Mouse Models of Innate Immunity: Methods and Protocols [2nd ed.] 978-1-4939-9166-2, 978-1-4939-9167-9

Table of contents :
Front Matter ....Pages i-xi
CRISPR/Cas9-Assisted Genome Editing in Murine Embryonic Stem Cells (Artiom Gruzdev, Greg J. Scott, Thomas B. Hagler, Manas K. Ray)....Pages 1-21
Genome Editing in Mouse Embryos with CRISPR/Cas9 (Greg J. Scott, Artiom Gruzdev)....Pages 23-40
Derivation of Macrophages from Mouse Bone Marrow (Beckley K. Davis)....Pages 41-55
Bone Marrow-Derived Dendritic Cells (Kelly Roney)....Pages 57-62
Quantification and Visualization of Neutrophil Extracellular Traps (NETs) from Murine Bone Marrow-Derived Neutrophils (Linda Vong, Philip M. Sherman, Michael Glogauer)....Pages 63-73
In Vitro Differentiation of Effector CD4+ T Helper Cell Subsets (Kaitlin A. Read, Michael D. Powell, Bharath K. Sreekumar, Kenneth J. Oestreich)....Pages 75-84
Generation and Culture of Mouse Embryonic Fibroblasts (Yee Sun Tan, Yu L. Lei)....Pages 85-91
Isolation of Tumor-Infiltrating Lymphocytes by Ficoll-Paque Density Gradient Centrifugation (Yee Sun Tan, Yu L. Lei)....Pages 93-99
Depletion and Reconstitution of Macrophages in Mice (Lisa K. Kozicky, Laura M. Sly)....Pages 101-112
Microfluidic Platform to Quantify Neutrophil Migratory Decision-Making (Brittany P. Boribong, Amina Rahimi, Caroline N. Jones)....Pages 113-122
A Vector Suite for the Overexpression and Purification of Tagged Outer Membrane, Periplasmic, and Secreted Proteins in E. coli (Michael A. Casasanta, Daniel J. Slade)....Pages 123-138
Mouse Model of Staphylococcus aureus Skin Infection (Natalia Malachowa, Scott D. Kobayashi, Jamie Lovaglio, Frank R. DeLeo)....Pages 139-147
Systemic Listeria monocytogenes Infection as a Model to Study T Helper Cell Immune Responses (Veronica M. Ringel-Scaia, Michael D. Powell, Kaitlin A. Read, Irving C. Allen, Kenneth J. Oestreich)....Pages 149-160
Endotoxin-Induced Uveitis in Rodents (Umesh C. S. Yadav, Kota V. Ramana)....Pages 161-168
Using Klebsiella pneumoniae to Model Acute Lung Inflammation in Mice (Dylan K. McDaniel, Irving C. Allen)....Pages 169-180
Assessment of Survival and Replication of Brucella spp. in Murine Peritoneal Macrophages (Clayton C. Caswell)....Pages 181-189
Influenza-Mediated Lung Infection Models (Charles E. McGee, Christopher J. Sample, Brita Kilburg-Basnyat, Kristin A. Gabor, Michael B. Fessler, Kymberly M. Gowdy)....Pages 191-205
Adoptive Transfer Colitis (Kristin Eden)....Pages 207-214
The Azoxymethane/Il10−/− Model of Colitis-Associated Cancer (CAC) (Aaron Rothemich, Janelle C. Arthur)....Pages 215-225
Modeling Autism-Related Disorders in Mice with Maternal Immune Activation (MIA) (Catherine R. Lammert, John R. Lukens)....Pages 227-236
Toxoplasma gondii as a Model of In Vivo Host-Parasite Interactions (Sheryl Coutermarsh-Ott)....Pages 237-247
Sepsis Induced by Cecal Ligation and Puncture (Wei Gong, Haitao Wen)....Pages 249-255
Back Matter ....Pages 257-258

Citation preview

Methods in Molecular Biology 1960

Irving C. Allen Editor

Mouse Models of Innate Immunity Methods and Protocols Second Edition

Methods

in

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

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

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

Mouse Models of Innate Immunity Methods and Protocols Second Edition

Edited by

Irving C. Allen Department of Biomedical Sciences, Virginia-Maryland Regional College, Blacksburg, VA, USA

Editor Irving C. Allen Department of Biomedical Sciences Virginia-Maryland Regional College Blacksburg, VA, USA

ISSN 1064-3745     ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-4939-9166-2    ISBN 978-1-4939-9167-9 (eBook) https://doi.org/10.1007/978-1-4939-9167-9 Library of Congress Control Number: 2019932790 © 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. Cover Figure: (Figure 6F from Chapter 1) Post-natal day 14 chimeric F0 founder mice generated following CRISPR/ Cas9 genetic modification. Contributed by Artiom Gruzdev and colleagues. 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 Humans are exposed to millions of potentially dangerous pathogens on a daily basis. Our innate and adaptive immune systems work together to evade infection and minimize the impact of contact, ingestion, and inhalation of these agents. While the adaptive immune system is critical for providing a pathogen-specific immune response and long-lasting memory, the innate immune system provides the initial response during the critical first hours and days following exposure to new pathogens. The innate immune system is evolutionarily conserved, with elements found across vertebrates, invertebrates, and plant species. This is particularly true in terms of pattern recognition receptors and their respective signaling cascades that are responsible for the recognition of specific pathogen-associated or damageassociated molecular patterns. These receptors drive the molecular and cellular responses that enable effective host defense against extracellular and intracellular pathogens, damage, and cellular stress. The ultimate goal of biomedical research is to improve the health and well-being of human patients. Because many of the elements associated with the innate immune system are evolutionarily conserved across species in terms of both structure and function, mouse models have become a preferred human surrogate or complementary model for clinical studies. The readily available assortment of genetically modified mouse strains provides potent tools to define the complex interactions associated with innate immune system function and host–microbe interactions. Indeed, advances in mouse models have occurred in unison with progress in human clinical studies, which together have had tremendous impacts on our understanding of the innate immune system across species. In this second edition of Mouse Models of Innate Immunity, we have again assembled a highly diverse and well-regarded group of active researchers with extensive experience in immunology, microbiology, genetics, and animal models. Similar to the other volumes in the Methods in Molecular Biology series, these contributors have provided a unique group of highly detailed protocols to aid in the design and execution of experiments to fully evaluate essential elements associated with the innate immune response. The emphasis of this second edition has been placed on in vitro, ex vivo, and in vivo mouse models that accurately mimic physiologically and clinically relevant disease processes. The first half of the book contains protocols that focus on assessments of specific cells and/or pathogens that are critical for understanding innate immune function. These in  vitro and ex  vivo studies are designed to provide simplified model systems to evaluate novel hypotheses and deduce mechanistic insight, without the complexity and confounding factors commonly associated with in vivo studies. Building beyond these simplified models, the second half of the book provides robust protocols that are highly useful to evaluate innate immune system function and characterize phenotypes in complex in  vivo model systems. These in vivo protocols describe methods to evaluate innate immune function in the skin, eye, lung, gut, and systemically. In addition to our focus on infectious disease models, in this

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second edition we have also included several new protocols associated with autism, cancer, microfluidics platforms, and CRISPR systems. Thus, it is my genuine hope and expectation that the second edition of Mouse Models of Innate Immunity will benefit the biomedical research community by p ­roviding expert insight and protocols that will allow investigators at all levels to rigorously plan, implement, and assess mechanisms associated with the innate immune response. Blacksburg, VA, USA

Irving C. Allen

Contents Preface. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .   v Contributors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .   ix   1 CRISPR/Cas9-Assisted Genome Editing in Murine Embryonic Stem Cells���������  1 Artiom Gruzdev, Greg J. Scott, Thomas B. Hagler, and Manas K. Ray   2 Genome Editing in Mouse Embryos with CRISPR/Cas9 �����������������������������������  23 Greg J. Scott and Artiom Gruzdev   3 Derivation of Macrophages from Mouse Bone Marrow ��������������������������������������� 41 Beckley K. Davis   4 Bone Marrow-Derived Dendritic Cells����������������������������������������������������������������� 57 Kelly Roney   5 Quantification and Visualization of Neutrophil Extracellular Traps (NETs) from Murine Bone Marrow-Derived Neutrophils������������������������������������������������� 63 Linda Vong, Philip M. Sherman, and Michael Glogauer   6 In Vitro Differentiation of Effector CD4+ T Helper Cell Subsets������������������������� 75 Kaitlin A. Read, Michael D. Powell, Bharath K. Sreekumar, and Kenneth J. Oestreich   7 Generation and Culture of Mouse Embryonic Fibroblasts ����������������������������������� 85 Yee Sun Tan and Yu L. Lei   8 Isolation of Tumor-Infiltrating Lymphocytes by Ficoll-­Paque Density Gradient Centrifugation��������������������������������������������������������������������������������������� 93 Yee Sun Tan and Yu L. Lei   9 Depletion and Reconstitution of Macrophages in Mice �������������������������������������� 101 Lisa K. Kozicky and Laura M. Sly 10 Microfluidic Platform to Quantify Neutrophil Migratory Decision-Making �������� 113 Brittany P. Boribong, Amina Rahimi, and Caroline N. Jones 11 A Vector Suite for the Overexpression and Purification of Tagged Outer Membrane, Periplasmic, and Secreted Proteins in E. coli�������������������������������������� 123 Michael A. Casasanta and Daniel J. Slade 12 Mouse Model of Staphylococcus aureus Skin Infection������������������������������������������ 139 Natalia Malachowa, Scott D. Kobayashi, Jamie Lovaglio, and Frank R. DeLeo 13 Systemic Listeria monocytogenes Infection as a Model to Study T Helper Cell Immune Responses�������������������������������������������������������������������������������������� 149 Veronica M. Ringel-Scaia, Michael D. Powell, Kaitlin A. Read, Irving C. Allen, and Kenneth J. Oestreich 14 Endotoxin-Induced Uveitis in Rodents �������������������������������������������������������������� 161 Umesh C. S. Yadav and Kota V. Ramana 15 Using Klebsiella pneumoniae to Model Acute Lung Inflammation in Mice���������� 169 Dylan K. McDaniel and Irving C. Allen

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16 Assessment of Survival and Replication of Brucella spp. in Murine Peritoneal Macrophages������������������������������������������������������������������������������������������������������ 181 Clayton C. Caswell 17 Influenza-Mediated Lung Infection Models�������������������������������������������������������� 191 Charles E. McGee, Christopher J. Sample, Brita Kilburg-Basnyat, Kristin A. Gabor, Michael B. Fessler, and Kymberly M. Gowdy 18 Adoptive Transfer Colitis������������������������������������������������������������������������������������ 207 Kristin Eden 19 The Azoxymethane/Il10 −/− Model of Colitis-Associated Cancer (CAC) ������������ 215 Aaron Rothemich and Janelle C. Arthur 20 Modeling Autism-Related Disorders in Mice with Maternal Immune Activation (MIA)������������������������������������������������������������������������������������������������ 227 Catherine R. Lammert and John R. Lukens 21 Toxoplasma gondii as a Model of In Vivo Host-Parasite Interactions�������������������� 237 Sheryl Coutermarsh-Ott 22 Sepsis Induced by Cecal Ligation and Puncture�������������������������������������������������� 249 Wei Gong and Haitao Wen Index������������������������������������������������������������������������������������������������������������������������ 257

Contributors Irving C. Allen  •  Department of Biomedical Sciences, Virginia-Maryland Regional College, Blacksburg, VA, USA Janelle C. Arthur  •  Department of Microbiology and Immunology, University of North Carolina at Chapel Hill, Chapel Hill, NC, USA; Lineberger Comprehensive Cancer Center, University of North Carolina at Chapel Hill, Chapel Hill, NC, USA; Center for Gastrointestinal Biology and Disease, University of North Carolina at Chapel Hill, Chapel Hill, NC, USA Brittany P. Boribong  •  Genetics, Bioinformatics, and Computational Biology, Virginia Tech, Blacksburg, VA, USA Michael A. Casasanta  •  Department of Biochemistry, Virginia Polytechnic Institute, State University, Blacksburg, VA, USA Clayton C. Caswell  •  Department of Biomedical Sciences and Pathobiology, Center for One Health Research, Virginia Maryland College of Veterinary Medicine, Virginia Tech, Blacksburg, VA, USA Sheryl Coutermarsh-Ott  •  Department of Biomedical Sciences and Pathobiology, Virginia-Maryland College of Veterinary Medicine, Virginia Tech, Blacksburg, VA, USA Beckley K. Davis  •  Department of Biology, Franklin & Marshall College, Lancaster, PA, USA Frank R. DeLeo  •  Laboratory of Bacteriology, Rocky Mountain Laboratories, National Institute of Allergy and Infectious Diseases, National Institutes of Health, Hamilton, MT, USA Kristin Eden  •  Department of Basic Science Education, Virginia Tech Carilion School of Medicine, Roanoke, VA, USA Michael B. Fessler  •  Immunity, Inflammation, and Disease Laboratory, National Institute of Environmental Health Sciences, National Institutes of Health, Research Triangle Park, NC, USA Kristin A. Gabor  •  Immunity, Inflammation, and Disease Laboratory, National Institute of Environmental Health Sciences, National Institutes of Health, Research Triangle Park, NC, USA Michael Glogauer  •  Faculty of Dentistry, University of Toronto, Toronto, ON, Canada Wei Gong  •  Department of Hepatobiliary Surgery and Liver Transplantation, Shandong Provincial Hospital Affiliated to Shandong University, Jinan, China Kymberly M. Gowdy  •  Department of Pharmacology and Toxicology, Brody School of Medicine, East Carolina University, Greenville, NC, USA Artiom Gruzdev  •  Knockout Mouse Core, Reproductive and Developmental Biology Laboratory, National Institute of Environmental Health Sciences, National Institutes of Health, Research Triangle Park, NC, USA Thomas B. Hagler  •  Knockout Mouse Core, Reproductive and Developmental Biology Laboratory, National Institute of Environmental Health Sciences, National Institutes of Health, Research Triangle Park, NC, USA

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Caroline N. Jones  •  Department of Biological Sciences, Virginia Tech, Blacksburg, VA, USA Brita Kilburg-Basnyat  •  Department of Pharmacology and Toxicology, Brody School of Medicine, East Carolina University, Greenville, NC, USA Scott D. Kobayashi  •  Laboratory of Bacteriology, Rocky Mountain Laboratories, National Institute of Allergy and Infectious Diseases, National Institutes of Health, Hamilton, MT, USA Lisa K. Kozicky  •  Division of Gastroenterology, Department of Pediatrics, BC Children’s Hospital Research Institute, University of British Columbia, Vancouver, BC, Canada Catherine R. Lammert  •  Department of Neuroscience, Center for Brain Immunology and Glia, University of Virginia, Charlottesville, VA, USA; Graduate Program in Neuroscience, University of Virginia, Charlottesville, VA, USA Yu L. Lei  •  Department of Periodontics and Oral Medicine, University of Michigan School of Dentistry, Ann Arbor, MI, USA; University of Michigan Rogel Cancer Center, Ann Arbor, MI, USA; Department of Otolaryngology—Head and Neck Surgery, University of Michigan Health System, Ann Arbor, MI, USA Jamie Lovaglio  •  Rocky Mountain Veterinary Branch, Rocky Mountain Laboratories, National Institute of Allergy and Infectious Diseases, National Institutes of Health, Hamilton, MT, USA John R. Lukens  •  Department of Neuroscience, Center for Brain Immunology and Glia, University of Virginia, Charlottesville, VA, USA; Graduate Program in Neuroscience, University of Virginia, Charlottesville, VA, USA Natalia Malachowa  •  Laboratory of Bacteriology, Rocky Mountain Laboratories, National Institute of Allergy and Infectious Diseases, National Institutes of Health, Hamilton, MT, USA Dylan K. McDaniel  •  Department of Biomedical Sciences and Pathobiology, Virginia-­ Maryland College of Veterinary Medicine, Virginia Tech, Blacksburg, VA, USA Charles E. McGee  •  Duke Human Vaccine Institute, Duke University Medical Center, Durham, NC, USA Kenneth J. Oestreich  •  Virginia Tech Carilion Research Institute, Roanoke, VA, USA; Department of Biomedical Sciences and Pathobiology, Virginia-Maryland College of Veterinary Medicine, Virginia Tech, Blacksburg, VA, USA; Virginia Tech Carilion School of Medicine, Roanoke, VA, USA Michael D. Powell  •  Virginia Tech Carilion Research Institute, Roanoke, VA, USA; Graduate Program in Translational Biology, Medicine, and Health, Virginia Tech, Blacksburg, VA, USA Amina Rahimi  •  Department of Biochemistry, Virginia Tech, Blacksburg, VA, USA Kota V. Ramana  •  Department of Biochemistry and Molecular Biology, University of Texas Medical Branch, Galveston, TX, USA Manas K. Ray  •  Knockout Mouse Core, Reproductive and Developmental Biology Laboratory, National Institute of Environmental Health Sciences, National Institutes of Health, Research Triangle Park, NC, USA Kaitlin A. Read  •  Virginia Tech Carilion Research Institute, Roanoke, VA, USA Veronica M. Ringel-Scaia  •  Department of Biomedical Sciences and Pathobiology, Virginia-Maryland College of Veterinary Medicine, Virginia Tech, Blacksburg, VA, USA; Graduate Program in Translational Biology, Medicine, and Health, Virginia Tech, Blacksburg, VA, USA

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Kelly Roney  •  Rho, Chapel Hill, NC, USA Aaron Rothemich  •  Department of Microbiology and Immunology, University of North Carolina at Chapel Hill, Chapel Hill, NC, USA Christopher J. Sample  •  Duke Human Vaccine Institute, Duke University Medical Center, Durham, NC, USA Greg J. Scott  •  Knockout Mouse Core, Reproductive and Developmental Biology Laboratory, National Institute of Environmental Health Sciences, National Institutes of Health, Research Triangle Park, NC, USA Philip M. Sherman  •  Cell Biology Program, Research Institute, Hospital for Sick Children, University of Toronto, Toronto, ON, Canada; Faculty of Dentistry, University of Toronto, Toronto, ON, Canada Daniel J. Slade  •  Department of Biochemistry, Virginia Polytechnic Institute, State University, Blacksburg, VA, USA Laura M. Sly  •  Division of Gastroenterology, Department of Pediatrics, BC Children’s Hospital Research Institute, University of British Columbia, Vancouver, BC, Canada Bharath K. Sreekumar  •  Virginia Tech Carilion Research Institute, Roanoke, VA, USA; Graduate Program in Translational Biology, Medicine, and Health, Virginia Tech, Blacksburg, VA, USA Yee Sun Tan  •  Department of Periodontics and Oral Medicine, University of Michigan School of Dentistry, Ann Arbor, MI, USA; University of Michigan Rogel Cancer Center, Ann Arbor, MI, USA Linda Vong  • Cell Biology Program, Research Institute, Hospital for Sick Children, University of Toronto, Toronto, ON, Canada Haitao Wen  •  Department of Microbial Infection and Immunity, The Ohio State University College of Medicine, Columbus, OH, USA; Infectious Disease Institute, The Ohio State University College of Medicine, Columbus, OH, USA Umesh C. S. Yadav  •  Department of Biochemistry and Molecular Biology, University of Texas Medical Branch, Galveston, TX, USA

Chapter 1 CRISPR/Cas9-Assisted Genome Editing in Murine Embryonic Stem Cells Artiom Gruzdev, Greg J. Scott, Thomas B. Hagler, and Manas K. Ray Abstract The study of gene function in normal human physiology and pathophysiology is complicated by countless factors such as genetic diversity (~98 million SNPs identified in the human genome as of June 2015), environmental exposure, epigenetic imprinting, maternal/in utero exposure, diet, exercise, age, sex, socioeconomic factors, and many other variables. Inbred mouse lines have allowed researchers to control for many of the variables that define human diversity but complicate the study of the human genome, gene/ protein function, cellular and molecular pathways, and countless other genetic diseases. Furthermore, genetically modified mouse models enable us to generate and study mice whose genomes differ by as little as a single point mutation while controlling for non-genomic variables. This allows researchers to elucidate the quintessential function of a gene, which will further not only our scientific understanding, but provide key insight into human health and disease. Recent advances in CRISPR/Cas9 genome editing have revolutionized scientific protocols for introducing mutations into the mammalian genome. The ensuing chapter provides an overview of CRISPR/Cas9 genome editing in murine embryonic stem cells for the generation of genetically modified mouse models. Key words Genetically modified mice, Embryonic stem cells, Gene-targeting, CRISPR/Cas9, Embryonic stem cell transfection, Embryonic stem cell screening, Blastocyst isolation, Microinjection, Chimeric mice, Germline transmission

1  Introduction Gene targeting, whether to disrupt a gene, generate a genetic variant, or to create a completely novel locus, has revolutionized mammalian research. Genetically modified mouse models are now so widely utilized in scientific research that it is easy to forget that only 30 years ago, the first “knockout” mice were generated from genetically modified embryonic stem (ES) cells [1, 2]. In 2001, Mario Capecchi, who shared the 2007 Nobel Prize in Physiology or Medicine with Oliver Smithies and Martin Evans, summarized the decades of research that led to the first “knockout mouse” [3, 4].

Irving C. Allen (ed.), Mouse Models of Innate Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1960, https://doi.org/10.1007/978-1-4939-9167-9_1, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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Soon after the first genetically modified mouse lines were generated, a conceptual blueprint was followed by other research groups to generate targeted mutations in the genome to study gene/protein function and the noncoding genome and model human diseases with underlying genetic causes [5]. The general process can be summarized into six distinct steps: (a) identify the gene/locus of interest with corresponding genomic nucleotide sequence; (b) generate a targeting vector/plasmid with sequence homologous to the endogenous locus flanking the desired exogenous genetic payload, including a positive selection marker; (c) introduce the desired mutation into the genome of the ES cells by transfecting them with linearized targeting vector; (d) inject the genetically modified ES cells into blastocyst-stage embryos and transfer them into pseudopregnant surrogate female mice; (e) breed resulting F0 chimeric founders to identify stable germline transmission of the ES cells and the genetically modified allele of interest; and (f) breed the F1 germline stable offspring as necessary to generate the desired experimental cohort for phenotypic study. In 2002, the first high-quality draft of the mouse genome was published by the Mouse Genome Sequencing Consortium [6]. Online genome databases greatly simplified the first step: identifying the target genomic sequence. Over the next 5 years, countless research groups generated targeted mutations within the mouse genome in ES cells and created mouse models to study the in vivo phenotypic consequences. By 2007, the International Knockout Mouse Consortium was launched to individually disrupt every gene in the mouse genome and to make those lines available to researchers worldwide [7]. The ease and efficiency of gene targeting continued to improve. Commercial availability of robust high-fidelity polymerases and recombineering with commercially available bacterial artificial chromosomes (BACs) containing large ~100–200 kb fragments of the mouse genome simplified the subcloning steps necessary to generate traditional targeting vectors [8, 9]. Specialized culture conditions supplemented with recombinant leukemia inhibitory factor (LIF) and dual inhibition of MEK (PD0325901) and GSK3 (CHIR99021) improved ES cell quality and increased germline transmission rates [10, 11]. However, the underlying principal, and limitations, remained largely the same; exogenous linear DNA was transfected into ES cells to undergo homologous recombination, essentially a repair of the double-stranded break of the linear targeting vector [12]. The rarity of on-target homologous recombination events required the use of a genome integrated positive selection marker, such as neomycin phosphotransferase (NeoR; G418/Geneticin resistance) or puromycin N-acetyltransferase (PAC; puromycin resistance). Targeting efficiency was further improved by negative selection markers like thymidine kinase or diphtheria toxin A fragment, but random genome integration of

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the targeting vector remained common, and, in practice, the negative selection markers conveyed only a modest enrichment for ESC clones with on-target homologous recombination [13]. Zinc finger nuclease (ZFN), transcription activator-like effector nucleases (TALEN), and clustered regularly interspaced short palindromic repeats (CRISPR) are all targetable nuclease systems that have shifted the targeting aspect from the exogenous linearized targeting vector/plasmid to the endogenous target locus [14–16]. Although ZFNs and TALENs predate the use of CRISPR in eukaryotic cells by several years, it was not until CRISPR/Cas9 in 2013 that mouse genome editing significantly shifted from the traditional paradigm of gene targeting of the late 1980s to utilizing targetable nucleases for genome editing (see Note 1). In this chapter, we summarize CRISPR/Cas9-assisted ES cell genome editing techniques involved in generating genetically modified ES cells and mice. We highlight key similarities and differences between traditional gene targeting and CRISPR/Cas9-­ assisted targeting. In addition, since the protocols for virtually all other CRISPR-related techniques are rapidly evolving, we reference some of the latest promising trends related to genome editing in ES cells.

2  Materials 2.1  Genome Editing Vectors

1. Genomic DNA from ES cell line. 2. Oligonucleotide/primers screening.

for

vector

generation

and

3. High-fidelity PCR reagents and thermal cycler. 4. Subcloning reagents. 5. Amplicon cloning vector, such as pBlueScript, pUC57, or pCR2.1. 6. Chemical or electro-competent DH5α E. coli cells. 7. Gel electrophoresis apparatus and reagents (agarose, ethidium bromide, DNA molecular markers, 1× TAE, or 1× TBE). 8. Plasmid Mini- and Midi-prep kit. 9. DNA quantification instrument (e.g., UV spectrophotometer, Thermo Fisher NanoDrop™, or Qubit fluorometer). 10. Sanger sequencing. 2.2  Embryonic Stem Cells Transfection and Selection

1. ES cells with dominant coat color marker. 2. CRISPR/Cas9 delivery plasmids; single plasmid such as pSpCas9(BB)-2A-Puro (PX459) v2 (Addgene #62988) for transient co-expression of Cas9 guide RNA, Cas9 nuclease,

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and puromycin resistance. Alternatively, multiple plasmids will be co-transfected to express the same three components. 3. ES cell transfection method by either lipofection (Lipofectamine™ 2000) or electroporation (Bio-Rad Gene Pulser Xcell™ System). 4. Tissue culture hood and humidified incubator (37 °C, 5% CO2). 5. Inverted microscope. 6. Gelatinized tissue culture plates (96-well, 6-well, 60 mm, and 100 mm). 7. MEF feeder cell medium: 88% DMEM, 10% FBS (ES cell certified), 2 mM l-glutamate, penicillin (100 U/mL), and streptomycin (100 μg/mL). 8. ES cell medium: 80% DMEM, 15% FBS (ES cell certified), 2 mM l-glutamate, 100 mM 2-mercaptoethanol, 0.1 mM MEM NEAA (nonessential amino acids), 1 mM sodium pyruvate, 1000 U/mL leukemia inhibitory factor (LIF), penicillin (100 U/mL), and streptomycin (100 μg/mL). Optional: PD0325901 (1 μM final concentration) and CHIR99021 (3 μM final concentration). 9. Trypsin-EDTA (0.25%). 10. Positive selection agent; Puromycin for PX459 v2 (see Note 2 for G418). 11. Mitomycin C (MMC)-treated antibiotic-resistant MEF feeder cells (e.g., DR4 MEF feeder cells; Applied StemCell ASF-1024). 12. ES cell cryopreservation protectant medium: 20% DMSO, 80% ES cell medium. 13. Liquid nitrogen safe cryovials (SARSTEDT, Micro tube 2 mL, PP). 14. Paraffin film (e.g., Parafilm® Film). 15. Slow cooling cryo-container (Nalgene™ Cryo 1 °C Freezing Container). 2.3  Embryonic Stem Screening

1. High-fidelity polymerase (e.g., New England Biolabs Phusion® Polymerase). 2. Genomic DNA from individual ES cell clones. 3. Cell lysis buffer: 10 mM Tris HCl, pH 8.0, 1 mM EDTA, 100 mM NaCl, 1% SDS, and 0.8 mg/mL proteinase K (added to lysis buffer right before use). 4. DNA precipitation buffer: 75 mM NaCl in 100% ethanol. 5. 70% Ethanol.

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2.4  Pseudopregnant Surrogate Mice Preparation

1. Female Swiss Webster or CD-1 mice.

2.5  Mouse Blastocyst Isolation

1. Albino C57BL/6 male and female mice (e.g., Jackson Labs: B6(Cg)-Tyr/J or Charles River: B6N-Tyrc-Brd/ BrdCrCrl).

2. Vasectomized male Swiss Webster or CD-1 mice.

2. Optional: Pregnant mare serum gonadotropin (PMSG) and human chorionic gonadotropin (hCG); intraperitoneal injection of 5 IU per mouse. 3. Mouth pipette apparatus (mouthpiece, 30–40 cm flexible tubing, in-line syringe filter (e.g., 0.22 μM Millex®-GS sterile filter, MilliporeSigma), and pulled glass transfer pipette. 4. Dissecting microscope and dissection tools. 5. Blastocyst collection medium (BCM): 89% DMEM, 10% FBS, 1% 200 μM glutamine. 2.6  Blastocyst ES Cell Microinjection

1. Inverted microscope with mechanical stage and movable objectives (e.g., Leica DM IRB Inverted Microscope). 2. Micromanipulator (e.g., Leica Micromanipulator) and microinjector (e.g., Eppendorf CellTram Air). 3. Prefabricated microinjection needle and embryo/blastocyst holder; Eppendorf TransferTip® (ES) and VacuTip. 4. Glass bottom injection dish (e.g., MatTek P50G-0-30-F, MatTek Corporation). 5. Buffered microinjection medium; blastocyst collection medium (above) with 20 mM HEPES buffer.

2.7  Embryo Transfer to Recipient Surrogates

1. Pseudopregnant Swiss/CD-1 female mice.

2.8  Germline Transmission

1. Mouse facility approved by the Association for Assessment and Accreditation of Laboratory Animal Care International (AAALAC).

2. Nonsurgical embryo transfer (NSET™) device (ParaTechs).

2. Wild-type females of appropriate genetic background to detect germline transmission of ES cell coat color marker (see Note 3). 3. IACUC-approved mouse biopsy method for genotyping.

3  Methods 3.1  Genome Editing Vectors

1. Traditional gene targeting and CRISPR/Cas9-assisted genome editing differ with regard to design, screening, and vector construction. To avoid discussing these differences in the abstract,

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we will describe a knock-in Glis2-eGFP fusion allele that was successfully generated by CRISPR/Cas9-assisted ES cell targeting. In this example, the genetic payload introduced is the ORF of eGFP, which will disrupt the endogenous stop codon of Glis2 and the Cas9 target site and replace that with an inframe eGFP ORF. 2. Once the gene is identified (such as Glis2), the specific nucleotide sequence for the target locus can be obtained from a variety of online mouse genome databases such as Ensembl (www.ensembl.org/Mus_musculus), National Center for Biotechnology Information (NCBI: www.ncbi.nlm.nih.gov/ genome), or the UCSC Genome Browser (genome.ucsc. edu) [17, 18]. 3. In traditional gene targeting, the linearized targeting vector carrying a positive selection marker is integrated into the genome of the ES cells, and only ES cells with genomic integration of the positive selection marker will survive the positive selection agent (such as G418 or puromycin). CRISPR/Cas9-­ assisted targeting method requires transient co-delivery of Cas9 nuclease, locus-specific guide RNA, and a positive selection marker. The processes is reliant on unique double-stranded break created in the endogenous locus by the targeted Cas9 nuclease, so prior to generation of the replacement plasmid (targeting vector equivalent), the Cas9 delivery system must be designed. There are many CRISPR/Cas9 delivery plasmids available from various commercial and nonprofit sources. The lab of Feng Zhang at MIT, one of the pioneers in CRISPR/ Cas9 genome editing, has generated a variety of CRISPR/ Cas9 delivery plasmids that are available from the nonprofit Addgene plasmid repository. For CRISPR/Cas9 ES cell targeting, a single plasmid, pSpCas9(BB)-2A-Puro (PX459) V2.0, can be utilized that will transiently express the single chimeric gRNA, Cas9 nuclease, and puromycin resistance in transfected ES cells (Fig. 1) [19, 20]. 4. Picking the Cas9 guide sequence requires careful forethought on the consequences of the resulting double-stranded break (DSB). The target site sequence should be unique, since the repair plasmid with the desired genetic payload will serve as repair template only for the DSB at on-target locus. Any other genomic locations that are targeted by the guide RNA will either be mutated via nonhomologous end joining (NHEJ) during the repair of the Cas9-generated DSB or will potentially be subjected to excessive DSBs in the genome resulting in senescence of the ES cell and no subsequent ES colony formation. Additionally, the approximate Cas9 target location should be analyzed for repetitive elements or regions with high homology to other locations in the genome, which can be

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Fig. 1 pSpCas9(BB)-2A-Puro (PX459) V2.0 CRISPR/Cas9 delivery plasmid. RNA Pol III U6 promoter (black arrow) expresses the chimeric guide RNA scaffold, sgRNA (black square). RNA Pol II Chicken β Hybrid (CBh) promoter (dark gray arrow) expresses SpCas9-T2A-PuromycinR “self-cleaving” fusion protein (gray and white arrows). Transfection of ES cells with circular/episomal PX459V2 will result in transient expression of Cas9 guide RNA, Cas9 nuclease, and puromycin resistance. The Cas9 nuclease is targeted to the endogenous locus and will cleave the genomic DNA at the sequence homologous to the sgRNA

readily accomplished by BLAST analysis of the region of interest. With approximately 5% of the mouse genome still unmapped, even the best online CRISPR design tools cannot predict off-target sites in unknown genomic sequence. Once high homology/repetitive element sequences are identified and excluded, then a variety of online CRISPR design tools can be used to pick the best Cas9 guides sequences, including CRISPOR (crispor.tefor.net) and Zhang Lab’s CRISPR Design (crispr.mit.edu). Once the Cas9 targeting sequence is selected, complementary oligos with BbsI-compatible overhangs can be synthesized for subcloning into pSpCas9(BB)2A-Puro (PX459) V2 or any other Cas9 guide expressing plasmid (Fig. 1) [19]. 5. In traditional gene targeting, homologous recombination between the targeting vector and the locus of interest required relatively large arms of homology and co-integration of a positive selection marker alongside the desired genetic payload. An absolute consensus has never been reached on the optimal length of total homology between the endogenous locus and the targeting vector. This is at least partially due to locus sequence variability including chromatin accessibility in the ES cell. Anecdotal published evidence and some locus-specific studies suggest that total homology between 7 and 10 kb, split between two homology arms, was generally sufficient to target a specific locus in mouse ES cells [21, 22]. With the Glis2-eGFP knock-in allele example, a traditional targeting vector with 7 kb of homology will be approximately 13.5 kb

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Fig. 2 Glis2-eGFP targeting scheme comparison. (a) Traditional targeting vector with 7 kb homology arms, a 2.4 kb genetic payload (eGFP ORF and FRT-flanked puromycin resistance), and MC1-DTA-negative selection marker. Following successful targeting, FLP recombinase-mediated recombination will excise the FRT-flanked-­ positive selection cassette leaving behind a single FRT site in the 3′ UTR. (b) CRISPR/Cas9-assisted targeting scheme utilizes the co-transfection of two circular/episomal plasmids: (1) PX459 to transiently express Cas9 nuclease with guide RNA and puromycin resistance and (2) a repair template with 1.8 kb homology arms flanking the eGFP ORF genetic payload and no positive selection marker. Following homology-directed repair of the Cas9-generated double-stranded break in the 3′ UTR of the endogenous Glis2 locus, the Glis2-eGFP allele is generated. Importantly, without genomic integration of the positive selection marker, there is nothing to excise via recombinase activity from the locus that might negatively affect expression or prevent targeting another locus in series since the final CRISPR/Cas9-targeted ESC clones remain puromycin sensitive

in length (Fig. 2a). There are many good reviews/protocols available that include thorough step-by-step instructions in designing and generating traditional targeting vectors [23]. With CRISPR/Cas9, the endogenous locus is targeted by the guide RNA-Cas9 nuclease complex, and the resulting doublestranded break (DSB) in the target locus must be repaired for

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the ES cell to survive. Therefore the “targeting vector” or more appropriately named the “repair template” in CRISPR/ Cas9 targeting does not require large arms of homology nor a positive selection marker to be included in its design. The most straightforward way to generate virtually any repair template for CRISPR/Cas9-assisted ES cell targeting is to first PCR amplify a 1–2 kb amplicon flanking the target Cas9 sequence from ES cell genomic DNA and subclone it into any generic plasmid (e.g., pBlueScript or pCR2.1). Then in utilizing restriction-free cloning/site-directed mutagenesis, disrupt the Cas9 target sequence in the newly cloned plasmid, replacing it with either the desired genetic payload or unique restriction enzyme sites into which a desired genetic payload may be subsequently subcloned [24] (Fig. 2b). This targeting scheme can be exploited to make many other types of loci, such as a specific SNP, a large genomic disruption, a conditional allele, or a knock-in allele. The only limitation is the cloning capacity of the repair template plasmid. 3.2  Embryonic Stem Cells Transfection and Selection

1. ES cell culture and transfection: Frozen ES cells are thawed at 37 °C in a heated water bath and suspended in ES medium. The cells are centrifuged and resuspended in ES medium to remove residual freezing medium DMSO. Cells are plated onto MMC-­treated feeder cells in 100 mm plates. ES cells are incubated (37 °C, 5% CO2) and allowed to grow until they reach 70–80% confluency. 2. Lipofection (recommended): One hour before lipofection, trypsinize one 60 or 100 mm plate containing 70–80% confluent ES cells. Neutralize the trypsin with serum containing ES medium, and wash twice with D-PBS. Count ES cells, and plate 2.5 × 105 ES cells per well into a 6-well gelatinized plate with MMC-­treated feeders in 2 mL of ES medium. Within 1 h of plating, add lipofection mixture (total volume 250 μL with 6 μg circular plasmid DNA (6:1 molar ratio of repair template to Cas9 delivery plasmid) and 9 μL Lipofectamine 2000 in Gibco Opti-­Mem™). Transfect one well without any positive selection marker plasmid (negative control). Keep one well un-­ transfected as a positive control for ES cell growth. Approximately 14–18 h after lipofection, wash and trypsinize the transfected wells and split the ES cells between two and three 100 mm tissue culture plates with MMC-treated feeders. See Note 4 about circular versus linear plasmid DNA transfection. 3. Electroporation (alternative to lipofection): On the day of electroporation, trypsinize one 100 mm plate containing 70–80% confluent ES cells. Neutralize the trypsin using ES cell medium, and wash the cells twice in D-PBS. Electroporate 1 × 107 cells

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Fig. 3 ES cell lipofection with circular plasmid for transient puromycin resistance. Twenty-four hours after lipofection, ES cells are treated with puromycin (1 μg/ mL) for 48 h and then returned to standard ES cell medium until macroscopic colony formation approximately 5–6 days later. If puromycin treatment is extended, selection for ESC clones with random genomic integration of the puromycin resistance cassette occurs rapidly. Therefore, it is critical to utilize a puromycin treatment dose and duration that will allow the survival of transiently puromycin-resistant clones, while killing non-transfected ES cells and without selecting for ESC clones with random integration of the puromycin resistance cassette

in the presence of 20–30 μg of circular plasmid DNA (6:1 molar ratio of repair template to Cas9 delivery plasmid) using a Bio-­Rad Gene Pulser set at 230 V and 500 μF. Following ­electroporation, plate the cells onto 3–4 100 mm tissue culture plates with MMC-treated feeders. 4. Positive selection: Begin positive selection with 1 μg/mL puromycin 4–6 h after lipofected cells were transferred into the 100 mm plate (or approximately 24 h after electroporation) and changed once again after 24 h. After 48 h of puromycin treatment, replace the puromycin containing medium with standard ES cell medium. Macroscopic colony formation should be visible within 5–9 days (see Fig. 3 for time course). 5. Colony picking: After transfection of ES cells and subsequent selection in the presence of puromycin, the surviving colonies growing on a 100 mm feeder plate are picked (usually 7–10 days post-transfection) and trypsinized before plating onto a new 96-well feeder (DR-4) plate. Cells are allowed to grow until 70–80% confluency (generally 4–5 days).

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1. Once the majority of wells reach 70–80% confluency, the clones are trypsinized for replicate plating. Half of the cell population will be used to cryopreserve a master plate that will be stored at −80 °C during ESC clonal screening. For freezing the 96-well master plate, the trypsinized ES cells are suspended in equal volumes (100 μL) of ES medium and pre-chilled ES medium containing 20% DMSO and mixed well with a multichannel pipette. The plate is then securely sealed with paraffin film, placed into a plastic sleeve, and horizontally positioned in a room temperature insulated foam container prior to being transferred to −80 °C for controlled speed freezing. 2. The other half of the trypsinized ES cells are plated onto a 0.1% gelatinized 96-well plate and allowed to grow to confluence for DNA extraction and subsequent PCR screening. Majority of the wells will reach 80–90% confluency in about 4–8 days after plating. Potential differentiation of ESC clones on the replicate plate grown for screening is not a concern, since genomic DNA and PCR screening will not be affected by differentiation.

3.4  Embryonic Stem Cell Screening

1. Medium is removed and cells are washed once with 1× PBS. Cell lysis buffer (50 μL) is added to each well and incubated overnight at 55 °C. DNA is precipitated by adding 100 μL salt-­ethanol mixture, and the 96-well plate is stored at 4 °C for at least 6 h (or overnight to maximize the yield). Precipitated genomic DNA will be readily visible on the bottom of each well. DNA is washed with 70% ethanol, allowed to dry at room temperature, and resuspended with DNase/ RNase-free water for screening. 2. Clonal PCR screening. With circular plasmid transfection, random integration of the repair template (“targeting vector”) is greatly reduced in CRISPR/Cas9-assisted targeting as compared to traditional homologous recombination targeting. However, it remains critical to be able to differentiate between an on-target homology directed repair allele and a complete (or partial) random genomic integration of the repair template. Therefore, up to a total of three PCR assays are required to genotype the ESC clones: (1) a 5′ screen spanning the 5′ homology arm (endogenous locus 5′ of homology arm to repair template genetic payload), (2) a 3′ PCR screen spanning the 3′ homology arm (genetic payload to endogenous locus 3′ of the 3′ homology arm), and (3) wild-type screen within the repair template homology arms flanking the Cas9-target site (Fig.  4). Depending on total homology and genetic payload length, sometimes a single PCR screen can be designed to coamplify the endogenous wild-type allele and the HDR allele. Since the homology arms of the repair template are significantly

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Fig. 4 Screening ESC clones for homology-directed repair (HDR) allele. (a) Example of PCR-based ES cell screening strategy (Glis2-eGFP allele). Three separate PCR assays are utilized to genotype the ESC clones. (b and c) 5′ and 3′ screen with primers outside of the homology arm spanning from the endogenous locus to the genetic payload (eGFP ORF). (d) Wild-type screen flanking the Cas9 target sequence to confirm zygosity of targeting. If genetic payload allows, the wild-type screen can be designed to confirm zygosity (presence or absence of the wildtype) as well as reconfirm the 5′ and 3′ screen. Important to note, the wild-type screen alone will not differentiate random genomic integration of the repair template from an on-target HDR allele. (b–d) Assessed genotype: wildtype (clones: 7, 10, and 11), heterozygous (clones: 1, 3, 5, 8, 9, and 12), homozygous (clones: 2 and 6), and partial (clone: 4). The primary purpose of the wild-type screen is to determine zygosity of the targeting; therefore, the PCR design needs to be focused on robust amplification of the wild-type locus rather than co-­ amplification of the wild-type and HDR alleles, and the presence of an HDR amplicon in the wild-type assay should be used as optional/non-exclusionary confirmation of the 5′ and 3′ screens

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shorter than traditional targeting, long-range PCR is not necessary to screen the ESC clones. Targeting efficiency rates with this protocol are significantly higher than traditional targeting; therefore, homozygous targeted ESC clones are common; therefore, the wild-type screen is instrumental in confirming ESC clone zygosity (see Note 5 about ESC clone screening). 3. Upon identification of correctly targeted ESC clones, the frozen master plate (96-well) will be cryorecovered to expand specific genetically modified clones. The sealed frozen master plate (96-well) is thawed at 37 °C in water (or bead) bath for approximately 10 min, and the specific ESC clones are transferred to a properly labeled 24-well feeder plate containing 2 mL ES cell medium and incubated overnight at 37 °C. To remove residual freezing medium DMSO, replace the ES cell medium the next morning and continue incubating until 80–90% confluent (4–6 days). ESC clones are then transferred onto a 6-well feeder plate for further expansion. In about 2–3 days, ES cells are transferred to 60 mm feeder dish. In parallel, a small (100 μL from 1 mL suspension) aliquot is also transferred to a 24-well dish (gelatin-coated without feeders) for genotype verification (as described above). In about 2–3 days, the ES cells in 60 mm dishes should be 80–90% confluent and be ready to be frozen down. The 60 mm dish is trypsinized and spun down, and cells are suspended in 600 μL ES medium, which is further mixed with equal volume of chilled freezing medium and then equally split between four cryovials per clone (300 μL per vial). The cryovials are placed into a cryo-container to control freeze rate and placed overnight in −80 °C for slow cooling. Next day, vials are transferred into liquid nitrogen for long-term storage. 3.5  Pseudopregnant Surrogate Mice Preparation

1. Modified blastocysts are transferred to pseudopregnant mice at the equivalent stage of e2.5. Proper planning is required to ensure blastocysts are harvested the same day that pseudopregnant mice are ready. Mice intended to be pseudopregnant are set up 1 day later than female mice used for blastocyst collection. 2. Forty-five female mice are set up with vasectomized males. Three female mice are set up with one male. The following morning, females are checked for post-copulation plug, and those positive for plug are separated from the male and cohoused with other pseudopregnant mice. Female mice with clear post-­copulation plug are critical, as embryos transferred to a non-­pseudopregnant mouse will not implant. 3. Pseudopregnant mice are housed for 2 days until nonsurgical blastocyst transfer.

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3.6  Mouse Blastocyst Isolation

1. All animal procedures must be approved by Institutional Animal Care and Use Committee (IACUC) and animal facility veterinary staff. 2. To generate sufficient blastocysts, 60–65 female mice are naturally bred. Two female mice are set up with one male. Alternatively, superovulation of ten female mice can be used to generate larger numbers of embryos per mouse if colony size is limited. Blastocyst quality is noted to diminish when using superovulation method. Female mice are checked for a postcopulation plug the following morning, e0.5 (embryo day 0.5). Female mice positive for plugs are separated from males and cohoused for 3 days. Plug rate averages around 25%. 3. Prepare a 4-well dish with 500 μL blastocyst medium per well. Multiple 60 mm dishes with 10 mL blastocyst medium (approximately 2 dishes for every 5 donors). Dishes are kept at 37 °C until needed and should be prepared within 24 h of blastocyst isolation. 4. After 3 days, e3.5, female mice are euthanized with CO2 to collect uteri. 5. Place mouse with ventral side facing up, and spray with 70% ethanol to minimize fur contamination. To surgically remove the uterus of each female, cut through the abdominal wall to expose the reproductive organs (Fig. 5). 6. With forceps, hold the adipose tissue adjacent to one ovary while severing connective tissue and additional adipose tissue

Fig. 5 Donor blastocyst collection from donors. Once the abdominal cavity of donor females is dissected, the uterus from productive copulation plugs is engorged with blood (black arrows) and readily differentiated from the pale white uterus observed following nonproductive copulation plugs (white arrows). Both uterine horns can be removed as a single piece from ovary to ovary (stripped arrow) and a single cut at the cervix

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along the entire uterus. Cut through the cervix to obtain both uterine horns in one piece as they are still connected to the cervix. Place collected tissue in 60 mm dish of blastocyst medium pre-incubated at 37 °C. 7. Under a dissection microscope, place one uterus at a time into a new 60 mm dish. Use a 25-gauge needle attached to a 10 mL syringe filled with blastocyst medium at 37 °C to flush the blastocysts from the uterus into the dish. With fine forceps, grab the distal end of one uterus, adjacent to ovary, and insert needle and expel 1 mL blastocyst medium while holding the uterus to the needle with fine forceps. Repeat process on opposite side of same uterus. Medium will flow out of the cervix and carry blastocysts into the dish. Five uteri are generally flushed before switching to a new dish, with collection dishes placed in the incubator until all uteri are flushed. 8. Collect the embryos from the dish using a mouth pipette apparatus tube assembly with an attached capillary pipette, and place blastocysts into fresh droplet of blastocyst collection medium. At this stage, embryos can be separated that appear atypical or are not at the correct stage of development. Ideal embryos will have a clearly visible blastocoel cavity and an intact zona pellucida. 9. Good quality embryos can be transferred into a 4-well dish containing blastocyst medium and incubated at 37 °C until ready for injection. 3.7  Blastocyst ES Cell Microinjection

1. Trypsinized ES cells are prepared and stored on ice. Injection medium consists of buffered M2 medium or blastocyst medium supplemented with 20 mM HEPES to buffer the medium outside of the incubator. 2. Place glass bottom injection dish onto stage of inverted microscope, and cover bottom of dish with injection medium. Attach holder and microinjection pipettes to microinjector, and bring into field of view at low magnification (4× objective). 3. Add few droplets of media containing ES cells to the dish on the stage, and allow cells to settle to the bottom of the dish, generally 10–15 min. 4. Using the mouth pipette apparatus, gather 20–40 blastocysts from 4-well dish and transfer blastocysts to the middle of the dish on the stage. Under low magnification (4× objective), aggregate all the embryos into a manageable grouping with the holder pipette. 5. Switch to high magnification (20× objective) to view blastocysts and ES cells. Only blastocysts that appear normal with good morphology under high magnification should be used (Fig. 6a). Blastocysts at the wrong developmental stage or that have died

Fig. 6 ES cell microinjection into donor wild-type blastocysts. (a) Timing variability of copulation/fertilization for each individual donor female yields blastocysts at slightly varying stages of development, including early blastocysts (e3.25), optimal mid blastocysts (e3.5), and late blastocyst (e4). (b–e) Morphologically normal genetically modified ES cells are microinjected into the blastocoel of wild-type donor embryos and subsequently transferred to surrogate pseudopregnant females. (f) By postnatal day 14, level of chimerism in the potential F0 founders is readily assessed via dominant coat/eye color expression, which are good surrogate markers for the level of chimerism in the F0 founder’s germline

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can be separated and excluded from injection. Using the holder pipette, apply vacuum to a blastocyst and position it so it is just touching the bottom of the dish for stability. Bring the zona pellucida into focus [25]. 6. While remaining in the same field, gather 15–20 good quality ES cells with the injection needle from the dish. Try to eliminate medium in between the cells so the overall volume going into the blastocyst is reduced. Using the micromanipulator, bring the injection needle close to the blastocyst and adjust the Z-axis of the injection needle to bring the tip of the needle into focus, along with the zona pellucida. 7. With a swift motion, penetrate the blastocyst with the injection needle, aiming for the blastocoel. Dispense approximately 15 ES cells into the blastocoel, and remove injection needle from the embryo (Fig. 6b–e). Just before exiting the embryo, drawing some media back into the needle to reduce the internal pressure can prevent ES cells from being pushed out. 8. Repeat process for all good quality blastocysts, and return to new well of the 4-well dish. Keep modified embryos incubating at 37 °C until ready to transfer to pseudopregnant surrogate. 3.8  Embryo Transfer to Recipient Surrogates

1. With the advent of nonsurgical embryo transfer (NSET), blastocyst transfer to pseudopregnant surrogate females has greatly improved blastocyst implantation while reducing animal surgery [26]. The technique utilizes a plastic transcervical device that allows for the pipette transfer of embryos directly to the uterus without the need of anesthesia, as the procedure poses no more distress to a mouse than regular handling. There are several commercial vendors available for mouse NSET devices, all of which provide detailed video procedures for the proper use of their devices, which in combination with the IACUC and veterinary support staff can readily establish a SOP for nonsurgical embryo transfer.

3.9  Germline Transmission

1. Approximately 18.5 days after nonsurgical embryo transfer, the chimeric founder mice (i.e., “chimeras”) are born (Fig. 6f). Chimeric mice are comprised of both genetically modified ES cells and wild-type donor blastocyst-derived cells. Most of the commonly used ES cell lines are XY male; therefore, only the male chimeric mice are likely to transmit the genetically modified allele in their germline. Germline transmitting X0 female chimeras have been reported, but the incidence of this is extremely rare, and is generally not worth pursuing from a logistical standpoint [27]. The level of chimerism is visually assessed by the eye color and coat color distribution, which serve as a surrogate for the reproductive system (see Note 6

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about chimeric founders). Germline transmission rate can also be assessed via copulation plug genotyping, where by the ­copulation plugs from timed mating of male chimeras and wild-type females is genotyped for the presence of the genetically modified allele [28]. 2. At approximately 8 weeks of age, the male chimeras are mated with wild-type females of a recessive coat color (see Note 3) to detect transmission of the dominant coat color of the ES cells and the genetically modified allele. If coat color transmission is observed without transmission of the genetically modified allele, there are several plausible explanations: (1) incorrectly genotyped or mixed up ESC clones were used for blastocyst microinjection; (2) ESC clone was mosaic/chimeric resulting in nontargeted ES cells to contribute to male chimera germline without transmission of the genetically modified allele; (3) the genetically modified allele is conveying a deleterious phenotype at a time point between haploid gamete development in the chimera and birth of the chimera’s offspring to the wild-type female mice. In all instances, troubleshooting of coat color transmission without mutant allele transmission begins with re-­genotyping the expanded ESC clones.

4  Notes 1. Cas9 guide efficiency continues to be a controversial topic, with numerous algorithms generated to determine on-target efficiency of Cas9 guides [29, 30]. However, it has become increasingly clear that in vitro Cas9 restriction digest of a plasmid or PCR amplicon containing the guide target sequence will only confirm that the guide is not degraded. A guide/ Cas9 complex should absolutely cut a plasmid or amplicon in vitro similar to any other restriction enzyme. However, little insight can be gained about Cas9 guide targeting efficiency by comparing in vitro digest of non-genomic DNA and in vivo targeting efficiency in the context of the entire genome. 2. Following transfection, the Cas9 nuclease and puromycin resistance are expressed for approximately 72 h, after which point the episomal plasmids are either degraded or diluted out during cell division. Therefore, positive selection for transient resistance can only occur during 24–72 h post-transfection. Further positive selection pressure will begin to select for clones with random genomic integration of the puromycin resistance cassette. For a single Cas9/Puro delivery plasmid like PX459v2 that will include the random integration of the Cas9 nuclease as well. 48 h of puromycin treatment should be sufficient to kill all ES cells in the no PX459v2 control transfection. G418, even at

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elevated doses, is not potent enough to kill all nonresistant cells in 48 h; therefore, G418 is not suitable for transient positive selection. See Fig. 4 for ES cell survival following lipofection and transient expression of puromycin resistance. 3. Dominant coat color marker and germline transmission detection. Dominant eye color for all listed is black. 4. The CRISPR/Cas9-assisted gene targeting protocol described here relies on transient co-expression of puromycin resistance, Cas9 nuclease, and the guide. It is critical to maintain the Cas9 delivery and repair template plasmids in circular form to minimize random genomic integration. The following steps, although anecdotal in evidence, have helped decrease linear/ nicked plasmid DNA for ES cell transfection: (1) collect bacteria in mid-stationary phase (12–16 h of growth), rather than logarithmic or death phase; (2) minimize vortexing and excessive pipetting to reduce mechanical shear; (3) minimize alkaline lysis during plasmid isolation; (4) treat final prep as if grown in EndA+ bacteria and perform “optional” washes (if applicable with kit being used); and (5) store final plasmid preps at 4 °C rather than −20 °C prior to ES cell transfection. DNA is very stable at 4 °C, and signs of plasmid degradation at 4 °C are indicative of a problem with the plasmid isolation method. ES cell (background); Dominant coat color; Germline detection breeding. AB2.2 (129S); Agouti; Standard or Albino B6. C2 (C57BL/6N); Black; Albino B6. E14 (129P2); Chinchilla; Standard or Albino B6. G4 (129B6F1); Black and Agouti; Standard or Albino B6 (optimal). If bred to standard B6, all offspring must be tested for germline transmission of the mutant allele once germline transmission of agouti coat color is detected from a chimera founder. JM8A (C57BL/6N-A); Agouti and Black; Standard or Albino B6 (optimal) 5. PCR screening of CRISPR/Cas9-targeted ESC clones allows for rapid identification of ESC clones carrying the genetically modified allele. Additionally, with shorter homology arms (compared to traditional targeting) and lack of integrated positive selection marker, the entire genetic payload sequence in the targeted locus can be readily confirmed by sequencing the screening PCR amplicons. Since targeting efficiency is drastically increased, high stringency criteria can be used to select ESC clones for expansion from the master plate, avoiding clones with shifted size amplicons, (potentially) off-target

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amplicons, faint on-target bands, or any clones for which the zygosity cannot be definitively determined (i.e., lack of an amplicon only guarantees that the PCR reaction did not work). 6. The ultimate purpose of a F0 chimera founder is to germline transmit the genetically modified allele of the ESC clone. Dominant coat and eye color expression have been shown to be good surrogates for level of chimerism in the germline, but ultimately, they are only estimates. The only true way to confirm germline chimerism level is through breeding, and many mouse lines have been successfully generated from germline chimeras that superficially have low level (visual) chimerism. References 1. Doetschman T, Gregg RG, Maeda N et al (1987) Targeted correction of a mutant HPRT gene in mouse embryonic stem cells. Nature 330(6148):576–578. https://doi.org/ 10.1038/330576a0 2. Thomas KR, Capecchi MR (1987) Site-­ directed mutagenesis by gene targeting in mouse embryo-derived stem cells. Cell 51(3): 503–512 3. Capecchi MR (2001) Generating mice with targeted mutations. Nat Med 7(10):1086– 1090. https://doi.org/10.1038/ nm1001-1086 4. Koller BH, Hagemann LJ, Doetschman T et al (1989) Germ-line transmission of a planned alteration made in a hypoxanthine phosphoribosyltransferase gene by homologous recombination in embryonic stem cells. Proc Natl Acad Sci U S A 86(22):8927–8931 5. Snouwaert JN, Brigman KK, Latour AM et al (1992) An animal model for cystic fibrosis made by gene targeting. Science 257(5073): 1083–1088 6. Mouse Genome Sequencing C, Waterston RH, Lindblad-Toh K et al (2002) Initial sequencing and comparative analysis of the mouse genome. Nature 420(6915):520–562. https://doi. org/10.1038/nature01262 7. Ringwald M, Iyer V, Mason JC et al (2011) The IKMC web portal: a central point of entry to data and resources from the international knockout Mouse consortium. Nucleic Acids Res 39(Database issue):D849–D855. https:// doi.org/10.1093/nar/gkq879 8. Lundberg KS, Shoemaker DD, Adams MW et al (1991) High-fidelity amplification using a thermostable DNA polymerase isolated from Pyrococcus furiosus. Gene 108(1):1–6 9. Tsyrulnyk A, Moriggl R (2008) A detailed protocol for bacterial artificial chromosome

recombineering to study essential genes in stem cells. Methods Mol Biol 430:269–293. https://doi.org/10.1007/978-1-59745182-6_19 10. Williams RL, Hilton DJ, Pease S et al (1988) Myeloid leukaemia inhibitory factor maintains the developmental potential of embryonic stem cells. Nature 336(6200):684–687. https:// doi.org/10.1038/336684a0 11. Ying QL, Wray J, Nichols J et al (2008) The ground state of embryonic stem cell selfrenewal. Nature 453(7194):519–523. https:// doi.org/10.1038/nature06968 12. Folger KR, Wong EA, Wahl G et al (1982) Patterns of integration of DNA microinjected into cultured mammalian cells: evidence for homologous recombination between injected plasmid DNA molecules. Mol Cell Biol 2(11):1372–1387 13. Yagi T, Ikawa Y, Yoshida K et al (1990) Homologous recombination at c-fyn locus of mouse embryonic stem cells with use of diphtheria toxin A-fragment gene in negative selection. Proc Natl Acad Sci U S A 87(24): 9918–9922 14. Connelly JP, Barker JC, Pruett-Miller S et al (2010) Gene correction by homologous recombination with zinc finger nucleases in primary cells from a mouse model of a generic recessive genetic disease. Mol Ther 18(6):1103–1110. https://doi.org/10.1038/mt.2010.57 15. Christian M, Cermak T, Doyle EL et al (2010) Targeting DNA double-strand breaks with TAL effector nucleases. Genetics 186(2): 757–761. https://doi.org/10.1534/ genetics.110.120717 16. Cong L, Ran FA, Cox D et al (2013) Multiplex genome engineering using CRISPR/Cas systems. Science 339(6121):819–823. https:// doi.org/10.1126/science.1231143

CRISPR/Cas9-Assisted Genome Editing in Murine Embryonic Stem Cells 17. Zerbino DR, Achuthan P, Akanni W et al (2018) Ensembl 2018. Nucleic Acids Res 46(D1):D754–D761. https://doi. org/10.1093/nar/gkx1098 18. Casper J, Zweig AS, Villarreal C et al (2018) The UCSC genome browser database: 2018 update. Nucleic Acids Res 46(D1):D762–D769. https://doi.org/10.1093/nar/gkx1020 19. Ran FA, Hsu PD, Wright J et al (2013) Genome engineering using the CRISPR-Cas9 system. Nat Protoc 8(11):2281–2308. https://doi.org/10.1038/nprot.2013.143 20. Hsu PD, Lander ES, Zhang F (2014) Development and applications of CRISPRCas9 for genome engineering. Cell 157(6):1262–1278. https://doi. org/10.1016/j.cell.2014.05.010 21. Hasty P, Rivera-Perez J, Bradley A (1991) The length of homology required for gene targeting in embryonic stem cells. Mol Cell Biol 11(11):5586–5591 22. Lu ZH, Books JT, Kaufman RM et al (2003) Long targeting arms do not increase the efficiency of homologous recombination in the beta-globin locus of murine embryonic stem cells. Blood 102(4):1531–1533. https://doi. org/10.1182/blood-2003-03-0708 23. Zimmermann AG, Sun Y (2013) Conventional murine gene targeting. Methods Mol Biol 1031:1–18. https://doi.org/10.1007/9781-62703-481-4_1 24. van den Ent F, Lowe J (2006) RF cloning: a restriction-free method for inserting target

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genes into plasmids. J Biochem Biophys Methods 67(1):67–74. https://doi. org/10.1016/j.jbbm.2005.12.008 25. Scott GJGA, Hagler TB, Ray MK (2018) Trans–inner cell mass injection of embryonic stem cells leads to higher chimerism rates. J Vis Exp. https://doi.org/10.3791/56955 26. Bin Ali R, van der Ahe F, Braumuller TM et al (2014) Improved pregnancy and birth rates with routine application of nonsurgical embryo transfer. Transgenic Res 23(4):691–695. https://doi.org/10.1007/s11248-0149802-3 27. Deng JM, Satoh K, Wang H et al (2011) Generation of viable male and female mice from two fathers. Biol Reprod 84(3):613–618. https://doi.org/10.1095/biolreprod.110. 088831 28. Wilson S, Sheardown SA (2011) Identification of germline competent chimaeras by copulatory plug genotyping. Transgenic Res 20(2):429–433. https://doi.org/10.1007/ s11248-010-9413-6 29. Moreno-Mateos MA, Vejnar CE, Beaudoin JD et al (2015) CRISPRscan: designing highly efficient sgRNAs for CRISPR-Cas9 targeting in vivo. Nat Methods 12(10):982–988. https://doi.org/10.1038/nmeth.3543 30. Doench JG, Fusi N, Sullender M et al (2016) Optimized sgRNA design to maximize activity and minimize off-target effects of CRISPR-­ Cas9. Nat Biotechnol 34(2):184–191. https:// doi.org/10.1038/nbt.3437

Chapter 2 Genome Editing in Mouse Embryos with CRISPR/Cas9 Greg J. Scott and Artiom Gruzdev Abstract Transgenic mouse models can be subdivided into two main categories based on genomic location: (1) targeted genomic manipulation and (2) random integration into the genome. Despite the potential confounding insertional mutagenesis and host locus-dependent expression, random integration transgenics allowed for rapid in vivo assessment of gene/protein function. Since precise genomic manipulation required the time-consuming prerequisite of first generating genetically modified embryonic stem cells, the rapid nature of generating random integration transgenes remained a strong benefit outweighing various disadvantages. The advent of targetable nucleases, such as CRISPR/Cas9, has eliminated the prerequisite of first generating genetically modified embryonic stem cells for some types of targeted genomic mutations. This chapter outlines the generation of mouse models with targeted genomic manipulation using the CRISPR/Cas9 system directly into single cell mouse embryos. Key words Genetically modified mice, Gene targeting, CRISPR/Cas9, Microinjection, Germline transmission, Nonhomologous end joining (NHEJ), Homology-directed repair (HDR)

1  Introduction Genetically modified and transgenic mouse models have become ubiquitous in modern biomedical research and have expanded our scientific understanding of countless biological processes, molecular pathways, genetics, toxicology, and disease pathogenesis. Transgenic mouse models can be subdivided into two primary sub-­ types based on the genomic destination of the transgenic element: (1) random genomic integration or (2) targeted integration. The protocols for generating random integration transgenics have remained largely unchanged since first published [1, 2]. In general, a linear DNA fragment is microinjected into a pronucleus of a single cell embryo, which is surgically transferred to a pseudopregnant surrogate female. The microinjected DNA fragment can be integrated into the embryo’s genome via several mechanisms including non-random microhomology-driven integration and nonhomologous end joining (NHEJ) [3–5]. The allelic result is unpredictable and therefore requires extensive characterization of Irving C. Allen (ed.), Mouse Models of Innate Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1960, https://doi.org/10.1007/978-1-4939-9167-9_2, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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multiple transgenic sublines to identify the line of interest. Despite the potential for insertional mutagenesis, concatemer integration, multiple integration sites per genome, partial transgene integration, and unknown chromatin effects of the site of integration, pronuclear microinjection has been successfully used to generate countless transgenic mouse lines over the last 30 years. The advent of targeted transgenics, made possible by homologous recombination targeting in ES cells (the focus of Chapter 1), only increased the demand for random integration transgenes [6]. Despite the potential caveats, the time frame for in vivo phenotyping of a transgene directly microinjected into a single cell embryo remained substantially faster than first generating transgenic embryonic stem cells and subsequent germline chimeric founders. The time frame to phenotyping targeted mutations continued to lag random integration transgenes until 2010, when the first targetable nucleases were used in single cell embryos [7]. Although zinc finger nucleases (ZFNs) were first utilized in single cell mouse embryos, it was the ease of use and accessibility of CRISPR/Cas9 that finally made targeted genomic manipulation/transgenesis a timely alternative to random integration transgenesis [8, 9]. In this chapter, we describe the use of the CRISPR/ Cas9 system in single cell embryos to rapidly generate genetically modified mouse lines with locus-specific targeting. CRISPR/Cas9 genome editing is a rapidly evolving field at this time; therefore, our focus here will be a design, implementation, and screening protocol that may be readily applied to any genomic location.

2  Materials 2.1  CRISPR/Cas9 Genome Editing Reagents

1. (Embryo Microinjection) Polyadenylated Cas9 mRNA with 7-methylguanylate cap, 2 μg per procedure, 20 μL at 100 ng/ μL (commercial or in vitro transcribed with m7G and poly(A) tail from Cas9 containing plasmid such as pCAG-T3-hCAS-pA (Addgene #48625)) [10]. 2. (Embryo Electroporation) Recombinant Cas9 protein with nuclear localization signal (NLS): 10 μg per procedure, 20 μL at 0.5 μg/μL. 3. Locus-specific Cas9 single guide RNA (sgRNA; chimeric scaffold), 2 μg (electroporation) or 400 ng (microinjection set) (commercial synthesis or in vitro transcribed without m7G cap or poly(A) tail). 4. In vitro transcription kits: T7 for sgRNA (Lucigen AmpliScribe™ T7 High Yield Transcription Kit), T3 for m7Gcapped poly(A)-tailed Cas9 mRNA (mMESSAGE mMACHINE™ T3 Transcription Kit and Poly(A) Tailing Kit, Thermo Fisher Scientific), or T7 for m7G-capped poly(A)-

25

Genome Editing in Mouse Embryos with CRISPR/Cas9

tailed Cas9 mRNA (HiScribe™ T7 ARCA mRNA Kit (with tailing), New England Biolabs). 5. Repair template; single-stranded DNA (see Note 1). 6. Nuclease-free TE and ddH2O. 2.2  Pseudopregnant Surrogate Recipient Females

1. Swiss or CD-1 vasectomized male mice (see Note 2).

2.3  Single Cell Embryo Collection from Donor Mice

1. Male and female mice of donor strain (see Note 3).

2. Swiss female mice, 6–9 weeks of age. 3. Blunt, curved end forceps for copulation plug detection.

2. Low dead-space syringes (e.g., standard insulin syringes). 3. Sterile injectable saline. 4. Pregnant mare serum gonadotropin (PMSG) and human chorionic gonadotropin (HCG) (Lee Biosolutions) (see Note 4). 5. Time-mated female mice (e0.5, morning of copulation plug). 6. Dissection instrument (sharp scissors, sharp forceps). 7. 60 mm, 100 mm, and 4-well dishes. 8. Embryo culture MilliporeSigma).

medium

(e.g.,

EmbryoMax®

KSOM

9. Mouth pipette apparatus (mouthpiece, 30–40 cm flexible tubing, in-line syringe filter [e.g., 0.22 μM Millex®-GS sterile filter, MilliporeSigma]), P1000 pipette tip, and pulled glass transfer pipette. 10. 70% ethanol. 11. Hyaluronidase in M2 Medium (e.g., EmbryoMax® M2 Medium with hyaluronidase, MilliporeSigma). 2.4  CRISPR/Cas9 Reagent Preparation

1. Nuclease-free ddH2O. 2. (Embryo Microinjection) 10× nuclease-free phosphate-­ buffered saline (PBS). 3. (Embryo Microinjection) PVDF non-sterile centrifugal filter unit with a pore size of 0.1 μM (e.g., Ultrafree-MC VV Centrifugal Filter, MilliporeSigma). 4. (Embryo Electroporation) Nuclease-free 10× Tris-EDTA (TE) (pH 7–8). 5. (Embryo Electroporation) Opti-MEM I Reduced Serum Medium.

2.5  CRISPR/Cas9 Embryo Microinjection

1. Inverted microscope (e.g., Leica DM IRE2). 2. Micromanipulators (e.g., Leica, Sutter). 3. Injection control device (e.g., Eppendorf FemtoJet® 4×). 4. Holding microinjector (e.g., Eppendorf air-tram).

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5. Injection medium (M2 or other suitably buffered medium). 6. Injection dish Corporation).

(e.g.,

MatTek

P50G-0-30-F,

MatTek

7. Embryo-holding pipette (Eppendorf VacuTip). 8. Injection needle; prefabricated (e.g., Eppendorf Femtotips) or custom made from borosilicate glass with filament and micropipette puller (Sutter Instruments). For pulling glass injection needles, please consult your puller’s manual. Every puller, element, and glass stock are different and will require different settings. 9. Embryo-safe oil; silicone-based or similar (e.g., Sigma DMPS2X) (see Note 5). 10. Injection needle Microloader). 2.6  CRISPR/Cas9 Embryo Electroporation

microloader

(Eppendorf

Femtotips

1. Electroporation unit capable of square wave, 30 V, 0.1 s interval pulses (e.g., Bio-Rad Pulser Xcell™). 2. Either 1 mm gap electroporation cuvette (Bio-Rad) or 1 mm gap electroporation slide (BTX Harvard Apparatus). 3. Opti-MEM I Reduced Serum Medium.

2.7  Surgical Embryo Transfer to Pseudopregnant Recipients

1. IACUC-approved anesthetic and analgesic (see Note 6). 2. Animal hair clipper. 3. Iodine swab, alcohol wipe, sterilized lab wipes, sterile gloves. 4. Sterilized surgical instruments: sharp scissors, two sharp forceps, blunt forceps, micro serrefine, wound clips (with applicator/ removal tool). 5. Epinephrine (for vasoconstriction during oviduct embryo transfer). 6. Insulin syringe. 7. Mouth pipette apparatus. 8. Stereo dissecting microscope X2 (e.g., Leica MZ8). 9. Warming pad.

2.8  CRISPR F0 Founder Screening Strategies and Tips

1. Genomic DNA from F0 founders. 2. High-fidelity polymerase (e.g., Phusion, New England Biolabs). 3. Thermocycler and PCR reagents. 4. Agarose electrophoresis apparatus and reagents. 5. PCR amplicon gel extraction method. 6. Sanger sequencing.

Genome Editing in Mouse Embryos with CRISPR/Cas9

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3  Methods 3.1  CRISPR/Cas9 Genome Editing Reagents

1. For direct embryo microinjection, Cas9 mRNA can be readily generated using commercial in vitro transcription kits with m7G-cap and poly(A) tail. T3 promoter and Cas9 ORF containing plasmids such as pCAG-T3-hCAS-pA (Addgene #48625) can be linearized with a restriction enzyme 3′ of the Cas9 ORF (Sph I for pCAG-T3-hCAS-pA) and used as template for T3 in vitro RNA transcription in the presence of 7-methyl guanosine cap to generate a virtually limitless supply of m7G-capped and poly(A)-tailed Cas9 mRNA [10]. Alternatively, Cas9 mRNA is readily available from numerous commercial sources. 2. For embryo electroporation, recombinant Cas9 protein is available from numerous commercial sources. Electroporation with Cas9 mRNA has yielded highly variable results; therefore, Cas9 protein is recommended for embryo electroporation. 3. Cas9 sgRNA can be readily generated via T7 in vitro RNA transcription kits utilizing a DNA template generated from extended primers without any sub-cloning (Fig. 1). 4. Although any DNA can serve as a repair template for a Cas9-­ generated double-stranded break, in single cell embryos,

Fig. 1 Cloning-free Cas9 sgRNA synthesis. Primers are annealed and extended to generate the DNA template for T7 in vitro RNA transcription to generate the target-­specific Cas9 sgRNA. Locus-specific forward primer: 5′-GAAAT TAATA CGACT CACTA TAggc caccc tggac gacga cgGTT TTAGA GCTAG AAATA GCAAG-3′; Universal Reverse Primer: 5′-AAAAG CACCG ACTCG GTGCC ACTTT TTCAA GTTGA TAACG GACTA GCCTT ATTTT AACTT GCTAT TTCTA GCTCT AAAAC-3′. Extended primers will generate a template with a T7 promoter (underlined), 20 bp Cas9 targeting sequence (lowercase), and chimeric scaffold (uppercase)

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efficiency is critical. Every embryo injected is a potential founder mouse that must be housed and screened, and subsequently bred, which is considerably more expensive than screening a 96-well plate of ES cells (as described in the previous chapter). Circular plasmid DNA is too inefficient for routine use in single cell embryos, while double-stranded DNA has the risk of random genomic integration as well as being cytotoxic to the embryo. Single-stranded DNA has been shown to be the most robust option for targeted mutation in single cell embryos. 200 bp single-stranded donor oligonucleotides (ssODN) work especially well and are readily and inexpensively available from DNA synthesis companies. With a recommended minimum of 100 bp of total homology, the singlestranded 200 bp ssODN can introduce SNPs, protein tags, recombinase sites, premature stop codons, splice disruptions, or other small genomic manipulation (10,000 × g). Aspirate the supernatant. Lyse cells in 100 μl of lysis buffer with protease inhibitors for 30 min on ice. Spin lysed cells for 15 min at maximum speed (>10,000 × g) at 4 °C. Transfer lysate to a new tube and quantitate protein concentration by bicinchoninic acid assay or Bradford protein assay. Load 20–100 μg of total cell lysate on an SDS-PAGE gel (see Note 28). Gels can be transferred to nitrocellulose or PVDF membranes for immunoblotting. We used standard immunoblotting techniques for the detection of cytokines, cell surface receptors, and intracellular signaling molecules. 7. Functional assays such as phagocytosis, reactive oxygen or nitrogen species generation, and migration are generally performed 24–72 h after polarization.

3.7  Polarization to Alternatively Activated Macrophages (AAM or M2a) (See Note 23)

1. Culture macrophages for 6–8 days. 2. Add 0.5–1.0 × 106 bone marrow-derived macrophages in 1 ml of media to each well in a 6-well tissue culture plate. Add 10–20 U/ml of recombinant mouse IL-4 or IL-13 for 18 h (see Note 24).

Reverse primer 5-CTGGAAGACTCCTCCCAGGGTATAT 5-GGCTGTCAGAGCCTCGTGGCTTTGG-3 5-GTGGAGCAGCAGATGTGAGTGGCT-3 5-CAGATATGCAGGGAGTCACC-3 5-CACCTCTTCACTGCAGGGACAGTTGGCAGA-3 5-TGTCTAGGTCCTGGAGTCCAGCAGACTCAA-3 5-GGCAGTCATGTCCGGTGATG-3 5-CCAGCATCACCCCATTAGAT-3

Forward primer

5-CAGCCTCTTCTCATTCCTGCTTGTC-3

5-CCCTTCCGAAGTTTCTGGCAGCAGC-3

5-ATGGCCATGTGGGAGCTGGAGAAAG-3

5-CAGAAGAATGGAAGAGTCAG-3

5-GGTCCCAGTGCATATGGATGAGACCATAGA-3

5-CCAGTTTTACCTGGTAGAAGTGATG-3

5-ACAGCAGTGTGCAGTTGATGA-3

5-GCACTTGGCAAAATGGAGAT-3

Gene

Tnfa

iNos

Il12p40

Arg1

Fizz1

Il-10

SK-1

Gapdh

Table 1 RT-PCR primers

50 Beckley K. Davis

Bone Marrow-Derived Macrophages

51

3. Stimulate cells with 1–100 ng ultrapure LPS and incubate (see Note 29). 4. For gene induction studies measuring transcription of proinflammatory cytokines, 2–6 h of stimulation with LPS works well. Briefly, cells are washed in 1× DPBS, and cells are removed via physical scraping. Total RNA can be isolated using standard techniques. 5. 1 μg of total RNA is reverse transcribed using Superscript III with oligo dT16–18 primer following the manufacturer’s suggested protocol. We use 2 μl of cDNA reaction for amplification with Phusion™ DNA polymerase (see Note 25) to amplify the following genes: Arg1, Fizz1, and Gapdh (see Table 1). 6. It has been reported in the literature that activity and levels of Arginase I can be measured in alternatively activated macrophage lysates [23, 24]. 3.8  Polarization to Regulatory Macrophages (M2b/ c/d) (See Note 23)

Regulatory macrophages might represent a heterogenous population of macrophages that arise from different stimulation/polarization protocols. Indeed, there have been regimens that produce “regulatory” macrophages that include immune complexes (M2b), glucocorticoids plus IL-10 (M2c), and others (M2d) [11, 13, 15]. Here, we will focus on regulatory macrophages generated in the presence of immune complexes. 1. Add 0.5–1.0 × 106 bone marrow-derived macrophages in 1 ml of media to each well in a 6-well tissue culture plate. 2. Prepare ovalbumin immune complexes by adding 20 μl of 1 mg/ml of EndoFit™ (see Note 30) to 500 μl of DMEM. Add 75 μl of 4 mg/ml rabbit anti-ovalbumin IgG drop wise. Nutate for 30–60 min at room temperature to allow complexes to form. 3. Stimulate macrophages with 1–50 ng of ultrapure LPS and with 100 μl endotoxin-free ovalbumin-IgG complexes as prepared above. Control stimulations (including unstimulated, LPS only, OVA only, and OVA specific IgG only) should be done in parallel. 4. Incubate the macrophages for 18–24 h in a 37 °C incubator. 5. For gene induction studies measuring transcription of proinflammatory cytokines, 2–6 h of stimulation with LPS works well. Briefly, cells are washed in 1× DPBS, and cells are removed via physical scraping. Total RNA can be isolated using standard techniques. 6. 1 μg of total RNA is reverse transcribed using Superscript III with oligo dT16–18 primer following the manufacturer’s suggested protocol.

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7. We use 2 μl of cDNA reaction for amplification with Phusion® DNA polymerase (see Note 25) to amplify the following genes: Il10, Il12p40 SK-1, and Gapdh (see Table 1). 8. Collect cell-free supernatants for ELISA measurement of IL-12 p40, IL-10, and either TNF-α or IL-6 (see Note 27).

4  Notes 1. It is imperative that all solutions remain sterile and pyrogen-­ free. Bone marrow-derived macrophages are exceptionally sensitive to bacterial moieties. If possible, all manipulations should be carried out in a laminar flow hood using aseptic techniques. The generation of bone marrow-derived macrophages from novel, transgenic, or gene ablation mice may require individual optimization. We have successfully used these protocols using wild-type mice and several novel mouse strains [19, 25–27]. 2. We have used TLR agonists from InvivoGen; other vendors such as Sigma and Invitrogen provide similar products. 3. We use C57Bl/6 mice to derive macrophages; however, other groups have used commonly available inbred strains using similar protocols. In all cases we exclusively use mice housed in specific pathogen-free facilities to minimize the activation status of macrophages. 4. Disposable dissecting trays can be fashioned out of Styrofoam. 5. In addition to bone marrow-derived cells, tissue-specific macrophages can be harvested in parallel. Tissue-derived cells can be harvested from diverse sources, including the spleen, liver, lung, and intestine. Additional immunologically relevant tissues such as lymph nodes and thymus can also be harvested at this time to assay different cellular components, making full use of the experimental animal. 6. Tibia bones are also a source for bone marrow and can be processed in an analogous manner to increase the yield per mouse of bone marrow precursor cells. The tibias should be separated at the ankle joint. 7. We have used 20–27 gauge needles to irrigate femurs. Smaller gauge needles will be easier to use for tibial bone marrow evacuations. 8. We have used different isotonic solutions (1× DBPS, DMEM, and HBSS) to irrigate the bone marrow cavity with no decrease in cell numbers, viability, or biological function. 9. We have used both treated and non-treated tissue culture plasticware to cultivate bone marrow-derived macrophages. We prefer to use treated plasticware to avoid possible confusion

Bone Marrow-Derived Macrophages

53

while growing different cell types. As a result of using treated tissue culture plasticware, bone marrow-derived macrophages adhere tightly to these dishes and may require physical dissociation with a cell scraper or prolonged treatment with trypsin-­ EDTA solution. 10. We have noticed slight variability in bone marrow-derived macrophage growth and maturation, possibly due to variability of growth factors (M-CSF) in L929-conditioned media. 11. Daily inspection of cells allows for visual confirmation of cell growth, adherence, and the possibility of contamination. 12. Bone marrow macrophages can adhere tightly to tissue culture-­ treated plasticwear and may require either increased incubation time with 0.05% trypsin-EDTA, increased concentration of trypsin-EDTA solution, or mechanical detachment with a cell scraper. 13. For higher throughput analysis, 96-well round bottom tissue culture plates can be used in place of 1.5 ml tubes. 14. We have used many different fluorophores and antibody sources. It is imperative that the fluorophores not overlap in emission spectra and are compatible with flow cytometer lasers. 15. Differential staining by Romanowsky staining (Diff-Quik™) can provide an easy means by which cells can be identified and their relative percentages obtained. 16. The day before the experiment, harvest the bone marrow-­ derived macrophages, and plate in either 6-, 12- or 24-well tissue culture plates at densities of 0.5–1.0 × 106 in 3 ml, 3–5 × 105 in 2 ml, or 0.5–2 × 105 cells in 1 ml of media, respectively. 17. We have seen sufficient cytokine secretion (IL-1β, IL-6, TNF-­ α, and IFN-β) in response to TLR stimulation in bone marrow-­ derived macrophages. Assays may require additional incubation times depending upon the stimulus and the biological readout. Also, it may be necessary to dilute the samples in order to fall within the linear range of the assay. 18. Different substrates can be used to effectively determine relative phagocytosis indexes. Light microscopy with differential staining can be used instead of fluorescent microscopy to determine phagocytosis of either bacteria or yeast. 19. The rate of phagocytosis may be variable depending on individual preparations of bone marrow-derived macrophages, their activation status, and the substrates used. 20. Other methods of fixation can be used. Aldehydes are the most common fixative. Care must be used when dealing with either paraformaldehyde or glutaraldehyde as both chemicals are

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suspected carcinogens. 100% ice-cold methanol precipitation can be used with satisfactory results. 21. Permeabilization is accomplished by the addition of detergent; we have used other detergents, such as saponin, with similar results. 22. Flow cytometry can also be used to analyze the relative amount of phagocytosis of bacteria [7, 19]. 23. After 6–8 days in culture, macrophages can be polarized into one of the three main populations: classically activated, alternatively activated, or regulatory macrophages. 24. Commercial sources of recombinant growth factors such as interleukins are typically expressed in E. coli. These preparations have varying amounts of microbial contaminants (i.e., LPS) that may alter macrophage function. Source and lot variation should be evaluated. 25. Other DNA polymerases can be used (e.g., Takara LA™ Taq). We have had success with Phusion™ using different source material and amounts, primer sets, and amplifying conditions. 26. Different effector molecules have different kinetic secretion profiles. Initial time point experiments will better define the appropriate stimulation periods. 27. A series of four twofold serial dilutions of supernatants will allow for experimental values to fall within the linear range of the assay. 28. The level of sensitivity of analyte will depend on the reagents used. Optimization with different detection antibodies and lysate concentrations might be necessary. 29. We have seen lot, source, and experimenter variability with LPS preparations. Single use aliquots should be stored at −80 °C and quality assured before experimentation. 30. Commercial preparations of ovalbumin contain varying amounts of LPS. We have used EndoFit™ ovalbumin; other sources of ovalbumin should be tested for LPS before experimentation. References 1. Varol C, Mildner A, Jung S (2015) Macrophages: development and tissue specialization. Annu Rev Immunol 33:643–675 2. Karp CL, Murray PJ (2012) Non-canonical alternatives: what a macrophage is 4. J Exp Med 209:427–431 3. Mege JL, Mehraj V, Capo C (2011) Macrophage polarization and bacterial infections. Curr Opin Infect Dis 24:230–234

4. Bloemen K, Verstraelen S, Van Den Heuvel R et al (2007) The allergic cascade: review of the most important molecules in the asthmatic lung. Immunol Lett 113:6–18 5. Chawla A, Nguyen KD, Goh YP (2011) Macrophage-mediated inflammation in metabolic disease. Nat Rev Immunol 11:738–749 6. Biswas SK, Chittezhath M, Shalova IN et al (2012) Macrophage polarization and plasticity

Bone Marrow-Derived Macrophages in health and disease. Immunol Res 53: 11–24 7. Sica A, Mantovani A (2012) Macrophage plasticity and polarization: in vivo veritas. J Clin Invest 122:787–795 8. DeFalco T, Potter SJ, Williams AV et al (2015) Macrophages contribute to the spermatogonial niche in the adult testis. Cell Rep 12: 1107–1119 9. Hulsmans M, Clauss S, Xiao L et al (2017) Macrophages facilitate electrical conduction in the heart. Cell 169:510–22 e20 10. McArdle S, Mikulski Z, Ley K (2016) Live cell imaging to understand monocyte, macrophage, and dendritic cell function in atherosclerosis. J Exp Med 213:1117–1131 11. Ginhoux F, Jung S (2014) Monocytes and macrophages: developmental pathways and tissue homeostasis. Nat Rev Immunol 14:392–404 12. Geissmann F, Manz MG, Jung S et al (2010) Development of monocytes, macrophages, and dendritic cells. Science 327:656–661 13. Yona S, Kim KW, Wolf Y et al (2013) Fate mapping reveals origins and dynamics of monocytes and tissue macrophages under homeostasis. Immunity 38:79–91 14. Mildner A, Yona S, Jung S (2013) A close encounter of the third kind monocyte-derived cells. Adv Immunol 120:69–103 15. Martinez FO, Gordon S (2014) The M1 and M2 paradigm of macrophage activation: time for reassessment. F1000Prime Rep 6:13 16. Gordon S (2007) The macrophage: past, present and future. Eur J Immunol 37(Suppl 1): S9–S17 17. Gordon S, Martinez FO (2010) Alternative activation of macrophages: mechanism and functions. Immunity 32:593–604 18. McElvania Tekippe E, Allen IC, Hulseberg PD et al (2010) Granuloma formation and host defense in chronic mycobacterium tuberculosis

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infection requires PYCARD/ASC but not NLRP3 or caspase-1. PLoS One 5:e12320 19. Wen H, Lei Y, Eun SY et al (2010) Plexin-­A4-­ semaphorin 3A signaling is required for tolllike receptor- and sepsis-induced cytokine storm. J Exp Med 207:2943–2957 20. Mesquita FS, Thomas M, Sachse M et al (2012) The salmonella deubiquitinase SseL inhibits selective autophagy of cytosolic aggregates. PLoS Pathog 8:e1002743 21. Sharif O, Bolshakov VN, Raines S et al (2007) Transcriptional profiling of the LPS induced NF-kappaB response in macrophages. BMC Immunol 8:1 22. Selinummi J, Ruusuvuori P, Podolsky I et al (2009) Bright field microscopy as an alternative to whole cell fluorescence in automated analysis of macrophage images. PLoS One 4:e7497 23. Edwards JP, Zhang X, Frauwirth KA et al (2006) Biochemical and functional characterization of three activated macrophage populations. J Leukoc Biol 80:1298–1307 24. Wynn TA, Barron L, Thompson RW et al (2011) Quantitative assessment of macrophage functions in repair and fibrosis. Curr Protoc Immunol. Chapter 14: Unit14 22 25. Allen IC, Wilson JE, Schneider M et al (2012) NLRP12 suppresses colon inflammation and tumorigenesis through the negative regulation of noncanonical NF-kappaB signaling. Immunity 36:742–754 26. Allen IC, TeKippe EM, Woodford RM et al (2010) The NLRP3 inflammasome functions as a negative regulator of tumorigenesis during colitis-associated cancer. J Exp Med 207: 1045–1056 27. Allen IC, Moore CB, Schneider M et al (2011) NLRX1 protein attenuates inflammatory responses to infection by interfering with the RIG-I-MAVS and TRAF6-NF-kappaB signaling pathways. Immunity 34:854–865

Chapter 4 Bone Marrow-Derived Dendritic Cells Kelly Roney Abstract Dendritic cells are a specialized type of antigen-presenting cell that bridges both innate and adaptive immune system function. While much is understood about dendritic cells and their role in the immune system, the study of these cells is critical to gain a more complete understanding of their function. The isolation and culture of dendritic cells from mouse tissues can be challenging, due in part to the low number of cells isolated. The following protocol outlines methods to optimize the isolation and culture of large numbers of dendritic cells from mouse bone marrow to facilitate a broad range of downstream experimental applications. Key words Dendritic cell, Mouse, Bone marrow, GM-CSF, IL-4

1  Introduction Dendritic cells are located throughout the body, including circulating in the bloodstream, and localized in tissues such as the skin, spleen, lung, and lymph nodes. It is critical to understand mechanisms impacting dendritic cell function due to their broad roles in a variety of immune system processes, including bridging innate and adaptive immune system activation, antigen processing and presentation, and T cell activation [1–3]. Due in part to the small number of cells that can be isolated, ex vivo dendritic cell studies can be difficult. To circumvent this limitation, methods have been optimized to enable the isolation and culture of high numbers of dendritic cells derived from mouse bone marrow [4]. The protocol described in this chapter presents one of these validated and highly efficient methods [5–7].

2  Materials 1. 1× Phosphate buffered saline (PBS). 2. 70% ethanol. 3. Roswell Park Memorial Institute media (RPMI)-1640. Irving C. Allen (ed.), Mouse Models of Innate Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1960, https://doi.org/10.1007/978-1-4939-9167-9_4, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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4. Complete RPMI: 500 ml of RPMI-1640, 5.5 × 105 mol/l of 2-mercaptoethanol, 25 mM/l of HEPES, 100 U/ml of penicillin, 100 μg/ml of streptomycin sulfate, 50 ml of heat-­ inactivated fetal bovine serum (use serum of sufficient grade for primary cell culture). 5. Buffered ammonium chloride (ACK) lysis buffer: Add 4.14 g of NH4Cl, 0.5 g of KHCO3, 18.8 mg of Na2 EDTA into 300 ml of double distilled water (or equivalent), and stir to dissolve. Bring the solution up to 500 ml using double distilled water (or equivalent) and adjust the pH to 7.2–7.4 with 1 N HCl. Filter the solution with a 70 μm nylon filter, and store at room temperature. 6. Mouse granulocyte-macrophage colony-stimulating factor (GM-CSF). 7. Mouse Interleukin-4 (IL-4). 8. Mouse tumor necrosis factor (TNF alpha). 9. 100 mm × 20 mm culture plate without tissue culture treatment. 10. 6 well culture plate without tissue culture treatment. 11. Sterilized 4 in. (or similar size) forceps. 12. Sterilized 4 in. (or similar size) scissors. 13. Sterile paper towels or other absorbent material. 14. Sterile needles: 22 gauge × 1 in. (see Note 1). 15. 1 ml sterile syringe (see Note 1). 16. 50 ml conical sterile polypropylene tubes. 17. 15 ml conical polypropylene tubes.

3  Methods 3.1  Bone Marrow Isolation (See Note 2)

1. Euthanize the mouse according to local animal care, and use committee guidelines and regulations. Bone marrow should be harvested immediately following euthanasia for best yield. 2. Remove the skin from the abdomen down. Remove the hind legs by blunt dissection. Separate the femur from the tibia by cutting the connection point with scissors (see Note 3). 3. Remove the muscle and as much connective tissue as possible from the femurs and tibias by firmly grasping the top of the bone with two sets of forceps held in an upside down “V” while resting the bottom of the bone on sterilized paper towels. Slide the forceps down the bone while firmly grasping the bone, removing the muscle and tissue. The muscle and tissue remaining at the bottom of the bone may be removed with scissors.

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4. Place the harvested bone into a 100 mm non-tissue culture treated plate on ice filled with PBS until all bones are harvested (see Note 4). 5. While grasping the bone with forceps, remove the epiphyses by cutting the tips of the bone with scissors (see Note 5). 6. Fill a 100 mm non-tissue culture treated plate with cold complete RPMI (enough to cover the bottom). Prepare a 1 ml syringe by fitting with a 27 gauge needle and filing with cold complete RPMI. 7. Using the prefilled syringe, remove the bone marrow from the bone by placing the tip of the needle in the bone while grasping the bone with forceps and injecting complete RPMI to flush out the marrow into the petri dish containing complete RPMI (see Note 6). 8. Completely remove the bone marrow from all four leg bones and combine into a single petri dish. Using the same syringe used to flush the bones, gently disperse any clumps of bone marrow by pulling the media and bone marrow into the syringe and releasing. 9. Place a 70 μm cell strainer atop a 50 ml conical tube. Pipette the RPMI/bone marrow cell solution from the petri dish though the cell strainer. 10. Remove the plunger from the syringe, and use the rubber tip of the plunger to disperse any bone marrow or tissue clumps left in the strainer. Rinse the cell strainer with complete RPMI (see Note 7). 11. Spin the media/cell mixture at 13,523 × g at 4 °C for 5 min to pellet the cells. 12. Discard the media by gently tilting the tube and pouring off the media into a waste disposal beaker. Recap the tube and gently tap to break up the cell pellet. Place the tube on ice. 13. Add 1.0 ml of cell lysis solution to the cells and gently tap the tube with your finger to mix the lysis solution for 30–60 s. 14. Immediately add 50 ml of ice cold PBS to dilute the lysis buffer. 15. Centrifuge the PBS/cell mixture at 13,523 × g at 4 °C for 5 min to pellet the cells. Remove the PBS by gently tilting the tube and pouring off the PBS into a waste disposal beaker. Recap and gently tap the tube with your finger to break up the cell pellet. 16. Resuspend the cell pellet in 50 ml of ice cold PBS to wash the cells. Centrifuge the PBS/cell mixture at 13,523 × g at 4 °C for 5 min to pellet the cells. 17. Resuspend the cells in 10 ml of cold complete RPMI, and place on ice. Remove 100 μl of the cell suspension, and count the cells.

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Bone marrow harvested from two tibias and two fibias should yield approximately 70 × 106 cells. Health, age, and transplant status of the mouse may affect bone marrow yields [8]. 3.2  Dendritic Cell Culture

1. Day 0: Plate 2 × 106 cells in 20 ml of media with 20 ng/ml of GM-CSF onto a 100 mm non-tissue culture treated plate (see Notes 8 and 9). 2. Day 3: Add 20 ml of media with GM-CSF to bring the final concentration of GM-CSF for the whole culture (a total of 40 ml) to 10 ng/ml. 3. Day 6: Remove 20 ml from the plate. Centrifuge to recover the cells, discard the supernatant, and resuspend the pellet in 20 ml of complete RPMI. Add GM-CSF and IL-4 so that the final concentration of the entire culture is 10 ng/ml of GM-CSF and 10 ng/ml of IL-4 (see Note 10). 4. Day 8: Repeat step 3, only use 5 ng/ml of GM-CSF and 10 ng/ml of IL-4 for the total culture concentration. 5. Day 10: Harvest the dendritic cells, which will be floating or lightly adherent in the culture. To harvest, remove the media containing the DCs and transfer to a 50 ml conical tube. Plates can be rinsed gently with warmed PBS to remove the lightly adherent cells. Avoid harvesting the adherent cells. 6. Dendritic cells may be used at Day 10 or may be further matured (see Note 11).

3.3  Maturation

1. Day 10: Wash the cells twice in cold PBS and count. Resuspend cells in complete RPMI media at a concentration of 1 × 106 cells/ml. Add 1 ml of cells to each well of a 6-well bacterial culture plate (see Note 11). 2. Add 1 ml of complete RPMI to each well so that the concentration of the culture is 5 ng/ml of GM-CSF, 5 ng/ml of IL-4, and 20 ng/ml of TNF (see Note 11). 3. Day 10 + 1: The following day add 1.0 ml of media with TNF to each well so that the concentration in the entire culture is 10 ng/ml. Harvest the cells for use the following day (see Note 11). 4. Day 10 + 2: The mature dendritic cells will be loosely adherent. To harvest, remove the media from the culture and transfer to a 15 ml conical tube. Add cold PBS to the culture plate, and gently loosen cells from the plate with a cell scraper. Add the cold PBS cell mixture to the media in the 15 ml conical tube, pellet the cells by centrifugation, and wash in cold PBS to remove residual cytokines (see Note 12).

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4  Notes 1. Needle length and gauge may be varied to meet individual preferences. A 5 ml or 10 ml syringe may also be used instead of the 1 ml syringe. 2. All procedures should be carried out at room temperature unless otherwise specified. All procedures except centrifugation should be performed in a biological safety cabinet. Tissue culture techniques should be utilized throughout the procedure. 3. A beaker filled with 70% ethanol can be used to dip forceps and scissors. This technique is also useful to remove hair or tissue from the tools during bone marrow isolation. 4. Bones may be sterilized by submerging in cold 70% ethanol for 2 min in a 100 mm bacterial culture plate and then rinsing two times by submerging bones in a dish with cold PBS for 2 min to rinse away the ethanol. We have not found this step necessary if good tissue culture techniques are used for bone marrow isolation. 5. If bone marrow material is a limiting factor, the epiphyses can be cut into pieces in compete media in a petri dish. The resulting bone marrow cells can be isolated by passing the complete media and bone through a 70 μm size cell strainer. 6. The bone should turn a brighter white and more translucent as the marrow is flushed. Removal of the syringe to the opposite end of the bone or an additional flush of complete media is often necessary. If necessary, the syringe can be refilled from the media in the marrow collection petri dish. 7. Cells can be placed on ice at this stage for 1–2 h if complete media is used. 8. Count bone marrow cells carefully. Over-plating may result in over-proliferated cells that will not differentiate into dendritic cells. 9. If a smaller number of cells are desired from a single plate, cells may be grown in100 × 20 mm non-tissue culture treated plates in 10 ml of media. Adjust all steps forward to a starting amount of 10 ml of media. 10. For best results, warm the resuspended cells to 37 °C before returning to the culture plate to prevent cooling of the cell culture plate. 11. Dendritic cells that are in a more immature state may be better for some experiments, such as antigen uptake assays, whereas other experiments may require a more mature cell that has received more cytokine stimulation.

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12. Alternative Day 10 for larger cell cultures: Suspend cells at 1 × 106 cells per ml and plate 5 ml on a 100 mm non-tissue culture treated plate. Then add 5 ml of media. The final cytokine concentrations should be 5 ng/ml of GM-CSF, 5 ng/ml of IL-4, and 20 ng/ml of TNF. Alternative Day 10 + 1: Add 10 ml of media with 10 ng/ml of TNF for the entire culture. Harvest cells on alternative D10 + 2. References 1. Altfeld M, Fadda L, Frleta D et al (2011) DCs and NK cells: critical effectors in the immune response to HIV-1. Nat Rev Immunol 11:176–186 2. Bousso P (2008) T-cell activation by dendritic cells in the lymph node: lessons from the movies. Nat Rev Immunol 8:675–684 3. Roy RM, Klein BS (2012) Dendritic cells in antifungal immunity and vaccine design. Cell Host Microbe 11:436–446 4. Inaba K, Inaba M, Romani N et al (1992) Generation of large numbers of dendritic cells from mouse bone marrow cultures supplemented with granulocyte/macrophage colonystimulating factor. J Exp Med 176:1693–1702 5. Lutz MB, Kukutsch N, Ogilvie AL et al (1999) An advanced culture method for generating

large quantities of highly pure dendritic cells from mouse bone marrow. J Immunol Methods 223:77–92 6. Van Deventer HW, Serody JS, Mckinnon KP et al (2002) Transfection of macrophage inflammatory protein 1 alpha into B16 F10 melanoma cells inhibits growth of pulmonary metastases but not subcutaneous tumors. J Immunol 169:1634–1639 7. Wong AW, Brickey WJ, Taxman DJ et al (2003) CIITA-regulated plexin-A1 affects T-celldendritic cell interactions. Nat Immunol 4:891–898 8. Lutz MB, Rossner S (2007) Factors influencing the generation of murine dendritic cells from bone marrow: the special role of fetal calf serum. Immunobiology 212:855–862

Chapter 5 Quantification and Visualization of Neutrophil Extracellular Traps (NETs) from Murine Bone Marrow-Derived Neutrophils Linda Vong, Philip M. Sherman, and Michael Glogauer Abstract Neutrophils are some of the first leukocytes to respond to inflammatory stimuli. Once recruited, these cells are equipped with an assortment of proteolytic enzymes and antimicrobial factors that disarm and degrade pathogens. Neutrophils employ a highly novel mechanism to contain and trap bacteria in the local inflammatory microenvironment, termed neutrophil extracellular traps (NETs). During NET formation, neutrophils eject weblike structures of chromatin, which captures and immobilizes invading pathogens. In this chapter, we describe protocols to isolate bone marrow-derived neutrophils from mice. We further describe in vitro methods to spectrophotometrically quantify, immunolabel, and visualize NET structures. Key words Neutrophil extracellular trap, Bone marrow neutrophil, Nucleic acid stain, Histone H3, Elastase, Fluorescence microscopy

1  Introduction Neutrophil maturation occurs in the bone marrow and involves the synthetization and packaging of enzymes and antimicrobial proteins into specialized granules [1]. Neutrophils are highly recruited to sites of inflammation and have several unique functions in response to pathogen exposure, including the formation of neutrophil extracellular traps (NETs). NET formation is a novel type of cell death that is characterized by the rapid externalization of weblike strands of decondensed chromatin, which includes both DNA and histones. These strands are highly decorated with a diverse assortment of antimicrobial and proteolytic enzymes [2–4]. Together, these defined characteristics can be utilized in experimental settings to specifically label and characterize NETs. Mouse bone marrow contains a large number of functionally competent neutrophils. These bone marrow-derived neutrophils survive much longer ex vivo compared to blood neutrophils [5].

Irving C. Allen (ed.), Mouse Models of Innate Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1960, https://doi.org/10.1007/978-1-4939-9167-9_5, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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In the following protocol, we describe methods to collect mature mouse bone marrow-derived neutrophils (BMDN) from the tibia and femur. These cells are purified by discontinuous Percoll density gradient centrifugation [6]. BMDNs collected from the 80/65% Percoll interface contain >85% BMDN, which is confirmed by FACS analysis using anti-GR-1 antibodies. To robustly quantify NET formation, BMDNs were incubated with Sytox Green, which is a cell-­ impermeable fluorescent DNA dye. The use of Sytox Green ensures that the measurement of extracellular DNA is specific to cells where membrane integrity has been compromised, which is typical during the formation of NETs, rather than cells that are viable with intact membranes. Fluorescence emission is monitored following a 3 h incubation with phorbol 12-myristate 13-acetate (PMA), which is a potent NET inducer [7]. The percentage of NET formation can be determined by subtracting the background fluorescence and dividing by the maximal fluorescence signal detected from lysed BMDN. NETs can also be visualized directly. Here, BMDNs can be plated onto poly-l-lysine-­coated coverslips and then stained with antibodies for the NET components DNA, histone H3 [7], or elastase [8]. Together, the protocols described in this chapter provide complementary approaches to quantify and visualize NETs.

2  Materials 2.1  Bone Marrow Neutrophil Isolation

1. 8–9-week-old male or female C57BL/6 mice. 2. Laminar flow hood. 3. Small dissection scissors. 4. Forceps. 5. Lint-free wipes. 6. Sterile polyethylene disposable transfer pipettes. 7. 60 × 15 mm sterile polystyrene petri dishes. 8. 50 mL conical tubes. 9. 15 mL conical tubes. 10. 10 mL syringes. 11. 25G5/8 needles. 12. 20G needles. 13. 70% ethanol. 14. Ice-cold deionized water. 15. Ice, ice bucket. 16. MEM alpha cell culture medium 1× (Gibco). Store at 4 °C. 17. 1× Phosphate-buffered saline (PBS), pH 7.4 without calcium chloride/magnesium chloride. Store at 4 °C.

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18. Hanks balanced saline solution (HBSS) with calcium chloride/ magnesium chloride. Store at 4 °C. 19. Percoll density gradients: Prepare 100% Percoll stock by mixing 90 mL of Percoll (pH 8.5–8.9) with 10 mL of 10× Dulbecco’s phosphate-­ buffered saline. In a 50 mL conical tube, prepare 80% (mix 40 mL of 100% Percoll with 10 mL of 1× PBS), 65% (mix 32.5 mL of 100% Percoll with 17.5 mL of 1× PBS), and 55% (mix 27.5 mL of 100% Percoll with 22.5 mL of 1× PBS) Percoll gradient solutions. Store at 4 °C. 20. 3.6% (w/v) NaCl: Dissolve 3.6 g of NaCl in 100 mL deionized water. Store at 4 °C. 21. Turk’s solution: Dissolve 0.1% crystal violet in 3% acetic acid (prepared in sterile water). Shake vigorously. Store at room temperature. 2.2  Quantification of Neutrophil Extracellular DNA

1. Bone marrow-derived neutrophils (BMDN, 1 × 106 cells/ mL). 2. Fluorescence microplate reader equipped with filters to detect excitation/emission maxima: 485/520 nm. 3. Humidified CO2 incubator. 4. Black 96-well microplate. 5. 96-well microplate lids. 6. Microplate sealing tape. 7. Hanks balanced saline solution (HBSS) with calcium chloride/ magnesium chloride. Store at 4 °C. 8. 5 mM Sytox Green nucleic acid stain (stock). Protect from light and store at −20 °C. Just prior to addition to wells, prepare a 10× working solution (50 μM) by diluting 5 mM stock solution 1:100 with HBSS, into a foil-wrapped conical tube. 9. DNase 1 (RNase-free), 2 Units/μL. Store at −20 °C. 10. 1 mM Phorbol 12-myristate 13-acetate (PMA) (stock): Dissolve 1 mg of PMA in 1.62 mL dimethyl sulfoxide). Aliquot and store at −20 °C. 11. 10% Triton X-100 (stock).

2.3  Immunofluore­ scence Visualization of NETs (Fig. 1)

1. Bone marrow-derived neutrophils (BMDN, 1 × 106 cells/ mL). 2. Epi-fluorescence or confocal microscope equipped with filters to detect excitation/emission maxima: 358/461 nm (DAPI), 550/570 nm (TRITC), and 495/519 nm (Alexa Fluor 488). 3. Humidified CO2 incubator. 4. Sterile 12-well cell culture plates. Store at room temperature.

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Fig. 1 Visualization of murine bone marrow-derived NETs by immunofluorescence microscopy. Resting BMDN or BMDN activated with PMA (100 μM, 3 h at 37 °C) was fixed and immunostained for DNA (DAPI; panels a, e), elastase (panels b, f), or histone H3 (panels c, g). Overlay of the three channels is shown in panels (d) and (h) for resting and PMA-activated BMDN, respectively. Scale bar = 100 μm

5. 12 mm round poly-l-lysine-coated glass coverslips. Store at 4 °C. 6. 75 × 25 × 1 mm microscope slides. Store at room temperature. 7. 1 mL microcentrifuge tubes. 8. Ice-cold methanol. Store at −20 °C. 9. 1× phosphate-buffered saline (PBS), pH 7.4. Store at 4 °C. 10. Phosphate-buffered saline supplemented with Tween-20 (PBS-­Tween). Mix 1 L PBS with 0.5 mL Tween-20. Store at room temperature. 11. 1 mM Phorbol 12-myristate 13-acetate (PMA) (stock): Dissolve 1 mg of PMA in 1.62 mL of dimethyl sulfoxide. Aliquot and store at −20 °C. Just prior to use, prepare a 1 μM working stock solution by diluting 1:1000-fold into HBSS. Store on ice until ready for use. 12. Fluorescent mounting medium. Store at 4 °C. 13. 5 mg/mL 4′,6-diamidino-2-phenylindole, diacetate (DAPI) (stock): Dissolve 10 mg in 2 mL of deionized water. Solution may take some time to dissolve completely and may require sonication. For long-term storage, aliquot and store at −20 °C. For short-term storage, store at 4 °C (stable for at least 6 months). 14. Histone H3 (D1H2) XP rabbit monoclonal antibody. Store at −20 °C. 15. Neutrophil elastase polyclonal antibody. Aliquot and store at −20 °C. 16. Goat anti-rabbit TRITC secondary antibody. Aliquot and store at −20 °C.

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17. Goat anti-rabbit Alexa Fluor 488 secondary antibody. Store at 4 °C. 18. Blocking buffer: 3% bovine serum albumin (BSA) prepared in PBS. Dissolve 0.3 g BSA in 10 mL of PBS. Store at 4 °C.

3  Methods 3.1  Bone Marrow-­ Derived Neutrophil Isolation

1. Sacrifice mouse by cervical dislocation (alternate methods such as CO2 asphyxiation may also be utilized—refer to institutional guidelines). 2. In a laminar flow hood, spray the front (ventral) side of the mouse with 70% ethanol, and make a lateral incision at the midline. Strip away the fur to expose the lower abdomen, soft tissue, and bone of the hind limbs (see Note 1). 3. Use scissors to make a cut above the hip joint, detach, and transfer intact hind limb to a 50 mL conical tube containing 20 mL of MEM alpha medium. Repeat with the second hind limb. 4. Gently cut away and remove soft tissue from the tibia and femur using scissors and lint-free wipes (see Note 2). Separate the tibia from the femur and transfer to a 60 × 15 mm petri dish containing MEM alpha medium. Repeat with the second hind limb. 5. Transfer tibias and femurs to a second petri dish containing 70% ethanol. Soak the bones for ~30 s, and then allow them to dry. 6. Use scissors to cut the proximal and distal ends off the tibia/ femur, and flush the marrow into a third petri dish using a 10 mL syringe (containing 8 mL of MEM alpha medium) with a 25G5/8 needle attached (see Note 3). 7. Using a fresh syringe, attach a 20G needle, and very gently aspirate the bone marrow to separate any clumps. This process should be repeated approximately 4–5 times. Repeat this procedure with the remaining bones. 8. Transfer to a 15 mL conical tube, and centrifuge at 400 × g for 10 min at room temperature. 9. Gently pour off the supernatant, and resuspend the cell pellet with 1 mL of PBS (without calcium chloride/magnesium chloride). 10. Prepare a Percoll density gradient. In a 15 mL conical tube, carefully add 4 mL of 80% Percoll. Gently overlay this first layer with 3 mL of 65% Percoll, followed by 3 mL of 55% Percoll. Care should be taken to avoid mixing or disturbing the gradient solutions as they are added to the tube (see Note 4). Allow the Percoll tubes to stand for 5 min, and then carefully add the cell suspension, prepared in step 9, to the top of the density

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gradient. Centrifuge at 1000 × g for 30 min at room temperature, without braking. 11. Remove the centrifuge tube and visually inspect the gradient. The bone marrow neutrophils will have separated into the 80%/65% Percoll interface. 12. Gently dispose of the uppermost serum and 55% Percoll layers using a disposable sterile transfer pipette. In a fresh 15 mL conical tube, collect the upper portion of the 65% Percoll gradient and cells at the 80%/65% gradient interface. Top-up volume to 14 mL with 1 mL of 1× PBS (without calcium chloride/magnesium chloride) and centrifuge at 400 × g for 10 min at 4 °C. 13. Pour off the supernatant, and lyse the remaining red blood cells by gently resuspending the cell pellet in 3 mL of ice-cold deionized water. Leave undisturbed for 30 s to 1 min. Add 1 mL of 3.6% NaCl and mix gently. Centrifuge at 400 × g for 5 min at 4 °C. 14. Pour off the supernatant and gently resuspend the bone marrow-­ derived neutrophil cell pellet with 1 mL of HBSS (with calcium chloride/magnesium chloride). Determine the concentration using a Neubauer hemocytometer. Mix 10 μL of cell suspension with 90 μL of HBSS and 5 μL of Turk’s solution. Load 10 μL onto a hemocytometer. 15. Dilute bone marrow neutrophils to a concentration of 1 × 106 cells/mL using HBSS. Typically, ~6 × 106 bone marrowderived neutrophils can be harvested per mouse. 3.2  Quantification of Extracellular DNA

The following protocol outlines the procedures for measuring neutrophil extracellular DNA, which is an index for the formation of NETs. BMDN (see Subheading 3.1) or neutrophils from other sources (such as cell lines or peripheral blood) are plated at a density of 1 × 105 cells per well and activated with PMA (Table 1).

Table 1 Treatment conditions used for the quantification of neutrophil extracellular DNA (NETs)

HBSS (μL)

BMDN (1 × 106/mL) (μL)

Triton X-100 (10%) (μL)

PMA (1 μM) (μL)

DNase 2 U/μL

Sytox Green (50 μM) (μL)

BMDN

170

100

–

–

–

30

BMDN+Triton X-100

160

100

10

–

–

30

BMDN+PMA (100 nM)

140

100

–

30

–

30

BMDN+PMA (100 nM) + DNase

137.5

100

–

30

2.5

30

BMDN+DNase

167.5

100

–

–

2.5

30

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The cell-­ impermeable DNA-binding dye, Sytox Green, is then added and the resulting fluorescence quantified on a fluorescence microplate reader. This protocol can be modified to incorporate other cell activators or additional inhibitors, and measurements made at variable time points to monitor the kinetics of NET formation. 1. In a black 96-well microplate (see Note 5), prepare duplicates of the treatment wells shown in Table 1. 2. Using a pipette, add HBSS, 1 × 105 BMDN, and Triton X-100 (to determine total DNA content) or PMA (an activator of NET formation) to their respective wells. Cover the microplate with a lid, and transfer to a humidified incubator (37 °C, 5% CO2). 3. After 2 h, add DNase to wells that require DNA degradation (based on experimental design), and transfer back to the humidified incubator (37 °C, 5% CO2) (see Note 6). 4. After a further 45 min, carefully add 30 μL of Sytox Green (10× working stock; 50 μM) to each well, mix, and transfer back to the humidified incubator. Allow to stand for a further 15 min, after which the plate can be sealed with microplate sealing tape and fluorescence quantified on a fluorescence microplate reader (see Note 7). 5. To quantify the amount of extracellular DNA (as a percentage of total DNA), subtract the fluorescence intensity of the DNase-­ containing wells from the comparative control, and divide by the fluorescence intensity emitted from “BMDN+Triton X-100” wells (total DNA present). Example Percentage total DNA ( induced by PMA ) =

Fluorescence intensity ( BMDN + PMA ) - Fluorescence intensity ( BMDN + PMA + DNase ) ntensity ( BMDN + Triton X　 100 ) Fluorescence in

3.3  Immuno­ fluorescence Visualization of NETs

Visualization of NETs by immunofluorescence offers a complementary measure of extracellular DNA quantification, Subheading3.2. While there are many markers for NETs, this protocol uses DAPI (to stain for DNA), histone H3, and the serine protease elastase to label bone marrow NETs. 1. Place one poly-l-lysine-coated glass coverslip into each well of a sterile 12-well cell culture plate (see Note 8). 2. Plate BMDN onto the center of the poly-l-lysine-coated coverslips at a density of 1 × 105 cells, by gently pipetting 100 μL of BMDN cell suspension (1 × 106 cells/mL). To induce NETs with PMA (100 nM), mix 100 μL of BMDN cell suspension

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(1 × 106 cells/mL) with 15 μL of 10× PMA working stock solution (1 μM) and 35 μL of HBSS, in a separate microcentrifuge tube, before plating onto poly-l-lysine-coated coverslips. Replace the lid of the cell culture plate, and transfer to a humidified incubator (37 °C, 5% CO2) (see Note 9). 3. Incubate for 3 h. 4. Gently wash away cells that have not adhered with PBS. Aspirate with a pipette tip and discard (see Note 10). 5. Transfer the cell culture plate, containing coverslips, to a fume hood and fix adherent cells by adding 400 μL of neutral-buffered formalin. Allow to stand for 15 min at room temperature. 6. Gently aspirate neutral-buffered formalin using a pipette tip, discard, and replace wells with 800 μL of PBS. 7. Repeat a further three times. 8. Transfer coverslips containing adherent BMDN to a new cell culture plate, containing 800 μL of PBS per well. Store at 4 °C (overnight) until ready to perform immunofluorescence labeling. 9. For immunofluorescence labeling of NETs, permeabilize adherent cells by transferring the prepared coverslips to a new cell culture plate, and add enough ice-cold 100% methanol to sufficiently cover the surface (to a depth of 3.5 mm, ensuring the cells do not dry out). Transfer to a −20 °C freezer and incubate for 10 min. 10. Aspirate with a pipette tip, and gently wash coverslips with PBS for 5 min. 11. Block non-specific binding sites with blocking buffer (3% BSA prepared in PBS) for 1 h at room temperature (see Note 11). 12. Aspirate with a pipette tip, and gently wash coverslips with PBS for 5 min. 13. To stain for histone H3, prepare primary histone H3 antibody (1:100 dilution) in blocking buffer supplemented with 0.3% Triton X-100. Allow for 200 μL of diluted primary antibody per coverslip. For example, for 1 mL of diluted primary antibody, mix the following: 10 μL of histone H3 antibody, 33 μL of 10% Triton X-100 (prepared with deionized water), and 957 μL of blocking buffer (3% BSA prepared in PBS). 14. Transfer coverslips to a humidified chamber and incubate at 4 °C overnight, with gentle rotation (see Note 12). 15. Aspirate with a pipette tip, and gently wash coverslips with PBS-­Tween for 5 min. Repeat three times. 16. Prepare goat anti-rabbit TRITC secondary antibody (1:400 dilution) in blocking buffer supplemented with 0.3% Triton

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X-100. Pipette diluted antibody onto coverslips and incubate in a humidified chamber (protected from light) for 1 h at room temperature. For all subsequent steps, protect coverslips from light. 17. Aspirate with a pipette tip, and gently wash coverslips with PBS-­Tween for 10 min. Repeat three times. 18. Incubate coverslips with blocking buffer (3% BSA prepared in PBS) for 1 h at room temperature. 19. To stain for elastase, prepare primary elastase antibody (1:200 dilution) in blocking buffer supplemented with 0.3% Triton X-100. Pipette diluted antibody onto coverslips, and incubate in a humidified chamber (protected from light) for 1 h at room temperature. 20. Aspirate with a pipette tip, and gently wash coverslips with PBS-­Tween for 5 min. Repeat three times. 21. Prepare goat anti-rabbit Alexa Fluor 488 secondary antibody (1:400 dilution) in blocking buffer supplemented with 0.3% Triton X-100. Transfer coverslips to a humidified chamber (protected from light), and incubate for 1 h at room temperature. 22. Aspirate with a pipette tip, and gently wash coverslips with PBS-­Tween for 10 min. Repeat three times. 23. To counterstain for DNA, incubate coverslips with 1:12,500 dilution of DAPI (mix 0.4 μL of DAPI stock solution with 5 mL of PBS), 5 min at room temperature. 24. Aspirate with a pipette tip, and gently wash coverslips with PBS. Repeat several times. 25. Mount coverslips using a small drop of fluorescent mounting medium per microscope slide (see Note 13). Allow the preparation to dry overnight by storing the slides in a slide holder at room temperature, protected from light. Thereafter, transfer to 4 °C for storage. 26. Visualize the DNA, histone H3, and elastase staining on a confocal or epi-fluorescence microscope, equipped with filters suitable for DAPI (excitation/emission: 358/461 nm), TRITC (excitation/emission: 550/570 nm), and Alexa Fluor 488 (excitation/emission: 495/519 nm).

4  Notes 1. Spraying with ethanol helps to reduce the amount of fur that sticks to exposed tissue. Make a shallow cut or incision with scissors so as to not pierce the intestinal tract, and continue cutting laterally to completely remove skin from the lower abdomen to hind paws.

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2. Hold the tibia/femur between your thumb and forefinger, and use lint-free wipes (providing more friction) to remove the surrounding soft tissue. 3. Flush the bone marrow out, using about 2 mL of media per tibia/femur. The bones should appear transparent afterwards. 4. Percoll gradients should be slowly added to the conical tube, with the tip of disposable transfer pipette touching the tube wall. If added too quickly, the separation of bone marrow cells between the discontinuous gradients will be less effective. 5. Black microplates are used for fluorescence assays as they reduce the level of auto fluorescence and therefore background signal. 6. DNase can be added to the assay 60 min before quantification of extracellular DNA. If increasing or reducing the overall incubation period (i.e., from 3 h), adjust accordingly. 7. Sytox Green is added to the assay 15 min before the quantification of extracellular DNA. If increasing or reducing the overall incubation period (i.e., from 3 h), adjust the time point at which the DNA stain is added accordingly. Prior to measurement, briefly shake the plate to mix the well contents. This option is available on most microplate readers. 8. 12-well cell culture plates are used to contain the coverslips, although other holders such petri dishes, etc. can also be used. Ensure that, if using 24-well plates, the coverslips can be removed with forceps without breaking. 9. At least 2–3 replicates of each treatment should be prepared, as the cell suspension occasionally leaks from the coverslip (onto the cell culture plate surface) and can then no longer be used. 10. NETs are very fragile and can easily be dislodged. Sufficient care should be taken during all washes—only use a pipette, and gently aspirate. 11. Blocking buffer can be prepared the day before and stored at 4 °C for at least a week. 12. An easy alternative to commercially available but expensive chambers is to line a shallow plastic container (large enough to hold the tissue culture plate, containing coverslips) with moistened paper towels and replace the lid. 13. To mount the coverslips, place a small drop of fluorescent mounting medium onto the surface of a glass microscope slide. Use forceps to pick up the coverslip, and gently dry the underside on paper towel or lint-free wipes. Place one edge of the coverslip just outside of the mounting medium, and lower until it comes into contact with the medium. Release the remainder of the coverslip to allow the mounting medium to distribute evenly.

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References 1. Borregaard N (2010) Neutrophils, from marrow to microbes. Immunity 33:657–670 2. Brinkmann V, Reichard U, Goosmann C et al (2004) Neutrophil extracellular traps kill bacteria. Science 303:1532–1535 3. Papayannopoulos V, Metzler K, Hakkim A et al (2010) Neutrophil elastase and myeloperoxidase regulate the formation of neutrophil extracellular traps. J Cell Biol 191:677–691 4. Urban C, Ermert D, Schmid M et al (2009) Neutrophil extracellular traps contain calprotectin, a cytosolic protein complex involved in host defense against Candida albicans. PLoS Pathog 5:e1000639

5. Boxio R, Bossenmeyer-Pourie C, Steinckwich N et al (2004) Mouse bone marrow contains large numbers of functionally competent neutrophils. J Leukoc Biol 75:604–611 6. Chervenick PA, Boggs DR, Marsh JC et al (1968) Quantitative studies of blood and bone marrow neutrophils in normal mice. Am J Phys 215:353–360 7. Fuchs TA, Abed C, Goosmann R et al (2007) Novel cell death program leads to neutrophil extracellular traps. J Cell Biol 176:231–241 8. Lim MB, Kuiper JW, Katchky A et al (2011) Rac2 is required for the formation of neutrophil extracellular traps. J Leukoc Biol 90:771–776

Chapter 6 In Vitro Differentiation of Effector CD4+ T Helper Cell Subsets Kaitlin A. Read, Michael D. Powell, Bharath K. Sreekumar, and Kenneth J. Oestreich Abstract CD4+ T “helper” cells are key orchestrators of adaptive immune responses. Upon activation, naïve CD4+ T cells are capable of differentiating into a number of effector subsets that perform distinct immune functions. These subsets include T helper 1 (TH1), TH2, TH9, TH17, TH22, T follicular helper (TFH), and regulatory T cell (TREG) populations. The differentiation of these subsets is dependent, in large part, on the coordinated interplay between signals from the extracellular cytokine environment and downstream transcriptional networks. The use of in vitro T helper cell culture systems has been extensively employed to aid in the elucidation of the molecular mechanisms that govern the differentiation of each effector subset. Here, we provide a detailed summary of the differentiation conditions that are utilized to generate effector CD4+ T cell populations in vitro. Key words Adaptive immunity, CD4+ T helper cell, Cytokines, Effector CD4+ T cell differentiation, T helper 1, T helper 2, T helper 9, T helper 17, T regulatory cells

1  Introduction For more than 30 years, it has been recognized that naïve CD4+ T helper cells differentiate into distinct effector subsets responsible for performing divergent immune functions [1]. The initial description of TH1 and TH2 cells by Mosmann and Coffman, along with the subsequent characterization of additional populations such as TH9, TH17, TH22, TFH, and TREG populations, has challenged researchers to identify the regulatory cues that govern the differentiation of each subset [2–11]. The development of highly tractable culture systems that allow for the differentiation of many of these subsets in vitro has aided in the elucidation of lineage-specific regulatory requirements and has enhanced our overall understanding of T helper cell biology [12–15]. Initially, T cell populations were expanded in vitro via co-­ culture with both antigen and antigen-presenting cells (APCs). Irving C. Allen (ed.), Mouse Models of Innate Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1960, https://doi.org/10.1007/978-1-4939-9167-9_6, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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However, in 1985, Dennis Carson’s laboratory established that T cells could be activated in vitro through the use of monoclonal antibodies to the T cell receptor (TCR; αCD3) in the absence of APCs [16]. Since that time, in vitro stimulation conditions have been developed that utilize both αCD3 antibodies and those specific for the co-stimulatory receptor CD28 to mimic the APC/T helper cell interaction [17]. This simplified culture system circumvents the need for antigen-specific activation and is the basis for the in vitro culture conditions described in this chapter. In addition to signals derived from the T cell and co-­stimulatory receptors, effector cells also require signals from extracellular cytokines to dictate cellular phenotype. Cytokine signals are received through multimeric receptors and propagated largely through Janus kinase/signal transducer activator of transcription (JAK/ STAT) signaling pathways [18, 19]. STAT transcription factors subsequently regulate the expression of so-called lineage-defining transcription factors that are required for the differentiation of each effector subset (Fig. 1).

Clearance of intracellular pathogens (e.g. bacteria, viruses), anti-tumor responses

TH1

TFH

T-bet

Bcl-6

IL-12, IFN-γ, IL-2 Clearance of parasites (e.g. helminthic worms)

TH2 IL-4 IL-2

GATA3

IL-6 IL-21

Naïve CD4+ T cell

TH22

TH17

TGF-β IL-6 IL-1β IL-23

RORγt

Clearance of extracellular bacteria, mucosal immunity

IL-4 TGF-β IL-2

IL-6 IL-23

Clearance of extracellular pathogens, epidermal immunity

Provide help to B cells, promotion of high-affinity antibody production

TGF-β IL-2

TH9

AHR

PU.1

TREG Foxp3

Clearance of parasites, anti-tumor responses

Control of inflammation; production of antiinflammatory cytokines

Fig. 1 Schematic depicting known regulatory factors that govern effector T helper cell differentiation. Differentiation decisions of T helper cell subsets are regulated by signals from extracellular cytokines, which induce the expression and/or activity of intracellular lineage-defining transcription factors. Key cytokines that induce the differentiation of each subset are shown in gray. Recognized functions of each subset are indicated adjacent to each cell type. Lineage-defining transcription factors are shown inside the shaded box of each effector cell

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The specific cytokine signaling pathways and lineage-defining transcription factors that regulate individual T helper cell subset differentiation decisions have been expertly and thoroughly reviewed elsewhere [19–23]. Here, we provide a detailed summary of current culturing methods used to generate effector T helper cell subsets in vitro. These methods, and derivations thereof, will continue to serve as a valuable set of tools, alongside in vivo approaches, to aid researchers interested in obtaining a greater understanding of the regulatory mechanisms that govern the differentiation of T helper cell populations.

2  Materials 2.1  Disposables

1. 24-well tissue culture plates. 2. Sterile 15 mL conical tubes. 3. Sterile frosted-end microscope slides. 4. Sterile petri dishes. 5. Parafilm. 6. Additional tissue culture supplies as specified by manufacturer of T cell isolation kit.

2.2  Naïve Murine CD4+ T Cell Isolation Reagents

1. Sterile ultrapure water. 2. Sterile 1× DPBS without calcium chloride or magnesium chloride. 3. Sterile red blood cell (RBC) lysis buffer: 0.84% NH4Cl and complete IMDM media containing 10% FBS, 1% Pen/Strep, 50 μM β mercaptoethanol (see Note 1). 4. Naïve murine CD4+ T Cell Isolation Kit (see Note 2).

2.3  T Helper Cell Culture and Differentiation Reagents

1. Antibodies for T helper cell stimulation: α-CD3ε (clone 145-­ 2C11), α-CD28 (clone 37.51). 2. TH1 cell differentiation: α-IL-4 (clone 11B11), recombinant murine IL-12, recombinant human IL-2. 3. TH2 cell differentiation: α-IFNγ (clone XMG1.2), recombinant murine IL-4, recombinant human IL-2 (see Note 3). 4. TH9 cell differentiation: α-IFNγ, recombinant murine IL-4, recombinant human TGF-β1. 5. TH17 cell differentiation: α-IL-4, α-IFNγ, recombinant murine IL-6, recombinant human TGF-β1 (see Note 4). 6. iTreg cell differentiation: α-IL-4, α-IFNγ, ##ανδ recombinant human TGF-β1.

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3  Methods 3.1  Isolation of Naïve CD4+ T cells

1. One day prior to CD4+ T cell isolation, coat an appropriate number of wells (see Note 5) in a 24-well tissue culture plate with 500 μL/well 1× PBS containing α-CD3ε (5 μg/mL) and α-CD28 (10 μg/mL) (see Note 6). Wrap the plate with parafilm and incubate at 4 °C overnight. 2. On the day of isolation, prepare 15 mL conical tubes containing 5 mL ice-cold cIMDM for each spleen and pooled lymph node sample (see Note 7). Euthanize 5–8-week-old wild-type mice according to individual institutional guidelines, and create a sterile working surface by spraying thoroughly with 70% ethanol. In a tissue culture hood, harvest spleen and lymph nodes, and place into the prepared tubes containing media. From this point on, samples should be kept on ice as much as possible. 3. Transfer the first lymph node sample, including media, to a sterile petri dish. Use the textured ends of frosted microscope slides to generate a single-cell suspension of the lymph nodes (see Note 8). Rinse the end of the slides in media to ensure maximal recovery. Using a serological pipet, add an additional 5 mL of cIMDM to the cell suspension and rinse the surface of the petri dish thoroughly. Transfer the cell suspension back to the 15 mL conical tube and keep on ice while you prepare the remainder of the samples. 4. Once a single-cell suspension has been prepared for each spleen and lymph node sample, centrifuge the cells at 600 × g for 5 min at 4 °C. 5. Aspirate the supernatant from each sample and discard. Flick-­ mix each cell pellet thoroughly, and add an appropriate volume of RBC lysis buffer to each sample (4 mL for spleen, 2 mL for lymph nodes). Invert to mix and incubate at room temperature for exactly 3 min, beginning when the first sample is resuspended in RBC lysis buffer. 6. Following the 3-min incubation, add an appropriate volume of cIMDM to each sample (5 mL for spleen, 4 mL for lymph nodes). Invert to mix and incubate at room temperature for 6 min. 7. Following the above incubation, debris will have collected at the bottom of each 15 mL conical tube. Without inverting, transfer spleen and lymph node samples for the same biological replicate to a single, fresh 15 mL conical tube. Centrifuge the pooled sample at 600 × g for 5 min at 4 °C. Aspirate and discard the supernatant. 8. The cell pellet is now ready for naïve CD4+ T cell isolation according to manufacturer’s instructions for the negative selection kit chosen.

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3.2  Differentiation of CD4+ T Cell Subsets

79

1. Prepare stimulation wells by aspirating the coating mixture and rinsing each well twice with sterile 1× PBS. Following isolation, naïve CD4+ T cells should be plated at 3–5 × 105 cells/ well (see Note 5). Incubate cells for 3–4 days at 37 °C with 5% CO2, in the following polarizing conditions (see Note 9). 2. TH1 polarizing conditions: 10 μg/mL α-IL-4, 5 ng/mL IL-12, 250 U/mL IL-2. 3. TH2 polarizing conditions: α-IFN-γ (), IL-4 (10 ng/mL), IL-2 (250 U/mL). 4. TH9 polarizing conditions: 10 μg/mL α-IFN-γ, 10 ng/mL IL-4, 2 ng/mL TGF-β. 5. TH17 polarizing conditions: 10 μg/mL α-IL-4, 10 μg/mL α-IFN-γ, 50 ng/mL IL-6, 2 ng/mL TGF-β1. 6. iTREG polarizing conditions: 10 μg/mL α-IL-4, 10 μg/mL α-IFN-γ, 2 ng/mL TGF-β. 7. TFH-like polarizing conditions (see Note 10).

3.3  Expansion of Differentiated CD4+ T Cells

1. Following 3–4 days of differentiation under polarizing conditions, CD4+ T cells can be harvested for use in downstream applications or expanded in the absence of stimulation, depending on experimental applications (Fig. 2). 2. To expand differentiated T helper cell populations, remove cells from stimulation wells via thorough pipetting and transfer to a 15 mL conical tube. If multiple subsets are being expanded simultaneously, collected cells should be kept on ice as much as possible.

Harvest of murine spleen and lymph nodes

Day 0

Naïve CD4+ T cells

Stimulation (α-CD3, α-CD28) Subset-specific cytokine and blocking antibody conditions

1. Harvest naïve primary murine CD4+ T cells from spleens and lymph nodes using negative selection. 2. Stimulate using α-CD3 (T cell receptor agonist) and α-CD28 (co-receptor agonist) antibodies. 3. Polarize using subset-specific cytokine conditions.

CD4+ effector T cells

Harvest for analysis/ downstream applications Expand into fresh media; polarizing conditions

Day 3-4 Option 1: Harvest differentiated CD4+ effector cell populations for analysis and downstream applications. Option 2: Remove effector cells from stimulation and expand to reduce cell density; place cells in fresh cytokine conditions and/or further modulate cytokine environment.

CD4+ effector T cells

Harvest for analysis/ downstream applications

Re-stimulation

Day 5-7 Option 1: Harvest differentiated cell populations for analysis and downstream applications. Option 2: Re-stimulate differentiated cell populations with α-CD3 or PMA/ Ionomycin to evaluate cell phenotype/ cytokine production.

Fig. 2 Schematic outlining the general procedures involved in the differentiation of CD4+ T helper cell subsets. Key time points are shown for steps involved in the differentiation and analysis of effector subsets. Briefly, naïve CD4+ T cells are isolated, activated with α-CD3/α-CD28, and subjected to subset-specific cytokine skewing conditions for 3–4 days. Subsequently, cells can either be harvested for analyses or expanded for continued culture

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3. Centrifuge cell samples at 600 × g for 5 min at 4 °C. 4. Aspirate and discard supernatant. Plate cells at ~5 × 105 cells/well under subset-specific cytokine conditions as described above. At this point, IL-2 may be added to promote cell proliferation/survival in the absence of stimulation (see Note 11). 5. Incubate expanded cells for up to 4 days at 37 °C with 5% CO2 (see Note 12).

4  Notes 1. Some laboratories use supplemented RPMI medium for T helper cell culture, as an alternative to IMDM [6, 24–27]. 2. Our laboratory has successfully utilized negative selection-­ based kits from both R&D Systems and BioLegend for the isolation of naïve CD4+ T cells. Additional kits are available at the discretion of individual laboratories. 3. Some laboratories routinely include IL-12 blocking antibodies for the culture of TH2 cells [28]. It is imperative this step be included when using culture conditions that utilize antigen-­ presenting cells for T helper cell stimulation, as these cells are capable of producing IL-12. 4. A variety of culture conditions have been used to generate both “standard” and “pathogenic” TH17 cells. For these approaches, additional cytokines and/or blocking antibodies include α-IL-2 (clone JES6-1A12) [25], IL-1β [24, 29, 30], IL-23 [29, 31], or TGF-β3 [32]. 5. Our laboratory routinely utilizes 10 wells in a 24-well plate to culture naïve CD4+ T cells isolated from 2 mice, which provides a plating density of ~3–5 × 105 cells/well. Other laboratories utilize a range of plating densities, which should be optimized for individual experiments. 6. While α-CD3ε is typically plate-bound, other laboratories have described the use of both plate-bound and soluble α-CD28. Beads that are conjugated to α-CD3ε and α-CD28 are also commonly used for T cell activation. Additionally, a range of concentrations has been described for both α-CD3ε and α-CD28, depending on the target differentiation conditions (Table  1). These aspects should be considered for individual experiments. 7. If multiple mice will be utilized for a single biological replicate, their spleens and lymph nodes, respectively, may be pooled at this step. Using the kits described in Note 1, we recommend that cells from no more than two mice be pooled for a single isolation reaction.

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Table 1 Published concentration ranges for T helper subset polarizing conditions TH1 (refs. 24–27, 30, 36) Stimulation reagents

Concentration range(s)

α-CD3 (clone 145-2C11)

1–5 μg/mL

α-CD28 (clone 37.51)

0.5–10 μg/mL

Blocking antibodies α-IL-4 (clone 11B11)

1–10 μg/mL

Cytokines IL-12

5–30 ng/mL

IL-2

30–250 U/mL; 10–20 ng/mL

*

TH2 (refs. 24–28, 30) Stimulation reagents

Concentration range(s)

α-CD3 (clone 145-2C11)

1–5 μg/mL

α-CD28 (clone 37.51)

0.5–10 μg/mL

Blocking antibodies α-IFN-γ (clone XMG1.2)

1–10 μg/mL

α-IL-12* (clone AMC0122)

10 μg/mL

Cytokines IL-4

10–30 ng/mL

IL-2*

30–250 U/mL; 10–20 ng/mL

TH17 (refs. 6, 24, 25, 27, 29, 30, 32) Stimulation reagents

Concentration range(s)

α-CD3 (clone 145-2C11)

1–5 μg/mL

α-CD28 (clone 37.51)

0.5–10 μg/mL

Blocking antibodies α-IFN-γ (clone XMG1.2)

1–10 μg/mL

α-IL-4 (clone 11B11)

1–10 μg/mL

α-IL-2* (JES6-1A12)

1 μg/mL

Cytokines IL-6

20–100 ng/mL (continued)

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Table 1 (continued) IL-1β*

10 ng/mL

TGF-β1

1–5 ng/mL

IL-23*

50 ng/mL

iTREG (refs. 24, 25, 27) Stimulation reagents

Concentration range(s)

α-CD3 (clone 145-2C11)

1–5 μg/mL

α-CD28 (clone 37.51)

0.5–10 μg/mL

Blocking antibodies α-IFN-γ (clone XMG1.2)

1–10 μg/mL

α-IL-4 (clone 11B11)

1–10 μg/mL

Cytokines TGF-β1

2–15 ng/mL

IL-2

30 U/mL

*

Items marked with an asterisk are not included in all differentiation protocols for a given subset

8. Take care during the homogenization of spleen and lymph node tissue not to exert excessive pressure on the sample, as this may result in diminished yield. Slides should be rinsed regularly using the media present in the petri dish to remove suspended cells. 9. While the concentrations provided in Subheading 3 are representative of successful polarization conditions, a range of concentrations have been described for each T helper cell subset (see Table 1). Thus, individual laboratories should optimize these conditions for their applications. 10. The differentiation of bona fide TFH cells requires interactions with B cells in the follicles of lymphoid tissues. However, several laboratories have developed in vitro culture systems that result in varying degrees of expression of the TFH cell phenotype. Using α-CD3ε and α-CD28 stimulation, a population of TFH-like cells can be generated in the presence of IL-6, IL-21, and neutralizing antibodies for IL-4, IFN-γ, and TGF-β [24, 33]. Similar conditions have also been described utilizing TCR-transgenic T cells in the presence of antigen and antigen-­ presenting cells, with the addition of IL-12 neutralizing antibodies [34]. Additionally, TH1 cells removed from stimulation and exposed to a low IL-2 environment containing IL-12 have been shown to upregulate the expression of the TFH cell lineage-defining factor Bcl-6, along with key aspects of the TFH

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gene program [35–37]. As the conditions that result in these cells vary greatly, we have opted not to include TFH populations in the current review. 11. Care should be taken to ensure that an appropriate concentration of IL-2 is utilized depending on the desired T helper cell subset. While survival signals from IL-2 is helpful to ensure T helper cell viability in the absence of stimulation, signals from IL-2 have been shown to have disparate effects on the differentiation of distinct T helper cell subsets. For example, IL-2 signaling is known to be refractory for the development of both TH17 and TFH subsets, so utilizing an optimized concentration that permits cell survival but does not repress the TH17 or TFH gene program is an important experimental consideration. 12. Differentiated cells can be analyzed via a number of techniques to examine expression of lineage-defining transcription factors or cell surface markers, as well as cytokine production (e.g., qRT-PCR, immunoblot, flow cytometry, and ELISA).

Acknowledgments The authors would like to thank members of the Oestreich Lab for thoughtful discussions and critical reading of the manuscript. We apologize to those whose work could not be adequately cited and discussed due to length restrictions. This work was supported by funds from the Virginia-Maryland College of Veterinary Medicine and by National Institutes of Health grants, R56AI127800 and R01AI134972. References 1. Mosmann TR, Cherwinski H, Bond MW et al (1986) Two types of murine helper T cell clone. I. Definition according to profiles of lymphokine activities and secreted proteins. J Immunol 136:2348–2357 2. Breitfeld D, Ohl L, Kremmer E et al (2000) Follicular B helper T cells express CXC chemokine receptor 5, localize to B cell follicles, and support immunoglobulin production. J Exp Med 192:1545–1552 3. Dardalhon V, Awasthi A, Kwon H et al (2008) IL-4 inhibits TGF-beta-induced Foxp3+ T cells and, together with TGF-beta, generates IL-9+ IL-10+ Foxp3(−) effector T cells. Nat Immunol 9:1347–1355 4. Duhen T, Geiger R, Jarrossay D et al (2009) Production of interleukin 22 but not interleukin 17 by a subset of human skin-homing memory T cells. Nat Immunol 10:857–863

5. Eyerich S, Eyerich K, Pennino D et al (2009) Th22 cells represent a distinct human T cell subset involved in epidermal immunity and remodeling. J Clin Invest 119: 3573–3585 6. Ivanov II, McKenzie BS, Zhou L et al (2006) The orphan nuclear receptor RORgammat directs the differentiation program of proinflammatory IL-17+ T helper cells. Cell 126:1121–1133 7. Kim CH, Rott LS, Clark-Lewis I et al (2001) Subspecialization of CXCR5+ T cells: B helper activity is focused in a germinal center-­localized subset of CXCR5+ T cells. J Exp Med 193: 1373–1381 8. Langrish CL, Chen Y, Blumenschein WM et al (2005) IL-23 drives a pathogenic T cell population that induces autoimmune inflammation. J Exp Med 201:233–240

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9. Sakaguchi S, Sakaguchi N, Asano M et al (1995) Immunologic self-tolerance maintained by activated T cells expressing IL-2 receptor alpha-chains (CD25). Breakdown of a single mechanism of self-tolerance causes various autoimmune diseases. J Immunol 155:1151–1164 10. Schaerli P, Willimann K, Lang AB et al (2000) CXC chemokine receptor 5 expression defines follicular homing T cells with B cell helper function. J Exp Med 192:1553–1562 11. Veldhoen M, Uyttenhove C, van Snick J et al (2008) Transforming growth factor-beta ‘reprograms’ the differentiation of T helper 2 cells and promotes an interleukin 9-producing subset. Nat Immunol 9:1341–1346 12. O'Shea JJ, Paul WE (2010) Mechanisms underlying lineage commitment and plasticity of helper CD4+ T cells. Science 327:1098–1102 13. Oestreich KJ, Weinmann AS (2011) Encoding stability versus flexibility: lessons learned from examining epigenetics in T helper cell differentiation. Curr Top Microbiol Immunol 356: 145–164 14. Oestreich KJ, Weinmann AS (2012) Master regulators or lineage-specifying? Changing views on CD4(+) T cell transcription factors. Nat Rev Immunol 12:799–804 15. Zhu J, Paul WE (2010) Peripheral CD4+ T-cell differentiation regulated by networks of cytokines and transcription factors. Immunol Rev 238:247–262 16. Tsoukas CD, Landgraf B, Bentin J et al (1985) Activation of resting T lymphocytes by anti­CD3 (T3) antibodies in the absence of monocytes. J Immunol 135:1719–1723 17. Trickett A, Kwan YL (2003) T cell stimulation and expansion using anti-CD3/CD28 beads. J Immunol Methods 275:251–255 18. O'Shea JJ, Lahesmaa R, Vahedi G et al (2011) Genomic views of STAT function in CD4+ T helper cell differentiation. Nat Rev Immunol 11:239–250 19. Zhu J, Yamane H, Paul WE (2010) Differentiation of effector CD4 T cell populations. Annu Rev Immunol 28:445–489 20. Basu R, Hatton RD, Weaver CT (2013) The Th17 family: flexibility follows function. Immunol Rev 252:89–103 21. Crotty S (2014) T follicular helper cell differentiation, function, and roles in disease. Immunity 41:529–542 22. Kaplan MH, Hufford MM, Olson MR (2015) The development and in vivo function of T helper 9 cells. Nat Rev Immunol 15:295–307 23. Rudensky AY (2011) Regulatory T cells and Foxp3. Immunol Rev 241:260–268

24. Awe O, Hufford MM, Wu H et al (2015) PU.1 expression in T follicular helper cells limits CD40L-dependent germinal center B cell development. J Immunol 195:3705–3715 25. Sekiya T, Yoshimura A (2016) In vitro Th differentiation protocol. Methods Mol Biol 1344:183–191 26. Szabo SJ, Sullivan BM, Stemmann C et al (2002) Distinct effects of T-bet in TH1 lineage commitment and IFN-gamma production in CD4 and CD8 T cells. Science 295:338–342 27. Flaherty S, Reynolds JM (2015) Mouse naive CD4+ T cell isolation and in vitro differentiation into T cell subsets. J Vis Exp 98:e52739 28. Makar KW, Perez-Melgosa M, Shnyreva M et al (2003) Active recruitment of DNA methyltransferases regulates interleukin 4 in thymocytes and T cells. Nat Immunol 4:1183–1190 29. Chung Y, Chang SH, Martinez GJ et al (2009) Critical regulation of early Th17 cell differentiation by interleukin-1 signaling. Immunity 30:576–587 30. Plank MW, Kaiko GE, Maltby S et al (2017) Th22 cells form a distinct Th lineage from Th17 cells in vitro with unique transcriptional properties and Tbet-Dependent Th1 plasticity. J Immunol 198:2182–2190 31. Rumble J, Segal BM (2014) In vitro polarization of T-helper cells. Methods Mol Biol 1193:105–113 32. Lee Y, Awasthi A, Yosef N et al (2012) Induction and molecular signature of pathogenic TH17 cells. Nat Immunol 13:991–999 33. Zeng H, Cohen S, Guy C et al (2016) mTORC1 and mTORC2 kinase signaling and glucose metabolism drive follicular helper T cell differentiation. Immunity 45: 540–554 34. Lu KT, Kanno Y, Cannons JL et al (2011) Functional and epigenetic studies reveal multistep differentiation and plasticity of in vitro-­ generated and in vivo-derived follicular T helper cells. Immunity 35:622–632 35. McDonald PW, Read KA, Baker CE et al (2016) IL-7 signalling represses Bcl-6 and the TFH gene program. Nat Commun 7:10285 36. Oestreich KJ, Mohn SE, Weinmann AS (2012) Molecular mechanisms that control the expression and activity of Bcl-6 in TH1 cells to regulate flexibility with a TFH-like gene profile. Nat Immunol 13:405–411 37. Read KA, Powell MD, Baker CE et al (2017) Integrated STAT3 and Ikaros Zinc finger transcription factor activities regulate Bcl-6 expression in CD4+ Th cells. J Immunol 199: 2377–2387

Chapter 7 Generation and Culture of Mouse Embryonic Fibroblasts Yee Sun Tan and Yu L. Lei Abstract In addition to leukocytes, a variety of cells also participate in the innate immune response, including endothelial cells, epithelial cells, and fibroblasts. Thus, the study of these cells is highly relevant in broadening our understanding of mechanisms that modulate innate immunity. With the rise of genetically engineered animals, it is now common to confirm in vitro data acquired using immortalized cell lines with more physiologically relevant primary cells from these animals ex vivo. Indeed, many studies exploring innate immune system function employ mouse embryonic fibroblasts (MEFs). These cells are relatively simple to generate and are a powerful tool to explore regulatory networks, examine biochemical profiling of protein complexes, and investigate novel signaling pathways associated with innate immune system signaling. Here, we provide a robust protocol to isolate, maintain, and store primary MEFs. This protocol is designed for users with minimal experience using mouse models. We have also added precautions and common pitfalls associated with these procedures. Key words Innate immunity, Mouse embryonic fibroblasts, MEF

1  Introduction The use of gene deletion and transgenic mouse models have led to significant advances across all fields of biomedical research, especially in immunology. Indeed, the use of primary cells from genetically manipulated mice has become standard practice to verify and confirm in vitro data generated from immortalized cell lines. In the innate immunity field, use of these animal models resulted in the exponential expansion of discoveries associated with novel families of pattern recognition receptors that sense pathogen-associated molecular patterns, identifying critical adaptor molecules that modulate immune system signaling, expanding our understanding of intricate regulatory networks underlying innate immune system responses, and defining novel co-stimulatory/co-inhibitory pathways [1–3]. Innate immunity is an evolutionarily conserved system shared by a wide spectrum of cells, including cells that are derived from non-hematopoietic compartments. These include, but are not limited to, cells such as epithelial cells and fibroblasts that are Irving C. Allen (ed.), Mouse Models of Innate Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1960, https://doi.org/10.1007/978-1-4939-9167-9_7, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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associated with the primary physical barriers that protect the host from microbial or environmental insults. Due to their anatomic location and physiologic functions, the cells that make up these barriers are equipped with an assortment of extracellular and cytoplasmic sensors that are linked to the necessary innate immune signaling pathways within the cell to initiate host-pathogen defense mechanisms. Consistent with these features, mouse embryonic fibroblasts (MEFs) are commonly used to study aspects of the innate immune system. Indeed, MEFs are extensively utilized in biochemical identification and functional characterization studies of novel genes and proteins [4–9]. These cells are functionally relevant and relatively simple to generate from mice. The following chapter provides a standard protocol for the isolation and culture of MEFs.

2  Materials 2.1  Mice

1. At least two breeding cages of mice for each genotype need to be tested. Each breeding cage should include one male and two female mice, 6–12 weeks of age (see Note 1). Prior to the initiation of any study involving animals, users must attain approval by the appropriate institutional regulatory body (i.e., the Institutional Animal Care and Use Committee (IACUC)). All breeding and care for mice must also follow official institutional guidelines.

2.2  Reagents

1. 70% (v/v) ethanol. 2. Phosphate-buffered saline (1× PBS), pH 7.4. 3. Dulbecco’s Modified Eagle Medium (DMEM). 4. Fetal bovine serum (FBS). 5. 100× penicillin-streptomycin (10,000 units of penicillin and 10,000 μg of streptomycin/ml). 6. 0.25% trypsin-EDTA solution. 7. Freezing medium: 10% dimethyl sulfoxide (DMSO), 20% FBS, 70% DMEM high glucose, and 1% penicillin-streptomycin (see Note 2).

2.3  Supplies and Equipment

1. 60 mm and 100 mm tissue culture dishes. 2. 15 ml and 50 ml conical tubes. 3. 2 ml screw cap cryovials. 4. Laminar flow hood (see Note 3). 5. Sterile 40 μm cell strainers. 6. Surgical instruments: dissecting forceps, scissors, and single-­ edge razor/scalpel blades (see Note 4).

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7. Examination gloves. 8. Freezing containers. 9. Inverted microscope. 10. 37 °C tissue culture incubator with 5% CO2.

3  Methods 3.1  Establish Timed Breeding Cages (See Note 5)

1. Place one male mouse with two female mice. 2. Examine the female mice for the presence of copulation plugs each morning until the plug is identified. 3. Once the plug is identified, transfer the female mice to a new cage and note the time of embryo development as 0.5 days. 4. Allow the embryos to develop for 13.5–15.5 days. The most widely accepted ages for the embryos used in MEF generation are between 13.5 and 15.5 days.

3.2  Presurgical Preparations

1. Pregnant mice should be euthanized as specified in an approved animal protocol. The most widely used methods include the use of carbon dioxide, isoflurane overdose, or avertin overdose. The primary method should be followed by a secondary physical method, such as cervical dislocation. 2. Spray the entire mouse with 70% ethanol. 3. The dissection should be performed in a laminar flow hood (see Note 6). 4. In a clean area of the hood, label PBS-filled 100 mm tissue culture plates with the genotype of each mouse and a unique number to identify each embryo (see Note 7).

3.3  Dissect the Embryos

1. Lift the abdominal wall with blunt forceps, and make a shallow 3 mm transverse incision. Next, make a longitudinal incision to expose the abdominal wall. At this point, the uterus with embryos should be easily visible. Take all precautions to keep the surgical site uncontaminated. If the surgical instruments are contaminated by hair, fully rinse the instrument in 70% ethanol or use a new set of instruments. 2. Expose the embryos by opening the abdominal wall. Take special precautions to avoid damaging the internal organs in the peritoneal cavity, especially the small intestine and colon. Damage to these organs increases the risk of contaminating the embryos with bacteria. 3. Carefully transfer each embryo, with intact yolk sac, to the tissue culture plates prepared in Subheading 3.2, step 4.

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3.4  Prepare the Fetuses

1. Tease apart the yolk sac with a pair of fine forceps and discard the yolk sac. 2. Carefully remove the fetus head, and keep it in a separate PBS-­ filled conical tube labeled with its genotype and identifying number. This identification should match the respective 100 mm tissue culture plate. The fetal remains that are not used for MEF generation, including the head, are used for genotyping. 3. Carefully remove the heart and liver, which are both dark red tissues, and discard. 4. Wash each prepared fetus in PBS and transfer it to a new 60 mm tissue culture dish that has been identically labeled as the original plate.

3.5  Prepare the MEF Suspension

1. Add 3 ml of pre-chilled 0.25% trypsin-EDTA to each dish. 2. Mince the fetus tissue thoroughly with two razor or scalpel blades. This step is critical for enhancing the final yield of MEFs. It is important that this step be thoroughly conducted and not rushed. 3. Gently pipet all of the tissue fragment suspensions up and down ten times. 4. Incubate each tissue culture dish at 37 °C for 10 min. Do not expose cells to extended trypsin treatment. Extensive exposure to trypsin will markedly reduce cell viability. 5. Transfer the suspension to a 40 μm cell strainer placed on a 50 ml conical tube, and allow cell suspension to flow through the cell strainer with gravity. 6. Spin the cells down at 500 × g and 4 °C for 5 min. Wash the cell pellets once with pre-chilled complete DMEM medium containing 10% FBS to deactivate and remove any residual trypsin.

3.6  Passage 0 MEFs Culture

1. Cell pellets from the previous step are resuspended in complete culture media. 2. Add 11 ml of culture medium to each of the 100 mm culture dishes or 75 mm2 culture flasks. 3. Culture the cells in a standard humidified 37 °C, 5% CO2 incubator for 24 h. If the all of the previous steps are followed properly, the cells should be close to confluence. These cells are considered to be at passage 0 (see Note 8).

3.7  Harvest MEFs (See Note 9)

1. Following the initial passage, split cells when they reach approximately 70% confluence. 2. Remove culture medium and gently wash cells two times with PBS.

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3. Add 4 ml of 0.05% trypsin-EDTA solution to the culture dish, and incubate at 37 °C for 3 min. Shake 1–2 times during the incubation. 4. Deactivate the trypsin and resuspend the cells with 15 ml complete culture medium. Spin down the cell pellets at 500 × g and 4 °C for 5 min. 5. Wash the pellets one time with PBS and resuspend in 15 ml complete culture medium for passage. 3.8  Freeze and Thaw MEFs

1. Resuspend 1.0 × 106 MEFs in 1 ml of freezing medium, and transfer to a pre-labeled cryovial. 2. Place cryovials in the controlled rate freezer for 24 h, and then transfer them to liquid nitrogen for long-term cryopreservation (see Note 10). 3. When thawing cells from frozen stocks, remove cells from liquid nitrogen and immediately place the cryovials in a 37 °C water bath. 4. Transfer the cell suspension to a 50 ml conical tube, and add pre-warmed complete culture media dropwise into the suspension. Then spin down the cells and proceed with Subheadings 3.6 and 3.7.

4  Notes 1. Although the use of wild-type mice from the same genetic background is commonly accepted as controls for MEF generation, we highly recommend generating littermate control cells for better accuracy. If gene manipulation does not result in breeding failure or death at early embryonic stages, then the use of heterozygous breeding pairs is recommended. The male to female mouse ratio should be 1:2 in the breeding cages. 2. Be very careful when handling DMSO, which is a known carcinogenic chemical and could penetrate intact skin. 3. We recommend using different hoods for generating MEFs and regular cell culture to minimize the chances of contamination. 4. All instruments should be sterilized by autoclaving prior to experiments. 5. Timed breeding is critical for the successful preparation of MEFs. The most widely used approach to determine the starting date of embryo development is the identification of a copulation plug (semen plug) in the vagina. Although it is usually identifiable before 8:00 am, it should be noted that the copulation plug can be hard to recognize 12 h after mating.

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In addition, it is more difficult to identify the plug in very young mice. Hence, copulation plugs need to be examined early in the morning every day until it is identified. The time when a copulation plug is identified will be marked as day 0.5 for embryo development. All female mice should then be transferred to a new cage for the next 13–15 days. 6. Due to the thick hair, it can be challenging to keep the dissection process sterile. Thus, it is of critical importance to take all available precautions to reduce the potential for contamination. It is advisable to separate relatively “clean areas” and relatively “dirty areas” in the hood after transferring mice into the working space. Embryo dissection should be confined to the “dirty area,” and all subsequent procedures should be performed in the “clean area.” 7. Clear numbering of the plates will greatly benefit subsequent genotyping and matching the results to each of the dissected embryos. 8. Primary MEFs maintain active proliferation for about four passages. The proliferation rates will considerably drop after five passages. Hence, only early passage cells are appropriate for downstream applications. Active MEFs typically double in cell number every 48 h. Hence, it is recommended to freeze passages 0 and 1 MEFs for the convenience of further experimentation. 9. The culture and maintenance of MEFs are similar to other fibroblast cell lines. 10. There are a number of affordable controlled rate freezers available from different vendors. The principle is to decrease the ambient temperature gradually. Do not directly place cryovials into liquid nitrogen. This will significantly reduce cell viability.

Acknowledgment This work is supported by NIH grants R00 DE024173 (YLL), R01 DE026728 (YLL), and Rogel Cancer Center Fund for Discovery (YLL). References 1. Davis BK, Wen H, Ting JP (2011) The inflammasome NLRs in immunity, inflammation, and associated diseases. Annu Rev Immunol 29: 707–735 2. Ting JP, Duncan JA, Lei Y (2010) How the noninflammasome NLRs function in the innate immune system. Science 327:286–290

3. Wen H, Lei Y, Eun SY, Ting JP (2010) Plexin-­ A4-­ semaphorin 3A signaling is required for Toll-like receptor- and sepsis-induced cytokine storm. J Exp Med 207:2943–2957 4. Ishikawa H, Barber GN (2008) STING is an endoplasmic reticulum adaptor that facilitates innate immune signalling. Nature 455:674–678

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vmediated type I interferon induction. Nat 5. Seth RB, Sun L, Ea CK, Chen ZJ (2005) Immunol 6:981–988 Identification and characterization of MAVS, a mitochondrial antiviral signaling protein that 8. Meylan E, Curran J, Hofmann K et al (2005) activates NF-kappaB and IRF 3. Cell 122: Cardif is an adaptor protein in the RIG-I antivi669–682 ral pathway and is targeted by hepatitis C virus. Nature 437:1167–1172 6. Yoneyama M, Kikuchi M, Natsukawa T et al (2004) The RNA helicase RIG-I has an essential 9. Allen IC, Moore CB, Schneider M et al (2011) function in double-stranded RNA-induced innate NLRX1 protein attenuates inflammatory antiviral responses. Nat Immunol 5:730–737 responses to infection by interfering with the RIG-I-MAVS and TRAF6-NF-kappaB signaling 7. Kawai T, Takahashi K, Sato S et al (2005) IPS-­ pathways. Immunity 34:854–865 1, an adaptor triggering RIG-I- and Mda5-­

Chapter 8 Isolation of Tumor-Infiltrating Lymphocytes by Ficoll-­Paque Density Gradient Centrifugation Yee Sun Tan and Yu L. Lei Abstract With the renewed enthusiasm in immuno-oncology, characterization of the tumor immune microenvironment constitutes an essential and unique aspect to the assessment of therapeutics. The isolation of tumor-­ infiltrating lymphocytes (TILs) is a desirable approach toward the understanding of antitumor immune response. This chapter provides an effective protocol to mechanically dissociate tumor tissue and generate single-cell suspension from excised tumors. TILs are then isolated by Ficoll-Paque density gradient centrifugation. This protocol is applicable to both human and experimental tumors in immunocompetent murine models. Key words Tumor-infiltrating lymphocytes, Tumor microenvironment, Immunotherapy, Immune escape, Mechanical dissociation

1  Introduction Immune checkpoint receptors (ICRs) play a major role in the regulation of T-cell function and responses and have been demonstrated to be effective targets in several cancers [1]. However, the response rates to these ICR inhibitors are generally less than 35% among cancer patients, such as those with melanoma or head and neck squamous cell carcinoma [2, 3]. In addition to human tumors, the use of immunocompetent mouse models is instrumental in testing new combinatorial therapies to overcome tumor resistance to ICR blockade [4–7]. Therefore, the isolation of tumor-­ infiltrating lymphocytes (TILs) from human and mouse tumors and downstream analysis, such as flow cytometry and effector function characterization, provide critical insight into strategies that can prime hypoimmunogenic cold cancers. Ficoll-Paque, which consists of Ficoll PM400 (synthetic polymers of sucrose and epichlorohydrin) and sodium diatrizoate, has been widely used in the clinic to isolate lymphocytes from blood, bone marrow, and umbilical cord blood through density Irving C. Allen (ed.), Mouse Models of Innate Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1960, https://doi.org/10.1007/978-1-4939-9167-9_8, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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gradient centrifugation [8–11]. As the densities of mononuclear cells such as lymphocytes and monocytes are less than that of Ficoll-Paque (e.g., 1.077 g/ml), these cells form a layer at the interface during density gradient centrifugation and thus can be readily collected. Ficoll-Paque has also been used to isolate mononuclear cells from other species including mice. In this chapter, we describe a protocol that mechanically dissociates tumors and derives single-cell suspensions from human specimens and experimental tumors in immunocompetent mice. TILs and other mononuclear cells are purified using Ficoll-Paque density gradient centrifugation. Tumor dissociation methods mainly comprise enzymatic digestion (e.g., collagenase, trypsin, dispase), mechanical disruption of stroma, or a combination of both. Key considerations during this step are how to preserve viability, function, and phenotype of the TILs. Although mechanical dissociation of tumor may be more time-consuming, we found this method to be highly reproducible. The use of enzymatic digestions may result in higher cell yield, but there is a need for careful optimization of enzymatic treatments among different tumor models to retain TIL phenotypes because enzymes may reduce lymphocyte surface markers important for downstream analysis, such as flow cytometry [12–14]. Although there are other centrifugation media available, such as Percoll, this protocol uses Ficoll-Paque which is broadly adaptable for human and mouse samples.

2  Materials 2.1  Tumors

1. Fresh tumor samples from human or mice are needed for TIL separation. Formalin or paraformaldehyde fixation renders the samples incompatible with this protocol. 2. Fresh tumor samples need to be hydrated and placed in either phosphate-buffered saline (PBS) solution or culture media. Dehydration of tumor samples will substantially decrease the TIL yield and viability.

2.2  Reagents

1. 70% (v/v) ethanol in sterile water. 2. 1× PBS, pH 7.4 formulated without calcium and magnesium. 3. RPMI-1640. 4. Fetal bovine serum (FBS). 5. 100× penicillin-streptomycin (10,000 units/ml of penicillin and 10,000 μg/ml of streptomycin). 6. Ficoll-Paque PLUS (see Note 1). 7. Dimethyl sulfoxide (DMSO). 8. Complete RPMI media: RPMI-1640 with 10% FBS and 1× penicillin-streptomycin.

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1. Sterile dissecting forceps and scissors. 2. Single-edged razor blades. 3. Tissue culture hood. 4. Examination gloves. 5. Sterile 50 ml conical tubes. 6. 5, 10, 25 ml sterile serological pipets. 7. Pipet aid with speed settings. 8. 1 ml sterile pipet tips. 9. 100 mm sterile petri dishes. 10. Syringes with rubber plungers. 11. Sterile 70 μm cell strainers. 12. Weighing scale. 13. Centrifuge with swing-bucket rotors. 14. 2 ml screw cap cryovials. 15. Freezing containers.

3  Methods 3.1  Preparations Before Extracting Tumors

1. Media and Ficoll-Paque should be prewarmed to room temperature of 18–20 °C before use (see Note 2).

3.2  Mechanical Dissociation of Tumor to Obtain Single-Cell Suspensions (See Note 3)

1. For mouse tumors, first euthanize mouse according to institution-­approved protocol, and spray the tumor area with 70% ethanol. Excise the tumor using sterile scissors and forceps. 2. Using sterile forceps, place the excised human or mouse tumor in a lid of a sterile 100 mm petri dish containing 5 ml of RPMI-­ 1640 media at room temperature. Once tumor is extracted, keep it in media to prevent dehydration and loss of TILs (see Note 4). 3. Mince the tumor into small pieces using two single-edged razor blades. To obtain smaller pieces of 1–2 mm, hold two single-­edged razor blades in one hand and mince the tumor. Although this may be time-consuming, it is critical at this step to ensure that all tissue is minced up. 4. Place a 70 μm cell strainer in the other half of the 100 mm dish. 5. Cut a 1 ml pipet tip using a razor blade. Using this cut tip, transfer the tumor tissue to the cell strainer. If there are pieces of tissue left on the lid, add another 5 ml of RPMI-1640 media, and transfer them to cell strainer.

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6. Use the rubber plunger of a syringe to mesh the tissue through the cell strainer. The media in the dish will be cloudy due to dissociation of cells. Small pieces of tissue will remain on the strainer. 7. Place a new 70 μm cell strainer onto a sterile labeled 50 ml conical tube. 8. Pass the cell suspension through the strainer. Add more RPMI-­ 1640 media to the plate, and pipet up and down several times to resuspend all remaining cells. Pass the media from the dish through the cell strainer. 9. Top up the media to 30 ml with RPMI-1640 media at room temperature. Keep samples at room temperature of 18–20 °C (see Note 5). 3.3  Isolation of Mononuclear Cells

1. Immediately before the addition of Ficoll-Paque media, use a 25 ml pipet to mix the single-cell suspension. This ensures that cells do not pellet at the bottom or form clumps. 2. Thoroughly mix the Ficoll-Paque media by inverting the container (see Note 6). 3. Set the speed of pipet aid to slow release. Pipet 10 ml of Ficoll-­ Paque media, and carefully insert the pipet to the bottom of the 50 ml conical tube containing 30 ml of single-cell suspensions (Fig. 1a).

Fig. 1 Separation of mononuclear cells using Ficoll-Paque media and density gradient centrifugation. (a) 10 ml of Ficoll-Paque is added to 30 ml of single-cell suspension using a 10 ml serological pipet and pipet aid. Slow and gentle release (indicated by the dashed arrows) of the Ficoll-Paque will result in a distinct separation between the two layers. (b) Before density gradient centrifugation, the layer of single-cell suspension is above the Ficoll-Paque media. (c) After centrifugation, TILs and monocytes are found in the interface between the media and Ficoll-Paque, while the pellet consists of tumor cells and the erythrocytes

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4. Slowly release the Ficoll-Paque media to form a layer of Ficoll-­ Paque below the cell suspensions. This reduces mixing of the two solutions. A clear distinct separation between Ficoll-Paque and cell solution is seen (Fig. 1b). 5. Weigh samples to ensure proper balance of the rotors (see Note 7). 6. Centrifuge at 1025 × g for 20 min at 20 °C with slow acceleration and brakes turned off. 7. Carefully remove the tube from the rotor to avoid any disturbance to the layers formed during centrifugation (Fig. 1c). Pipet 20 ml of the upper layer of media to a waste bottle. 8. Transfer the layer of mononuclear cells to a sterile labeled 50 ml conical tube using a sterile pipet, along with the remainder of the media above the Ficoll-Paque. Minimize transfer of the Ficoll-Paque where possible. 3.4  Washing the Isolated Tumor-­ Infiltrating Lymphocytes

1. Fill the 50 ml conical tubes to 40 ml with complete RPMI media, and gently mix using a 25 ml pipet (see Note 8). 2. Centrifuge at 650 × g for 10 min at 20 °C. 3. Remove the supernatant, and gently resuspend cells in 30 ml of complete RPMI media. 4. Centrifuge at 650 × g for 10 min at 20 °C. 5. Remove the supernatant and depending on downstream application resuspend the cells accordingly (see Note 9).

4  Notes 1. There are several Ficoll-Paque media available, and users should decide which to use depending on the species and the cells needed. Ficoll-Plaque PLUS (1.077 g/ml) is widely used for the isolation of human mononuclear cells, and we have had success with it to isolate TILs from immunocompetent mice bearing syngeneic mouse tumors. However, lymphocytes from mouse are denser than that of human [8], and thus the use of Ficoll-Paque PREMIUM 1.084 is recommended if TILs are to be isolated from mouse tumors. 2. Ficoll-Paque densities are inversely correlated to temperature. Factors which contribute to the temperature include the temperature of single-cell suspension media, Ficoll-Paque media, and temperature of the centrifuge, and this may affect the yield of mononuclear cells obtained from density gradient separations. Therefore, it is important to ensure all media used are prewarmed to room temperature of 18–20 °C and that the centrifugation is done at 18–20 °C.

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3. Upon obtaining the tumor, all subsequent steps are performed in a tissue culture hood to minimize contamination. 4. When mincing the tumors, the media used is RPMI-1640 without any FBS. This is to prevent any clumping of cells due to FBS. After density gradient centrifugation, complete RPMI media is used to wash the cells. 5. Cell viability may be affected if temperature is too high. If needed, place tubes containing cell suspension briefly on a rack on ice. 6. Protect the Ficoll-Paque media from light as sodium diatrizoate, which helps to maintain optimal density, is light-­sensitive. It is also important to use aseptic techniques to maintain the sterility of the media. 7. Ensure that the rotors are properly balanced before starting the centrifuge as vibrations due to imbalance may lead to mixing of two layers and reduce yield and purity. Balance any differences in weight using RPMI-1640 media by gently adding the media to the top of the single-cell suspension. If a balance tube is needed for centrifugation, use a tube with similar weight, not volume. 8. The washing steps help to remove any contaminating Ficoll-­ Paque media, resulting in a highly purified and viable population of lymphocytes for downstream analyses. 9. If cells are to be frozen for future use, resuspend cells in freshly made freezing media of 90% FBS and 10% DMSO, and add 1 ml of freezing media to each sample. Transfer to sterile labeled cryovials. Cryovials are kept in a freezing container (i.e., Mr. Frosty containing fresh isopropyl alcohol) in −80 °C freezer overnight prior to being stored in liquid nitrogen.

Acknowledgments This work is supported by NIH grants R00 DE024173 (YLL) and R01 DE026728 (YLL), Rogel Cancer Center Fund for Discovery (YLL). References 1. Zou W, Wolchok JD, Chen L (2016) PD-L1 (B7-H1) and PD-1 pathway blockade for cancer therapy: mechanisms, response biomarkers, and combinations. Sci Transl Med 8(328): 328rv324. https://doi.org/10.1126/scitranslmed.aad7118 2. Ribas A, Hamid O, Daud A et al (2016) Association of pembrolizumab with tumor

response and survival among patients with advanced melanoma. JAMA 315(15):1600– 1609. https://doi.org/10.1001/jama.2016. 4059 3. Ferris RL, Blumenschein G Jr, Fayette J et al (2016) Nivolumab for recurrent squamous-cell carcinoma of the head and neck. N Engl J Med

Isolation of Tumor-Infiltrating Lymphocytes by Ficoll-Paque Density Gradient… 375(19):1856–1867. https://doi.org/ 10.1056/NEJMoa1602252 4. Leach DG, Dharmaraj N, Piotrowski SL et al (2018) STINGel: controlled release of a cyclic dinucleotide for enhanced cancer immunotherapy. Biomaterials 163:67–75. https://doi. org/10.1016/j.biomaterials.2018.01.035 5. Judd NP, Allen CT, Winkler AE et al (2012) Comparative analysis of tumor-infiltrating lymphocytes in a syngeneic mouse model of oral cancer. Otolaryngol Head Neck Surg 147(3):493–500. https://doi.org/ 10.1177/0194599812442037 6. Sun ZJ, Zhang L, Hall B et al (2012) Chemopreventive and chemotherapeutic actions of mTOR inhibitor in genetically defined head and neck squamous cell carcinoma mouse model. Clin Cancer Res 18(19):5304–5313. https://doi.org/10.1158/1078-0432. CCR-12-1371 7. Lei Y, Xie Y, Tan YS et al (2016) Telltale tumor infiltrating lymphocytes (TIL) in oral, head & neck cancer. Oral Oncol 61:159–165. https://doi.org/10.1016/j. oraloncology.2016.08.003 8. Boyum A, Lovhaug D, Tresland L et al (1991) Separation of leucocytes: improved cell purity by fine adjustments of gradient medium density and osmolality. Scand J Immunol 34(6):697–712

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9. Li J, Jie HB, Lei Y et al (2015) PD-1/SHP-2 inhibits Tc1/Th1 phenotypic responses and the activation of T cells in the tumor microenvironment. Cancer Res 75(3):508–518. https://doi. org/10.1158/0008-5472.CAN-14-1215 10. Kansy BA, Concha-Benavente F, Srivastava RM et al (2017) PD-1 status in CD8(+) T cells associates with survival and anti-PD-1 therapeutic outcomes in head and neck cancer. Cancer Res 77(22):6353–6364. https://doi. org/10.1158/0008-5472.CAN-16-3167 11. Boyum A (1977) Separation of lymphocytes, lymphocyte subgroups and monocytes: a review. Lymphology 10(2):71–76 12. Abuzakouk M, Feighery C, O'Farrelly C (1996) Collagenase and Dispase enzymes disrupt lymphocyte surface molecules. J Immunol Methods 194(2):211–216 13. Mulder WM, Koenen H, van de Muysenberg AJ et al (1994) Reduced expression of distinct T-cell CD molecules by collagenase/DNase treatment. Cancer Immunol Immunother 38(4):253–258 14. Autengruber A, Gereke M, Hansen G et al (2012) Impact of enzymatic tissue disintegration on the level of surface molecule expression and immune cell function. Eur J Microbiol Immunol (Bp) 2(2):112–120. https://doi. org/10.1556/EuJMI.2.2012.2.3

Chapter 9 Depletion and Reconstitution of Macrophages in Mice Lisa K. Kozicky and Laura M. Sly Abstract Macrophages are innate immune cells, which have important roles in the inflammatory response to infections or tissue injury, and have an equally important role in the resolution of inflammation. Macrophages play a key part in directing the innate immune response and subsequent adaptive immune response. They can acquire a variety of distinct but also overlapping activation states, depending on the local microenvironment, in order to perform these functions. Stimuli, such as IFNγ and LPS, can promote an inflammatory activation state, which is associated with the production of reactive oxygen species, and pro-inflammatory cytokines and chemokines. Immune complexes and LPS can promote an anti-inflammatory activation state to prevent damage to the host, which is associated with the production of high levels of the anti-­ inflammatory cytokine IL-10 and low levels of pro-inflammatory cytokines. Wound-healing macrophages can be activated by IL-4 or IL-13 and have roles in tissue remodeling and the resolution of inflammation. Macrophages are present in nearly every tissue of the body and are important for maintaining homeostasis, but their dysfunction can also lead to diseases, such as inflammatory bowel disease. To study the role macrophages play in a complex in vivo environment, depletion and reconstitution experiments can be utilized. Clodronate liposomes are an effective and versatile way to deplete macrophages in vivo; they can allow selective depletion from tissues of interest and can be used on transgenic mice. However, clodronate liposomes deplete all types of macrophages as well as dendritic cells, so other strategies are required in parallel to determine whether macrophages or macrophages of a particular activation state are required. Reconstitution of macrophages by adoptive transfer can be performed, with or without prior depletion, to further suggest that the observed effect is macrophage dependent. Macrophages activated ex vivo or macrophages from transgenic mice can be adoptively transferred during disease models to determine whether a specific protein or activation state affects disease outcome. Macrophage contribution to health and disease can be effectively studied using depletion with clodronate liposomes and by macrophage reconstitution, as demonstrated in this chapter. Key words Clodronate liposomes, PBS liposomes, Bone marrow-derived macrophages (BMDMs), Macrophage activation, Macrophage colony-stimulating factor (MCSF), PKH26

1  Introduction Macrophages are phagocytic innate immune cells, which have an important role in directing the innate and acquired immune responses. These critical immune cells participate in all steps of the immune response, which include recognition, response, and Irving C. Allen (ed.), Mouse Models of Innate Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1960, https://doi.org/10.1007/978-1-4939-9167-9_9, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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r­ esolution of inflammation. A key feature of macrophages is their plasticity or their ability to respond to local environmental cues and change their phenotype in order to mount an appropriate response. Thus, macrophages are heterogeneous cells, which have several distinct but potentially overlapping activation states, which makes them difficult to study in vivo [1]. There are three well-studied macrophage activation states that can be modeled in vitro by treating macrophages with exogenous stimuli. Macrophages primed with interferon γ (IFNγ) and activated with TLR agonists, such as bacterial lipopolysaccharide (LPS), defined as M(IFNγ + LPS), secrete pro-inflammatory cytokines and chemokines, which are critical in host defense against a pathogen [2]. Macrophages have an equally important role in turning off the immune response, to prevent damage to host tissues [3]. Examples of this immunosuppressive activation state in vitro are macrophages primed with immune complexes (Ic) or the pooled polyclonal IgG drug intravenous immunoglobulin (IVIg), M(Ic), or M(IVIg), respectively. They produce very high amounts of the anti-inflammatory cytokine IL-10 and produce very low amounts of pro-inflammatory cytokines, in response to normally inflammatory stimuli such as LPS [4, 5]. To resolve tissue damage from an inflammatory response, macrophages acquire a wound-healing phenotype, characterized in vitro by activation with IL-4 or IL-13, M(IL-4) or M(IL-13) [6]. In response to TLR activation, wound-healing macrophages produce relatively lower amounts of pro-inflammatory cytokines and higher amounts of IL-10, relative to M(IFNγ + LPS) [7]. However, M(Ic) are distinct from M(IL-4), as they do not contribute to the production of extracellular matrix and do not express key macrophage M(IL-4) protein markers, such as arginase 1 (Arg1) or Ym1 [8]. Macrophages can have protective or pathogenic functions depending on their activation state. For example, inflammatory macrophages can be cytotoxic to tumors but can also promote tumor growth through wound-healing or immunosuppression [9]. In the intestine, macrophages sample luminal contents, promoting a tolerogenic response to intestinal bacteria [10]. Conversely, macrophages can promote an excessive inflammatory response to normal intestinal bacteria and contribute to the inflammation associated with inflammatory bowel disease (IBD), whereas an excessive healing response can result in fibrosis associated with IBD [10]. To study the function of macrophages in mice, depletion experiments can be performed with liposomes loaded with clodronate. Clodronate is a small hydrophilic bisphosphonate molecule and when encapsulated in liposomes is internalized by professional phagocytes, including macrophages and some dendritic cells [11]. Once internalized, lysosomal phospholipases cause the release of clodronate into the cell and results in macrophage “suicide” or apoptosis [12]. Clodronate has low toxicity for non-phagocytic cells, as it cannot

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easily escape the liposomal phospholipid bilayer [11]. Depletion of macrophages using clodronate liposomes has advantages over other methods; they are relatively inexpensive, easy to administer, effective, and can be used with transgenic mice. Macrophages can also be depleted using transgenic mice expressing CD11b-DTR or CD11cDTR, which depletes CD11b- or CD11c-expressing cells when mice are exposed to the diphtheria toxin. In contrast to clodronate liposome-mediated macrophage depletion, this technique is expensive and time-consuming, especially if mice have to be crossed with other genetically modified mice for experiments. Clodronate liposomes have been used in vitro in cultured slices of brain, to show that microglia are important in homeostasis of the CNS, to promote the death of developing neurons involved in synaptogenesis [13]. They have also been used in the AOM-DSS model of colorectal cancer in mice, which showed that macrophages can promote tumorigenesis and alter the gut microbiota [14]. Our laboratory has used clodronate liposomes in SHIP deficient mice, to demonstrate that macrophages contribute to the development of spontaneous ileal inflammation in these mice [15]. Interestingly, clodronate liposomes are currently being tested in humans for diseases such as rheumatoid arthritis, to transiently suppress synovial macrophage activity [16]. Although the role of macrophages can be determined in specific tissues during homeostasis and disease, the contribution of macrophage activation states cannot be determined with clodronate liposome depletion [17]. The route of administration, dose given, and timing of injections can affect macrophage depletion. Different levels of macrophage depletion can be obtained, depending on route of administration. For example, clodronate liposomes can be used intravenously to deplete nearly 90% of all macrophages in the lamina propria of the intestine or can be used intraperitoneally to deplete nearly 50% of macrophages [18–20]. Tissue-specific depletion can be achieved when clodronate liposomes are used locally, such as to deplete specific resident macrophages intratracheally, intrarectally, or within the testes [12]. The dose of clodronate can be increased in order to get higher depletion rates, and if necessary, intraperitoneal rather than intravenous injections can be given in order to maximize the amount of liposomes given per injection. Multiple injections may be needed for maximal or longer-term macrophage depletion, which is particularly important in prolonged disease models. To deplete resident and infiltrating macrophages in an inflammatory disease model, such as DSS-induced colitis, injections should be given prior to and during the onset of inflammation [17]. Both PBS and PBS liposomes should be given as controls, although in some experimental conditions PBS liposomes have also been reported to deplete macrophages [18, 20]. PBS liposomes may also temporarily inhibit phagocytosis, so the inclusion of PBS liposomes as an appropriate control must be

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determined and evaluated experimentally [11]. Verification of macrophage depletion must be performed, either histologically or by flow cytometry, to determine extent of depletion with clodronate liposomes compared to controls, PBS liposomes and PBS. Since clodronate liposomes deplete all professional phagocytes, not only macrophages, macrophage reconstitution experiments can be used in parallel to ensure that the effect is macrophage specific. Reconstitution of macrophages by adoptive transfer of bone marrow-derived macrophages can also be used to test whether a specific macrophage activation state can influence outcome of disease, as clodronate liposomes effectively deplete all types of macrophages [17]. For example, adoptive transfer of M(IL-4), but not untreated macrophages or M(IFNγ) ameliorate inflammation during DSSinduced colitis [17]. A benefit of reconstitution experiments is that they can be done with transgenic mice, to determine whether expression of a protein is required for the effect. Using this strategy, our laboratory has reported that Arg1 expression is required for M(IL-4)-mediated amelioration of inflammation in PI3Kp110δ deficient mice during DSS-induced colitis [21]. We, and others, have reconstituted macrophages by intravenous injection, but other routes, such as intraperitoneal injection, have been used successfully [21, 22]. Adoptively transferred macrophages can be tracked using a florescent dye, such as PKH26, or by using a macrophages derived from a GFP reporter mouse [21, 23]. Since macrophages are responsive to cues from the microenvironment, the ex vivo skewing of macrophage activation states can be affected once the transferred macrophages encounter a new environment. Despite that, macrophages can be transferred prior to the onset of a disease model to determine whether they can prevent or reduce disease development, or macrophages can be transferred during disease development to determine whether they can treat or impact the natural development of disease. Macrophages can also be transferred into mice after depletion of resident and/or recruited macrophages to promote engraftment of the transferred macrophages. Herein, we describe effective approaches to study macrophage function in mice. In this chapter, we demonstrate the depletion of macrophages with clodronate liposomes and the derivation and reconstitution of bone marrow macrophages. The protocols here can be applied to a variety of models of disease and can be used with a variety of transgenic mice.

2  Materials 2.1  Depletion of Macrophages Using Clodronate Liposomes

1. Sterile laminar flow hood. 2. 1 mL syringe. 3. 26-gauge needle.

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4. Sterile phosphate buffered saline (PBS) (pH 7.4). 5. 5 mg/mL PBS liposomes (www.clodronateliposomes.com). 6. 5 mg/mL clodronate liposomes (www.clodronateliposomes. com) (see Notes 1 and 2). 7. 8–12-week-old C57BL/6 or desired mouse strain (mice should be housed and euthanized according to institutional requirements). 8. Foam board. 9. Forceps. 10. Scissors. 11. Formalin. 12. 70% ethanol. 13. Tissue cassettes for preparing histological sections. 2.2  Derivation of Bone Marrow Macrophages

1. Sterile laminar flow hood. 2. Foam board. 3. Forceps. 4. Scissors. 5. 70% ethanol. 6. 6-well tissue culture-treated, flat bottom plate. 7. Iscove’s modified Dulbecco’s medium (IMDM). 8. Fetal bovine serum (FBS). 9. Bone flush medium: IMDM, 10% FBS. 10. 50 mL conical tubes. 11. 10 mL syringe. 12. 26-gauge needle. 13. 75 cm2 tissue culture-treated flask. 14. Incubator set at 37 °C and 5% CO2. 15. Recombinant murine macrophage colony-stimulating factor (MCSF). 16. Penicillin-streptomycin. 17. Monothioglycerol (MTG). 18. MCSF culture medium: IMDM, 10% FBS, 5 ng/mL MCSF, 100 U/Ml penicillin/streptomycin, and 150 μM MTG. 19. Bright-field microscope. 20. Cell dissociation buffer (Enzyme-free, Gibco-BRL). 21. Cell scraper. 22. Sterile phosphate buffered saline (PBS) pH 7.4.

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2.3  Reconstitution of Macrophages

1. 8–12-week-old C57BL/6 or desired mouse strain (mice should be housed and euthanized according to institutional requirements). 2. Sterile laminar flow hood. 3. 26- or 27-gauge needle. 4. 1 mL syringe. 5. Electric heating pad. 6. Mouse restrainer with tail access. 7. Alcohol swab. 8. Gauze.

3  Methods 3.1  Depletion of Macrophages Using Clodronate Liposomes

1. Store liposomes at 4 °C. Remove clodronate liposomes, PBS liposomes (liposome control), and sterile PBS (injection control) from the 4 °C refrigerator, 2 h prior to injection to allow them to acclimate to room temperature (18 °C) (see Note 3). 2. In a sterile cabinet (see Note 4), invert each of the liposome tubes 8–10 times to ensure a homogeneous solution, as liposomes sediment during storage. Load 200 μL of clodronate liposomes, PBS liposomes, and sterile PBS (pH 7.4) each into 1 mL syringes with a 26-gauge needle attached. Remove any air bubbles from syringe. 3. To inject intraperitoneally (see Note 5), scruff the mouse with one hand by grabbing its skin at the back of the neck with your thumb and index finger and holding its tail and hind legs with your remaining fingers. Tilt its body toward the ground so that its internal organs move away from the site of injection. 4. Invert the syringe with liposomes six times to resuspend the solution before injection (see Notes 6 and 7). 5. Place the needle at a 30–40° angle relative to the mouse’s abdomen, in the lower right quadrant with the bevel up, insert the needle, and inject 200 μL of clodronate liposomes, PBS liposomes, or PBS (see Note 8). 6. When using an induced inflammation model, such as DSS-­ induced colitis, resident macrophages can be depleted by injecting mice 4 days prior to onset of inflammation. Newly infiltrating macrophages can be depleted by injecting every 2 days during the protocol. 7. Euthanize mice, using CO2 asphyxiation, according to institution-­approved protocols. 8. Remove tissues, as planned for experimental analysis, along with tissues from site of interest to confirm depletion of

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F4/80

Scale bars = 100mm

Clod lip

800 F4/80+ cells per field

PBS lip

PBS

*

600

*

400 200 0

PBS

PBS lip Clod lip

Fig. 1 Clodronate liposomes deplete F4/80+ colonic macrophages in vivo. C57BL/6 mice were given 2.5% DSS for 7 days and injected intraperitoneally 4 days prior to experiment and on days 1, 3, and 5 with either a PBS (injection control), PBS liposomes (PBS lip), or clodronate liposomes (Clod lip). Colons were removed, fixed, and stained for mature macrophages (F4/80) by immunohistochemistry. F4/80+ cells were quantified at 40x magnification from five sections/mouse, by two individuals blinded to the experimental conditions. n = 6 mice/ group from n = 3 independent experiments. *p 95% positive for F4/80 and Mac-1 by flow cytometry. 16. For example, macrophages can be activated with IFNγ (10 ng/mL) or IL-4 (10 ng/mL) from day 7 until day 10 or with IVIg (30 mg/mL) + LPS (10 ng/mL) for 24 h. 17. If activating macrophages prior to reconstitution, cells should be plated for phenotypic analysis (1 × 106 cells/mL). Leave activated macrophages either unstimulated or stimulated with LPS (10 ng/mL), in duplicate for 24 h (37 °C, 5% CO2). Collect supernatants and remove any particulate matter by spinning at 10,000 × g for 5 min. Collect and save clarified cell supernatants at −80 °C for analysis of IL-10 and IL-12/23p40 production by enzyme-linked immunosorbent assay (ELISA). M(IFNγ + LPS) will produce high amounts of IL-12/23p40 and low amounts of IL-10 [24]. M(IL-4 + LPS) will produce higher amounts of IL-10 and lower amounts of IL-12/23p40 than M(IFNγ + LPS) and will constitutively produce the enzymes Arg1 and Ym1 that are detectable by Western blotting [24]. M(IVIg + LPS) will produce very high amounts of IL-10 and very low amounts of IL-12/23p40, and supernatants can be collected from 75 cm2 flask after activation [5]. 18. Apply gentle pressure as you scrape. The scraper should glide across the surface smoothly. Alternatively, the flask can be gently tapped on a hard surface to detach cells. 19. Slowly pipette up and down five times with 1 mL initially, to de-clump pelleted cells, then add and resuspend with another 4 mL of PBS to count macrophages. 20. 100 μL can be comfortably and safely injected into mice via tail vein. This concentration allows for 1 × 106 macrophages to be injected. 21. Macrophages must be well suspended in PBS. Clumps of macrophages, when injected intravenously, can restrict blood flow and cause death. Macrophages precipitate out of PBS quickly, so prepare syringes just prior to injections. 22. PBS is given as an injection control, and untreated macrophages are given as a control, to determine whether the addition of resting bone marrow-derived macrophages affects disease.

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23. Macrophage cell membranes can be linked with PKH26 florescent dye, as per kit instructions (Sigma), or cells can be derived from GFP+ reporter strain mice. PKH26+ or GFP+ adoptively transferred macrophages can be detected histologically or by flow cytometry. References 1. Murray PJ, Wynn TA (2011) Protective and pathogenic functions of macrophage subsets. Nat Rev Immunol 11:723–737 2. Martinez FO, Gordon S (2014) The M1 and M2 paradigm of macrophage activation: time for reassessment. F1000Prime Rep 6:13 3. Fleming BD, Mosser DM (2011) Regulatory macrophages: setting the threshold for therapy. Eur J Immunol 41:2498–2502 4. Sutterwala FS, Noel GJ, Salgame P et al (1998) Reversal of proinflammatory responses by ligating the macrophage Fcgamma receptor type I. J Exp Med 188:217–222 5. Kozicky LK, Zhao ZY, Menzies SC et al (2015) Intravenous immunoglobulin skews macrophages to an anti-inflammatory, IL-10-­ producing activation state. J Leukoc Biol 98: 983–994 6. Gordon S (2003) Alternative activation of macrophages. Nat Rev Immunol 3:23–35 7. Murray PJ, Allen JE, Biswas SK et al (2014) Macrophage activation and polarization: nomenclature and experimental guidelines. Immunity 41:14–20 8. Edwards JP, Zhang X, Frauwirth KA et al (2006) Biochemical and functional characterization of three activated macrophage populations. J Leukoc Biol 80:1298–1307 9. Mosser DM, Edwards JP (2008) Exploring the full spectrum of macrophage activation. Nat Rev Immunol 8:958–969 10. Heinsbroek SE, Gordon S (2009) The role of macrophages in inflammatory bowel diseases. Expert Rev Mol Med 11:e14 11. Van Rooijen N, Sanders A (1994) Liposome mediated depletion of macrophages: mechanism of action, preparation of liposomes and applications. J Immunol Methods 174:83–93 12. van Rooijen N, Hendrikx E (2010) Liposomes for specific depletion of macrophages from organs and tissues. Methods Mol Biol 605: 189–203 13. Marín-Teva JL, Dusart I, Colin C et al (2004) Microglia promote the death of developing Purkinje cells. Neuron 41:535–547 14. Bader JE, Enos RT, Velázquez KT et al (2018) Macrophage depletion using clodronate lipo-

somes decreases tumorigenesis and alters gut microbiota in the AOM/DSS mouse model of colon cancer. Am J Physiol Gastrointest Liver Physiol 314:G22–G31 15. Ngoh EN, Weisser SB, Lo Y et al (2016) Activity of SHIP, which prevents expression of interleukin 1β, is reduced in patients with Crohn’s disease. Gastroenterology 150:465–476 16. Barrera P, Blom A, van Lent PL et al (2000) Synovial macrophage depletion with clodronate-­ containing liposomes in rheumatoid arthritis. Arthritis Rheum 43:1951–1959 17. Weisser SB, van Rooijen N, Sly LM (2012) Depletion and reconstitution of macrophages in mice. J Vis Exp 66:4105 18. Weisser SB, Brugger HK, Voglmaier NS et al (2011) SHIP-deficient, alternatively activated macrophages protect mice during DSS-­induced colitis. J Leukoc Biol 90:483–492 19. Smith P, Mangan NE, Walsh CM et al (2007) Infection with a helminth parasite prevents experimental colitis via a macrophage-­mediated mechanism. J Immunol 178:4557–4566 20. Qualls JE, Kaplan AM, van Rooijen N et al (2006) Suppression of experimental colitis by intestinal mononuclear phagocytes. J Leukoc Biol 80:802–815 21. Weisser SB, Kozicky LK, Brugger HK et al (2014) Arginase activity in alternatively activated macrophages protects PI3Kp110δ deficient mice from dextran sodium sulfate ­ induced intestinal inflammation. Eur J Immunol 44:3353–3367 22. Potts BE, Hart ML, Snyder LL et al (2008) Differentiation of C2D macrophage cells after adoptive transfer. Clin Vaccine Immunol 15:243–252 23. McNeill E, Iqbal AJ, Jones D et al (2017) Tracking monocyte recruitment and macrophage accumulation in atherosclerotic plaque progression using a novel hCD68GFP/ ApoE−/− reporter mouse-brief report. Arterioscler Thromb Vasc Biol 37:258–263 24. Weisser SB, McLarren KW, Kuroda E et al (2013) Generation and characterization of murine alternatively activated macrophages. Methods Mol Biol 946:225–239

Chapter 10 Microfluidic Platform to Quantify Neutrophil Migratory Decision-Making Brittany P. Boribong, Amina Rahimi, and Caroline N. Jones Abstract Neutrophils are the most abundant leukocytes in blood, serving as the first line of host defense in tissue damage and infections. Upon activation by chemokines released from pathogens or injured tissues, neutrophils migrate through complex tissue microenvironments toward sites of infections along the chemokine gradients, in a process named chemotaxis. However, current methods for measuring neutrophil chemotaxis require large volumes of blood and are often bulk, endpoint measurements. To address the need for rapid and robust assays, we engineered a novel dual gradient microfluidic platform that precisely quantifies neutrophil migratory decision-making with high temporal resolution. Here, we present a protocol to measure neutrophil migratory phenotypes (velocity, directionality) with single-cell resolution. Key words Microfluidics, Chemotaxis, Immune cell, Neutrophil migration phenotype, Migratory decision-making

1  Introduction Microfluidics is an enabling technology for studying chemotaxis of neutrophils in vitro with high temporal and spatial resolution [1, 2]. In this chapter, we describe a protocol for probing murine neutrophil migratory decision-making using a dual gradient microfluidic device. The same devices can be used to study chemotaxis of other migratory cell types isolated from small animals or humans (Table 1). Traditional migration assays, such as the transwell assay or Boyden chamber [11], are not ideal to measure chemotaxis in cells isolated from mouse samples as they require large numbers of cells (>100,000 per condition). These assays also cannot be used to monitor transient changes in neutrophil chemotaxis in small laboratory animals, such as mice, because the volume of blood needed for neutrophil isolation allows for only one sample and often even requires pooling of blood from multiple animals for a single assay. For instance, a study involving multiple conditions and treatments over multiple time-points could potentially require thousands of Irving C. Allen (ed.), Mouse Models of Innate Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1960, https://doi.org/10.1007/978-1-4939-9167-9_10, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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Table 1 Microfluidic chemotaxis assays

Cell type

Microfluidic migration channel dimensions

Reference

Mouse neutrophils

12 μm × 3 μM

[3]

Human neutrophils

10 μm × 5 μm

[4, 5]

Human monocytes

10 μm × 10 μm

[6]

Mouse dendritic cells

28 μm × 28 μm

[7]

Human T cells

10 μm × 10 μm

[8]

Human cancer cells

20 μm × 10 μm

[9, 10]

mice using current chemotaxis assays. This restricts the basic biological research that can be done to understand the complex dynamics of immune function in the context of injury, infection, or inflammatory disorders often studied in murine models. Significantly, the microfluidic platform described here can measure migratory phenotypes from as little as 500 cells, and the incorporation of the microfluidic device in a 12- or 24-well plate facilitates the screening of multiple mediators of murine neutrophil chemotaxis simultaneously [12]. Transwell assays are bulk end-point measurements of cells migrating to a single chemoattractant, thus do not allow for the measurements of cell velocity or directionality. To address the need for a neutrophil functional assay that is rapid, robust, while requiring minimal blood volume, we have developed a microfluidic platform that allows for the formation of two, opposing chemoattractant gradients at a high temporal and single-cell resolution. The microfluidic chemotaxis platform presented in this chapter allows quantification of neutrophil migratory decision-making with single-cell resolution. The device has a central loading channel, in which the cells are loaded, connected to two, separate chemoattractant reservoirs via ten migration channels (Fig. 1). Once chemoattractant is loaded into the reservoirs, it will diffuse down toward the center-loading channel, creating a linear gradient of chemoattractant for the cells to migrate toward. The migration channels were designed to measure 6 μm wide by 6 μm tall to accommodate mouse neutrophils (~10 μm in diameter). This geometry allows the neutrophil to squeeze through the channel, which increases migratory persistence as it mimics the endothelial transmigration neutrophils would undergo in vivo [13]. Also included on the migration channels are cell mazes, which allow for the measurement of directionality (Fig. 2). The concentration of chemoattractant in the perpendicular migration channel maze is less than the concentration of chemoattractant in the straight c­ hannels. Thus, if a cell enters a cell maze, it is following a weaker chemoattractant signal. In healthy

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Fig. 1 Microfluidic device design. (a) 3D schematic. (b) Overlay microscopy image take on Nikon TiE (10×). Chemoattractants have been labeled with fluorescent dextran (FITC and TRITC) to allow visualization of the gradient formation. Mouse neutrophil nuclei are labeled with Hoechst stain (DAPI). Scale bar = 250 μM. Before the chemotaxis experiment, (i) chemoattractant is primed into the two separate chemoattractant reservoirs, and (ii) cells are loaded into the cell loading chamber. (iii) A gradient is formed along the 10 migration channels (ii) toward the cell loading chamber. Within the mazes (iv), the chemoattractant concentration is lower than the concentration within the parallel migration channels, allowing measurement of directional migration

Fig. 2 Quantification of directional neutrophil migration along a chemoattractant gradient. (a) Neutrophils displaying directional migration within a migration channel. (b) Neutrophils displaying nondirectional migration and are “lost” within a cell maze (10×). Scale bar = 250 μM

neutrophils, cells will follow the steeper gradient >90% of the time [3]. The microfluidic platform enables the study of two, opposing chemoattractants (Table 2). It can also be used to study the chemoattractive and chemorepulsive properties of a single chemoattractant [13]. Preparation of cells for the microfluidic assays can come from various sources. This chapter will detail neutrophil isolation from bone marrow, but previous studies have isolated neutrophils from intraperitoneal (IP) injection and whole blood [3, 12]. The on-chip assay enables the measurement of several different migratory phenotypes, including cell numbers, velocity, and directionality (Table 3). Importantly, we can also measure the chemoat-

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Table 2 Validated chemoattractants to probe cell migration via microfluidic assay Chemoattractant

Description

Reference

Leukotriene B4 (LTB4)

Leukotriene involved in inflammation

[3]

N-Formyl-Met-Leu-Phe (fMLP)

Prototypical representative of the N-fomylated oligopeptide family of chemotactic factors released by bacteria

[3]

Macrophage inflammatory Involved in promoting proprotein 1-alpha inflammatory responses (MIP-1α)

[14]

Macrophage inflammatory Involved in promoting proprotein 1-beta inflammatory responses (MIP-1β)

[15]

Complement component 5a (C5a)

Initiates accumulation of leukocytes

[3]

Interleukin 8 (IL-8)

Induces chemotaxis in granulocytes

[16]

Table 3 Migration phenotypes quantified using a microfluidic platform Migration Phenotype Definition

Units

Cell count in chemoattractant reservoir

Percentage of cells that migrated fully toward a specific chemoattractant

Cell percentage

Cell count in Percentage of cells that entered a migration channels migration channel toward a specific chemoattractant

Cell percentage

Velocity

Speed in which a cell migrated toward a specific chemoattractant

μm/min

Directionality

Number of cells that can migrate Cell number straight across a migration channel toward a specific chemoattractant (directional) vs. the number of cells that enter the maze (nondirectional)

tractant preference of a neutrophil in the presence of two opposing gradients. This chapter will detail how to isolate and prepare neutrophils from bone marrow for the migration assay, fabricate the microfluidic device, prime the microfluidic device, run the assay, and analyze the resulting migration data.

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2  Materials 2.1  Bone Marrow-­ Derived Neutrophil Isolation

1. Laminar flow hood. 2. 70% ethanol. 3. Mouse femurs. 4. Small dissection scissors. 5. Forceps. 6. Sterile 6-well plates. 7. 50 mL conical tubes. 8. 15 mL conical tubes. 9. 1.5 mL microcentrifuge tubes. 10. 1 mL syringes. 11. 26 G needles. 12. 70 μM cell strainers. 13. Ice, ice bucket. 14. RPMI 1640 Medium with HEPES buffer, l-glutamine, and penicillin-streptomycin supplemented with 10% fetal bovine serum (FBS). Store at 4 °C. 15. Hank’s Balanced Saline Solution (HBSS) with sodium bicarbonate, without phenol red, calcium chloride, and magnesium sulfate. 16. 5 mL Round-Bottom Polystyrene Tubes. 17. RoboSep Buffer (STEMCELL Technologies). 18. EasySep™ mouse neutrophil Enrichment Kit (STEMCELL Technologies). 19. EasySep™ Magnet (STEMCELL Technologies). 20. Centrifuge. 21. Recombinant murine granulocyte colony-stimulating factor (G-CSF). 22. Humidified CO2 incubator.

2.2  Microfluidic Device Fabrication

1. AutoCAD computer software and LinkCAD. 2. Clean room. 3. Oxygen Plasma Machine (Nordson MARCH). 4. Hot Plate. 5. Oven. 6. Ultrahigh purity nitrogen (99.999%). 7. Oxygen.

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8. Clean room tape. 9. 100 mm silicon wafer. 10. Polydimethylsiloxane (PDMS) and initiator (184) (Sylgard). 11. Large plastic weighing tray. 12. Plastic fork. 13. Vacuum desiccator. 14. Deionized water. 15. Tweezers. 16. Scalpels. 17. 0.75 mm biopsy punch. 18. 1.0 mm biopsy punch. 19. 8.0 mm biopsy punch. 20. 6-well glass bottom plates. 2.3  Microfluidic Assay Preparation

1. RPMI 1640 Medium with HEPES buffer, l-glutamine, and penicillin-streptomycin supplemented with 10% fetal bovine serum (FBS). Store at 4 °C. 2. Recombinant murine granulocyte colony-stimulating factor (G-CSF). 3. Gel loading pipette tips. 4. Microfluidic devices bonded to 6-well glass bottom plates. 5. Mouse fibronectin. 6. Vacuum desiccator. 7. Bone marrow-derived neutrophils in complete media (RPMI+10% FBS) (1.0 × 107 cells/mL). (Option to stain with Hoechst 33342 Fluorescent Stain at concentration of 1:1000). 8. Leukotriene B4 (LTB4) (Cayman Chemical). 9. N-Formyl-Met-Leu-Phe (fMLP) (Sigma Aldrich).

2.4  Microscopy

1. Primed microfluidic devices on a 6-well glass bottom plate. 2. 5% CO2. 3. Biochamber to maintain constant temperature and humidity levels (Okolab). 4. Temperature unit (Okolab). 5. Gas chamber to hold standard cell culture well plate and contain humidified gas. 6. Gas controllers to control levels of CO2 and O2 (Okolab). 7. Inverted microscope equipped with motorized stage and perfect focus system (PFS).

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3  Methods 3.1  Sample Preparation (See Note 1)

1. Isolate femurs from mouse legs. 2. Flush out the bone marrow using a 26 G needle and syringe at least five times or until the bone becomes translucent. 3. Remove excess tissue and bone residue using a 70 μM cell strainer. 4. Isolate immune cells following STEMCELL Technology’s negative selection protocol. Ensure that isolation occurs at room temperature (off ice) and without lysing of red blood cells, as this will negatively affect migration. 5. Incubate isolated immune cells with Hoechst fluorescent stain at a concentration of 1:1000 for 10 min at 37 °C at 5% CO2 to fluorescently stain DNA in the nucleus. 6. Resuspend immune cells in complete media supplemented with 100 nM of G-CSF to a concentration of 1.0 × 107 cells/ mL.

3.2  Microfluidic Device Fabrication

1. Design microfluidic device using AutoCAD and LinkCad Software. 2. Order mask transparency (FineLine Imaging). 3. Fabricate master mold wafer using standard photolithography techniques [17]. Pattern the first 6-μm-thin epoxy-based negative photoresist layer to define the migration channels according to the instructions from the manufacturer. Pattern the second 50-μm-thick layer to define the cell-loading channel and chemoattractant reservoirs. 4. Mix 20 g of polydimethylsiloxane (PDMS) and 2 g of initiator using a plastic fork in a plastic weighing tray. Mix for at least 5 min, until the PDMS is opaque with bubbles. 5. Pour PDMS mixture onto the wafer to create PDMS devices. 6. Placed poured PDMS mold in a vacuum desiccator for 4 h to degas the PDMS. 7. Place microfluidic devices in an oven set to 65 °C for at least 6 h to bake and cure the PDMS. 8. Punch out two cell-loading inlets using a 1.0 mm biopsy punch. 9. Punch out two chemoattractant reservoir inlets using a 0.75 mm biopsy punch. 10. Punch out microfluidic devices using an 8.0 mm biopsy punch. 11. Remove dust or PDMS particles from microfluidic devices by using nitrogen and clean room tape. 12. Rinse a glass bottom 6-well plate with deionized water and dry with nitrogen.

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13. Oxygen plasma treat the 6-well plate twice: once alone and once again with six microfluidic devices. 14. Using tweezers, carefully place devices in the center of the wells of the plate to bond devices to glass. Press down on devices to ensure the complete surface of the microfluidic devices are bonded to the glass. Do not attempt to move or adjust device position after initial bonding. 15. Bake the plate with bonded devices on a hot plate set to 80 °C for 10 min. 3.3  Microfluidic Assay Preparation (See Note 2)

1. Create fibronectin solution by mixing mouse fibronectin [stock solution 1 mg/mL] and complete media to create 11 μg/mL fibronectin solution (see Note 3). 2. Place a droplet of fibronectin (~50 μL) on top of the device, ensuring all four punches are covered. 3. Place plate in vacuum desiccator for 10 min to allow fibronectin to fill channels within the device, and remove air from the PDMS. 4. Remove plate from desiccator and confirm fibronectin has filled the device (see Note 4). 5. Allow the fibronectin to evaporate at room temperature for 30 min. 6. Create LTB4 chemoattractant solution by serial diluting stock chemoattractant with complete media until it reaches a concentration of 100 nM (see Note 5). 7. Create fMLP chemoattractant solution by serial diluting stock chemoattractant with complete media until it reaches a concentration of 100 nM (see Note 5). 8. Fill wells with 3 mL of complete media supplemented with 100 nM of G-CSF, until the tops of devices are submerged under media. 9. Using a gel loading pipette tip, slowly load 10 μL of chemoattractant #1 (i.e., LTB4) into one chemoattractant reservoir. When chemoattractant is expelling from the center-loading channel inlets, release the pipette tip. 10. Repeat previous step with chemoattractant #2 (fMLP or media for a negative control) in the second reservoir. 11. Slowly rinse the center-loading channel with 10 μL complete media. This will remove any chemoattractant in the loading channel as well as create a gradient in the cell migratory channels from the reservoir to the center channel (see Note 6). 12. Using a gel loading pipette tip, slowly load 10 μL of cells in the center-loading channel. When cells are expelling from the opposite inlet, release the pipette tip.

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13. Remove the surrounding media and add 3 mL of complete media supplemented with 100 nM G-CSF, warmed to 37 °C (see Note 7). 3.4  Microscopy and Image Analysis

1. Set biochamber to 37 °C, 100% humidity, and 5% CO2. 2. Image the entire device using time-lapse imaging on a microscope at a magnification of 10× or higher. 3. Image the device with both TRANS and DAPI channels. 4. Image at intervals no greater than 3 min per frame. 5. Migration is quantified as follows: (1) percentage of cells migrating fully toward chemoattractant reservoirs, (2) velocity of migration, and (3) directionality of migration. 6. Calculate cell counts using ImageJ software [18]. 7. Calculate cell velocity and directionality using the FIJI [19] plug-in, TrackMate [20].

4  Notes 1. Neutrophils can also be obtained from whole blood or intraperitoneal (IP) injection, but must not be lysed of red blood cells as to not affect their migratory abilities. 2. Microfluidic devices can be prepared several weeks in advanced. However, microfluidic devices that are primed immediately after oxygen plasma treatment will be hydrophilic, which helps promote the priming of channels within the device. 3. Fibronectin allows the neutrophils to adhere to a surface as opposed to migrating on a flat, glass surface. Adhesion to a component of an extracellular matrix resembles adhesion to endothelial cells before transmigration into the tissue. 4. Fibronectin has filled the chambers, and there are no bubbles present in the device. 5. In this chapter, we focus specifically on LTB4 and fMLP, but any chemoattractant can be replaced in this step (Table 1). 6. To confirm gradient within the microfluidic device, add 5 μL of fluorescent dextran to chemoattractant solution to visualize chemoattractant in device. Image the fluorescent channel on the microscope and using ImageJ, calculate fluorescent intensity along migration channel to confirm gradient formation. 7. Placing the top, plastic lid to the 6-well plate in an incubator set to 37 °C before imaging will prevent condensation forming on the lid during the assay.

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References 13(11):1063–1071. https://doi.org/10.1038/ 1. Wu J, Wu X, Lin F (2013) Recent developnmat4062 ments in microfluidics-based chemotaxis studies. Lab Chip 13(13):2484–2499. https://doi. 11. Boyden S (1962) The chemotactic effect of org/10.1039/c3lc50415h mixtures of antibody and antigen on polymorphonuclear leucocytes. J Exp Med 115: 2. Irimia D, Ellett F (2016) Big insights from 453–466 small volumes: deciphering complex leukocyte behaviors using microfluidics. J Leukoc Biol 12. Jones CN, Hoang AN, Dimisko L et al (2014) 100(2):291–304. https://doi.org/10.1189/ Microfluidic platform for measuring neutrojlb.5RU0216-056R phil chemotaxis from unprocessed whole blood. J Vis Exp 88. https://doi.org/ 3. Jones CN, Hoang AN, Martel JM et al (2016) 10.3791/51215 Microfluidic assay for precise measurements of mouse, rat, and human neutrophil chemotaxis 13. Boneschansker L, Yan J, Wong E et al (2014) in whole-blood droplets. J Leukoc Biol Microfluidic platform for the quantitative anal100(1):241–247. https://doi.org/10.1189/ ysis of leukocyte migration signatures. Nat jlb.5TA0715-310RR Commun 5:4787. https://doi.org/10.1038/ ncomms5787 4. Hamza B, Irimia D (2015) Whole blood human neutrophil trafficking in a microfluidic 14. Ramos CD, Canetti C, Souto JT et al (2005) model of infection and inflammation. Lab MIP-1alpha[CCL3] acting on the CCR1 recepChip 15(12):2625–2633. https://doi.org/ tor mediates neutrophil migration in immune 10.1039/c5lc00245a inflammation via sequential release of TNFalpha and LTB4. J Leukoc Biol 78(1):167–177. 5. Sackmann EK, Berthier E, Young EW et al https://doi.org/10.1189/jlb.0404237 (2012) Microfluidic kit-on-a-lid: a versatile platform for neutrophil chemotaxis assays. 15. Taub DD, Conlon K, Lloyd AR et al (1993) Blood 120(14):e45–e53. https://doi. Preferential migration of activated CD4+ and org/10.1182/blood-2012-03-416453 CD8+ T cells in response to MIP-1 alpha and MIP-1 beta. Science 260(5106):355–358 6. Jones CN, Dalli J, Dimisko L et al (2012) Microfluidic chambers for monitoring leuko- 16. Singer M, Sansonetti PJ (2004) IL-8 is a key cyte trafficking and humanized nano-­ chemokine regulating neutrophil recruitment proresolving medicines interactions. Proc in a new mouse model of Shigella-induced coliNatl Acad Sci U S A 109(50):20560–20565. tis. J Immunol 173(6):4197–4206 h t t p s : / / d o i . o r g / 1 0 . 1 0 7 3 / 17. Becker H, Gartner C (2000) Polymer micropnas.1210269109 fabrication methods for microfluidic analytical 7. Schwarz J, Bierbaum V, Merrin J et al (2016) A applications. Electrophoresis 21(1):12–26. microfluidic device for measuring cell migrahttps://doi.org/10.1002/ tion towards substrate-bound and soluble che(sici)1522-2683(20000101)21:13.0.co;2-7 doi.org/10.1038/srep36440 18. Schindelin J, Rueden CT, Hiner MC et al 8. Jain NG, Wong EA, Aranyosi AJ et al (2015) (2015) The ImageJ ecosystem: an open platMicrofluidic mazes to characterize T-cell exploform for biomedical image analysis. Mol ration patterns following activation in vitro. Reprod Dev 82(7–8):518–529. https://doi. Integr Biol (Camb) 7(11):1423–1431. https:// org/10.1002/mrd.22489 doi.org/10.1039/c5ib00146c 19. Schindelin J, Arganda-Carreras I, Frise E et al 9. Chen YC, Allen SG, Ingram PN et al (2015) (2012) Fiji: an open-source platform for Single-cell migration chip for chemotaxis-­based biological-­ image analysis. Nat Methods microfluidic selection of heterogeneous cell 9(7):676–682. https://doi.org/10.1038/ populations. Sci Rep 5:9980. https://doi. nmeth.2019 org/10.1038/srep09980 20. Tinevez JY, Perry N, Schindelin J et al (2017) 10. Wong IY, Javaid S, Wong EA et al (2014) TrackMate: an open and extensible platform for Collective and individual migration following single-particle tracking. Methods 115:80–90. the epithelial-mesenchymal transition. Nat Mater https://doi.org/10.1016/j.ymeth.2016.09.016

Chapter 11 A Vector Suite for the Overexpression and Purification of Tagged Outer Membrane, Periplasmic, and Secreted Proteins in E. coli Michael A. Casasanta and Daniel J. Slade Abstract Outer membrane and secreted proteins in Gram-negative bacteria constitute a high percentage of virulence factors that are critical in disease initiation and progression. Despite their importance, it is often difficult to study these proteins due to challenges with expression and purification. Here we present a suite of vectors for the inducible expression of N-terminally 6His-tagged outer membrane, periplasmic, and secreted proteins in E. coli and show this system to be capable of producing milligram quantities of pure protein for downstream functional and structural analysis. This system can not only be used to purify recombinant virulence factors for structural and functional studies but can also be used to create gain-of-­function E. coli for use in phenotypic screens, and examples of each are provided herein. Key words OmpA, TamA, SufI, Periplasm, Protein secretion, Sec apparatus, Twin-arginine translocation, Membrane protein, Protein purification, Protein expression

1  Introduction Protein secretion from the cytoplasm to the periplasm, outer membrane, and extracellular environment is an important mechanism for bacterial pathogenesis [1–3]. While a complete overview of all multicomponent protein secretion systems in bacteria is beyond the scope of this chapter [2], we will provide background and techniques used in Gram-negative bacteria for the initial export of proteins through the inner membrane. The two main systems for the export of proteins from the cytoplasm to the periplasm are the general secretion route termed the Sec pathway (unfolded proteins) and the twin-arginine translocation (Tat) pathway (folded proteins) [4]. Both the Sec and Tat systems rely on translocated proteins to have an N-terminal signal peptide (~15–40 AA) that is used for initial binding and initiation of inner membrane translocation. These signal peptides can be bioinformatically identified by online servers such as SignalP or PREDTAT (see Note 1). Irving C. Allen (ed.), Mouse Models of Innate Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1960, https://doi.org/10.1007/978-1-4939-9167-9_11, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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E. coli is the most common bacterial strain for recombinant protein expression, and this bacterium contains both Sec and Tat systems for protein translocation. Our experience is that while a protein from a non-E. coli genus of bacteria might be predicted to have a signal sequence, it is often not efficiently recognized by the E. coli machinery, which results in a lack of robust recombinant protein expression. To rectify this gap, we have developed a suite of vectors (see Fig. 1, Table 1, and Note 2) with signal sequences from highly expressed E. coli proteins, thereby facilitating the translocation of recombinant proteins through the inner membrane using either the Sec or Tat pathways. A similar method was previously used for the successful production of highly pure p ­ rotein

Fig. 1 Overview of E. coli expression vectors and multiple cloning sites (MCS) for the production of recombinant proteins secreted to extracytoplasmic spaces. (a) Overview showing features of vectors created from a pET-­ 16b base vector. (b) Overview showing features of vectors created from a pACYCduet-1 base vector. (c) Representation of the protein products created by both vectors. (d) Table describing vector-specific features for each of the five vectors described in this chapter

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Table 1 Addgene ID for plasmids Plasmid

Addgene ID

pDJSVT86

108965

pDJSVT87

109302

pDJSVT89

109303

pDJSVT90

109304

pDJSVT91

109305

for the structure determination of EstA, a Pseudomonas aeruginosa outer membrane protein of the Type 5 secreted autotransporter family [5]. To facilitate Sec secretion, we used the signal sequences from E. coli OmpA (residues 1–27) and TamA (residues 1–27), and for the TAT system, we used SufI (FtsP) (residues 1–30). To add flexibility to our vectors, we created versions that have variations of ampicillin or chloramphenicol antibiotic resistance and multiple origins of replication (p15A, ColE1). All versions of these vectors result in proteins that contain an N-terminal 6His tag for purification and detection. These vectors provide flexibility by allowing replication in a variety of E. coli hosts and, in addition, will allow for the simultaneous expression of two proteins by concurrently transforming compatible plasmids of interest. After predicted cleavage of the signal sequence by signal peptidase I (SPaseI) as proteins are transported to the bacterial extracytoplasmic space, [6] recombinant proteins will retain a minimal N-terminal extension and a 6His tag before the amino acids of your gene of interest. For vectors pDJSVT86 and pDJSVT87 which contain an OmpA 1–27 tag, the residues will be N-APKDNTHHHHHHxxxxxx, where xxxxxx begins the amino acids from the gene the user cloned into the vector. For vectors pDJSVT89 and pDJSVT90 which contain a TamA 1–27 tag, the residues will be N-ANVRLQHHHHHHxxxxxx. For vector pDJSVT91, which contains a SufI 1–30 tag, the residues will be N- AGQHHHHHHxxxxxx. For a graphical summary describing protein products, please refer to Fig. 1. All five vectors have been submitted to the Addgene nonprofit plasmid repository and can be obtained by requesting the ID number in Table 1.

2  Materials All solutions should be made using high-quality deionized water. Unless otherwise stated, store all solutions at room temperature. No sodium azide is added to any reagent. All E. coli should be properly handled by following BSL-1 specific guidelines.

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2.1  Instrumentation

Thermocycler, refrigerated centrifuge (capable of 15,000 RCF), refrigerated ultracentrifuge (capable of 100,000 RCF), swinging bucket centrifuge capable of spinning 12-well plates, cell sonifier or pressure cell (EmulsiFlex-C3, Avestin) for bacterial lysis, DNA and protein electrophoresis apparatus, Western blot transfer apparatus, glass column for protein purification, exponential decay electroporator, fast-protein liquid chromatography (FPLC) system (ӒKTA pure system, GE Healthcare, USA), fluorescent microscope with 63–100× objective (GFP and DAPI filters), 500 mL and 4 L baffled bottom flasks, stir plate, cold room or cold box for protein purification, stirred ultrafiltration cell, heating and cooling shaking incubator.

2.2  PCR and Cloning

1. 50× TAE buffer: 242 g Tris base, 100 mL of 500 mM EDTA (pH 8.0) solution, 57.1 mL glacial acetic acid in a final volume of 1 L. 2. Agarose gels: 1 g of agarose per 100 mL of 1× TAE buffer (1% gel) with 0.5 μg/mL ethidium bromide. 3. High-fidelity polymerase with proofreading activity to ensure accurate cloning of your target gene. 4. T4 DNA ligase (NEB). 5. Synthetic oligonucleotides for PCR. 6. EZ-10 Spin Column Plasmid DNA Miniprep Kit (BioBasic). 7. EZ-10 Spin Column PCR Products Purification Kit (BioBasic).

2.3  Growth Medium and Supplements

1. LB broth: 10 g tryptone (1% w/v), 5 g yeast extract (0.5% w/v), 10 g NaCl (1% w/v) per liter of media. Autoclave for sterility. Use baffled bottom shaking flasks at ≤25% occupancy to ensure proper aeration during growth. 2. LB agar: add 15 g/L Bacto agar to LB broth prior to autoclaving. LB agar plates should contain an appropriate antibiotic for transformant selection. Add the antibiotic once the agar has cooled to 50 °C prior to pouring plates. 3. Isopropyl β-d-1-thiogalactopyranoside (IPTG) for the induction of protein expression under T7 promoters. The final concentration of IPTG should be 25–250 μM in LB media. 4. ZYP-5052 autoinduction medium: 1% tryptone, 0.5% yeast extract, 50 mM Na2HPO4, 50 mM KH2PO4, 25 mM (NH4)2SO4, 0.5% glycerol, 0.05% glucose, 0.2% α-lactose, 2 mM MgSO4, 0.2× trace elements (please see the following reference: Studier, 2005, Protein Expression and Purification) [7]. 5. Carbenicillin (ampicillin) and chloramphenicol at 100 μg/mL and 25 μg/mL final concentrations, respectively. Store at −20 °C.

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1. DH5α or TOP10 E. coli for cloning and stable storage of plasmid backbones and clones. 2. BL21(DE3) and ArcticExpress (DE3) RIL E. coli for protein expression (Agilent Technologies). 3. Liquid nitrogen to freeze extra aliquots of electrocompetent cells. 4. 10% glycerol (sterile filtered). 5. SOC broth medium: 20 g/L tryptone, 5 g/L yeast extract, 0.5 g/L NaCl. Adjust to pH 7.0 by addition of 5 N NaOH, sterilize by autoclaving, and then add 20 mL of sterile 1 M glucose (final concentration of 20 mM). 6. Optional: Competent cell preparation using a Mix & Go! E. coli Transformation Kit (Zymo Research). 7. 1 mM gap electroporation cuvettes.

2.5  Secreted Protein Purification

1. 1 M Tris pH 7.5. 2. 4 M NaCl. 3. Stirred ultrafiltration cell with pump for pressure-based sample concentration. 4. Ultracel® 10 kDa ultrafiltration discs. 5. Chelating sepharose fast flow resin charged with nickel chloride for purification of 6His-tagged proteins. 6. Econo-Column® Chromatography Columns, 2.5 × 10 cm. 7. Ni-sepharose wash buffer: 20 mM Tris pH 7.5, 50 mM imidazole, 400 mM NaCl. 8. Ni-sepharose elution buffer: 20 mM Tris pH 7.5, 250 mM imidazole, 50 mM NaCl. 9. Optional ion exchange chromatography columns for additional protein purification. 10. Size-exclusion column: HiPrep 16/60 Sephacryl S-200 HR. 11. Size-exclusion chromatography buffer: 20 mM Tris pH 7.5, 150 mM NaCl, 10% glycerol. 12. Amicon Ultra centrifugal filter units.

2.6  Membrane Protein Purification

1. Sarkosyl (additional detergents can be screened to optimize purification). 2. E. coli lysis and membrane fraction dilution buffer: 20 mM Tris pH 7.5, 20 mM imidazole, 400 mM NaCl, 1 mM PMSF. 3. Membrane pellet resuspension buffer: 20 mM Tris pH 7.5, 2% sarkosyl. 4. Chelating sepharose fast flow resin charged with nickel chloride for purification of 6His-tagged proteins.

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5. Econo-Column® Chromatography Columns, 2.5 × 10 cm. 6. Ni-sepharose membrane wash buffer: 20 mM Tris pH 7.5, 50 mM imidazole, 400 mM NaCl, 0.2% sarkosyl. 7. Ni-sepharose membrane elution buffer: 20 mM Tris pH 7.5, 250 mM imidazole, 50 mM NaCl, 0.2% sarkosyl. 8. Size-exclusion column: HiPrep 16/60 Sephacryl S-200 HR. 9. Size-exclusion chromatography buffer: 20 mM Tris pH 7.5, 150 mM NaCl, 10% glycerol, 0.2% sarkosyl. 10. Amicon Ultra centrifugal filter units. 2.7  Immunoblotting

1. Precast 4–20% acrylamide precast gels for SDS-PAGE analysis of protein expression and purification. 2. Color Prestained (11–245 kDa).

Protein

Standard,

Broad

Range

3. Polyvinylidene difluoride (PVDF) (preferred) or nitrocellulose membranes. 4. Western blot transfer buffer (Towbin buffer): 25 mM Tris, 192 mM glycine, pH 8.3, 20% methanol (vol/vol). 5. Tris-buffered saline (10× TBS): 1.5 M NaCl, 0.1 M Tris–HCl, pH 7.4. 6. TBST: 1× Tris-buffered saline with 0.1% Tween-20. 7. Methanol. 8. Blocking and diluent solution: TBST with 3% BSA. 9. THE™ His Tag Antibody, mAb, mouse. 10. Anti-mouse m-IgGκ BP-HRP secondary antibody. 11. ECL Western blotting substrate. 12. Lucent Blue X-ray film. Optional: Digital gel and blot imaging system to replace exposure on film. 2.8  Immunofluores­ cence Microscopy

1. 10 mm round #1.5 thickness coverslip glass. 2. Standard glass microscopy slides. 3. 0.1 mg/mL poly-l-lysine. 4. PBS with 0.2% gelatin pH 7.5. 5. PBS with 3.2% paraformaldehyde. 6. THE™ His Tag Antibody, mAb, mouse. 7. Goat anti-Rabbit IgG (H + L) secondary antibody, Alexa Fluor 488. 8. ProLong Gold Antifade mountant. 9. 1000× (100 μg/mL) DAPI (4′,6-diamidino-2-phenylindole, dihydrochloride) 10. Nail polish to seal coverslip to the slide.

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3  Methods The methods outlined describe the use of the described suite of vectors to clone target genes for expression (see Note 2), purification, characterization, and development of gain-of-function E. coli strains. For simplicity, all of the results shown in Fig. 2 have used

Fig. 2 Expression of recombinant, extracytoplasmic proteins in E. coli using a suite of vectors described herein. (a) Overview of protein export and the potential sub- and extracellular location of proteins. (b) Description of vectors and the secretion systems used by each. (c) Detection by 6His Western blot and purification of two secreted proteins directly from the media of an overnight autoinduction growth. (d) Immunoblot detection and purification of the Fusobacterium nucleatum outer membrane protein FplA and detection of its expression on the surface of E. coli as detected by immunofluorescence using the same FplA antibody used in the immunoblot

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plasmid pDJSVT86, [8] which contains an OmpA signal sequence, ampicillin resistance, T7 promoter, and the lac operator DNA sequence for IPTG induction. 3.1  Target Gene Cloning

1. Standard molecular cloning methods for cohesive-end ligations should be used for inserting your gene of interest into the multiple cloning site (MCS), which contain the restriction enzyme sites NotI, KpnI, XhoI, and BamHI (Plasmids pDJSVT86, pDJSVT89, pDJSVT91) and NotI, KpnI, XhoI (pDJSVT87, pDJSVT90) (Fig. 1) (see Notes 3 and 4). In detail, you will generally include either a NotI or KpnI site in the design of your 5′ primer and either a KpnI, XhoI, or BamHI site in the 3′ reverse primer when amplifying your gene of interest by PCR (see Note 5). For example, to clone your gene of interest using NotI and XhoI restriction sites, design your primers as follows: Forward primer (NotI): 5′ GACTACGCGGCCGCg XXXXXXXXXXXXX 3′. Underlined: Extra bases added at the end of the primer to increase restriction enzyme cleavage efficiency. They don’t change the construct at all and will be cleaved by the enzyme and not incorporated into the plasmid. Bold: NotI restriction site. You need to add a single guanine nucleotide (g) after this site to keep your protein in frame for translation (see Fig. 1d). XXXXXXX: The sequence of your gene of interest. This should start with the first amino acid that comes after the predicted Sec or Tat signal sequence (see Note 1). We recommend making this sequence long enough to have a melting temperature of 50–60 °C. Do not include the restriction site or extra 5′ bases in the melting temperature determination when designing the primer. Reverse primer (XhoI): ** This primer should be ordered 5′ to 3′, but we will help you first design it in the 3′ to 5′ orientation for simplicity: 3′ XXXXXXXXXXXXXXX ­ TAGCTCGAGATCTAG 5′. XXXXXXX: The sequence of your gene of interest. This should be the final amino acids in the protein (or final residues you want to clone). We recommend making this sequence long enough to have a melting temperature of 50–60 °C. Do not include the restriction site or extra 5′ bases in the melting temperature determination when designing the primer. Italics: Include a stop codon after your final amino acid encoding nucleotides. Bold: XhoI restriction site. Underlined: Extra bases added at the end of the primer to increase restriction enzyme cleavage efficiency. They don’t change the construct at all and will be cleaved by the enzyme and not incorporated into the plasmid.

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For ordering from your supplier of choice, you need to reverse and complement the sequence, and the primer above would result in the following sequence: 5′ CTAGATCTCGAGCTAXXXXXXXXXXXXXXX 3′. 2. After polymerase chain reaction (PCR) with the designed primers and template DNA of your target gene, analyze 5 μL of the 50 μL reactions on a 1% agarose gel, and visualize with a UV light source if using ethidium bromide in the gel. If the desired products were obtained, purify the remaining 45 μL of PCR product using a DNA spin column specific for PCR product purification. 3. Both the purified base vector and PCR product of your gene of interest should be digested with the proper restriction enzymes for 1–16 h, and we recommend dephosphorylating the vector with Antarctic phosphatase as outlined in the manufacturer’s recommendations. Post digestion and dephosphorylation, vector and insert should be purified using a Plasmid DNA Miniprep Kit. Purified DNA can be quantified by either analysis on an agarose gel, via NanoDrop, or Qubit. 4. DNA is now ready for ligation with T4 DNA ligase and subsequent transformation into competent DH5α or TOP10 E. coli. These strains can be made electrocompetent by a simple and robust protocol [9]. In brief, this protocol grows E. coli in SOC media and uses ice cold 10% glycerol to produce competent cells. We have found an alternate method by using the Mix & Go! E. coli Transformation Kit. This has the advantage of not having to desalt ligations before transformation, as you need to do with electrocompetent cells. 5. After successful cloning of your construct and verification by restriction digest with enzymes used to insert the gene of interest, we recommend using Sanger sequencing to verify a correct insert with a proper in frame DNA sequence. This can be achieved by using T7 promoter and T7 terminator primers for plasmids pDJSVT86, 89, and 91 and ACYCDuetUP1 and T7 terminator for plasmids pDJSVT87 and 90. 3.2  Protein Expression

1. We have successfully used LB with IPTG, or ZYP-5052 autoinduction media for robust expression and purification of secreted and membrane proteins. E. coli have been used with success (BL21 (DE3)) for proteins secreted into the media. In addition, we found that outer membrane proteins expressed and purified well from ArcticExpress (DE3) RIL E. coli. 2. While it has not been tested, combining the vectors described herein with recently reported E. coli strains designed specifically for robust production of outer membrane proteins could increase target protein yield [10].

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3.3  Expression and Purification of Secreted Proteins

1. Transform DNA from positively cloned constructs into an E. coli expression strain of your choice; we recommend either BL21 (DE3) or ArcticExpress (DE3) RIL E. coli for protein expression. 2. A single colony was picked and transferred to a sterile tube containing 5 mL of LB media supplemented with carbenicillin (pDJSVT86, 89, 91) or chloramphenicol (pDJSVT87, 90) at 100 or 25 μg/mL, respectively, and grown overnight at 37 °C with 250 RPM shaking. 3. Overnight growths were used to inoculate 100 mL of autoinduction media in a 500 mL baffled bottom flask (1:100 inoculation) and then grown overnight (>16 h) at 37 °C with 250 RPM shaking. If the yield of your protein is low, you can scale up to 1 L of media in a 4 L baffled bottom flask and follow the same protocol. 4. If autoinduction of protein expression at 37 °C does not produce enough protein, we recommend using ArcticExpress (DE3) RIL E. coli for protein expression at 8 °C overnight with 25 μM IPTG added at an OD600 = 0.6 before lowering the temperature for >16 h. 5. The following morning, E. coli are harvested by centrifugation at 5 k RCF for 15 min to pellet cells. 6. Media was then poured off and sterile filtered using a 0.3 μm filter and a vacuum flask. 7. The 100 mL of collected media was adjusted to 20 mM Tris pH 7.5 and 400 mM NaCl before purification over Ni-­ sepharose beads. Optionally, if larger cultures are required to yield the desired amount of protein, the pH and salt-adjusted media can then be concentrated using a stirred ultrafiltration cell and the desired molecular weight cutoff filters (e.g., 10 kDa) to facilitate an easier purification. 8. Media is next incubated with 5 mL of packed Ni-sepharose beads while stirring at 4 °C for 1 h in a glass beaker. 9. Media and Ni-sepharose beads are then poured into a 2.5 × 10 cm glass purification column with an adjustable flow stop cock. Allow the beads to pack into a uniform resin bed, and then slowly pass the media containing your 6His-tagged protein back through the resin bed to ensure efficient protein capture. 10. Wash the beads with 100–300 mL of Ni-sepharose wash buffer, followed by elution of your target protein in 10–20 mL of Ni-sepharose elution buffer. 11. Purified protein can now be further purified using ion exchange or size-exclusion chromatography on a fast-protein liquid chromatography (FPLC) system. In addition, the initial

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affinity purification using Ni-sepharose resin can be run on an FPLC. 12. Post purification, check for yield and purity by SDS-PAGE and a bicinchoninic acid assay (BCA) or Qubit instrument. 13. To ensure the protein is your target and contains a 6His tag, a Western blot can be run using a 6His primary antibody as described below. 14. Protein can be further concentrated for downstream experiments using Amicon spin concentrators. 3.4  Expression and Purification of Outer Membrane Proteins (See Note 6)

1. Transform DNA from positively cloned constructs into an E. coli expression strain of your choice; we recommend either BL21 (DE3) or ArcticExpress (DE3) RIL E. coli. 2. A single colony was picked and transferred to a sterile tube containing 5 mL of LB media supplemented with carbenicillin (pDJSVT86, 89, 91) or chloramphenicol (pDJSVT87, 90) at 100 or 25 μg/mL, respectively, and grown overnight at 37 °C with 250 RPM shaking. 3. Overnight growths were used to inoculate 100 mL of autoinduction media in a 500 mL baffled bottom flask (1:100 inoculation) and then grown overnight (>16 h) at 37 °C with 250 RPM shaking. If the yield of your protein is low, you can scale up to 1 L of media in a 4 L baffled bottom flask and follow the same protocol. 4. If autoinduction of protein expression at 37 °C does not produce enough protein, we recommend using ArcticExpress (DE3) RIL E. coli for protein expression at 8 °C overnight with 25 μM IPTG added at an OD600 = 0.6 before lowering the temperature for >16 h. 5. The following morning, E. coli are harvested by centrifugation at 5 k RCF for 15 min to pellet cells. We recommend using these pellets immediately instead of freezing as your protein of interest could be on the surface and may be sensitive to freeze thaw cycles. 6. Resuspend the pellet in 10 mL of E. coli lysis buffer per 1 g of cell pellet, and lyse on a French press or pressure cell for 5–10 passes at maximum pressure of 20,000 psi. Alternatively, a sonifier could be used but is more harsh on proteins. 7. After lysis, centrifuge the lysate at 4 °C and 12,000 RCF for 15 min. 8. Remove the supernatant and transfer to ultracentrifuge tubes for centrifugation 4 °C and 100,000 RCF for 1 h. This will produce a pellet of total membranes. 9. The pellet of total membranes should be resuspended in 10 mL of membrane pellet resuspension buffer by pipetting to resuspend

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pellets in all tubes and recombining into a small beaker and stirring at 4 °C for 1 h to solubilize the membranes. 10. At this point, most protocols would have you recentrifuge this mixture to pellet the outer membrane proteins, but we have found that our outer membrane protein with soluble extracellular domains partially fractionate into the sarkosyl-containing buffer, a trait usually reserved for inner membrane proteins. Therefore, we then dilute the 10 mL of total membrane containing buffer with 90 mL of membrane fraction dilution buffer (the same at the initial lysis buffer), to bring the final concentration of sarkosyl to 0.2%. 11. Protein in membrane purification buffer is next incubated with 5 mL of packed Ni-sepharose beads while stirring at 4 °C for 1 h in a glass beaker. 12. Media and Ni-sepharose beads are then poured into a 2.5 × 10 cm glass purification column with an adjustable flow stop cock. Allow the beads to pack into a uniform resin bed, and then slowly pass the media containing your 6His-tagged protein back through the resin bed to ensure efficient protein capture. 13. Wash the beads with 100–300 mL of Ni-sepharose membrane wash buffer, followed by elution of your target protein in 10–20 mL of Ni-sepharose membrane elution buffer. 14. Purified protein can now be further purified using ion exchange or size-exclusion chromatography on a fast-protein liquid chromatography (FPLC) system. In addition, the initial affinity purification using Ni-sepharose resin can be run on an FPLC. 15. Post purification, check for yield and purity by SDS-PAGE and a bicinchoninic acid assay (BCA) or Qubit instrument. 16. To ensure the protein is your target and contains a 6His tag, a Western blot can be run using a 6His primary antibody as described below. 17. To further concentrate protein for downstream experiments, we recommend using Amicon Ultra centrifugal filter units. 3.5  Immunoblotting (See Note 7)

1. Separate the desired protein purification fractions or purified proteins by SDS-PAGE using a precast 4–20% gel. Load 5 μL of Color Prestained Protein Standard, Broad Range (11–245 kDa), to the gel to ensure that you can identify the molecular weight of your target protein by Western blot. 2. Transfer proteins to a methanol-activated polyvinylidene difluoride (PVDF) membrane using Western blot transfer buffer. 3. Post transfer, incubate the membrane in 10 mL of blocking buffer (TBST 3% BSA) in a plastic container (ensure the blot is

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submerged) for >2 h with gently rocking at room temperature or overnight at 4 °C. 4. Remove the blocking buffer, and add in 10 mL of a 1:1000 dilution of THE™ His Tag Antibody, mAb, mouse (Genscript) in blocking buffer. 5. Incubate for 1 h at room temperature with light rocking. 6. Discard of the primary antibody solution and wash extensively with TBST. 7. Add in 10 mL of a 1:5000 dilution of anti-mouse m-IgGκ BP-­ HRP secondary antibody (Santa Cruz) in blocking buffer. 8. Incubate for 1 h at room temperature with light rocking. 9. Discard the secondary antibody solution and wash extensively with TBST. 10. Remove the membrane from the TBST wash solution, and place on a flat surface covered in aluminum foil. 11. Add 3 mL of properly mixed ECL Western blotting substrate, and ensure that the entire blot is covered in liquid for ~3 min. 12. Carefully pick up the blot with tweezers, and transfer to a Western blot developing cassette. 13. Use either Lucent Blue X-ray film or a digital gel and blot imaging system to visualize 6His-tagged proteins on the gel. 3.6  Immunofluores­ cence Microscopy of Outer Membrane Proteins

We provide a protocol for visualizing 6His-tagged domains on the surface of E. coli using the commercially available THE™ His Tag Antibody, mAb, mouse (Genscript). 1. DNA from positively cloned constructs were transformed into either BL21 (DE3) or ArcticExpress (DE3) RIL E. coli for protein expression. 2. A single colony was picked and transferred to a sterile tube containing 5 mL of LB media supplemented with carbenicillin (pDJSVT86, 89, 91) or chloramphenicol (pDJSVT87, 90) at 100 or 25 μg/mL, respectively, and grown overnight at 37 °C with 250 RPM shaking. 3. Overnight growths were used to inoculate 100 mL of autoinduction media in a 500 mL baffled bottom flask (1:100 inoculation), and then grown overnight (≥16 h) at 37 °C with 250 RPM shaking. 4. If autoinduction of protein expression at 37 °C does not produce enough protein, we recommend using ArcticExpress (DE3) RIL E. coli for protein expression at 8 °C overnight with 25 μM IPTG added at an OD600 = 0.6 before lowering the temperature for >16 h. 5. The following morning, E. coli are back diluted to OD600 = 0.2, and 1 mL is centrifuged at 5000 RCF for 5 min.

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6. The pelleted bacteria are washed twice with 500 μL of PBS 0.2% gelatin pH 7.5 followed by centrifuging at 5000 RCF for 4 min. 7. Resuspend cells in 500 μL of PBS with 3.2% paraformaldehyde and fix for 20 min. 8. Concurrently, incubate round 10 mm #1.5 coverslips with 0.1 mg/mL poly-l-lysine for 20 min at room temperature. 9. After bacteria are fixed, transfer a single coverslip to a 12-well plate, and then place the 500 μL of bacteria into the well on top of the coverslip. 10. Add 2 mL of PBS to the well and swirl to make sure the coverslip is submerged. 11. Transfer the 12-well plate into a swinging bucket centrifuge, and pellet bacteria onto coverslips at 2000 RCF for 10 min. 12. Post-centrifugation, transfer coverslips to a fresh well in a 12-well plate, and wash the coverslip 2× with 2 mL of PBS 0.2% gelatin pH 7.5. 13. Prepare a 100 μL solution of 1:100 dilution of your antibody of choice (THE™ His Tag Antibody, mAb, mouse, Genscript) in PBS 0.2% gelatin pH 7.5. 14. Place 20 μL of the solution on a piece of parafilm, and invert the coverslip on the antibody solution for 1 h at room temperature. 15. To maintain moisture, place a small container over the coverslip with a wet sponge taped to the lid. 16. After incubation in primary antibody, transfer the coverslip bacteria up into a new well in a 12-well plate, and wash 2× with PBS 0.2% gelatin pH 7.5. 17. Add 1 mL of PBS 0.2% gelatin to each well, and add two drops from the dropper of Goat anti-Rabbit IgG (H + L) secondary antibody, Alexa Fluor 488. 18. Incubate at room temperature for 30 min, followed by two washes with 2 mL of PBS 0.2% gelatin pH 7.5 and two washes with 2 mL of PBS pH 7.5. 19. Add 4 μL of ProLong Gold Antifade mountant to a glass slide, and invert the cover slip on top. 20. Gently press down and remove excess liquid from the edges of a coverslip with a delicate task wipe. 21. Seal the coverslip to the slide with nail polish around the edges. 22. Store the slide in a dark case until you are ready to image. 23. Image the fluorescently green labeled bacteria using a fluorescent microscope and either a GFP or FITC filter under 63× or 100× magnification. Obtaining images using either brightfield or phase contrasts (better) will help to determine the percentage of cells that expressed your protein in the outer membrane.

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4  Notes 1. Web servers for signal peptide prediction: PREDTAT (http:// www.compgen.org/tools/PRED-TAT/submit), SignalP— We find the most accurate predictions use SignalP v 3.0 with the Hidden Markov model (HMM) option selected (http:// www.cbs.dtu.dk/services/SignalP-3.0/). 2. Plasmids presented in Fig. 1 are available through Addgene (www.addgene.org/Daniel_Slade/) as outlined in Table 1, and all sequences and details of plasmids and their development can be found on our Open Science Framework (https:// osf.io/n7tmj/) data repositories. 3. Note that these plasmids were created to maximize cloning efficiency for AT-rich genomes by utilizing the GC-rich restriction enzyme sites NotI (GCGGCCGC), KpnI (GGTACC), XhoI (CTCGAG) (see Subheading 3.1). 4. These vectors can also be used for expression of cytoplasmic proteins by cloning into the vector using the NcoI site located 5′ of the signal sequence DNA. The NotI, KpnI, or XhoI site can then be used as the 3′ restriction site. Note, this will also remove the 6His tag located in the vector. This will also partially revert the plasmids back to their parent vectors of pET-­16b (pDJSVT86, 89, 91) and pACYCDuet-1 (pDJSVT87, 90) (see Subheading 3.1). 5. We prefer to clone all of our genes through polymerase chain reaction (PCR), but users can also choose to synthesize genes if desired, which would allow for codon optimization in E. coli (see Subheading 3.1). 6. As with all membrane proteins, there are vast differences in optimal expression and purification conditions, and we recommend reading a comprehensive review by SM Smith [11] or other sources of your choice. For the following studies, we reference a paper that addressed the purification of membrane proteins in Campylobacter jejuni [12]. The researchers reported that purification of outer membrane proteins using N-­lauroylsarcosine (sarkosyl) produced the purest protein, and therefore we used a hybrid strategy for the purification of FplA, a Type 5d secreted outer membrane protein that contains a 50 kDa soluble portion extracellular to the membrane (Casasanta et al. [8]). This likely makes the protein act less like a true embedded outer membrane protein, which we address during the following methods (see Subheading 3.4). 7. To detect the production of secreted, periplasmic, or outer membrane proteins using the vectors described here, we recommend taking advantage of the N-terminal 6His tag that

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remains on the proteins after passing through the Sec or Tat secretion systems. In addition, you can also use antibodies specific for your protein of interest (see Subheading 3.5).

Acknowledgments Research was supported by start-up funding from the Department of Biochemistry at Virginia Tech, the Institute for Critical Technology and Applied Science (ICTAS) at Virginia Tech, and the National Institute for Food and Agriculture. References 1. Lee VT, Schneewind O (2001) Protein secretion and the pathogenesis of bacterial infections. Genes Dev 15:1725–1752 2. Green ER, Mecsas J (2016) Bacterial secretion systems: an overview. In: Kudva IT et al (eds) Virulence mechanisms of bacterial pathogens, 5th edn. American Society of Microbiology, pp 215–239 3. Dautin N, Bernstein HD (2007) Protein secretion in gram-negative bacteria via the autotransporter pathway. Annu Rev Microbiol 61:89–112 4. Natale P, Brüser T, Driessen AJM (2008) Secand tat-mediated protein secretion across the bacterial cytoplasmic membrane--distinct translocases and mechanisms. Biochim Biophys Acta 1778:1735–1756 5. van den Berg B (2010) Crystal structure of a full-length autotransporter. J Mol Biol 396: 627–633 6. Auclair SM, Bhanu MK, Kendall DA (2012) Signal peptidase I: cleaving the way to mature proteins. Protein Sci 21:13–25

7. Studier FW (2005) Protein production by auto-induction in high density shaking cultures. Protein Expr Purif 41:207–234 8. Casasanta MA, Yoo CC, Smith HB et al (2017) A chemical and biological toolbox for type Vd secretion: characterization of the phospholipase A1 autotransporter FplA from fusobacterium nucleatum. J Biol Chem. https://doi. org/10.1074/jbc.M117.819144 9. Woodall CA (2003) Electroporation of E coli. Methods Mol Biol 235:55–69 10. Meuskens I, Michalik M, Chauhan N et al (2017) A new strain collection for improved expression of outer membrane proteins. Front Cell Infect Microbiol 7(464) 11. Smith SM (2011) Strategies for the purification of membrane proteins. Methods Mol Biol 681:485–496 12. Hobb RI, Fields JA, Burns CM et al (2009) Evaluation of procedures for outer membrane isolation from campylobacter jejuni. Microbiology 155:979–988

Chapter 12 Mouse Model of Staphylococcus aureus Skin Infection Natalia Malachowa, Scott D. Kobayashi, Jamie Lovaglio, and Frank R. DeLeo Abstract Bacterial skin and soft tissue infections are abundant worldwide, and many are caused by Staphylococcus aureus. Indeed, S. aureus is the leading cause of skin and soft tissue infections in the USA. Here we describe a mouse model of skin and soft tissue infection induced by subcutaneous inoculation of S. aureus. This animal model can be used to investigate a number of factors related to the pathogenesis of skin and soft tissue infections, including strain virulence and the contribution of specific bacterial molecules to disease, and it can be employed to test the potential effectiveness of antibiotic therapies or vaccine candidates. Key words Skin infection, Abscess, Bacteria, Mouse, Staphylococcus aureus

1  Introduction Animal infection models are an integral part of host-pathogen research and are used to approximate processes that occur during human infections. Intact human skin is a unique and complex tissue that forms a physical barrier and serves as a frontline of defense against invading microorganisms. In addition to functioning as a mechanical barrier, skin plays an essential role in innate and adaptive immunity. For example, skin is comprised of numerous cell types including keratinocytes, Langerhans cells, and macrophages, each of which can release immunomodulatory cytokines and chemokines that amplify the inflammatory response during infection [1]. In turn, professional phagocytes such as neutrophils and monocytes are recruited from the bloodstream to infected tissues, where they ingest and kill invading microbes. Skin cells can also produce antimicrobial peptides, which provide an additional layer of defense against microbial pathogens [2]. Mechanically compromised skin and impaired skin function can lead to a wide range of dermatological diseases, such as atopic dermatitis and psoriasis, or skin cancers including melanomas. Nevertheless, the majority of Irving C. Allen (ed.), Mouse Models of Innate Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1960, https://doi.org/10.1007/978-1-4939-9167-9_12, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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skin diseases have an infectious etiology [3, 4]. Skin infections are caused by a taxonomic diversity of infectious microorganisms that include fungi, viruses [5], parasites [6–8], protozoa, and a variety of Gram-positive and Gram-negative bacterial species [9–11]. Staphylococcus aureus is one of the most prominent human pathogens and is a major cause of skin and soft tissue infections. In the USA, the community-associated methicillin-resistant S. aureus (CA-MRSA) strain USA300 is the predominant cause of skin and soft tissue infections [12–14]. The success of S. aureus as a human pathogen is dependent on several factors including the ability to adapt to environmental changes and produce a variety of molecules that contribute to virulence. Several mouse models of infection have been developed to increase our understanding of staphylococcal pathogenesis [15–20]. In this chapter, we describe a mouse model of S. aureus skin and soft tissue infection (SSTI) induced by subcutaneous inoculation of USA300. The methods described herein utilize commercially available reagents and can be reproduced by most standard laboratories.

2  Materials These materials are for use with S. aureus as the infectious agent and can be altered to fit the culture requirements for any bacterium of interest. 2.1  Bacterial Culture and Inoculum

1. Trypticase Soy Broth (TSB) and Trypticase Soy Agar (TSA). 2. Sterile Dulbecco’s phosphate-buffered saline (DPBS). 3. Sterile 3.5-mL clear screw cap septum vials. 4. S. aureus strain LAC (representative of the epidemic USA300 strain, NARSA # JE2, see Note 1).

2.2  Preparation of Mouse Skin for Infection

1. RC2 Rodent Circuit Controller isoflurane anesthesia machine and induction chamber (VetEquip, Livermore, CA). 2. Low profile anesthesia masks (Kent Scientific Corporation, Torrington, CT). 3. Isoflurane. 4. Ophthalmic ointment. 5. Crl:SKH1-E outbred, immunocompetent, hairless or Balb/c female mice 6–8 weeks old, 20–25 g (Charles River Laboratories International, Inc., Wilmington, MA) (see Note 2). Mice are group-housed and allowed food and water ad libitum. 6. Finisher® electric trimmer with narrow blade (Oster® Direct, McMinnville, TN). It is also possible to use a #50 clipper blade on standard electric clippers or a Wahl® Touch-Up & Stylique battery operated trimmer set.

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7. Nair® hair remover lotion (Church & Dwight Co. Inc., Princeton, NJ). 8. Non-woven, gauze sponges, 3″ × 3″. 2.3  Subcutaneous Inoculation of Bacteria and Analysis of Skin Lesions

1. OHAUS® Scout® SPX421 portable balance (OHAUS Corporation, Parsippany, NJ). 2. Rubber-tipped forceps (Mopec, Oak Park, MI). 3. 1-mL syringe with luer lock tip. 4. 25–27-gauge needles. 5. Digital calipers.

3  Methods 3.1  Preparation of Bacterial Inoculum

1. Inoculate S. aureus (from a frozen glycerol stock) into sterilized TSB media in a flask-to-media volume ratio of 5:1. 2. Grow the bacteria at 37 °C with rotary shaking for 16–18 h (late stationary-phase of growth, OD600~2.1). 3. Transfer bacteria from late stationary-phase culture to fresh TSB media (1:200 dilution), and incubate at 37 °C with rotary shaking until the optical density at 600 nm (OD600) of the culture reaches 0.75 (~2–2.5 h for USA300, which is mid-­ exponential phase of growth) (see Note 3). 4. Collect bacteria by centrifugation (3000 × g, 4 °C for 10 min). Bacteria are maintained at ~4 °C until ready for inoculation by incubation on ice (see Note 4). 5. Wash bacteria by suspending pellet in an equal volume of DPBS, and harvest by centrifugation at 3000 × g, 4 °C for 10 min. 6. Resuspend bacteria in sterile DPBS to attain a final concentration of 2 × 108 colony-forming units (CFU)/mL. 7. Transfer bacterial suspension to sterile septum vials, and keep on ice until injection (see Note 5). 8. Verify the concentration of the bacterial inoculum by plating a 10−6 dilution on TSA plates. 9. Incubate the plates for 24 h at 37 °C and enumerate CFUs. This is a retrospective verification.

3.2  Preparation of Mice for Infection

Hair removal from the inoculation site allows for enhanced visualization and accurate measurements of the abscess (see Note 6) and may occur the day prior to inoculation. All mouse procedures are performed under general anesthesia (see Note 7). Mice are maintained under constant anesthesia during experimental procedures by use of anesthesia masks. The procedures described in this

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chapter conform to the guidelines set forth by the National Institutes of Health (NIH) and were reviewed and approved by the Animal Care and Use Committee (ACUC) at Rocky Mountain Laboratories, National Institute of Allergy and Infectious Diseases, NIH (see Note 8). 1. Anesthetize mouse in an anesthesia induction chamber using the following recommended settings: 2–4% isoflurane and 1 L/min oxygen flow under standard atmospheric pressure (see Notes 9 and 10). 2. Remove mouse from anesthesia chamber, and use an anesthesia mask to connect to isoflurane. The recommended settings for isoflurane administration while using the anesthesia mask are 1–3% isoflurane and 0.5 L/min oxygen flow. 3. Place ophthalmic ointment in both eyes to protect the corneas. 4. Carefully shave the inoculation site, typically the right and/or left flank, using an electric clipper (Fig. 1a, b). 5. Cover the shaved area with Nair® for 3 min (see manufacturer instructions on the package for details) (see Note 11) (Fig. 1c). 6. Remove Nair® gently, yet thoroughly using non-woven gauze sponges and warm water (Fig. 1d–f) (see Note 12). 7. At this point, mice should be identified with tattoos, microchips, or ear punches to allow tracking and identification of individual animals. 8. Remove the mouse from isoflurane anesthesia, allow to recover in a warm environment and then return to the home cage. 3.3  Subcutaneous Inoculation

1. Weigh each mouse prior to inoculation (see Note 13). 2. Anesthetize mouse in an anesthesia induction chamber using the following recommended settings: 2–4% isoflurane and 1 L/min oxygen flow under standard atmospheric pressure (see Note 9). 3. Remove mouse from anesthesia chamber, and use an anesthesia mask to connect to isoflurane. The recommended settings for isoflurane administration while using the anesthesia mask are 1–3% isoflurane and 0.5 L/min oxygen flow. 4. Place ophthalmic ointment in both eyes to protect the corneas. 5. Disinfect the skin over the intended inoculation site with 70% ethanol. 6. Create a “tent” in the skin of the prepared inoculation site utilizing rubber-tipped forceps, and insert the needle into the skin at the “base” of the tent (Fig. 2a) (see Note 14).

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Fig. 1 Preparation of mice for infection. (a, b) Shaving a mouse with an electric trimmer. (c) Nair® is applied to remove remaining hair and delay regrowth. (d–f) Removing Nair® with gauze sponge and warm water. (g, h) Syringe loaded prior to expelling air bubbles (g) and after air bubbles were removed (h)

7. Inoculate the animal subcutaneously with 0.05 mL of 1 × 107 live S. aureus or sterile saline (Fig. 2b) (see Note 15). 8. Remove the mouse from isoflurane anesthesia, allow to recover in a warm environment, and then return to the home cage.

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Fig. 2 Subcutaneous inoculation and subsequent analysis of skin lesions. (a, b) Subcutaneous injection. Panel (a) demonstrates formation of the “tent” when pinching the skin. (c, d) Taking measurements of the abscess area with calipers. (d) The “green” arrow represents the length that is measured in the “head-to-tail” direction of the mouse and “red” is a width of the abscess or dermonecrosis area (dorsal–ventral). (e) Example of the abscess and dermonecrosis area on day 4 post-injection (Crl:SKH1-E hairless mice). (f) A selected area from picture (e) is enlarged to improve visualization of lesion sites 3.4  Monitoring the Course of Infection

1. The progression of disease, in this case abscess development, can be monitored by daily measurement of lesion dimensions (see Note 16). 2. Measure the abscess length (L) and width (W) with the calipers (Fig. 2c, d). The abscess length and width dimensions are used to calculate the abscess volume [V = 4/3π (L/2)2 × W/2] and area [A = π(L/2) × W/2] [19, 21]. 3. Euthanize the mouse when established endpoint criteria have been reached or at the end of the experiment (e.g., isoflurane overdose or CO2 asphyxiation followed by cervical dislocation) (see Note 17). 4. Euthanized mice are bagged, labeled, and incinerated. Cages and bedding are autoclaved before washing.

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4  Notes 1. The mouse SSTI infection model described in this chapter was developed for assessment of S. aureus virulence. However, this model can be adapted to test SSTI of other S. aureus strains and/or bacterial species. 2. Mouse strains are diverse and can vary by factors such as immunological features. Therefore, selection of the appropriate mouse strain should be considered carefully. For example, we frequently use Crl:SKH1-E hairless mice for the SSTI model, as animal preparation time is reduced (shaving and application of Nair® are omitted) and skin lesions are easily measured. 3. The bacterial growth parameters used to generate the inoculum can be varied to suit the experimental hypothesis. In addition, frozen bacteria can be used to standardize the inoculum between different experiments. Changes in the inoculum may affect experimental outcome and should be vetted in pilot studies. 4. The total volume of bacteria for the injections should be calculated based on number of animals used per group. In our studies, we typically use 15 animals per test group and five animals for DPBS control based on guidance from statistical power analysis. See also Note 5 for additional information. 5. For ease of loading syringes with the inoculum, we typically transfer three times the amount of bacterial inoculum that is required for all injections per test or control group to the septum vials. Excess inoculum is discarded following decontamination by autoclaving. Prolonged storage of the inoculum in DPBS on ice may reduce bacterial viability. 6. This step is unnecessary when using Crl:SKH1-E hairless mice. 7. All personnel entering the animal and procedure rooms must wear basic personal protective equipment, including laboratory coat, gloves, and facemask. Eye protection must be worn when working with potentially infectious and/or hazardous materials. 8. Animal care and use policies may vary by individual institution. Institutional policies and procedures regarding animal experimentation should be consulted prior to commencement of experimental design. 9. We recommend pre-oxygenating mice in the chamber for 3 min prior to turning on the isoflurane. 10. Depending on the experimental design, laboratory settings, or institutional requirements, other inhalant or injectable anesthetics such as sevoflurane, ketamine, or a combination of ketamine and xylazine may be substituted [22].

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11. Nair® is applied to remove remaining hairs and prevent their regrowth for more accurate measurement of experimental lesions. 12. Either prolonged treatment or incomplete removal of Nair® will cause pronounced skin irritation and should be avoided. 13. A vessel to contain the mouse is useful to facilitate manipulation of animals and to accurately assess weight. 14. The SSTI model can be modified to employ other modes of inoculation such as intradermal injection. 15. The optimal number of CFUs needed to achieve a reproducible SSTI should be determined in pilot experiments [18]. 16. Typically, the subcutaneous injection of USA300 results in formation of a measurable abscess or area of dermonecrosis starting on days 2–3 with a maximum size achieved on ~day 6, followed by resolution of infection toward day 14 (Fig. 2e, f). Of note, the formation of dermonecrotic lesions is S. aureus strain and/or virulence factor specific [18, 20]. 17. In addition to scheduled euthanasia time points, specific endpoint criteria should be established for each study to allow for early euthanasia of study animals. Endpoint criteria will vary based on experimental requirements and ACUC approval but may include size of the abscess, abscess rupture, abscess interfering with normal ambulation, hunched posture, reluctance to move, poor body condition score [23], weight loss, etc.

Acknowledgments The authors are supported by the Intramural Research Program of the National Institute of Allergy and Infectious Diseases, National Institutes of Health. The authors thank Anita Mora and Austin Athman (Visual & Medical Arts, RML/NIAID) for photography. References 1. Kupper TS, Fuhlbrigge RC (2004) Immune surveillance in the skin: mechanisms and clinical consequences. Nat Rev Immunol 4:211–222 2. Nizet V, Ohtake T, Lauth X et al (2001) Innate antimicrobial peptide protects the skin from invasive bacterial infection. Nature 414:454–457 3. Lowell BA, Froelich CW, Federman DG et al (2001) Dermatology in primary care: prevalence and patient disposition. J Am Acad Dermatol 45:250–255 4. Sari F, Brian B, Brian M (2005) Skin disease in a primary care practice. Skinmed 4:350–353

5. Weinberg JM, Mysliwiec A, Turiansky GW et al (1997) Viral folliculitis: atypical presentations of herpes simplex, herpes zoster, and molluscum contagiosum. Arch Dermatol 133:983–986 6. Mika A, Goh P, Holt DC et al (2011) Scabies mite peritrophins are potential targets of human host innate immunity. PLoS Negl Trop Dis 5:e1331 7. Hengge UR, Currie BJ, Jäger G et al (2006) Scabies: a ubiquitous neglected skin disease. Lancet Infect Dis 6:769–779 8. Feldmeier H (2012) Pediculosis capitis: new insights into epidemiology, diagnosis and

Mouse Model of Staphylococcus aureus Skin Infection treatment. Eur J Clin Microbiol Infect Dis 31:2105–2110 9. Saracino A, Kelly R, Liew D et al (2011) Pyoderma gangrenosum requiring inpatient management: a report of 26 cases with follow up. Australas J Dermatol 52:218–221 10. Shim TN, Lew TT, Preston PW (2012) Disseminated cutaneous Mycobacterium chelonae. Lancet Infect Dis 12:254 11. Pallin DJ, Espinola JA, Leung DY et al (2009) Epidemiology of dermatitis and skin infections in United States physicians’ offices, 1993–2005. Clin Infect Dis 49:901–907 12. Talan DA, Krishnadasan A, Gorwitz RJ et al (2011) Comparison of Staphylococcus aureus from skin and soft-tissue infections in US emergency department patients, 2004 and 2008. Clin Infect Dis 53:144–149 13. Tickler IA, Goering RV, Mediavilla JR et al (2017) Continued expansion of USA300-like methicillin-resistant Staphylococcus aureus (MRSA) among hospitalized patients in the United States. Diagn Microbiol Infect Dis 88:342–347 14. Carrel M, Perencevich EN, David MZ (2015) USA300 methicillin-resistant Staphylococcus aureus, United States, 2000–2013. Emerg Infect Dis 21:1973–1980 15. Watts A, Ke D, Wang Q et al (2005) Staphylococcus aureus strains that express serotype 5 or serotype 8 capsular polysaccharides differ in virulence. Infect Immun 73: 3502–3511 16. Hoebe K, Georgel P, Rutschmann S et al (2005) CD36 is a sensor of diacylglycerides. Nature 433:523–527

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17. Hume EB, Cole N, Khan S et al (2005) A Staphylococcus aureus mouse keratitis topical infection model: cytokine balance in different strains of mice. Immunol Cell Biol 83:294–300 18. Voyich JM, Otto M, Mathema B et al (2006) Is Panton-Valentine leukocidin the major virulence determinant in community-associated methicillin-resistant Staphylococcus aureus disease? J Infect Dis 194:1761–1770 19. Bunce C, Wheeler L, Reed G et al (1992) Murine model of cutaneous infection with gram-positive cocci. Infect Immun 60: 2636–2640 20. Kennedy AD, Wardenburg JB, Gardner DJ et al (2010) Targeting of alpha-hemolysin by active or passive immunization decreases severity of USA300 skin infection in a mouse model. J Infect Dis 202:1050–1058 21. Lukomski S, Montgomery CA, Rurangirwa J et al (1999) Extracellular cysteine protease produced by Streptococcus pyogenes participates in the pathogenesis of invasive skin infection and dissemination in mice. Infect Immun 67:1779–1788 22. Gaertner DJ, Hallman TM, Hankenson FC et al (2008) Anesthesia and analgesia for laboratory rodents. In: Richard EF, Marilyn JB, Peggy JD et al (eds) Anesthesia and analgesia in laboratory animals, 2nd edn. Academic Press, San Diego, pp 239–297 23. Ullman-Culleré MH, Foltz CJ (1999) Body condition scoring: a rapid and accurate method for assessing health status in mice. Lab Anim Sci 49:319–323

Chapter 13 Systemic Listeria monocytogenes Infection as a Model to Study T Helper Cell Immune Responses Veronica M. Ringel-Scaia, Michael D. Powell, Kaitlin A. Read, Irving C. Allen, and Kenneth J. Oestreich Abstract Listeria monocytogenes, a Gram-positive facultative intracellular pathogen, has been widely used as a model for studying the immune response. Here, we describe a protocol for the systemic infection of mice with L. monocytogenes, followed by isolation of lymphocytes from spleens and lymph nodes. We also include details on how to culture and store L. monocytogenes, as well as the specifics for fluorescence-activated cell sorting (FACS) for CD4+ cells in response to the systemic infection. This protocol can be adapted by changing the dosage of L. monocytogenes for a more or less aggressive infection and/or sorting for other immune cell subtypes of interest. Key words Listeria, Mouse, Lymphocytes, Flow cytometry, Fluorescence-activated cell sorting

1  Introduction Listeria monocytogenes is a Gram-positive facultative intracellular pathogen. An opportunistic food-borne pathogen, L. monocytogenes is found ubiquitously in soil, water, and decaying vegetation [1, 2]. Neonates, the elderly, immunocompromised individuals, and pregnant women are at particularly high risk for infection with L. monocytogenes. Most infections arise due to the consumption of contaminated food. The bacteria’s aptitude for growth in low temperature, high salt, and/or acidic conditions (settings often used to preserve foodstuff) and the absence of pasteurization for raw food and/or minimal processing of goods such as soft cheeses pose significant challenges regarding the regulation of Listeria contamination in the food industry [2]. L. monocytogenes infection results in gastroenteritis in healthy individuals, meningitis and septicemia in immunocompromised individuals, and chorioamnionitis in pregnant women [3, 4]. A favorable outcome following infection requires concerted innate and adaptive immune responses (for a recent review detailing the infection and pathogenesis of L. monocytogenes, see ref. 5). Irving C. Allen (ed.), Mouse Models of Innate Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1960, https://doi.org/10.1007/978-1-4939-9167-9_13, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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L. monocytogenes has been widely used experimentally to characterize host immune responses [1, 6, 7]. As such, L. monocytogenes infection can be used to assess the susceptibility of different mouse strains (e.g., knockout mice) to intracellular pathogens and to elucidate specific genetic components that affect host response to infection [7–13]. Animals with deficient immune responses to L. monocytogenes generally exhibit higher bacterial loads and/or delayed bacterial clearance relative to resistant mouse strains [6, 14]. Additionally, L. monocytogenes clearance and protection rely on a robust T cell response [1, 7, 8, 10]. The predictable T cell response to infection has long been used to the advantage of investigators to study the genesis and function of individual T cell populations during the course of an immune response. The methods described herein detail a straightforward technique to infect mice with a known concentration of L. monocytogenes in order to analyze CD4+ T cell populations generated in response to infection. By employing a sublethal dose of L. monocytogenes, this method limits the experimental variability and the pain or discomfort of the animals while still initiating a robust response, all with a relatively quick turnaround time. This protocol consists of six general methods: (1) culturing L. monocytogenes, (2) preparing bacteria inoculum for infection, (3) infecting mice intravenously, (4) preparation of a single-cell suspension, (5) analyzing CD4+ T cell populations generated in response to infection via flow cytometry, and (6) measuring bacterial load to confirm dosage. We have described methods for the analysis of CD4+ T cell populations from L. monocytogenes-infected mice, but this can be readily adapted to assess different immune cell subsets of interest.

2  Materials 2.1  Listeria monocytogenes

1. Culture Listeria monocytogenes from a frozen glycerol stock, agar stab, or freeze-dried pellet (see Note 1). The methods detailed here can be used with any strain of L. monocytogenes, independent of virulence (see Note 2). Care should be taken to ensure suitable bacteriological practice. Always begin with a single colony of bacteria to maintain clonality.

2.2  Media for Growing L. monocytogenes

1. Brain Heart Infusion (BHI) broth: 37 g Bacto Brain Heart Infusion in 1 L of purified water.

2.3  Solutions

1. Sterile ultrapure water.

2. BHI agar plates: 52 g BBL Brain Heart Infusion Agar in 1 L of purified water.

2. Sterile 1× DPBS without calcium chloride or magnesium chloride.

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3. Sterile red blood cell lysis buffer (RBC lysis buffer): 0.84% NH4Cl in sterile ultrapure water. 4. Complete IMDM media (cIMDM): 500 mL IMDM, 10% FBS, 1% Pen/Strep, 50 μM β-mercaptoethanol. 5. Trypan blue. 6. Sterile FACS buffer: 1× DPBS without calcium chloride or magnesium chloride, 2% FBS, 1% BSA. 7. Reagents for sample preparation and flow cytometry analysis required for T helper cell populations of interest (see Notes 12–23 associated with Subheading 3.5 for information regarding selection of reagents appropriate for individual applications). 2.4  Consumables

1. Sterile inoculating loops. 2. Sterile glass beads for spreading bacteria on agar plates. 3. Cuvettes for the spectrophotometer. 4. Sterile 1.5 mL microcentrifuge tubes. 5. Cryovials. 6. 1 mL syringes. 7. 27-gauge, 0.5-in.-long needles. 8. Clean WyPalls. 9. 60 mm petri dishes. 10. Sterile 50 mL conical tubes. 11. Sterile 15 mL conical tubes. 12. Sterile 5 mL, 10 mL, and 25 mL serological pipettes. 13. 24-well tissue culture plates. 14. Sterile frosted-end microscope slides. 15. Sterile petri dishes. 16. Sterile round-bottom flow cytometry tubes.

2.5  Equipment

1. Orbital shaker incubator. 2. Bacterial incubator. 3. Spectrophotometer. 4. Centrifuge with high-speed rotor. 5. Handheld homogenizer. 6. Dissection tools: blunt forceps, curved forceps, blunt scissors, sharp scissors. 7. Centrifuge with swing-bucket rotor for 15 mL conical tubes. 8. Vortex. 9. Hemacytometer or cell counter.

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10. Refrigerated bench-top centrifuge with rotor for 1.5 mL microcentrifuge tubes. 11. Flow cytometer (see Notes 12–23 associated with Subheading 3.5 for information regarding cytometer compatibility with chosen staining/analysis strategies).

3  Methods 3.1  Culturing L. monocytogenes

1. Obtain viable L. monocytogenes from a frozen glycerol stock, agar stab, or freeze-dried pellet. Using a sterile disposable or metal heat-sterilized inoculating loop, streak L. monocytogenes onto a pre-dried BHI agar plate in a method to obtain individual colonies. Incubate at 37 °C in a bacterial incubator overnight. 2. Pick a single colony from the overnight culture using a sterile inoculating loop, and place into 5 mL BHI broth in a 15 mL conical tube. Be sure not to screw the cap on completely. 3. Incubate the L. monocytogenes culture in an orbital shaker at 200–250 rpm at 37 °C overnight. (For long-term storage, see Note 3).

3.2  Preparing L. monocytogenes Infectious Dose for In Vivo Infection on Mice

1. Using a sterile disposable or metal heat-sterilized inoculating loop, streak L. monocytogenes onto a pre-dried BHI agar plate using any streaking technique that will obtain individual colonies. Incubate at 37 °C in a bacterial incubator overnight. 2. Pick a single colony from the overnight culture using a sterile inoculating loop, and place into 5 mL BHI broth in a 15 mL conical tube. Be sure not to screw the cap on completely. 3. Incubate the L. monocytogenes culture in an orbital shaker at 200–250 rpm at 37 °C overnight. 4. The following day, make a 1:100 dilution with the overnight liquid culture in 5–10 mL BHI broth. Incubate in an orbital shaker at 200–250 rpm at 37 °C until culture reaches the logarithmic phase, typically measured at an optical density reading at 600 nm (OD600) of approximately 0.4. Following a 1:100 dilution, this usually takes 3 h. 5. Determine the concentration of the L. monocytogenes culture from the OD600 value (see Note 4). Centrifuge in high-speed rotor at 10,000 × g for 10 min. Discard supernatant. Resuspend in sterile 1× DPBS. Dilute as needed in sterile 1× DPBS to achieve final infectious concentration (see Note 5) in a 1.5 mL microcentrifuge tube. 6. Keep the diluted L. monocytogenes in DPBS suspension on ice while preparing mice for injections.

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3.3  Intravenous Injection of Mice with  L. monocytogenes

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1. Before infection, ensure that mice are transferred to an appropriate protocol approved by the Institutional Animal Care and Use Committee (IACUC). 2. Restrain the mouse in a suitable restraining device for tail vein injections. 3. Ensure the L. monocytogenes suspension is mixed thoroughly, and load a sterile 1 mL syringe fitted with a 27-gauge needle with at least 100 μL volume. Take care to eliminate all air bubbles. 4. Gently warm the mouse or tail using an approved technique (i.e., heat lamp or vein illuminator restraint) to better visualize the lateral tail vein. Inject 50 μL of the bacterial suspension into the tail vein. Briefly apply pressure to the entry wound with a clean WyPall. Return the mouse to its cage. 5. Repeat until all mice have been injected with the dose of interest, and monitor the mice as appropriate following approved institutional IACUC protocols (see Note 6). 6. For uninfected controls, repeat tail vein injections following the same protocol as detailed above, but using sterile 1× DPBS (vehicle).

3.4  Preparation of a Single-Cell Suspension from Spleens and Lymph Nodes of Infected Mice

1. On the day of isolation (see Note 7), label a 15 mL conical tube for each spleen and pooled lymph node sample to be harvested, and add 5 mL of ice-cold cIMDM to each tube. Tubes should be kept on ice during the harvest. Work should be completed under Biosafety Level 2 (BSL-2)-appropriate conditions for all steps (including the use of a tissue culture hood, appropriate disinfecting solution(s), and personal protective equipment). 2. Euthanize L. monocytogenes infected mice (and uninfected controls, as needed) according to individual institutional guidelines. Working with one mouse at a time, spray thoroughly with 70% ethanol, and harvest spleen and lymph nodes. Place tissue samples into the prepared media tubes as they are harvested. Keep samples on ice as much as possible. 3. To begin T helper cell isolation, transfer the first spleen or lymph node sample, along with media, to a sterile petri dish. Use the textured ends of frosted microscope slides to homogenize lymph node tissue to generate a single-cell suspension (see Note 8). Throughout homogenization, rinse the end of the slides by dipping them into the media in the dish to ensure maximal recovery. Before discarding slides, rinse thoroughly, and wick remaining cell suspension/media off of the slide ends by pressing them against clean areas on the dish surface. 4. Add an additional 5 mL of cIMDM to the cell suspension using a 10 mL serological pipet, and rinse the surface of the

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petri dish thoroughly. Transfer the cell suspension back to the 15 mL conical tube (see Note 9). Keep cell suspensions on ice, while you prepare the remainder of the samples. 5. Once all cell suspensions have been prepared, centrifuge samples at 600 × g for 5 min at 4 °C. 6. Aspirate and discard the supernatant from each sample. Close the tubes tightly, and flick-mix each cell pellet to ensure even distribution in RBC lysis buffer. Add an appropriate volume of RBC lysis buffer to each sample (4 mL for spleen samples, 2 mL for lymph node samples). Invert gently to mix and incubate at room temperature for exactly 3 min (see Note 10). 7. Immediately following the 3-min incubation, add an appropriate volume of cIMDM to each sample to dilute the RBS lysis buffer (5 mL for spleen, 4 mL for lymph nodes). Invert gently but thoroughly to mix and incubate at room temperature for 6 min. 8. During the above incubation, debris from cell lysis will collect at the bottom of each 15 mL conical tube. Carefully transfer spleen and lymph node samples for the same biological replicate to a single, fresh 15 mL conical tube, without transferring debris. Centrifuge the pooled samples at 600 × g for 5 min at 4 °C. Aspirate and discard the supernatant. 9. The cell pellet is now ready for analysis via flow cytometry (see Note 11). 3.5  Fluorescence-­ Activated Cell Sorting (FACS) Analysis of CD4+ T Cells Generated in Response to Systemic L. monocytogenes Infection

1. Prepare cells as desired (see Note 12), and carefully design the staining panel (see Notes 13–15) to analyze target CD4+ T helper subsets. Ensure sufficient controls are included and that the chosen fluorophore panel is compatible with the flow cytometer you intend to use (see Notes 16 and 17). 2. Prepare at least 50 mL of FACS buffer as detailed above (see Note 18). 3. Calculate the number of cells to be stained (see Note 19). If possible, ensure equal cell numbers between treatment groups. 4. Harvest the appropriate number of cells in 1.5 mL microcentrifuge tubes (see Note 20). Pellet cells by centrifugation at 350 × g for 5 min at room temperature. 5. Carefully aspirate media without disrupting the cell pellet. Wash cells by resuspending in 1 mL of FACS buffer. Pellet cells by centrifugation at 350 × g for 5 min at room temperature. 6. Carefully aspirate media without disrupting the cell pellet. Resuspend cell pellets in FACS buffer at a concentration of 1 × 106 cells per 100 μL. Treatment groups may now be divided into separate 1.5 mL microcentrifuge tubes depending on desired staining setup.

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7. To each tube, add appropriate amounts of antibody (see Note 21) depending on experimental plan. Once antibodies are added, samples must be protected from light to prevent photobleaching of the fluorophores. 8. Incubate samples for 1 h at room temperature (see Notes 18 and 22). 9. Add an additional 1 mL of FACS buffer to each sample. Pellet cells by centrifugation at 350 × g for 5 min at room temperature. Carefully aspirate supernatant without dislodging cells. 10. Wash cells by resuspending in 1 mL of FACS buffer. Pellet cells by centrifugation at 350 × g for 5 min at room temperature. Carefully aspirate supernatant. 11. (Optional) Repeat step 10. 12. If only extracellular epitopes are being stained, resuspend samples in 0.3–1 mL of FACS buffer, and transfer to a 5 mL round-bottomed flow cytometry tube. These samples can now be analyzed as desired via flow cytometry. Alternatively, samples can be prepared for intracellular staining (see Note 23). 3.6  Measurement of Bacterial Load in Tissues from Infected Mice

1. No more than 24 h after intravenous injection with L. monocytogenes, euthanize mice (see Note 24). Remove the spleen and liver (see Note 25), and collect in a sterile 1.5 mL microcentrifuge tube. Place the tubes on wet ice. Dispose of animal carcasses as appropriate. 2. Add 1 mL sterile 1× DPBS to each organ. 3. Use a handheld homogenizer with a sterile shaft (see Note 26) to achieve a uniform mixture. 4. Make serial dilutions of the homogenate using the dilutions and dosage injected as a reference. 5. Add 25 μL of each dilution to the center of a BHI agar plate, and use sterile glass beads, a sterile inoculating loop, or another method to spread evenly across the plate. Incubate the plates overnight, and count the colonies the following day. Determine the concentration by using the following calculation: CFU/ mL = (number of colonies × dilution factor)/0.025.

4  Notes 1. Institutional Biosafety Committee (IBC) approval may be necessary before beginning work with L. monocytogenes, due to the infectious nature of the bacteria. 2. Several strains of L. monocytogenes with unique genomic components exist and may have unique benefits in this experimental setting [15–18].

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3. L. monocytogenes can be stored long term by the addition of equal volume of 80% (v/v) sterile glycerol to the overnight BHI culture. Aliquots of this final 40% (v/v) glycerol-BHI concentration can be stored at −80 °C. 4. In order to determine an accurate concentration in CFU/mL based on OD600 reading, use an inoculating loop to pick a single L. monocytogenes colony from a BHI agar plate that has been incubated overnight, and add to 25 mL BHI broth. Incubate the culture in an orbital shaker at 200–250 rpm at 37 °C. Periodically, remove 1 mL from the culture, and measure the OD600. At OD600 readings of (as close as possible to) OD600 = 0.2, 0.4, 0.6, 0.8, and 1.0, make serial dilutions of the culture, and add 25 μL to the center of a pre-dried BHI agar plate, and use sterile glass beads, a sterile inoculating loop, or another method to spread evenly across the plate. Incubate the plates overnight, and count the colonies the following day. Determine the concentration by using the following calculation: CFU/mL = (number of colonies × dilution factor)/0.025. Graph the CFU/mL at the OD600 readings with CFU/mL on the y-axis and OD600 on the x-axis, and calculate the linear equation y = mx + b. Use this equation to plug the OD600 value into the x-value in order to calculate the y-value for concentration (CFU/mL). 5. Our laboratory typically utilizes a dose of 100,000 CFU/ mL, which is equivalent to 5000 CFU/mouse when injected with 50 μL. 6. The IACUC protocol approved for our laboratory mandates monitoring the mice at least twice in the first 24 h following injections, daily for days 3–4 post-injection, and then three times weekly for the remainder of the experiment unless/until the clinical condition of any mouse worsens. Clinical condition is scored according to the following parameters: Weight, 0 = 0–5%, 1 = 5–8%, 2 = 9–12%, 3 = 13–15%, and 4 > 16%; respiration, 0 = normal, 1 = slightly decreased, 2 = moderately reduced, 3 = severely reduced (quantifiable by eye), and 4 = abdominal effort and extremely reduced (