which have been linked to efferocytosis activity including PPARγ, PPARδ, RXRα [8], and liver X receptors (LXRs) [9]. LXRs regulate the expression of the important efferocytosis receptor MerTK [10] and several other bridging and signaling molecules [11] directly involved in efferocytosis.

Matthew C. Gage and Ine´s Pineda-Torra (eds.), Lipid-Activated Nuclear Receptors: Methods and Protocols, Methods in Molecular Biology, vol. 1951, https://doi.org/10.1007/978-1-4939-9130-3_11, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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This protocol is based upon original methodology described by Marissa Nadolski and Ed Thorp [12] and has been optimized to provide a simple yet robust assay for measuring efferocytosis efficiency in a relatively small number of cells. This is an ideal feature of an assay, when the subject is a precious commodity such as primary bone marrow-derived macrophages from a rare genetically modified mouse [13], with which you may also want to perform additional experiments with. In this chapter the utilization of 8-well chamber slides is described, which allows for multiple conditions to be tested such as various time points and a range of agonists and concentrations. After addition of Jurkat cells undergoing apoptosis, the BMDMs are fixed and imaged, and the efficiency of efferocytosis is quantified using freely available software.

2

Materials Prepare all solutions using ultrapure water and analytical grade reagents. Prepare and store all reagents at 4
C or 20
C as indicated (unless otherwise noted). Follow all local waste disposal regulations when disposing of waste materials.

2.1 Jurkat Cell Culture

1. Glutamine: 200 nM. 2. Gentamycin: 10 mg/mL. 3. Complete RPMI media: (RPMI, 10% FBS, 1% glutamine, gentamycin 20 μg/mL). 4. Jurkat cells (see Notes 1 and 2). 5. T25 cm2 filter cap flasks. 6. Sterile polystyrene pipettes (5 mL, 10 mL, 25 mL). 7. 70% ethanol. 8. Cell incubator with programmable control of CO2 and temperature.

2.2 Bone MarrowDerived Macrophage Cell Culture

1. Murine bone marrow-derived macrophages (see Note 3). 2. Cell scrapers. 3. Sterile 8-well culture slides. 4. Cell incubator with programmable control of CO2 and temperature.

2.3 Preparation of Apoptotic Cells

1. Complete RPMI media: (RPMI, 10% FBS, 1% glutamine, gentamycin 20 μg/mL). 2. 10 cm sterile TC dish. 3. Calcein AM (1 mg/mL in DMSO) store at 20
C. 4. Warmed PBS 1 37
C.

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5. UV irradiator. 6. Sterile polystyrene pipettes (5 mL, 10 mL, 25 mL). 7. 70% ethanol. 8. Cell incubator with programmable control of CO2 and temperature. 9. 1% paraformaldehyde. 2.4 Efferocytosis Assay

1. Complete DMEM (DMEM, 10% FBS, 1% glutamine, gentamycin 20 μg/mL). 2. Apoptosing Jurkat cell solution (from Subheading 2.3). 3. BMDM cultured on 8-well culture slides. 4. Ice-cold PBS 1. 5. 1% paraformaldehyde. 6. 70% ethanol. 7. Single channel manual pipettes (1 mL, 200 μL). 8. Cell incubator with programmable control of CO2 and temperature.

2.5

Slide Preparation

1. 8-well culture slide removal tool (see Note 4). 2. Fluoromount aqueous mounting media (Sigma). 3. 22  50 mm glass cover slips. 4. Nail varnish.

2.6 Image Acquisition

1. Fluorescent microscope with 20 magnification and GFP filter (excitation wavelength 450–490 nm, emission wavelength 515–565 nm) and digital camera (see Note 5). 2. Image acquisition software (see Note 6).

2.7 Efferocytosis Quantification

3

1. ImageJ image analysis software with grid overlay plugin (https://imagej.nih.gov/ij/download.html).

Methods

3.1 Jurkat Cell Culture

This is your source of apoptotic cells. 1. Grow Jurkat cells in complete RPMI in T25 cm2 filter cap flasks. Count daily with a hemocytometer maintaining cell density at between 2  105 and 1  106 cells/mL (see Note 7). 2. Grow sufficient number of cells as required for the number of experiments you are performing.

3.2 BMDM Cell Culture

1. When BMDM reached 90% confluency (see Note 8), scrape the cells from the TC dish, and pipette up and down several times

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Condition 1: 3 wells per condition (time /agonist etc.)

Condition 2:

Controls BMDM Jurkats only only

Fig. 1 Schematic of 8 well culture slide experimental setup

in a 10 mL pipette to homogenize the cells into a single cell suspension. 2. Count BMDMs on a hemocytometer. 3. Centrifuge cells at 500  g for 10 min. 4. Resuspend cell pellet in complete DMEM to a final concentration of 2.5  105 cells/mL. 5. Seed resuspended BMDM in onto 8-well slides, 250 μL/well (see Note 9 and Fig. 1). 6. Incubate overnight at 37
C, 5% CO2. 3.3 Preparation of Apoptotic Jurkat Cells

1. Count Jurkat cells using a hemocytometer. 2. Centrifuge cells at 500  g for 5 min. 3. Resuspend pellet at 2  106 cells/mL, and plate 20  106 cells (10 mL) onto a 10 cm TC dish. 4. Add 10 μL of Calcein AM solution to the dish, and mix thoroughly by gently pipetting up and down several times using a 10 mL pipette. 5. Incubate for approximately 2 h at 37
C 5% CO2 (see Notes 10 and 11). 6. Collect cells in 15 mL conical tube. 7. Wash dish with 3 mL of Jurkat complete media, and add to the conical tube of Jurkat cells. 8. Centrifuge at 500  g for 5 min (see Note 12 and Fig. 2). 9. Wash cells by removing the supernatant, resuspending the cells pellet in 5 mL of warmed PBS 1X and centrifuging at 500  g for 5 min.

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Fig. 2 Green cell pellet of calcein-AM labelled Jurkat cells

Fig. 3 Calcein-AM labelled, UV-irradiated Jurkat cells with visible blebbing. Original magnification 40

10. Repeat step 9 twice more for a total of three PBS washes. 11. Resuspend the cell pellet in 10 mL of warmed (37
C) Jurkat complete media, and plate onto a 10 cm TC dish. 12. Irradiate cells for 5 min (see Notes 13 and 14). Swirl dish every 60 s to ensure homogeneous exposure of cells to UV light source.

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13. Incubate the irradiated cells at 37
C for approximately 2 h (see Note 15) until approximately 50% of the cells are visibly blebbing (Fig. 3). 14. Collect the cells into a 15 mL conical tube. 15. Centrifuge at 500  g for 5 min. Resuspend the apoptotic cells at 2.5  105 cells/mL in complete DMEM media. 3.4 Efferocytosis Assay

1. Remove complete DMEM media from chamber well slides with BMDM. 2. Add apoptotic Jurkats in a 1:1 ratio to the BMDM (250 μL). 3. Incubate BMDM and apoptotic Jurkats together for various periods of time ranging from 0 to 90 min (see Note 16). 4. Remove media, and immediately wash cells with 250 μL ice-cold PBS. 5. Remove PBS, and repeat ice-cold PBS washes twice more for a total of three washes (see Note 17). 6. Add 250 μL cold 1% PFA solution, and incubate at 4
C in the dark for 10 min. 7. Repeat one more PBS wash step. 8. Add 250 μL PBS to cover each well. Slides can now be stored wet overnight at 4
C in the dark, or proceed immediately to slide preparation.

3.5

Slide Preparation

1. Flick off PBS from the 8-well slides. 2. Remove the 8-well plastic frame using the supplied removal device in accordance with manufacturer’s instructions. 3. Add 3–4 drops of mounting media along the length of the slide. 4. Carefully place glass coverslip onto the slide, taking care not to introduce bubbles (see Note 18). 5. Gently brush a small amount of nail varnish on to the edges of the slide to seal it (see Note 19). 6. Allow varnished slides to dry in the dark at room temperature for a minimum of 1 h (see Note 20).

3.6 Image Acquisition

1. At the microscope, image slides at 20 magnification using the GFP filter in a darkened room. 2. Acquire a minimum of three images per well to ensure fair representation of the sample as shown in Fig. 1 (see Note 21).

3.7 Efferocytosis Quantification

1. Open image using ImageJ software on appropriate computer (see Note 22).

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2. Divide images into nine equal squares with grid overlay plugin as shown in Fig. 4. 3. Count the total number of cells (both the bright and dark adherent BMDMs) in a grid square as shown in Fig. 4. 4. Count the number of (1) bright fluorescent cells (which have engulfed labeled Jurkats) in the same grid square as shown in Fig. 5a, b. 5. Count the number of BMDM cells with attached Jurkats (2) in the same grid square as shown in Fig. 5a, b. 6. Calculate the proportion of total cells performing efferocytosis (efferocytosis efficiency) as % of total cells by using the following equation: ðTotal number of cells=ðNumber of bright cells þ Number of cells with attached JurkatsÞÞ  100

7. Repeat for the remaining four grids as shown in Fig. 4. 8. Calculate the mean efferocytosis efficiency (%) for this image by adding the five resulting percentages together and dividing by 5. 9. Repeat this for each of the three images you have for each well (see Note 23).

Fig. 4 Fluorescent image of bone marrow derived macrophages Image with super-imposed nine square grid. Original magnification 40

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(b)

(a)

Cells performing efferocytosis

Attached Macrophage

(Engulfment)

Calcein-AM labelled apoptotic cell

(Attached)

Fig. 5 (a) Bone marrow derived macrophages before, during and after engulfment of calcein-AM labelled apoptotic Jurkat cells. Original magnification 80, (b) schematic illustrating cells performing efferocytosis through engulfment or attachment

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Notes 1. Jurkats are an immortalized T lymphocyte cell line which are widely available commercially. Follow supplier’s instructions for storage and revival. 2. We have also applied this same apoptosis protocol to BMDMs to investigate the phagocytotic effects of BMDM capability on macrophages (BMDM), in addition to the Jurkat T cell model. 3. BMDM are a common resource. For isolation and culture and storage of BMDM, see ref. 13. 4. The 8-well culture slide removal tool should be supplied by the manufacturer with the 8-well culture slides. 5. We use a Zeiss Axio Vert.A1 fluorescent microscope with a Zeiss Axiocam 503 mono digital camera with Zen Pro acquisition software. 6. We use the freely available Zen lite v2.3 software for viewing our acquired images on different hardware which is available from www.zeiss.com/microscopy/int/products/microscopesofware/zen-lite. 7. Jurkats were counted by staining with Trypan Blue, 1 in 2 dilutions. Using the culture conditions described, Jurkat cells were diluted in fresh media 1 in 5, every 2 days to maintain a maximum cell density of 1  106 cells/mL in T25 flasks standing upright. 8. Using our complete DMEM growth media, this was typically on day 5. 9. Pharmacological treatment can be applied at this point (e.g., an LXR agonist).

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10. This incubation is to ensure maximum incorporation of Calcein AM dye. 11. Avoid bright light when using the Calcein AM (e.g., by switching off tissue culture hood light and covering cells in aluminum foil during transit) to preserve fluorescence. 12. Cell pellet should appear green (Fig. 2), demonstrating incorporation of dye into living cells. 13. For irradiation we use a Hug Flight UV sterilizer (model HF-15151-E). If using different equipment, please follow your local health and safety regulations for UV exposure protection. 14. Remove lid of TC dish so cells get full UV exposure. 15. Check cells every 30 min with a light microscope for blebbing, as the rate of apoptosis induction may depend on the power of your UV lamp and distance of cells from UV source. 16. Efferocytosis efficiency may be influenced by your mutation/ experimental condition—we have found it best to try a range of time points to establish 20–30% baseline efferocytosis and saturation at approximately 80%—which may not necessarily be solely measuring first-time uptake events. 17. We have found that if you are using multiple conditions and/or timepoints, it helps to have more than one experimentalist on hand to ensure the incubations, and washes are performed consistently in a timely fashion. 18. Using forceps, place coverslip gently from one end so that any bubbles that may form can travel through the mounting fluid, escaping to the edge and don’t become trapped. 19. Take care not to move the glass cover slip when applying the varnish, as not to dislodge the fixed cells. 20. For best results we image our slides within 24 h of fixation. 21. See Fig. 1 for our typical slide layout and depiction of image acquisition. 22. There are other commercially available software packages for image analysis which may also be able to quantify your efferocytosis images. However, ImageJ is freely available and ensures maximum usability and audience for this protocol. 23. Our image analysis is performed blindly by our investigators, and the data is grouped after efferocytosis efficiency has been analyzed for each image.

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Acknowledgments This work was supported by grants from British Heart Foundation Project Grant PG/13/10/30000 (IPT) and PG/16/87/32492 (MG). I would like to thank Ed Thorp for his insights into efferocytosis while optimizing this assay and Yu Zhang for image acquisition used in Figs. 4 and 5, and Ine´s Pineda-Torra for equipment use and critical revision of the manuscript. References 1. deCathelineau AM, Henson PM (2003) The final step in programmed cell death: phagocytes carry apoptotic cells to the grave. Essays Biochem 39:105–117 2. Sather S, Kenyon KD, Lefkowitz JB et al (2007) A soluble form of the Mer receptor tyrosine kinase inhibits macrophage clearance of apoptotic cells and platelet aggregation. Blood 109 3. McNelis JC, Olefsky JM (2014) Macrophages, Immunity, and Metabolic Disease. Immunity 41:36–48 4. Li Y, Gerbod-Giannone M-C, Seitz H et al (2006) Cholesterol-induced apoptotic macrophages elicit an inflammatory response in phagocytes, which is partially attenuated by the Mer receptor. J Biol Chem 281:6707–6717 5. Kojima Y, Weissman IL, Leeper NJ (2017) The role of efferocytosis in atherosclerosis. Circulation 135:476–489 6. Thorp E, Cui D, Schrijvers DM et al (2008) Mertk receptor mutation reduces efferocytosis efficiency and promotes apoptotic cell accumulation and plaque necrosis in atherosclerotic lesions of apoe/ mice. Arterioscler Thromb Vasc Biol 28:1421–1428 7. Mendis S, Puska P, Norrving B (2011) Global atlas on cardiovascular disease prevention and

control. World Heal Organ Collab with World Hear Fed World Stroke Organ, 155 8. Korns D, Frasch SC, Fernandez-Boyanapalli R et al (2011) Modulation of macrophage efferocytosis in inflammation. Front Immunol 2:57 9. Edwards PA, Kast HR, Anisfeld AM (2002) BAREing it all: the adoption of LXR and FXR and their roles in lipid homeostasis. J Lipid Res 43:2–12 10. A-Gonzalez N, Bensinger SJ, Hong C et al (2009) Apoptotic cells promote their own clearance and immune tolerance through activation of the nuclear receptor LXR. Immunity 31:245–258 11. Gage MC, Be´cares N, Louie R et al (2018) Disrupting LXRα phosphorylation promotes FoxM1 expression and modulates atherosclerosis by inducing macrophage proliferation. Proc Natl Acad Sci 15(28):E6556–E6565 12. Nadolski M, Thorp E (2009) Standardized tabas laboratory in-vitro efferocytosis engulfment assay. http://www.tabaslab.com/ protocols/ 13. Pineda-Torra I, Gage M, de JA et al (2015) Isolation, culture, and polarization of murine bone marrow-derived and peritoneal macrophages. Methods Mol Biol 1339:101–109

Chapter 12 Methods to Study Monocyte and Macrophage Trafficking in Atherosclerosis Progression and Resolution Ada Weinstock and Edward A. Fisher Abstract Monocytes are circulating cells imperative to the response against pathogens. Upon infection, they are quickly recruited to the affected tissue where they can differentiate into specialized phagocytes and antigenpresenting cells. Additionally, monocytes play a vital role in chronic inflammation, where they can promote and enhance inflammation or induce its resolution. There are two major subsets of monocytes, “inflammatory” and “nonclassical,” which are believed to have distinct functions. In atherosclerosis, both types of monocytes are constantly recruited to lesions, where they contribute to plaque formation and atherosclerosis progression. Surprisingly, these cells can also be recruited to lesions and promote resolution of atherosclerosis. Tracking these cells in various disease stages may inform about the dynamic changes occurring in the inflamed and resolving tissues. In this chapter we will discuss methods for differential labeling of the two monocyte subsets in order to examine their dynamics in inflammation. Key words Monocytes, Macrophages, Methods, Trafficking, Recruitment, Atherosclerosis resolution

1

Introduction The discovery of phagocytes is mainly attributed to Ilya Metchnikoff, who received the Nobel Prize in 1908 for his findings. Metchnikoff discovered that a subtype of white blood cells that were recruited to sites of inflammation was able to engulf foreign bodies in starfish larva. The main observation was that these cells, which were termed “phagocytes,” mediate bacterial clearance through their capabilities to surround and kill other cells [1, 2]. Monocytes are a subset of phagocytes, which are released from bone marrow and extramedulla sites of hematopoiesis, patrol the blood, and can be rapidly recruited to sites of inflammation, such as infection or tissue damage. In tissues, monocytes may differentiate into macrophages and dendritic cells [3, 4]. All of the above mentioned cell types participate in diverse functions, including

Matthew C. Gage and Ine´s Pineda-Torra (eds.), Lipid-Activated Nuclear Receptors: Methods and Protocols, Methods in Molecular Biology, vol. 1951, https://doi.org/10.1007/978-1-4939-9130-3_12, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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pathogen clearance, wound healing, removal of dead cells, and recruitment and activation of adaptive immune cells. In the blood of mice and humans, monocytes represent 4–10% of the leukocytes and can be distinguished from other blood leukocytes through their expression of CD115 (CSF-1R) [5]. Classically, monocytes are divided into two major subsets, which have been described in multiple species and have distinct characteristics. In mice, classical monocytes are defined as CCR2+CX3CR1+Ly6chi and are commonly referred to as “inflammatory monocytes,” whereas nonclassical monocytes are CCR2
CX3CR1++CCR5+Ly6clo [6]. In humans, the equivalent subpopulations are CD14++CD16
and CD14+CD16+, respectively [7]. The crucial roles of monocytes and monocyte-derived cells in homeostasis and inflammation prompted the interest in understanding their trafficking in vivo. Early studies used the intravenous injection of India ink, which was believed to be taken up primarily by monocytes. With that technique Ebert and Florey reported the recruitment of monocytes to sites of injury and their differentiation into macrophages in rabbit ears [8]. It was later shown that monocytes can be labeled with 3H-thymidine, a radioactive nucleotide that incorporates into newly synthesized DNA. In their kinetic studies, van Furth and Cohn demonstrated that 60–80% of blood monocytes can be radioactively labeled after either one intravenous or four intramuscular injections of 3H-thymidine in mice [9], with slightly different labeling kinetics; intravenous administration resulted in a peak of labeled blood monocytes after 48 h, whereas with intramuscular injections, labeling in the blood peaked 60 h postinjection. Radiolabeling was also performed on leukocytes ex vivo, after which the cells were introduced back to the model animals. In one such study, labeled cells were introduced back into pigeons, to examine the origin of cells in the atherosclerotic plaque [10]. This study showed that most of the cells in early lesions are of monocyte/macrophage origin, whereas mature lesions are more complex and contain smooth muscle cells, extracellular matrix, and adaptive immune cells. Adoptive transfer of cells was also performed later without the need for radiolabeling but with the use of genetic differences between the transferred cells and the recipient animal [11]. For instance, monocytes of male mice were transferred to female recipients in order to quantify the recruitment of monocytes to atherosclerotic lesions using a polymerase chain reaction (PCR) to the Y-chromosome gene sry. In the past three decades, many other genetic-based models have been introduced to track leukocytes and monocytes in particular. The use of the congenic pan-leukocyte markers CD45.1 and CD45.2 enabled the tracking of adoptively transferred hematopoietic cells in the recipient mouse using PCR or flow cytometry [12].

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An important advance in the field was the generation of mice that allow fluorescent tagging of monocytic populations, such as the CX3CR1-GFP [13] and the CCR2-RFP [14] mice. The CX3CR1-GFP mouse is widely used and enables the tracking of all cells that express CX3CR1, among them monocytes. However, since CX3CR1 is frequently downregulated in monocytes that are differentiating into macrophages, another reporter mouse was recently described: the CD68-GFP mouse [15]. Additionally, a CX3CR1 fate-mapping mouse was generated, enabling the constitutive and, in a later development, conditional labeling of cells that had expressed CX3CR1 at any given time [16]. To research the trafficking of specific monocyte populations without the need for genetic manipulations or cell transfer, researchers have taken advantage of the innate properties of these cells to take up a wide variety of easily-labeled substances. One approach to fluorescently label phagocytes is via the administration of liposomes containing fluorescent dyes, such as DiI [17, 18]. Liposomes are bilayered phospholipid spheres that are phagocytosed by specialized cells, such as macrophages, dendritic cells, and monocytes. Encapsulation of a fluorescent dye in liposomes results in the fluorescent labeling of its ingesting cell, thus preferentially marking phagocytes. The same cellular property is also utilized for the specific depletion of phagocytes, using liposomes loaded with clodronate [19], since phagocytes that ingest the clodronate-liposomes undergo apoptosis. Specifically, monocytes are eliminated from the circulation with maximal depletion at 18 h post intravenous injection of clodronate-liposomes. Following clodronate-liposome treatment, Ly6Chi cells begin to reappear in the blood after 2 days, while Ly6Clo monocytes reappear only after 7 days [17]. Sunderko¨tter and colleagues depleted all monocyte subsets with clodronateliposomes and reinjected the mice with DiI-liposomes after 2 days, to preferentially label the newly formed Ly6Chi cells. With this method they have showed that Ly6Chi monocytes can differentiate into Ly6Clo cells. We and others have used fluorescent latex bead injection following monocyte depletion with clodronate-liposomes to mark circulating monocytes [6, 20]. Interestingly, administration of these inert latex beads without prior depletion of circulating monocytes results in differential labeling of Ly6Clo monocytes [21]. Hence, depending on the time point after clodronate depletion of monocytes, the fluorescent beads will label preferentially Ly6Chi or Ly6Clo monocyte subsets, and we and others have applied these protocols to track monocyte entry and the subsequent fate of monocyte-derived cells in atherosclerotic plaques [6, 22, 23] (Fig. 1a and 1b). Differential labeling of Ly6chi monocytes can also be achieved by an approach that does not require clodronate treatment. Since

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A

EdU labeling efficiency (-72 hours)

Beads labeling efficiency (48 hours pre harvest)

Harvest baseline group and measure baseline EdU (0 hours)

EdU injection to label monocytes that accumulate during plaque formation (-96 hours)

Bead injection to label monocytes recruited during plaque resolution (72 hours pre harvest)

Atherosclerosis progression B

Beads

Harvest resolution groups and measure both EdU and beads

Resolution phase C

EdU

Fig. 1 Detection of beads and EdU in atherosclerotic plaques. (a) Suggested labeling timing with EdU and beads for examining monocyte trafficking during the resolution of atherosclerosis (modified from ref. 23). EdU is injected 4–5 days prior to the induction of resolution, to mark cells recruited during plaque formation. Comparing the abundance of EdU-positive cells between baseline and the resolution group will characterize the subset of monocytes that are leaving the plaque. Beads are injected 2–3 days prior to harvest, to label monocytes that are newly recruited specifically during lesion resolution. (b, c) Frozen sections of aortic arches were imaged in the (b) 488 channel to visualize latex beads or in the (c) 647 channel to visualize EdU (see Note 1)

their progenitors in the bone marrow are highly proliferative, this enables their labeling by compounds that intercalate into newly synthesized DNA, such as EdU or BrdU [24, 25] (Fig. 1c). Beginning at 6 h following BrdU treatment, labeled Ly6chi monocytes appear in the circulation and plateau at peak labeling 12–24 h posttreatment [25]. It should be noted that traditionally EdU and BrdU have been used to mark proliferating cells in tissues, highlighting the importance of the choice of the time point for analysis after injection of these nucleotide mimics. For example, if observations of macrophages in peripheral tissues, such as atherosclerotic plaques, are made within 6 h of injection, positively labeled macrophages will likely be the result of proliferation in situ. On the other hand, if the time point is later, positive cells in tissues will be the combination of newly recruited cells (with EdU or BrdU incorporation occurring in bone marrow precursors) and local proliferation. As noted in the laboratory procedures below, fortunately, it is relatively easy to determine if the observed labeling was

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from recruitment of circulating monocytes or from the proliferation of monocyte-derived cells in the tissue. This chapter will discuss methods of monocyte labeling and subsequent cell tracking (entry into and disappearance from) atherosclerotic plaques, focusing on results we have published (e.g., [20, 23, 26]). Specifically, techniques of labeling Ly6chi or Ly6clo monocytes distinctly and the subsequent disappearance of macrophages derived from these cells (taken to indicate cell exit from plaques) will be presented. An illustration of the value of these methods is our recent demonstration that resolution of atherosclerosis, a process referred to often as regression, requires the recruitment of Ly6chi monocytes in a CCR2-dependent manner [22]. These studies employed one of the protocols described below for the dual labeling of circulating monocytes at different time points with latex beads and EdU in order to differentiate between monocyte recruitment into and macrophage disappearance from plaques after the resolution of atherosclerosis was induced [23]. The ability to track monocytes and cells derived from them has provided major insights into the study of the innate immune system in various contexts such as inflammatory diseases, including atherosclerosis, and the homeostatic regulation of tissue macrophages. The protocols described are relatively easy to implement, and we expect continued progress in their application by refinements in existing approaches as well as by the development of new ones. Using the approach described herein, the EdU will label monocytes that will enter the plaque during the progression of the disease (Fig. 1a). This method allows the quantification of the EdUþ cell abundance in plaques, after which one can compare between baseline plaques (plaques not exposed to resolving conditions) and plaques that underwent atherosclerosis resolution. This comparison will indicate whether monocytes and cells derived from circulating monocytes (such as macrophages and dendritic cells) that were recruited during the progression phase had disappeared from the plaque during the resolution phase, as well as to quantitatively estimate the changes. Furthermore, with this method, because of when they are administered, fluorescent beads will label monocytes that are newly recruited to the plaque only in the phase of atherosclerosis resolution, which we know continues to occur even under conditions favorable to resolve plaques [27]. Importantly, alterations in the time of EdU or bead injection should be carefully considered, as discussed in the Introduction. For example, EdU injection