Phagocytosis and Phagosomes: Methods and Protocols [2 ed.] 1071633376, 9781071633373

Table of contents :
Preface
Contents
Contributors
Chapter 1: Bacterial Binding, Phagocytosis, and Killing Capacity: Measurements Using Colony Forming Units
1 Introduction
2 Materials
2.1 Bacteria and Microbiology Components
2.2 Macrophage Components
2.3 General Assay Components
3 Methods
3.1 Bacterial Binding Assay
3.2 Gentamicin Protection Assay for Bacterial Phagocytosis
3.3 Particle Binding Assay
3.4 Bacterial Killing Assay: Macrophage
4 Notes
References
Chapter 2: Analysis of Human and Mouse Neutrophil Phagocytosis by Flow Cytometry
1 Introduction
2 Materials
3 Methods
3.1 Phagocytosis by Human Blood Neutrophils
3.1.1 Preparation of pHrodo Particles
3.1.2 Acquisition of Blood Samples
3.1.3 Blood Neutrophil Stimulation
3.1.4 Fluorescent Labeling
3.1.5 Sample Acquisition
3.2 Mouse Peritoneal Recruitment and Phagocytosis
3.2.1 Preparation of pHrodo Particles and Intraperitoneal Injection
3.2.2 Peritoneal Lavage
3.2.3 Sample Fixation
3.2.4 Labeling
4 Notes
References
Chapter 3: Quantifying Phagocytosis by Immunofluorescence and Microscopy
1 Introduction
2 Materials
2.1 Cell Culture
2.2 Opsonizing Particles with IgG
2.2.1 Polystyrene Beads
2.2.2 Fluorescent Polystyrene Beads
2.2.3 Escherichia coli
2.2.4 Fluorescent Escherichia coli
2.2.5 Sheep Red Blood Cells
2.3 Inside-Outside Staining of Phagocytosed Opsonized Particles
2.4 Inside-Outside Staining of Phagocytosed Non-opsonized Particles
2.5 Microscopy and Quantification Phagocytosis
3 Methods
3.1 Cell Culture of RAW 264.7 Macrophage
3.2 Opsonization of Particles with IgG
3.2.1 Opsonization of Polystyrene Beads with IgG
3.2.2 Opsonization of E. coli with IgG
3.2.3 Opsonization of Sheep RBCs with IgG
3.3 Phagocytosis and Inside-Out Staining of Non-opsonized E. coli
3.4 Phagocytosis of Opsonized Fluorescent Particles
3.5 Phagocytosis and Inside-Out Staining of Opsonized Phagocytosed Particles
3.6 Quantification of Phagocytosis
4 Notes
References
Chapter 4: Methods for Quantitative Efferocytosis Assays
1 Introduction
2 Materials
2.1 Primary Human Macrophage Preparation
2.2 J774.2 Cell Culture and Transfection
2.3 Synthetic Efferocytic Targets
2.4 Preparation of Apoptotic Jurkat T Cells
2.5 Preparation of Apoptotic Neutrophils
2.6 Efferocytosis Assays
2.7 Immunostaining
2.8 Microscopy
3 Methods
3.1 Cell Culture and Cell Transfection
3.1.1 Primary Human Macrophage Preparation
3.1.2 Culture and Transfection of the J774.2 Macrophage Cell Line
3.2 Generation of Synthetic and Natural Efferocytic Targets
3.2.1 Synthetic Lipid-Coated Apoptotic Cell Targets
3.2.2 Synthetic Protein-Coated Apoptotic Cell Targets
3.2.3 Preparation of Apoptotic Jurkat T Cells
3.2.4 Preparation of Apoptotic Neutrophils
3.3 Efferocytosis Assays with Synthetic and Natural Targets
3.3.1 Synthetic Target-Based Efferocytosis Assay
3.3.2 Efferocytosis Assay Using Apoptotic Cells
3.3.3 Immunostaining
3.3.4 Fixed Sample Imaging and Quantification of Efferocytosis
3.3.5 Live-Cell Imaging
4 Notes
References
Chapter 5: Analysis of Efferocytic Receptor Dynamics and Synapse Formation in a Frustrated Efferocytosis Model
Abbreviations
1 Introduction
2 Materials
2.1 Cell Culture and THP-1 Differentiation
2.2 Preparation of Supported Lipid Bilayers
2.3 Immunostaining
2.4 Frustrated Efferocytosis and Image Acquisition
2.5 Image Analysis for Single Particle Tracking
3 Methods
3.1 Cell Culture and THP-1 Differentiation
3.2 Functionalization of Coverslips with Supported Lipid Bilayers
3.3 Immunostaining
3.4 Frustrated Efferocytosis and Image Acquisition
3.5 Single Particle Tracking Analysis
3.6 Morphological Analysis
4 Notes
References
Chapter 6: Super-Resolution Spinning-Disk Confocal Microscopy Using Optical Photon Reassignment (SoRa) to Visualize the Actin ...
1 Introduction
2 Materials
2.1 RAW 264.7 Murine Macrophage Cell Preparation
2.2 Transient Transfection of RAW Cells
2.3 Electroporation of RAW Cells
2.4 Opsonization of Polystyrene Beads
2.5 Phagocytosis Live Imaging and Microscopy
2.6 Immunofluorescence and Phalloidin Staining
2.7 Image Analysis
3 Methods
3.1 RAW 264.7 Murine Macrophage Cell Preparation
3.2 Transient Transfection of RAW Cells (See Note 14)
3.3 Electroporation of RAW Cells (See Note 14)
3.4 Opsonization of Polystyrene Beads
3.5 Phagocytosis Live Imaging and Microscopy
3.6 Immunofluorescence of Podosomes in Resting Macrophages
3.7 Image Analysis
4 Notes
References
Chapter 7: Filamentous Bacteria as Targets to Study Phagocytosis
1 Introduction
2 Materials
3 Methods
3.1 Obtaining Filamentous Bacterial Targets
3.1.1 Generating Filamentous Bacterial Targets of Escherichia coli or Salmonella typhimurium Using Antibiotics
3.1.2 Generating Filamentous Bacterial Targets of a Thermosensitive ftsZ Escherichia coli Mutant Strain
3.1.3 Engineering E. coli PAT84 to Express Fluorescent Proteins
3.1.4 Preparing Fixed Filaments of E. coli PAT84
3.1.5 Generating Filamentous Bacterial Targets of L. pneumophila in BCYE Agar
3.1.6 Generating Filamentous L. pneumophila in BYE Media
3.1.7 Preparing Fixed Filaments of L. pneumophila
3.2 Phagocytosis Assay Using Filamentous Bacterial Targets
3.3 Measuring Phagocytic Internalization Using Filamentous Bacteria
3.4 Remodeling and Maturation of Phagocytic Cups and Phagosomes
3.4.1 Phagocytic Cup Remodeling
3.4.2 Phagocytic Cup Maturation
3.4.3 Assessing the Luminal Environment of Phagocytic Cups and Phagosomes During Maturation
4 Notes
References
Chapter 8: The Derivation and Use of HoxB8-Driven Conditionally Immortalized Macrophages
1 Introduction
2 Materials
2.1 ER-HoxB8 DNA Isolation
2.2 ER-HoxB8 Lentivirus Production
2.3 Bone Marrow Isolation
2.4 ER-HoxB8 Lentiviral Transduction
2.5 Macrophage Differentiation
3 Methods
3.1 ER-HoxB8 and Lentiviral Packaging Constructs DNA Isolation
3.2 Generating ER-HoxB8 Lentivirus
3.2.1 Plating HEK293T Cells
3.2.2 Transfection
3.2.3 Virus Harvesting
3.2.4 Ultracentrifugation
3.3 Isolating Bone Marrow
3.4 Transduction with ER-HoxB8 Lentivirus
3.5 Macrophage Differentiation
4 Notes
References
Chapter 9: Quantitative Immunofluorescence to Study Phagosome Maturation and Resolution
1 Introduction
2 Materials
3 Methods
3.1 Cell Culture
3.2 Particle Preparation
3.2.1 mCherry-Escherichia coli (E. coli) Production and Fluorescent Protein Expression
3.2.2 Preparing IgG-Coated Particles
3.3 Phagocytosis, Phagosome Maturation, and Phagosome Resolution
3.3.1 Phagocytosis of IgG-Coated Beads
3.3.2 Phagocytosis of mCherry-E. coli and Fixation
3.4 Antibody Staining of Whole Phagocytes
3.5 Microscopy and Analysis of Whole Phagocytes
3.5.1 Maturation of Intact Phagosomes
3.5.2 Imaging and Quantification of Fragmented Phagosomes
3.6 Immunofluorescence and Flow Cytometry Analysis of Isolated Phagosomes
3.6.1 Cell Culture for Phagosome Isolation
3.6.2 Phagosome Isolation
3.6.3 LAMP1 and Clathrin Staining of Isolated Phagosomes
3.6.4 Microscopy and Analysis of Isolated Phagosomes
3.6.5 Flow Cytometry and Analysis of Isolated Phagosomes
4 Notes
References
Chapter 10: Approaches to Measuring Reductive and Oxidative Events in Phagosomes
1 Introduction
2 Materials
2.1 Preparation of Redox Reporter Beads
2.2 Cell Preparation and Handling
2.3 Performing Redox Measurements on Live Cells
3 Methods
3.1 Preparation of Redox Reporter Beads
3.1.1 Preparation of Dextran-Linked Reductase-Reporter Beads
3.1.2 Preparation of ROS-Reactive OxyBURST Beads
3.2 Cell Preparation and Handling
3.2.1 BMMØs and BMDCs
3.2.2 GB8-MØs
3.2.3 J774s
3.3 Performing Redox Measurements in Live Cells
3.3.1 Assessment of Phagosome-Specific Disulfide Reduction
3.3.2 Assessment of Phagosome-Specific ROS Production
3.3.3 Measurement of Extracellular H2O2
3.3.4 Measurement of Reactive Oxygen Species Production
4 Notes
References
Chapter 11: Measuring Phagosomal pH by Fluorescence Microscopy
1 Introduction
1.1 Instrumentation
1.2 Probe Selection
1.3 Calibration
2 Materials
2.1 Cell Lines
2.2 Reagents
2.3 Microscopy System Requirements
3 Methods
3.1 Labeling of Phagocytic Targets with pH Sensors
3.2 IgG Opsonization of Zymosan
3.3 Phagocytosis Assay
3.4 In Situ Calibration
4 Notes
References
Chapter 12: Multiplexed Phagosomal Assays for the Detection and Quantification of Bidirectional Exchange Between the Phagolyso...
1 Introduction
2 Materials
2.1 Cells, Reagents, and Buffers
2.2 Imaging Instrument and Analysis Software
3 Methods
3.1 Preparation of Experimental Reporter Particles
3.2 Preparation of Cellulase Reporter Beads
3.3 Preparation of DQ Green BSA and pHrodo Reporter Beads
3.4 Cell Preparation and Handling
3.5 Imaging System Setup
3.6 Acquisition of Images
3.7 Analysis
3.8 Using Spotfire DecisionSite to Visualize Data in Graphical Format
4 Notes
References
Chapter 13: Quantitative Spatio-temporal Analysis of Phagosome Maturation in Live Cells
1 Introduction
2 Materials
2.1 Cells, Buffers, and Solutions
2.1.1 Cells
2.1.2 Culture Media
2.1.3 Reagents for the Preparation of Phagocytic Targets
2.1.4 Labelling of Intracellular Organelles
2.1.5 Preparation of BMM
2.1.6 Preparation of HMDM
2.1.7 Preparation of Human and EB
2.2 Equipment
2.2.1 Preparation of BMM
2.2.2 Preparation of HMDM
2.2.3 Preparation of iPSDM
2.2.4 Live Cell Dishes
2.2.5 Microscope and Environmental Chamber
3 Methods
3.1 Preparation of Macrophages
3.1.1 Preparation of BMM
3.1.2 Preparation of HMDM (As Alternative to BMM)
3.1.3 Preparation of iPSDM (As Alternative to BMM or HMDM)
3.1.4 Preparing iPSDM for Experiments
3.2 Preparation of Phagocytic Targets
3.2.1 Polystyrene and Silica Beads
3.2.2 Preparation of Mycobacteria for Infection
3.3 Labelling of Intracellular Organelles
3.3.1 LysoTracker Delivery into Phagosomes
3.3.2 Cell Transfection
3.4 Live Cell Imaging
3.5 Image Analysis
3.5.1 Analysis of the Association of Various Markers to IgG-Coated Beads
3.5.2 Analysis of the Association of Various Markers to Mycobacteria
4 Notes
References
Chapter 14: Measurement of Salmonella enterica Internalization and Vacuole Lysis in Epithelial Cells
1 Introduction
2 Materials
3 Methods
3.1 Gentamicin Protection Assay
3.2 Chloroquine Resistance Assay
4 Notes
References
Chapter 15: Microscopy-Based Tracking and Quantification Methods to Study Phagosome Resolution
1 Introduction
2 Materials
3 Methods
3.1 Opsonization of Fixed Bacteria
3.2 Phagocytosis Assay
3.3 Live Cell Imaging
3.4 Fixation and Immunostaining
3.5 Determining the Total Volume of Phagosomes and Phagosome-Derived Vesicles (PDVs) per Cell
3.6 Determining the Co-occurrence of Various Membrane Markers with PDVs
4 Notes
References
Chapter 16: Isolation of Polystyrene Bead-Induced Phagosomes for Western Blotting
1 Introduction
2 Materials
2.1 Cell Culture
2.2 Induction of Phagocytosis and Cell Lysis
2.3 Phagosome Purification
2.4 SDS Polyacrylamide Gel Electrophoresis (SDS-PAGE)/Components
2.5 Immunoblotting
2.6 Immuno-detection
3 Methods
3.1 Induction of Phagocytosis
3.2 Cell Lysis
3.3 Phagosome Purification
3.4 Sample Preparation and SDS-PAGE Electrophoresis
3.5 Protein Transfer
3.6 Immunoblotting
3.7 Signal Detection
4 Notes
References
Chapter 17: Biochemically Reconstituted Fusion of Phagosomes with Endosomes and Lysosomes
1 Introduction
2 Materials
3 Methods
3.1 Preparation of BSA-Rhodamine
3.2 Coating of 1 μm Latex Beads with BSA
3.3 Preparation of Cytosol from J774E Macrophages
3.4 Preparation of Mowiol
3.5 Fluorescent and Paramagnetic Labeling of Lysosomes
3.6 Fluorescent and Paramagnetic Labeling of Late Endosomes
3.7 Fluorescent and Paramagnetic Labeling of Early Endosomes
3.8 Purification of Fluorescently and Paramagnetically Labeled Early Endosomes, Late Endosomes, or Lysosomes
3.9 Purification of Latex Bead Phagosomes
3.10 Cell-Free Fusion Between Phagosomes and Endosomes/Lysosomes
4 Notes
References
Chapter 18: An In Vitro System to Analyze Generation and Degradation of Phagosomal Phosphatidylinositol Phosphates
1 Introduction
2 Materials
2.1 Purification of Glutathione-S-Transferase (GST) or GST-fused PIP-Binding Proteins from E. coli BL21(DE3)
2.2 Detection of PIPs on Purified LBPs by Immunofluorescence Microscopy
3 Methods
3.1 Purification of Glutathione-S-Transferase (GST) or GST-fused PIP-Binding Proteins from E. coli BL21(DE3), Genotype E. coli...
3.2 Coating Latex Beads with BSA
3.3 Preparation of Cytosol from J774E Macrophages
3.4 Preparation of Mowiol
3.5 Purification of Latex Bead Phagolysosomes from J774E Macrophages
3.6 Incubation of Phagosomes with Lipid-Binding Probes
3.7 Fluorescence Microscopy Analysis and Quantification of the Amounts of Lipid-Binding Probes on Phagosomes
4 Notes
References
Chapter 19: Dissecting Phagosomal Pattern Recognition Receptor-Dependent Signaling and Antigen MHC-II Presentation from Phagos...
1 Introduction
2 Materials
2.1 Equipment
2.2 Mice
2.3 Reagents and Supplies
2.3.1 Cell Culture
2.3.2 Assays
Bead Preparation and Phagocytosis
PRR Signaling
Eα Ag Presentation
3 Methods
3.1 Cell Isolation or Differentiation
3.2 Signaling
3.2.1 Preparation of LPS-Coated Beads
3.2.2 DC Harvesting
3.2.3 DC Stimulation
3.3 Eα Antigen Presentation Assay
3.3.1 Eα-YFP-Coated Bead Preparation
3.3.2 Eα-YFP-Coated Bead Phagocytosis, Antigen Loading, and Presentation
3.3.3 Immunofluorescence
3.3.4 Flow Cytometry
4 Notes
References
Chapter 20: Examining the Kinetics of Phagocytosis-Coupled Inflammasome Activation in Murine Bone Marrow-Derived Dendritic Cel...
1 Introduction
2 Materials
2.1 Mice
2.2 DNA Constructs
2.3 Cell Culture
2.4 NLRC4/NLRP3 Inflammasome Stimuli
2.5 NLRP3 Inflammasome Stimuli
2.6 Other Reagents and Materials
2.7 Instruments and Software
3 Methods
3.1 Preparation of Reagents
3.1.1 Preparation of Toxin-Coated Beads
3.2 BM Isolation and BMDC Differentiation
3.3 Seeding and Splitting BM Cells
3.4 Transfecting Plat-E Cells and Seeding BM Cells for Retroviral Transduction
3.5 Salmonella Typhimurium Culture
3.6 Harvesting and Plating BMDCs
3.7 Cell Priming and Stimulation in 96-Well Plate
3.8 Caspase-11 Detection by TCA Precipitation
3.9 Cell Priming and Stimulation in Glass-Bottom p35 Dishes for ASC Speck Visualization by Fluorescence Microscopy or Live Cel...
3.10 ASC Speck Counting
3.11 Count Nuclei
4 Notes
References
Chapter 21: Analysis of LC3-Associated Phagocytosis and Antigen Presentation in Macrophages and B Cells
1 Introduction
2 Materials
2.1 Cell Culture
2.1.1 Isolation of Human Peripheral Blood Mononuclear Cells
2.1.2 Isolation and EBV Transformation of Human B Cells
EBV Production and Titration
2.1.3 Isolation of Human Macrophages
2.1.4 Isolation of Human Whole Blood CD4+ T Cells
2.2 CRISPR/Cas9 Knock-out of ATG4B in LCLs
2.3 Lentiviral Constructs of ATG4Bwt or ATG4BC78S Overexpression
2.4 Western Blot
2.4.1 Reagents
2.4.2 Buffers
2.5 Immunofluorescence and Confocal Microscopy
2.6 Role of LC3-Associated Phagocytosis During MHC Class II Antigen Presentation
2.7 Reagents
2.7.1 Cell Culture
2.7.2 Antibodies
2.7.3 Other Probes
3 Methods
3.1 Cell Culture
3.1.1 EBV Production and Titration
3.1.2 Isolation of Human Peripheral Blood Mononuclear Cells (PBMCs)
3.1.3 Isolation and EBV Transformation of Human B Cells
3.1.4 Isolation of Monocyte-Derived Macrophages
3.1.5 Isolation of Autologous CD4+ T Cells for Antigen Presentation
3.2 CRISPR/Cas9 Knock-out of ATG4B in LCLs
3.3 Lentivirus Transduction of Human Macrophages for ATG4 Overexpression
3.4 Trigger LC3-Associated Phagocytosis in Human Macrophages Using Candida albicans Extract
3.4.1 Candida albicans Extract Coated Beads
3.4.2 LAP Triggered with Candida albicans Extract Coated Beads
3.5 Western Blot to Validate ATG4 Knockout or Overexpression in Human Antigen Presenting Cells
3.6 An Immunofluorescence Protocol to Investigate the Role of ATG4 During LAP in Human Macrophages and Its Role During Autopha...
3.6.1 Immunofluorescence Protocol for Adherent Cells
3.6.2 Immunofluorescence Protocol for Cells in Suspensions
3.6.3 Image Acquisition and Analysis
3.7 Assess the Involvement of ATG4 in LAPosome Stabilization to Allow Macrophages to Sustain MHC Class II Antigen Presentation
4 Notes
References
Chapter 22: Visualizing Phagocytic Cargo In Vivo from Engulfment to Resolution in Caenorhabditis elegans
1 Introduction
2 Materials
2.1 Worm Strains and Maintenance
2.2 RNA Interference (RNAi) Reagents
2.3 Microscopy
3 Methods
3.1 Preparing Buffers and Media
3.1.1 Luria Broth (LB)
3.1.2 Nematode Growth Media (NGM) Plates
3.1.3 NGM Lite Plates
3.1.4 M9 Buffer
3.1.5 Egg Salts
3.1.6 Agarose Pads
3.2 Mounting Embryos for Time-Lapse Imaging
3.2.1 Mounting Embryos for Time-Lapse Imaging (Agarose Pad)
3.2.2 Mounting Embryos for Time-Lapse Imaging (Well)
3.3 Time-Lapse Imaging
3.4 Normalization of Time-Lapse Series
3.5 Cargo Signaling for Engulfment
3.6 Engulfment
3.7 Phagosome Maturation
3.8 Phagosome-Lysosome Fusion
3.9 Phagosome and Phagolysosome Acidification
3.10 Cargo Membrane Breakdown
3.10.1 Direct Membrane Breakdown Assay
3.10.2 Indirect Chromosome Assay for Membrane Breakdown
3.11 Phagolysosome Shrinkage
3.12 Phagolysosome Tubulation, Vesiculation, and Resolution
4 Notes
References
Chapter 23: Assessing the Phagosome Proteome by Quantitative Mass Spectrometry
1 Introduction
2 Materials
2.1 Equipment
2.2 Cell Lysis, Reduction, Alkylation, and Digestion
2.3 Sample Clean-up, TMT Labelling, and MS
2.4 Data Analysis
3 Methods
3.1 Cell Lysis, Reduction, Alkylation, and Digestion for Quantitative Proteomic Analysis
3.2 Quantitative Proteomics of Phagosomes Using TMTpro 16plex
3.3 Offline HPLC Fractionation for TMTpro 16plex Labelled Samples
3.4 Mass Spectrometry Analysis for TMT Samples
3.5 Mass Spectrometry Analysis for Label-Free Samples
3.6 Data Processing and Analysis
3.7 Quality Control
4 Notes
References
Chapter 24: Using Ion Substitution and Fluid Indicators to Monitor Macropinosome Dynamics in Live Cells
1 Introduction
2 Materials
2.1 Cell Culture
2.2 Primary Macrophage Derivation
2.3 Buffers
2.4 Chemicals
2.5 Confocal Microscope
3 Methods
3.1 Deriving BMDM
3.1.1 Preparation of M-CSF
3.1.2 Isolation of BMDMs from Murine Long Bones
3.2 Seeding BMDM onto Coverslips
3.3 Inducing Macropinocytosis
3.4 Imaging and Recording
3.5 Data Analysis
4 Notes
References
Index

Citation preview

Methods in Molecular Biology 2692

Roberto J. Botelho  Editor

Phagocytosis and Phagosomes Methods and Protocols Second Edition

METHODS

IN

MOLECULAR BIOLOGY

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

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

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

Phagocytosis and Phagosomes Methods and Protocols Second Edition

Edited by

Roberto J. Botelho Department of Chemistry and Biology, Toronto Metropolitan University, Toronto, ON, Canada

Editor Roberto J. Botelho Department of Chemistry and Biology Toronto Metropolitan University Toronto, ON, Canada

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-3337-3 ISBN 978-1-0716-3338-0 (eBook) https://doi.org/10.1007/978-1-0716-3338-0 © The Editor(s) (if applicable) and The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023 This work is subject to copyright. All rights are solely and exclusively licensed by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, expressed or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. This Humana imprint is published by the registered company Springer Science+Business Media, LLC, part of Springer Nature. The registered company address is: 1 New York Plaza, New York, NY 10004, U.S.A.

Preface I am pleased to present the second edition of Methods in Molecular Biology: Phagocytosis and Phagosomes that offers 24 chapters detailing experimental approaches used to investigate phagocytosis and phagosome maturation. The second edition includes both updated chapters found in the first edition, as well as entirely new methods for the study of phagocytosis and phagosome biology. Phagocytosis is the cellular engulfment of particulate matter, and it is of broad biological importance, being employed by unicellular organisms and by numerous cell types within multicellular organisms. Unicellular organisms, exemplified by hundreds of Protist species, employ phagocytosis to ingest food particles. In multicellular organisms, phagocytosis plays an essential function in tissue remodeling and tissue homeostasis by removing millions upon millions of apoptotic bodies, senescent cells, and cell fragments every single day. In addition, phagocytosis is a paramount immune weapon utilized by leukocytes to ingest and eliminate a plethora of potential pathogens and other foreign particulates. Internalization is only half the story. Once engulfed, the particle is now sequestered within a new organelle—the phagosome. The newly formed phagosome is an innocuous organelle, but it is equipped with molecular cues that guide its maturation into a phagolysosome, an acidic and hydrolytic organelle that ultimately degrades the particle. This transformation requires sequential fusion with endosomes and lysosomes. In recent years, we’ve come to recognize that once particulates are digested, the phagosome itself must be recycled through fragmentation; this process is referred to as phagosome resolution. Regardless of this, it is not surprising that microbes have evolved myriad mechanisms to control engulfment and phagosome maturation to hijack and replicate within hosts cells; these pathogenic microbes remain a large source of human morbidity globally. Phagocytosis and phagosome maturation are complex, diverse, and highly dynamic processes. Thus, a large array of complementary methods and tools have been developed and are required to study engulfment, maturation, resolution, and pathogen manipulation of phagocytes. In this volume, the reader will find detailed instructions and tips in the form of “Notes” on using many of these tools. For example, several chapters offer methodology to quantify uptake and maturation specific to certain phagocytes, particles, or pathogens, while other chapters offer methods that can be applied generically across the field. Methods are presented to study phagocytosis and phagosome maturation in vivo, in cellulo, and through in vitro analyses. Due to the dynamic and rapid nature of phagocytosis and maturation, these processes are especially amenable to analysis by microscopy and flow cytometry. Not surprisingly, many chapters in this volume detail the use of imaging methods to analyze these processes in fixed and in living cells. Overall, I believe this updated second edition will continue to be an important resource for both experts in the field and for those investigators delving into phagocytosis and phagosome maturation for the first time. As a last word, it is important to realize that phagocytosis is really a collection of distinct processes, rather than a single phenomenon. Indeed, the mechanics, outcomes, and accompanying events triggered by phagocytosis and

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Preface

phagosome maturation differ depending on the phagocyte, phagocytic receptors, and target particles engaged. This presents a significant challenge to our understanding of phagocytosis and phagosome maturation and is often a source of confusion in the literature. Therefore, one should resist pronouncing sweeping statements about these processes. Toronto, ON, Canada

Roberto J. Botelho

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

1 Bacterial Binding, Phagocytosis, and Killing Capacity: Measurements Using Colony Forming Units . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Kyle E. Novakowski, Dessi Loukov, and Dawn M. E. Bowdish 2 Analysis of Human and Mouse Neutrophil Phagocytosis by Flow Cytometry . . . Noah Fine, Oriyah Barzilay, and Michael Glogauer 3 Quantifying Phagocytosis by Immunofluorescence and Microscopy . . . . . . . . . . . Sierra Soffiaturo, Christopher Choy, and Roberto J. Botelho 4 Methods for Quantitative Efferocytosis Assays . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Nima Taefehshokr and Bryan Heit 5 Analysis of Efferocytic Receptor Dynamics and Synapse Formation in a Frustrated Efferocytosis Model . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Brandon H. Dickson and Bryan Heit 6 Super-Resolution Spinning-Disk Confocal Microscopy Using Optical Photon Reassignment (SoRa) to Visualize the Actin Cytoskeleton in Macrophages . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Freyja Verth and Gregory D. Fairn 7 Filamentous Bacteria as Targets to Study Phagocytosis . . . . . . . . . . . . . . . . . . . . . . Akriti Prashar, Maria Cecilia Gimenez, Serene Moussaoui, Iram Sobia Khan, and Mauricio R. Terebiznik 8 The Derivation and Use of HoxB8-Driven Conditionally Immortalized Macrophages . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Shranjit S. Lail, Neil McKenna, and Robin M. Yates 9 Quantitative Immunofluorescence to Study Phagosome Maturation and Resolution . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Me´lanie Mansat, Roya M. Dayam, and Roberto J. Botelho 10 Approaches to Measuring Reductive and Oxidative Events in Phagosomes . . . . . Shranjit S. Lail, Dale R. Balce, Johnathan Canton, and Robin M. Yates 11 Measuring Phagosomal pH by Fluorescence Microscopy . . . . . . . . . . . . . . . . . . . . Gerone A. Gonzales and Johnathan Canton 12 Multiplexed Phagosomal Assays for the Detection and Quantification of Bidirectional Exchange Between the Phagolysosomal Lumen and Extracellular Space . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jenny A. Nguyen, Catherine J. Greene, Samuel Cheung, and Robin M. Yates 13 Quantitative Spatio-temporal Analysis of Phagosome Maturation in Live Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Patricia Rosell Are´valo, Beren Aylan, and Maximiliano G. Gutierrez

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Measurement of Salmonella enterica Internalization and Vacuole Lysis in Epithelial Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jessica A. Klein, TuShun R. Powers, and Leigh A. Knodler Microscopy-Based Tracking and Quantification Methods to Study Phagosome Resolution . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Charlene E. Lancaster, Serene Moussaoui, Maria Cecilia Gimenez, and Mauricio R. Terebiznik Isolation of Polystyrene Bead-Induced Phagosomes for Western Blotting . . . . . . Benjamin B. A. Raymond, Joseph Inns, Andrew M. Frey, and Matthias Trost Biochemically Reconstituted Fusion of Phagosomes with Endosomes and Lysosomes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Andreas Jeschke and Albert Haas An In Vitro System to Analyze Generation and Degradation of Phagosomal Phosphatidylinositol Phosphates . . . . . . . . . . . . . . . . . . . . . . . . . . . . Andreas Jeschke Dissecting Phagosomal Pattern Recognition Receptor-Dependent Signaling and Antigen MHC-II Presentation from Phagosomes in Murine Dendritic Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Emilia Scharrig and Adriana R. Mantegazza Examining the Kinetics of Phagocytosis-Coupled Inflammasome Activation in Murine Bone Marrow-Derived Dendritic Cells . . . . . . . . . . . . . . . . . Daniel J. Netting and Adriana R. Mantegazza Analysis of LC3-Associated Phagocytosis and Antigen Presentation in Macrophages and B Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Svenja Luisa Nopper, Katarina Wendy Schmidt, Laure-Anne Ligeon, ¨ nz and Christian Mu Visualizing Phagocytic Cargo In Vivo from Engulfment to Resolution in Caenorhabditis elegans . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Gholamreza Fazeli, Julia Frondoni, Shruti Kolli, and Ann M. Wehman Assessing the Phagosome Proteome by Quantitative Mass Spectrometry. . . . . . . ˜ as, Jose´ Luis Marı´n-Rubio, Julien Peltier-Heap, Maria Emilia Duen Anetta Hartlova, and Matthias Trost Using Ion Substitution and Fluid Indicators to Monitor Macropinosome Dynamics in Live Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Guillermo A. de Paz Linares, Spencer A. Freeman, and Ruiqi Cai

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

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Contributors PATRICIA ROSELL ARE´VALO • Host-Pathogen Interactions in Tuberculosis Laboratory, The Francis Crick Institute, London, UK BEREN AYLAN • Host-Pathogen Interactions in Tuberculosis Laboratory, The Francis Crick Institute, London, UK DALE R. BALCE • Department of Comparative Biology and Experimental Medicine, Faculty of Veterinary Medicine, University of Calgary, Calgary, AB, Canada ORIYAH BARZILAY • Faculty of Dentistry, University of Toronto, Toronto, ON, Canada ROBERTO J. BOTELHO • Molecular Science Graduate Program, Toronto Metropolitan University, Toronto, ON, Canada; Department of Chemistry and Biology, Toronto Metropolitan University, Toronto, ON, Canada DAWN M. E. BOWDISH • Department of Medicine, McMaster University, Hamilton, ON, Canada RUIQI CAI • Program in Cell Biology, Peter Gilgan Centre for Research and Learning, The Hospital for Sick Children, Toronto, ON, Canada JOHNATHAN CANTON • Department of Comparative Biology and Experimental Medicine, Faculty of Veterinary Medicine, University of Calgary, Calgary, AB, Canada; Calvin, Joan and Phoebe Snyder Institute of Chronic Disease, Cumming School of Medicine, University of Calgary, Calgary, AB, Canada SAMUEL CHEUNG • Department of Comparative Biology and Experimental Medicine, Faculty of Veterinary Medicine, University of Calgary, Calgary, AB, Canada CHRISTOPHER CHOY • Molecular Science Graduate Program, Toronto Metropolitan University, Toronto, ON, Canada; Department of Chemistry and Biology, Toronto Metropolitan University, Toronto, ON, Canada; BlueRock Therapeutics, Toronto, ON, Canada ROYA M. DAYAM • Department of Chemistry and Biology and the Graduate Program in Molecular Science, Toronto Metropolitan University, Toronto, ON, Canada GUILLERMO A. DE PAZ LINARES • Department of Biochemistry, University of Toronto, Toronto, ON, Canada BRANDON H. DICKSON • Department of Microbiology and Immunology, and The Western Infection, Immunity and Inflammation Centre, The University of Western Ontario, London, ON, Canada MARIA EMILIA DUEN˜AS • Biosciences Institute, Newcastle University, Newcastle upon Tyne, UK GREGORY D. FAIRN • Department of Pathology, Dalhousie University, Halifax, NS, Canada; Department of Biochemistry and Molecular Biology, Dalhousie University, Halifax, NS, Canada GHOLAMREZA FAZELI • Imaging Core Facility, Biocenter, University of Wu¨rzburg, Wu¨rzburg, Germany NOAH FINE • Faculty of Dentistry, University of Toronto, Toronto, ON, Canada SPENCER A. FREEMAN • Department of Biochemistry, University of Toronto, Toronto, ON, Canada; Program in Cell Biology, Peter Gilgan Centre for Research and Learning, The Hospital for Sick Children, Toronto, ON, Canada

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ANDREW M. FREY • Laboratory for Biomedical Mass Spectrometry, Biosciences Institute, Newcastle University, Newcastle upon Tyne, UK JULIA FRONDONI • Department of Biological Sciences, University of Denver, Denver, CO, USA MARIA CECILIA GIMENEZ • Department of Biological Sciences, University of Toronto at Scarborough, Toronto, ON, Canada MICHAEL GLOGAUER • Faculty of Dentistry, University of Toronto, Toronto, ON, Canada GERONE A. GONZALES • Department of Biochemistry and Molecular Biology, Cumming School of Medicine, University of Calgary, Calgary, AB, Canada CATHERINE J. GREENE • Department of Biochemistry and Molecular Biology, Cumming School of Medicine, University of Calgary, Calgary, AB, Canada MAXIMILIANO G. GUTIERREZ • Host-Pathogen Interactions in Tuberculosis Laboratory, The Francis Crick Institute, London, UK ALBERT HAAS • Institute for Cell Biology, University of Bonn, Bonn, Germany ANETTA HARTLOVA • Institute of Biomedicine, Department of Microbiology and Immunology, the Sahlgrenska Academy/Faculty of Science, University of Gothenburg, Gothenburg, Sweden; Wallenberg Centre for Molecular and Translational Medicine, University of Gothenburg, Gothenburg, Sweden BRYAN HEIT • Department of Microbiology and Immunology, and The Western Infection, Immunity and Inflammation Centre, The University of Western Ontario, London, ON, Canada; Robarts Research Institute, London, ON, Canada JOSEPH INNS • Laboratory for Biomedical Mass Spectrometry, Biosciences Institute, Newcastle University, Newcastle upon Tyne, UK ANDREAS JESCHKE • Institute for Cell Biology, University of Bonn, Bonn, Germany IRAM SOBIA KHAN • Department of Biological Sciences, University of Toronto at Scarborough, Toronto, ON, Canada; Department of Cell and Systems Biology, University of Toronto, Toronto, ON, Canada JESSICA A. KLEIN • Paul G. Allen School for Global Health, College of Veterinary Medicine, Washington State University, Pullman, WA, USA LEIGH A. KNODLER • Paul G. Allen School for Global Health, College of Veterinary Medicine, Washington State University, Pullman, WA, USA; Department of Microbiology and Molecular Genetics, University of Vermont, Burlington, USA SHRUTI KOLLI • Department of Biological Sciences, University of Denver, Denver, CO, USA SHRANJIT S. LAIL • Department of Medical Science, Cumming School of Medicine, University of Calgary, Calgary, AB, Canada CHARLENE E. LANCASTER • Department of Biological Sciences, University of Toronto at Scarborough, Toronto, ON, Canada; Department of Cell and Systems Biology, University of Toronto, Toronto, ON, Canada LAURE-ANNE LIGEON • Viral Immunobiology, Institute of Experimental Immunology, University of Zu¨rich, Zu¨rich, Switzerland DESSI LOUKOV • Department of Medicine, McMaster University, Hamilton, ON, Canada ME´LANIE MANSAT • Department of Chemistry and Biology and the Graduate Program in Molecular Science, Toronto Metropolitan University, Toronto, ON, Canada ADRIANA R. MANTEGAZZA • Department of Microbiology and Immunology, Sidney Kimmel Medical College, Thomas Jefferson University, Philadelphia, PA, USA JOSE´ LUIS MARI´N-RUBIO • Biosciences Institute, Newcastle University, Newcastle upon Tyne, UK

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NEIL MCKENNA • Department of Comparative Biology and Experimental Medicine, Faculty of Veterinary Medicine, University of Calgary, Calgary, AB, Canada SERENE MOUSSAOUI • Department of Biological Sciences, University of Toronto at Scarborough, Toronto, ON, Canada; Department of Cell and Systems Biology, University of Toronto, Toronto, ON, Canada CHRISTIAN MU¨NZ • Viral Immunobiology, Institute of Experimental Immunology, University of Zu¨rich, Zu¨rich, Switzerland DANIEL J. NETTING • Department of Microbiology and Immunology, Sidney Kimmel Medical College, Thomas Jefferson University, Philadelphia, PA, USA JENNY A. NGUYEN • Department of Biochemistry and Molecular Biology, Cumming School of Medicine, University of Calgary, Calgary, AB, Canada SVENJA LUISA NOPPER • Viral Immunobiology, Institute of Experimental Immunology, University of Zu¨rich, Zu¨rich, Switzerland KYLE E. NOVAKOWSKI • Department of Medicine, McMaster University, Hamilton, ON, Canada JULIEN PELTIER-HEAP • Inoviv, London, UK TUSHUN R. POWERS • Paul G. Allen School for Global Health, College of Veterinary Medicine, Washington State University, Pullman, WA, USA AKRITI PRASHAR • Program in Cell Biology, Peter Gilgan Centre of Research and Learning, The Hospital for Sick Children, Toronto, ON, Canada BENJAMIN B. A. RAYMOND • Laboratory for Biomedical Mass Spectrometry, Biosciences Institute, Newcastle University, Newcastle upon Tyne, UK EMILIA SCHARRIG • Department of Microbiology and Immunology, Sidney Kimmel Medical College, Thomas Jefferson University, Philadelphia, PA, USA KATARINA WENDY SCHMIDT • Viral Immunobiology, Institute of Experimental Immunology, University of Zu¨rich, Zu¨rich, Switzerland SIERRA SOFFIATURO • Molecular Science Graduate Program, Toronto Metropolitan University, Toronto, ON, Canada; Department of Chemistry and Biology, Toronto Metropolitan University, Toronto, ON, Canada NIMA TAEFEHSHOKR • Department of Microbiology and Immunology, and The Western Infection, Immunity and Inflammation Centre, The University of Western Ontario, London, ON, Canada MAURICIO R. TEREBIZNIK • Department of Biological Sciences, University of Toronto at Scarborough, Toronto, ON, Canada; Department of Cell and Systems Biology, University of Toronto, Toronto, ON, Canada MATTHIAS TROST • Laboratory for Biomedical Mass Spectrometry, Biosciences Institute, Newcastle University, Newcastle upon Tyne, UK; Biosciences Institute, Newcastle University, Newcastle upon Tyne, UK FREYJA VERTH • Department of Pathology, Dalhousie University, Halifax, NS, Canada ANN M. WEHMAN • Department of Biological Sciences, University of Denver, Denver, CO, USA ROBIN M. YATES • Department of Biochemistry and Molecular Biology, Cumming School of Medicine, University of Calgary, Calgary, AB, Canada; Department of Comparative Biology and Experimental Medicine, Faculty of Veterinary Medicine, University of Calgary, Calgary, AB, Canada; Cumming School of Medicine, Snyder Institute of Chronic Disease, University of Calgary, Calgary, AB, Canada

Chapter 1 Bacterial Binding, Phagocytosis, and Killing Capacity: Measurements Using Colony Forming Units Kyle E. Novakowski, Dessi Loukov, and Dawn M. E. Bowdish Abstract Herein, we provide a colony forming unit (CFU)-based counting method for quantitating the bacterial binding, phagocytosis, and killing capacity of phagocytes. Although these functions can be measured by immunofluorescence- and dye-based assays, quantitating CFUs are comparatively inexpensive and easy to perform. The protocol described below is easily modified for use with different phagocytes (e.g., macrophages, neutrophils, cell lines), types of bacteria, or opsonic conditions. Key words Phagocytosis, Bacterial killing, Particle binding, Macrophage, Gentamicin protection, Phagocyte

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Introduction Essential functions of the innate immune system include the detection, engulfment, and destruction of bacteria, which is primarily performed by neutrophils and macrophages. Profiling the functions of professional phagocytes provides valuable insight into innate immune function and can be used in a wide range of biological applications. These include testing the effect of a chemical compound or characterizing the phenotype of a genetic knockout. Methods to quantify cellular association, phagocytosis, and killing capacity by enumeration of viable microbial units were first described in the 1930s and 1940s [1]. Since then, assays have diversified to include quantification of bacterial binding, uptake, and killing of fluorescently labeled bacteria by flow cytometry or fluorescence microscopy. However, viability-based assays remain the most cost-effective and versatile. To better understand the principles of this assay, a distinction must be made between binding and total cell association. Total cell association refers to bacteria that have been internalized by a phagocyte, in addition to bacteria that are bound, but remain

Roberto J. Botelho (ed.), Phagocytosis and Phagosomes: Methods and Protocols, Methods in Molecular Biology, vol. 2692, https://doi.org/10.1007/978-1-0716-3338-0_1, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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outside the cell. This distinction is important, because not all bound bacteria are phagocytosed. If, for example, the bacteria produce anti-phagocytic factors, or the presence of specific chemical inhibitors or gene deletions interferes with receptor expression, bacterial binding may occur without internalization [2–5]. One method to distinguish between bound and internalized bacteria is to differentially label internal and external bacteria using two different fluorescent dyes or to use a pH-sensitive dye which only fluoresces in the low pH environment of phagosome [6]. Unfortunately, these methods require expensive reagents and analysis equipment (e.g., flow cytometers, fluorescence microscopes). Furthermore, examination of samples by microscopy can be timeconsuming and is relatively low throughput. An alternative approach is the use of viability-based assays. These assays are simpler and more affordable alternatives that can be used with a wide range of bacteria, some of which may not be amenable to fluorescent labeling by dyes or fluorescent proteins. Bacterial binding can be quantified by incubating phagocytes and bacteria at cold temperatures, which allows for phagocyte-bacteria interaction but prevents internalization because processes such as actin polymerization and membrane trafficking do not occur at sub-physiological temperatures [7]. Total cell association can be quantified by incubating phagocytes and bacteria at physiologic temperatures (i.e., 37
C), which allows for both bacterial binding and internalization. In order to distinguish between bound and internalized bacteria, extracellular bacteria can be killed through the addition of antibiotics. In order to determine the rate at which phagocytosed bacteria are killed, one can measure bacterial numbers over time. We have successfully used this assay to characterize the role of a macrophage receptor in antipneumococcal immunity, to study the effects of age on macrophage killing capacity using human and mouse macrophages and to discover drugs that increased macrophage phagocytosis [8–11]. Herein, we describe methods to characterize three distinct phagocyte functions: binding of particles/bacteria, phagocytosis/internalization of bacteria, and killing of bacteria.

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Materials Reagents used in this assay are specific to assays performed using Streptococcus pneumoniae. When performing this assay with other bacteria, reagents such as liquid culture media and agar growth plates can be substituted for appropriate growth media. This protocol can also be adapted for use with inert particles (see Subheading 3.1, step 1).

Measurement of Bacterial Binding, Phagocytosis and Killing

2.1 Bacteria and Microbiology Components

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1. Early-to-mid-log phase bacteria (see Notes 1 and 2). 2. Appropriate liquid culture media (e.g., Luria-Bertani Broth or Tryptic Soy Broth). Follow manufacturer’s instructions for broth preparation. Autoclave to sterilize. Store at 4
C. Only open under sterile conditions (see Note 3). 3. Sterile water. 4. Hank’s Balanced Salt Solution (HBSS). 5. Gentamicin solution: dissolve gentamicin sulfate salt to a final concentration of 50 mg/mL in sterile water. Store at 20
C in aliquots (see Note 2). 6. Tryptic Soy Agar (TSA)/Sheep’s blood plates: dissolve 15 g of TSA in 500 mL deionized water. Autoclave to sterilize. Cool for 1.5 h in a 60
C water bath. Under sterile conditions, add 25 mL sterile, defibrinated sheep’s blood, and swirl gently to mix (see Note 3). 7. Spectrophotometer. 8. 15 mL polystyrene culture centrifuge tubes. 9. Incubator. The incubation conditions of plates for quantification of bacteria will vary depending on the bacteria being used.

2.2 Macrophage Components

1. Macrophages: any source of macrophages can be used, including bone marrow-derived, alveolar, resident or recruited peritoneal, blood monocyte-derived, etc. (see Note 4). 2. Phosphate buffered saline (PBS): warm PBS to 37
C prior to use. 3. Accutase cell detachment solution: store at 20
C in 10 mL aliquots. Warm to 37
C prior to use (see Note 5). 4. Cell lifters (see Note 6). 5. Trypan Blue, 0.4%. 6. Hemocytometer. 7. Light microscope with a minimum objective magnification of 20. 8. 50 mL conical tubes.

2.3 General Assay Components

1. 2 mL microcentrifuge tubes. 2. Vortex. 3. Microcentrifuge. 4. Serological pipettes (5, 10, 25 mL). 5. Micropipettes and micropipette tips. 6. Appropriate disposal reagents and equipment for liquid biological waste. Please follow institutional requirements. 7. Nutating Mixer.

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8. Black 96-well plate. 9. Spectrophotometer capable of reading fluorescence. 10. If performing assays for particle uptake: fluorescent polystyrene microspheres, 0.5–2 μM (Polysciences, Warrington, PA, USA).

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Methods

3.1 Bacterial Binding Assay

Carry out all procedures under sterile conditions in a biological safety cabinet. For the experiments using S. pneumoniae in Figs. 1, 2, and 3, TSB/Sheep’s blood plates were used. 1. Grow liquid cultures to early or mid-log phase (see Notes 1 and 7). While waiting, prepare and label tubes containing 900 μL sterile water and appropriate media plates for the experiments. 2. When the liquid bacterial culture reaches early to mid-log growth (measured using OD600; see Note 1), remove 1 mL into a 2 mL microcentrifuge tube and centrifuge at 12,000  g for 1 min.

Fig. 1 A schematic outlining the major steps to perform a CFU-based bacterial killing assay

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Fig. 2 A representative killing assay highlighting differences in kill curves with varying multiplicities of infection (MOI) using bone marrow-derived macrophages

Fig. 3 A representative killing assay highlighting differences in kill curves with varying preincubation times before assaying bacterial killing using bone marrow-derived macrophages

3. Carefully remove the supernatant and resuspend the pellet in 1 mL HBSS. Keep on ice to prevent additional bacterial growth. 4. Visually inspect macrophage culture under a light microscope to ensure high cell viability and that the culture is free of contamination. Viable macrophages typically have a dendriticlike morphology and should be strongly adherent. Floating or rounded cells generally indicate a loss of viability, which can be confirmed by Trypan Blue staining. 5. Remove macrophage culture media and gently wash with warm PBS. 6. Remove PBS and add 10 mL of warm Accutase cell detachment solution. Cover the plate and place in a 37
C incubator for 10 min. Visually inspect the plate under a light microscope to determine cell detachment. Some cells should be completely

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detached, and remaining adherent cells should be round in morphology. If cells still remain adherent, incubate for an additional 5–10 min. 7. Gently scrape the plate with a cell lifter, to remove any remaining attached cells (see Note 6). Pipette Accutase solution containing macrophages into a 50 mL conical tube. To maximize macrophage recovery, wash the plate with 10 mL warm PBS and add to the 50 mL conical tube. 8. Centrifuge for 10 min at 500  g. 9. Carefully remove all supernatant and gently resuspend the pellet in 1 mL warm PBS. Do not vortex. 10. Determine the cells/mL by diluting cells 1:1 with Trypan Blue, and pipette 10 μL into a hemocytometer. Count viable cells following standard usage of a hemocytometer. 11. Add up to 1  106 viable macrophages into a 2 mL microcentrifuge tube. Bring the volume of solution up to 900 μL by adding PBS. Repeat for each desired time point. For example, if performing time points of 0, 30, 60, and 90 min, you will require four tubes containing equal numbers of macrophages (see Note 8). 12. Briefly vortex bacteria and add the desired multiplicity of infection (MOI) to each tube. The MOI will vary depending on the bacteria used in the assay and number/type of macrophages but generally ranges from 10 to 100 (see Note 9, Fig. 2). In the case of Streptococcus pneumoniae, an MOI of 20 is optimal. The final volume of the macrophage/bacteria solution should be brought to 1 mL with PBS. 13. Incubate with gentle mixing at 4
C for 1 h. This incubation allows for cell to cell association. At time points 0, 20, 40, and 60 min, remove the tube and centrifuge at 500  g for 5 min. At low speeds, macrophages (and bound bacteria) will form a pellet, whereas unbound bacteria will remain in the supernatant. 14. Carefully remove the supernatant to remove unbound bacteria, and gently resuspend in 1 mL cold PBS. Centrifuge at 500  g, 4
C, for 5 min. Repeat wash two times. 15. Resuspend in 1 mL of PBS, and lyse macrophages by pipetting 100 μL of the macrophage solution into 900 μL sterile water (prepared in Step 1) (see Note 10). Briefly vortex. This tube is now a 101 dilution. 16. Perform a serial dilution by repeating Step 14 using 100 μL of the 101 dilution into 900 μL sterile water. Briefly vortex. This tube is now a 102 dilution. Repeat for 103 and 104 dilutions.

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17. Working from the most dilute to the most concentrated dilution, briefly vortex and pipette three separate 10 μL drops onto the agar plate. Repeat for all other dilutions. 18. Allow the plate to dry at room temperature for approximately 20 min or until the droplets are no longer visible on the agar plate. 19. Incubate plates inverted in a 37
C incubator overnight. 20. Count colonies for each dilution. This will quantitate all cellassociated bacteria. To determine CFU/mL or total number of internalized bacteria, multiply the number of colonies by the inverse of the dilution factor and by 102, to account for the 10 μL plating volume. For example, 17 colonies counted on a 103 droplet would be calculated as 17  103 ðdilution factorÞ  102 ðplating volumeÞ ¼ 1:7  106 CFU=mL ðsee Fig:1Þ: 3.2 Gentamicin Protection Assay for Bacterial Phagocytosis

Carry out all procedures under sterile conditions in a biological safety cabinet. This protocol has been optimized for use with Streptococcus pneumoniae but can be performed with other bacteria. Ensure that the bacteria are susceptible to this concentration of gentamicin prior to beginning the experiment (see Note 2). 1. Grow bacteria to early or mid-log phase in liquid culture (see Note 1). While waiting, prepare and label tubes containing 900 μL sterile water and TSB/Sheep’s blood plates to be used in Steps 16 and 18. 2. When the liquid bacterial culture reaches early to mid-log growth (as measured by OD600), remove 1 mL into a 2 mL microcentrifuge tube and centrifuge at 12,000  g for 1 min. For S. pneumoniae, early log phase (OD600 ¼ 0.5) is generally obtained from 1 to 4 h when prepared from a 1 mL freezer stock. Carefully remove the supernatant and resuspend the pellet in 1 mL HBSS. Keep on ice until use. 3. Visually inspect macrophage culture under a light microscope to ensure high cell viability and the culture is free of contamination. Macrophages typically have a dendritic-like morphology and should be strongly adherent. 4. Remove macrophage culture media and gently wash with warm PBS. 5. Remove PBS and add 10 mL of warm Accutase cell detachment solution. Cover the plate and place in a 37
C incubator for 10 min. Visually inspect the plate under a light microscope to determine whether cells have detached. If cells still remain adherent, incubate for an additional 5–10 min.

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6. Gently scrape the plate with a cell lifter to remove any loosely adherent cells (see Note 6). Pipette Accutase solution containing macrophages into a 50 mL conical tube. To maximize macrophage recovery, wash the plate with 10 mL warm PBS and add to the 50 mL conical tube. 7. Centrifuge for 10 min at  500 g. 8. Carefully remove all supernatant and gently resuspend the pellet in 1 mL of warm PBS. Do not vortex. 9. Determine the CFU/mL by diluting cells 1:1 with Trypan Blue, and pipette 10 μL into a hemocytometer. Count viable cells following standard usage of a hemocytometer. 10. Add up to 1  106 viable macrophages into a 2 mL microcentrifuge tube. Bring the volume of solution up to 900 μL by adding PBS. Repeat for each desired time point. For example, if performing time points of 0, 30, 60, and 90 min, you will require four tubes containing equal numbers of macrophages (see Note 8). 11. Briefly vortex bacteria and add the desired multiplicity of infection (MOI) to each tube (see Fig. 2). The MOI will vary depending on the bacteria used in the assay and number/type of macrophages. In the examples in Figs. 2 and 3, Streptococcus pneumoniae was added at an MOI of 20. The final volume of the macrophage/bacteria solution should be brought to 1 mL with PBS. 12. Incubate with gentle mixing at 37
C for 30 min to allow for bacterial binding and internalization (see Fig. 3). After this preincubation, at time points +0, +20, +40, and +60 min, remove the tube and centrifuge at 500  g for 5 min. 13. Carefully remove the supernatant to remove unbound bacteria, and gently resuspend in 1 mL HBSS + 50 μg/mL gentamicin. Incubate with gentle mixing at 37
C for 40 min (see Note 2). 14. Add 1 mL HBSS and centrifuge at 500  g for 5 min. Wash cells two additional times with 1 mL HBSS to completely remove gentamicin. 15. Following the final wash, resuspend in 1 mL HBSS, and lyse macrophages by pipetting 100 μL of the macrophage solution into 900 μL sterile water (prepared in Step 1) (see Note 9). Briefly vortex. This tube is now a 101 dilution. 16. Perform a serial dilution by repeating Step 16 using 100 μL of the 101 dilution into 900 μL sterile water. Briefly vortex. This tube is now a 102 dilution. Repeat for 103 and 104 dilutions.

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17. Working from the most dilute to the most concentrated dilution, briefly vortex and pipette three separate 10 μL drops onto the agar plate. Repeat for all other dilutions. 18. Allow the plate to dry at room temperature for approximately 20 min or until the droplets are no longer visible on the agar plate. 19. Incubate plates inverted in a 37
C incubator overnight. 20. Count colonies for each dilution. To determine CFU/mL or total number of internalized bacteria, multiply the number of colonies by the inverse of the dilution factor and by 102, to account for plating volume. For example, 17 colonies counted on a 103 droplet would be calculated as 17  103 ðdilution factorÞ  102 ðplating volumeÞ ¼ 1:7  106 CFU=mL ðsee Fig:1Þ: 3.3 Particle Binding Assay

This adapted protocol uses inert fluorescent particles such as polystyrene microspheres in place of bacteria. When using inert particles, attention should be given to particle size since some cell lines and phagocytes cannot internalize particles larger than 1–2 μm. In general, particles greater than 0.5 μm are internalized by phagocytosis, and those smaller than 0.5 μm are internalized by endocytosis [12]. Particles may be coated with ligands, such as bacterial cell wall components or antibodies either by passive adsorption or covalent coupling. Finally, it is advisable to perform a dose-response experiment to determine optimal particle/phagocyte ratios. 1. Visually inspect macrophage culture under a light microscope to ensure high cell viability and the culture is free of contamination. Macrophages typically have a dendritic-like morphology and should be strongly adherent. 2. Remove macrophage culture media and gently wash with warm PBS. 3. Remove PBS and add 10 mL of warm Accutase cell detachment solution. Cover the plate and place in a 37
C incubator for 10 min. Visually inspect the plate under a light microscope to determine whether cells have detached. If cells still remain adherent, incubate for an additional 5–10 min. 4. Gently scrape the plate with a cell lifter to remove any loosely adherent cells (see Note 6). Pipette Accutase solution containing macrophages into a 50 mL conical tube. To maximize macrophage recovery, wash the plate with 10 mL warm PBS and add to the 50 mL conical tube. 5. Centrifuge for 10 min at  500 g. 6. Carefully remove all supernatant and gently resuspend the pellet in 1 mL of warm PBS. Do not vortex.

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7. Determine the CFU/mL by diluting cells 1:1 with Trypan Blue, and pipette 10 μL into a hemocytometer. Count viable cells following standard usage of a hemocytometer. 8. Add up to 1  106 viable macrophages into a 2 mL microcentrifuge tube. Bring the volume of solution up to 900 μL by adding PBS. Repeat for each desired time point. For example, if performing time points of 0, 30, 60, and 90 min, you will require four tubes containing equal numbers of macrophages (see Note 8). 9. Vortex particles and add at the desired particle/macrophage ratio, and bring the total volume up to 1 mL with either PBS or cell culture media. 10. Incubate on with gentle agitation at 4
C for 1 h. At time points 0, 20, 40, and 60 min, remove the tube and centrifuge at 500  g for 5 min. 11. Carefully remove the supernatant to remove unbound particles and gently resuspend in 1 mL PBS or media. Centrifuge at 500  g for 5 min. Repeat wash twice more. 12. Resuspend in 200 μL PBS or media. 13. Add suspension to 1 well of a black 96-well plate. Read at the excitation and emission recommended by the particle manufacturer. 3.4 Bacterial Killing Assay: Macrophage

Carry out all procedures under sterile conditions in a biological safety cabinet. This protocol has been optimized for use with Streptococcus pneumoniae but can be performed with other bacteria. Please see additional notes for assay modifications. 1. Grow bacteria to early or mid-log phase in liquid culture (see Note 1). While waiting, prepare and label tubes containing 900 μL sterile water and TSB/Sheep’s blood plates to be used in Steps 16 and 18 (see Fig. 1 for experimental schematic). 2. When the liquid bacterial culture reaches the desired OD600, remove 1 mL into a 2 mL microcentrifuge tube, and centrifuge at 12,000  g for 1 min. 3. Carefully remove the supernatant and resuspend the pellet in 1 mL HBSS. Keep on ice to prevent additional bacterial growth. 4. Visually inspect the macrophage culture under a light microscope to ensure high cell viability and that the culture is free of contamination. Macrophages typically have a dendritic-like morphology and should be strongly adherent. 5. Remove macrophage culture media and gently wash with warm PBS. 6. Remove PBS and add 10 mL of warm Accutase cell detachment solution. Cover the plate and place in a 37
C incubator for

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10 min. Visually inspect the plate under a light microscope to determine cell detachment. Some cells should be completely detached, and the remaining adherent cells should be round in morphology. If cells still remain adherent, incubate for an additional 5–10 min. 7. Gently scrape the plate with a cell lifter, covering all surface area (see Note 6). Pipette Accutase solution containing macrophages into a 50 mL conical tube. To maximize macrophage recovery, wash the plate with 10 mL warm PBS and add to the 50 mL conical tube. 8. Centrifuge for 10 min at 500  g. 9. Carefully remove all supernatant and gently resuspend the pellet in 1 mL warm PBS. Do not vortex. 10. Determine the CFU/mL by diluting cells 1:1 with Trypan Blue, and pipette 10 μL into a hemocytometer. Count viable cells following standard usage of a hemocytometer. 11. Add up to 1  106 viable macrophages into a 2 mL microcentrifuge tube. Bring the volume of solution to 900 μL by adding PBS. 12. Briefly vortex bacteria and add the desired multiplicity of infection (MOI) to each tube (see Fig. 2). The MOI will vary depending on the bacteria used in the assay and number/type of macrophages. In the examples in Figs. 2 and 3, Streptococcus pneumoniae was added at an MOI of 20. The final volume of the macrophage/bacteria solution should be brought to 1 mL with PBS. 13. Incubate with gentle mixing at 37
C for 30 min to allow for bacterial binding and internalization (see Fig. 3). After this preincubation, remove the tube and centrifuge at 500  g for 5 min. 14. Carefully remove the supernatant to remove unbound bacteria, and gently resuspend in 1 mL HBSS. 15. Wash cells one additional time with 1 mL HBSS to completely remove unbound bacteria. 16. Following the final wash, resuspend in 1 mL HBSS. 17. Lyse macrophages by pipetting 100 μL of the macrophage solution into 900 μL sterile water (prepared in Step 1) (see Note 9). Briefly vortex. This tube is now a 101 dilution at time point +0 min. Return the original tube containing macrophages and bound/internalized bacteria to the mixer at 37
C for time points of +30, +60 and +90, and +120 and +180 min. 18. Perform a serial dilution using 100 μL of the 101 dilution into 900 μL sterile water for each time point. Briefly vortex. This tube is now a 102 dilution. Repeat for 103 and 104 dilutions.

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19. Working from the most dilute to the most concentrated dilution, briefly vortex and pipette three separate 10 μL drops onto the agar plate. Repeat for all other dilutions. 20. Allow the plate to dry at room temperature for approximately 20 min or until the droplets are no longer visible on the agar plate. 21. Repeat Steps 17–20 for time points of +30, +60 and +90, and +120 and +180 min. 22. Incubate plates inverted in a 37
C incubator overnight. 23. Count colonies for each dilution. To determine CFU/mL or total number of internalized bacteria, multiply the number of colonies by the inverse of the dilution factor and by 102, to account for plating volume. For example, 17 colonies counted on a 103 droplet would be calculated as 17  103 ðdilution factorÞ  102 ðplating volumeÞ ¼ 1:7  106 CFU=mL ðsee Fig:1Þ:

4

Notes 1. Accurate quantitation of the MOI is essential to ensure reproducibility. Therefore, it is critical to perform a growth curve for the strain of bacteria to be used in the assay in order to prepare the assay with an accurate MOI. Determine the CFU/mL when the culture is analyzed via a spectrophotometer at OD600. 2. The sensitivity of a given strain of bacteria to gentamicin should be titrated by performing a standard kill curve procedure in the presence of varying doses of gentamicin at 10 min time points for approximately 1 h. 3. The type of liquid culture media, agar plate, and incubation conditions can be modified depending on the strain of bacteria being used for the assay. 4. Permeability of the phagocytic cells should be given consideration when performing a gentamicin protection assay, as gentamicin can penetrate permeabilized membranes and kill internalized bacteria. This can occur, for example, when cells are forcefully washed or stored at incorrect temperatures. 5. Accutase is preferred over trypsin, as trypsin has been shown to cleave some phagocytic receptors. If assaying the relevance of specific receptors, it is advisable to treat cells with Accutase and check receptor expression via FACS or IF microscopy. Include a saline or untreated control. 6. Cell viability can be significantly enhanced by using cell lifters in place of cell scrapers.

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7. Before using this assay to quantify bacterial binding, ensure that the bacteria will survive at this temperature for the desired time course. To do so, grow bacteria to early or mid-log phase, perform a serial dilution, and plate droplets to determine CFU/mL. Then incubate bacteria at 4
C. At desired the time point(s), remove bacteria, perform serial dilutions, and plate. Following overnight incubation at optimal growth conditions for the desired bacteria, compare initial CFU/mL to CFU/mL at each time point. 8. A minimum of 1  105 macrophages are required to perform this assay. In our experience using 1  106 macrophages per condition increases reproducibility. 9. Bacteria should be titrated using serial dilutions in PBS, plated on appropriate solid agar medium to count CFUs. 10. Lysis of macrophages ensures that bacterial killing is immediately halted; however, plating should be performed immediately to ensure that bacterial viability doesn’t decrease. References 1. Cohn ZA, Morse SI (1959) Interactions between rabbit polymorphonuclear leucocytes and staphylococci. J Exp Med 110:419–443 2. Fox EN (1974) M proteins of group A streptococci. Bacteriol Rev 38:57–86 3. Cho K, Arimoto T, Igarashi T, Yamamoto M (2013) Involvement of lipoprotein PpiA of Streptococcus gordonii in evasion of phagocytosis by macrophages. Mol Oral Microbiol 28: 379–391. https://doi.org/10.1111/omi. 12031 4. Mukouhara T, Arimoto T, Cho K et al (2011) Surface lipoprotein PpiA of Streptococcus mutans suppresses scavenger receptor MARCO-dependent phagocytosis by macrophages. Infect Immun 79:4933–4940. https://doi.org/10.1128/IAI.05693-11 5. Dilworth JA, Hendley JO, Mandell GL (1975) Attachment and ingestion of gonococci by human neutrophils. Infect Immun 11:512– 516 6. Campbell PA, Canono BP, Drevets DA (2001) Measurement of bacterial ingestion and killing by macrophages. Curr Protoc Immunol Chapter 14:Unit 14.6. https://doi.org/10. 1002/0471142735.im1406s12 7. Underhill DM, Ozinsky A (2002) Phagocytosis of microbes: complexity in action. Annu Rev Immunol 20:825–852. https://doi.org/10. 1146/annurev.immunol.20.103001.114744

8. Dorrington MG, Roche AM, Chauvin SE et al (2013) MARCO is required for TLR2- and Nod2-mediated responses to Streptococcus pneumoniae and clearance of pneumococcal colonization in the murine nasopharynx. J Immunol 190:250–258. https://doi.org/10. 4049/jimmunol.1202113 9. Puchta A, Naidoo A, Verschoor CP et al (2016) TNF drives monocyte dysfunction with age and results in impaired antipneumococcal immunity. PLoS Pathog 12: e1005368. https://doi.org/10.1371/journal. ppat.1005368 10. Verschoor CP, Johnstone J, Loeb M et al (2014) Anti-pneumococcal deficits of monocyte-derived macrophages from the advanced-age, frail elderly and related impairments in PI3K-AKT signaling. Hum Immunol 75:1192–1196. https://doi.org/10.1016/j. humimm.2014.10.004 11. Perry JA, Koteva K, Verschoor CP et al (2015) A macrophage-stimulating compound from a screen of microbial natural products. J Antibiot (Tokyo) 68:40–46. https://doi.org/10.1038/ ja.2014.83 12. Mellman I (1996) Endocytosis and molecular sorting. Annu Rev Cell Dev Biol 12:575–625. https://doi.org/10.1146/annurev.cellbio.12. 1.575

Chapter 2 Analysis of Human and Mouse Neutrophil Phagocytosis by Flow Cytometry Noah Fine, Oriyah Barzilay, and Michael Glogauer Abstract Neutrophils are primary phagocytes that recognize their targets through surface chemistry, either through pattern recognition receptor (PPR) interaction with pathogen-associated molecular patterns (PAMPs) or through immunoglobulin (Ig) or complement mediated recognition. Opsonization can be important for target recognition and phagocytosis by neutrophils. Therefore, phagocytosis assays performed using neutrophils in whole blood, compared to isolated cells, will differ due to the presence of opsonizing blood serum components, as well as other blood components like platelets. Powerful and sensitive flow cytometry-based methods are presented to measure phagocytosis by human blood neutrophils and mouse peritoneal neutrophils. Key words Neutrophil, Phagocytosis, pHrodo, Peritonitis, Flow cytometry

1

Introduction Ingestion of inflammatory targets by neutrophils is an essential aspect of the innate immune response [1–3]. Sensitive measurement of this phenomenon is essential for a thorough understanding of the molecular and physiological mechanisms that control this process. pHrodo™ BioParticles® conjugates from Invitrogen allow for sensitive measurement of phagocytosis by flow cytometry [4]. pHrodo™ becomes highly fluorescent when exposed to acidic pH after internalization, and the process can be measured by conventional detection modalities including fluorescence imaging microscopy, fluorescent microplate reader, and flow cytometry. Here we describe protocols for flow cytometric analysis of phagocytosis using pHrodo™ Green E. coli BioParticles® conjugates. This method is easily adapted for use of zymosan or S. aureus bioparticle conjugates, and pHrodo™ Red conjugates, which are also available.

Roberto J. Botelho (ed.), Phagocytosis and Phagosomes: Methods and Protocols, Methods in Molecular Biology, vol. 2692, https://doi.org/10.1007/978-1-0716-3338-0_2, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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Neutrophils are among the fastest moving cells in the body, and their responses to inflammatory stimuli are nearly instantaneous. These characteristics, which are appropriate for the effectiveness of neutrophils in combating microbes, are detrimental with respect to the in vitro manipulation of these cells for experimental purposes. Because neutrophils can be easily activated by experimental manipulation [5–11], it is desirable to keep these interactions to a minimum if possible. We have developed a protocol whereby neutrophils in whole human blood or peritoneally elicited mouse neutrophils are fluorescently labeled with pHrodo™ BioParticles® and fixed in order to analyze phagocytosis by flow cytometry with a minimum amount of manipulation. For the in vitro blood neutrophil assay, the endpoint of the phagocytosis reaction is achieved by fixing the samples with paraformaldehyde (PFA), and samples are subsequently labeled for flow cytometric analysis. In the mouse, pHrodo™ BioParticles® are used to induce peritonitis, and neutrophils are recovered and immediately fixed with PFA. A combination of CD16 and SSC-A is sufficient to gate on neutrophils in whole human blood (unpublished results), while Ly6G+ve/F4/ 80-ve gating is used for mouse neutrophils. In addition to being simpler and faster than conventional approaches, we have found that our approach minimizes the activation of neutrophils that would otherwise occur during the purification of neutrophils from blood. Furthermore, fixation of neutrophils does not interfere with subsequent antibody labeling and multicolor flow cytometry analysis. Our protocol focuses on sample acquisition, preparation, labeling, flow cytometric acquisition, and analysis.

2

Materials 1. pHrodo™ Green E. coli BioParticles® conjugates from Invitrogen. 2. Phosphate-buffered saline (PBS): 137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 1.8 mM KH2PO4, pH 7.4. 3. Red blood cell lysis buffer such as BD Pharm lyse solution. 4. Hanks buffered salt solution (HBSS): 138 mM NaCl, 5.3 mM KCl, 0.34 mM Na2HPO4, 0.44 mM KH2PO4, 4.2 mM NaHCO3, 5.6 mM glucose, pH 7.4. 5. FACS buffer: HBSS with 2 mM EDTA, 1% BSA. 6. Vacutainers containing sodium citrate. 7. 16% PFA ampoule. 8. Rat serum: 1 mg/mL. 9. Mouse IgG: 1 mg/mL. 10. Mouse anti-human CD16-PE (Clone: 3G8).

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11. Mouse anti-human CD66a-APC (Clone: CD66a-B1.1). 12. Anti-mouse Ly6G-PerCP-Cy5.5 (Clone: 1A8). 13. Anti-mouse F4/80-BV421 (Clone: BM8). 14. Flow cytometry compensation beads (e.g., OneComp eBeads from eBiosciences). 15. Sonicator. 16. Coulter counter or hemocytometer. 17. Wild-type C57 black 6 mice (8–16 weeks old).

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Methods Unless noted otherwise, all samples are maintained on ice and centrifugations are at 4 °C.

3.1 Phagocytosis by Human Blood Neutrophils

1. Resuspend pHrodo™ fluorescent bioparticles (2 mg lyophilized per vial) in 2 mL of sterile PBS. 2. Sonicate on ice for 5 min in a glass tube (see Note 1).

3.1.1 Preparation of pHrodo Particles 3.1.2 Acquisition of Blood Samples

1. Blood should be drawn by a trained phlebotomist (see Note 2). 2. Blood samples are drawn from the median cubital vein in the crook of the elbow into a vacutainer containing 0.1 volumes of sodium citrate anticoagulant. 3. Mix the blood by gentle inversion and maintain at room temperature (see Note 3).

3.1.3 Blood Neutrophil Stimulation

1. Aliquot 50 μL of blood into flow cytometry tubes. 2. Add pHrodo™ (0.5, 2, 10 μL) or vehicle control to the blood and vortex the tubes gently to mix (Fig. 1). 3. Cover the tubes with paraffin wax and incubate them at 37 °C for the appropriate amount of time (see Note 4). 4. Add 1/10th volume of PFA (1.6% final) to each tube to stop the phagocytosis reaction (see Note 5). 5. Gently vortex each tube and incubate them on ice for 15 min. 6. Add 1 mL of PBS to dilute the fixative. 7. Centrifuge the tubes for 5 min at 1000 g at 4 °C. 8. Decant the supernatant. 9. Lyse the red blood cells (RBCs) by resuspending each pellet in 1 mL of red blood cell lysis buffer. 10. Incubate the tubes on ice for 15 min.

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Fig. 1 Phagocytosis by human blood neutrophils. (a) Cells passing through the cytometer in pairs (doublets) were excluded using side scatter height (SSC-H) × side scatter width (SSC-W), and neutrophils were gated in whole blood based on high expression of CD16 and CD66a. (b) Three dosages of pHrodo were added to 50 μL of human blood and incubated at 37 °C for the indicated times. At least 2 × 104 gated neutrophil events were acquired

11. Centrifuge the tubes for 5 min at 1000 g at 4 °C. 12. Decant the supernatant. 13. Repeat the RBC lysis steps (Steps 9–12) until pellets are white (see Note 6). 14. Resuspend the pellets in 1 mL of PBS. 15. Use a Coulter counter or hemocytometer to count the cells (see Note 7). 3.1.4 Fluorescent Labeling

1. Resuspend the cells in FACS buffer so that the final volume, including antibody, will be 50 μL. 2. To block Fc-receptors, add 1 μL rat serum and 2 μL mouse IgG to each tube, vortex gently, and incubate on ice for 20 min. 3. Prepare the mastermix of fluorescently conjugated antibodies (see Notes 8 and 9). 4. Do not use FITC-conjugated antibodies, since this is the same fluorescent channel as pHrodo Green bioparticles.

Analysis of Phagocytosis by Flow Cytometry

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5. Add the antibody mastermix, vortex gently, and incubate for 30 min on ice protected from light. 6. Wash each pellet with 1 mL of FACS buffer. 7. Vortex briefly and pellet the cells at 1000 g for 5 min at 4 °C. 8. Decant the supernatant. 9. Repeat the wash steps (steps 6–8) two additional times. 10. Resuspend the pellets in 250 μL of FACS buffer and vortex briefly. 11. Prepare single stained compensation beads controls according to manufacturers’ instructions. 12. Resuspend labeled compensation beads in 400 μL of FACS buffer and vortex briefly. 13. Cover the tubes with paraffin wax, and store in the fridge protected from the light. 3.1.5

Sample Acquisition

1. Perform instrument setup according to the protocols of your flow cytometry facility. 2. Vortex the first sample and load onto the cytometer. 3. Adjust the forward scatter (FSC) and side scatter (SSC) settings so that the population of interest (neutrophils) is displayed in the center of the scatterplot. 4. Ensure fluorescent signals are on scale. 5. Perform automated compensation using single stained beads controls and pHrodo-labeled cells. 6. Run each sample and acquire at least 2 × 104 gated events. 7. Export FCS files. 8. Data can be analyzed using FlowJo or other commercially available software.

3.2 Mouse Peritoneal Recruitment and Phagocytosis 3.2.1 Preparation of pHrodo Particles and Intraperitoneal Injection 3.2.2

Peritoneal Lavage

1. Resuspend pHrodo™ fluorescent bioparticles (2 mg lyophilized per vial) in 2 mL of sterile PBS. 2. Sonicate on ice for 5 min in a glass tube. 3. Inject 100 μL into the peritoneum using a 0.5cc syringe and a 25–27 gauge needle (see Note 10). 4. Inject between the midline and the right knee. 1. Carefully inject 3 mL of cold 1× PBS into the peritoneum using a 30 gauge needle to recover the cellular infiltrate from the peritoneal cavity (see Note 11), as follows. 2. Euthanize the mouse, and sterilize with ethanol. 3. Cut the skin to expose the peritoneal cavity, pinning down the skin to the sides.

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4. Gently massage the peritoneum for 40 s. 5. Carefully draw out up to 2.5 mL of peritoneal lavage using the same needle and syringe, and place it in a 15 mL conical centrifuge tube on ice. 3.2.3

Sample Fixation

1. Add PFA to the lavage solution at a final concentration of 1.6%. 2. Vortex each tube briefly and allow the cells to fix on ice for 15 min. 3. Top up the conical centrifuge tubes with cold 1×PBS to dilute the PFA, and centrifuge for 5 min at 1000 g. 4. Discard the supernatant. 5. If red blood cells are present, resuspend the pellet in 1 mL of 1× BD Pharm lyse solution. 6. Incubate for 5 min on ice. 7. Repeat the centrifugation step. 8. Resuspend the pellets in 1 mL of PBS. 9. Use a Coulter counter or hemocytometer to count the cells.

3.2.4

Labeling

1. Resuspend 0.5 × 106 cells in FACS buffer so that the final volume, including antibody, will be 50 μL, and aliquot to flow cytometry tubes. 2. To block the Fc-receptors, add 1 μL of rat serum and 2 μL of mouse IgG, and incubate for 20 min on ice. 3. Prepare a mastermix of fluorescently conjugated antibodies (see Notes 9 and 12). 4. Do not use FITC-conjugated antibodies, since this is the same fluorescent channel as pHrodo Green bioparticles. 5. Add the antibody mastermix, vortex gently, and incubate for 30 min on ice protected from light. 6. To wash, add 1 mL of FACS buffer. 7. Vortex briefly and pellet the cells at 1000 g for 5 min at 4 °C. 8. Decant the supernatant. 9. Repeat the wash steps (steps 6–8) two more times. 10. Resuspend cells in 250 μL of FACS buffer and vortex briefly. 11. Prepare single stained compensation beads controls according to manufacturers’ instructions. 12. Resuspend labeled compensation beads in 400 μL of FACS buffer and vortex briefly. 13. Cover the tubes with paraffin wax and store in the fridge protected from the light. 14. Perform flow cytometric acquisition and analysis as outlined in Subheading 3.1.5.

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Notes 1. Excess unused pHrodo™ can be frozen at -20 °C in aliquots; however, repeat freeze thawing was not tested. 2. Human studies need to obtain institutional Research Ethics Board (REB) approval to ensure safety and ethical considerations. Prior to inclusion, signed informed consent should be obtained from all participants. 3. Fresh blood is aliquoted immediately at room temperature and is not maintained on ice prior to stimulation at 37 °C. 4. Some degree of neutrophil stimulation occurs at 37 °C, and therefore we recommend a 20 min stimulation, or less, depending on the dosage of pHrodo™ bioparticles (Fig. 1b). This is sufficient to observe phagocytosis of E. coli bioparticles while minimizing metabolic activation of neutrophils at 37 °C. 5. One limitation of our approach is that flow cytometric assessment and exclusion of dead cells is not possible, since all of the neutrophils are dead after fixation with PFA. However, by cytological staining of human blood leukocytes and mouse peritoneal neutrophils, we found that only a small percentage (95% QE) camera or high-sensitivity confocal detector (e.g., GaAsP-PMT). 17. If the live-cell imaging chamber lacks CO2 perfusion, use RPMI 1640 or DMEM buffered with HEPES, in place of sodium bicarbonate, in order to maintain physiological pH in air. 18. Due to the heterogeneous nature of cellular responses, it is important to collect microscopy data in a manner consisted with good statistical practices. Sufficient replicates (individually imaged cells) and repeats (independent experiments) should be collected to ensure statistically representative samples [27]. Images should be analyzed by applying quantitative

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measures of efferocytic events (Subheading 3.3.4) and/or colocalization [28], followed by statistical analysis of the resulting data with the appropriate statistical test [29]. As with other data types, it is important to select a statistical test compatible with the number of experimental groups (single-group: t-Test or Mann-Whitney U test; multigroup: ANOVA or KruskalWallis test) and the distribution of the data (parametric: t-test or ANOVA; nonparametric: Mann–Whitney U test or KruskalWallis test). The application of a robust statistical approach to image analysis ensures that these data support objective conclusions based on reproducible and statistically validated results. 19. During staining the targets with streptavidin, anything exposed to the extracellular space will be stained, and anything inside the cell is inaccessible to staining. This will differentiate between targets that are entirely engulfed versus targets that are only tethered to the surface of the cell or partially engulfed. 20. Lipid solution must be kept in a tube flushed with an inert gas such as nitrogen to prevent lipid oxidation. Oxidation of lipids might engage scavenger receptor-mediated phagocytosis.

Acknowledgments This study was funded by a Canadian Institutes for Health Research (CIHR) Project Grant to BH. The funding agencies had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript. References 1. Yin C, Heit B (2021) Cellular responses to the efferocytosis of apoptotic cells. Front Immunol 12:631714 2. Santavanond JP, Rutter SF, Atkin-Smith GK et al (2021) Apoptotic bodies: mechanism of formation, isolation and functional relevance. Subcell Biochem 97:61–88 3. Wickman G, Julian L, Olson MF (2012) How apoptotic cells aid in the removal of their own cold dead bodies. Cell Death Differ 19:735– 742 4. Mai F-Y, He P, Ye J-Z et al (2019) Caspase-3mediated GSDME activation contributes to cisplatin- and doxorubicin-induced secondary necrosis in mouse macrophages. Cell Prolif 52: e12663

5. Cai B, Thorp EB, Doran AC et al (2017) MerTK receptor cleavage promotes plaque necrosis and defective resolution in atherosclerosis. J Clin Invest 127:1–5 6. Ismail OZ, Zhang X, Bonventre JV et al (2016) G protein α12(Gα12) is a negative regulator of kidney injury molecule-1-mediated efferocytosis. Am J Physiol Renal Physiol 310:F607– F620 7. Yin C, Vrieze AM, Rosoga M et al (2020) Efferocytic defects in early atherosclerosis are driven by GATA2 overexpression in macrophages. Front Immunol 11:594136 8. Yoon Y-S, Kim S-Y, Kim M-J et al (2015) PPARγ activation following apoptotic cell instillation promotes resolution of lung

Efferocytosis Assays inflammation and fibrosis via regulation of efferocytosis and proresolving cytokines. Mucosal Immunol 8:1031–1046 9. Peter C, Waibel M, Radu CG et al (2008) Migration to apoptotic “find-me” signals is mediated via the phagocyte receptor G2A. J Biol Chem 283:5296–5305 10. Suzuki J, Imanishi E, Nagata S (2016) Xkr8 phospholipid scrambling complex in apoptotic phosphatidylserine exposure. Proc Natl Acad Sci U S A 113:9509–9514 11. Segawa K, Kurata S, Yanagihashi Y et al (2014) Caspase-mediated cleavage of phospholipid flippase for apoptotic phosphatidylserine exposure. Science 344:1164–1168 12. Kinchen JM, Doukoumetzidis K, Almendinger J et al (2008) A pathway for phagosome maturation during engulfment of apoptotic cells. Nat Cell Biol 10:556–566 13. Yin C, Argintaru D, Heit B (2017) Rab17 mediates intermixing of phagocytosed apoptotic cells with recycling endosomes. Small GTPases 0:1–9 14. Ichimura T, Asseldonk EJPV, Humphreys BD et al (2008) Kidney injury molecule-1 is a phosphatidylserine receptor that confers a phagocytic phenotype on epithelial cells. J Clin Invest 118:1657–1668 15. Linger RMA, Keating AK, Earp HS et al (2008) TAM receptor tyrosine kinases: biologic functions, signaling, and potential therapeutic targeting in human cancer. Adv Cancer Res 100:35–83 16. Blackburn JWD, Lau DHC, Liu EY et al (2019) Soluble CD93 is an apoptotic cell opsonin recognized by αx β2. Eur J Immunol 49: 600–610 17. Finnemann SC, Nandrot EF (2006) MerTK activation during RPE phagocytosis in vivo requires alphaVbeta5 integrin. Adv Exp Med Biol 572:499–503 18. Rigotti A, Acton SL, Krieger M (1995) The class B scavenger receptors SR-BI and CD36 are receptors for anionic phospholipids. J Biol Chem 270:16221–16224

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19. Park D, Tosello-Trampont A-C, Elliott MR et al (2007) BAI1 is an engulfment receptor for apoptotic cells upstream of the ELMO/ Dock180/Rac module. Nature 450:430–434 20. Nandrot EF, Anand M, Almeida D et al (2007) Essential role for MFG-E8 as ligand for alphavbeta5 integrin in diurnal retinal phagocytosis. Proc Natl Acad Sci U S A 104:12005–12010 21. Evans AL, Blackburn JWD, Yin C et al (2017) Quantitative efferocytosis assays. Methods Mol Biol 1519:25–41 22. Ravichandran KS (2010) Find-me and eat-me signals in apoptotic cell clearance: progress and conundrums. J Exp Med 207:1807–1817 23. Leffell MS, Spitznagel JK (1975) Fate of human lactoferrin and myeloperoxidase in phagocytizing human neutrophils: effects of immunoglobulin G subclasses and immune complexes coated on latex beads. Infect Immun 12:813–820 24. Yeung T, Heit B, Dubuisson J-F et al (2009) Contribution of phosphatidylserine to membrane surface charge and protein targeting during phagosome maturation. J Cell Biol 185: 917–928 25. Flannagan RS, Harrison RE, Yip CM et al (2010) Dynamic macrophage “probing” is required for the efficient capture of phagocytic targets. J Cell Biol 191:1205–1218 26. Zhou D, Huang C, Lin Z et al (2014) Macrophage polarization and function with emphasis on the evolving roles of coordinated regulation of cellular signaling pathways. Cell Signal 26: 192–197 27. Vaux DL, Fidler F, Cumming G (2012) Replicates and repeats-what is the difference and is it significant? A brief discussion of statistics and experimental design. EMBO Rep 13:291–296 28. Bolte S, Cordelie`res FP (2006) A guided tour into subcellular colocalization analysis in light microscopy. J Microsc 224:213–232 29. Nayak BK, Hazra A (2011) How to choose the right statistical test? Int J Opthalmol 59: 85–86

Chapter 5 Analysis of Efferocytic Receptor Dynamics and Synapse Formation in a Frustrated Efferocytosis Model Brandon H. Dickson and Bryan Heit Abstract Efferocytes express multiple receptors that mediate the recognition and engulfment of apoptotic cells through a process known as efferocytosis. Ligation of these receptors induces the formation of a structured efferocytic synapse that mediates the engulfment of the apoptotic cell by the efferocyte. The lateral diffusion of these receptors allows for clustering-mediated receptor activation and is central for the formation of the efferocytic synapse. This chapter describes a single particle tracking protocol to analyze the diffusion of efferocytic receptors within a frustrated efferocytosis model. This enables high-resolution tracking of efferocytic receptors throughout synapse formation, allowing the user to simultaneously quantify synapse formation and the dynamics of receptor diffusion as the efferocytic synapse evolves. Key words Efferocytosis, Phagocytosis, Apoptosis, Single particle tracking, Frustrated efferocytosis, Diffusion

Abbreviations FBS PBS PE PMA PSF PtdCho PtdSer RPMI SLB SPT

Fetal bovine serum phosphate buffered saline phosphatidylethanolamine phorbol 12-myristate 13-acetate point-spread function phosphatidylcholine phosphatidylserine Roswell Park Memorial Institute Medium supported lipid bilayer single particle tracking

Roberto J. Botelho (ed.), Phagocytosis and Phagosomes: Methods and Protocols, Methods in Molecular Biology, vol. 2692, https://doi.org/10.1007/978-1-0716-3338-0_5, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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Introduction Efferocytosis is the phagocytosis (engulfment and destruction) of apoptotic cells [1]. Cells that perform efferocytosis are termed efferocytes, with macrophages representing the primary or sole efferocyte in many tissues [2]. Efferocytosis is required to rapidly clear dying cells in order to prevent secondary necrosis and the resulting uncontrolled release of alarmins and autoantigens [3, 4]. The recognition of apoptotic cells by efferocytes begins during apoptosis, where caspase-induced cleavage of lipid transporters leads to the exposure of phosphatidylserine (PtdSer) on the outer leaflet of the plasma membrane [5]. PtdSer is the universal “eat me” signal for dying cells and is recognized (either directly or indirectly via opsonins) by a diverse array of efferocytic receptors including MERTK, αvβ5 integrin, Stabilin-2, TIM-1/TIM-4, RAGE, BAI1, and others [6]. While these receptors may have unique signaling mechanisms, they converge on cellular pathways which restructure the actin cytoskeleton to form the efferocytic synapse – a structured membrane protrusion that surrounds the dying cell to direct its internalization [7]. This synapse between the efferocyte and the apoptotic cell can be visualized at highresolution in two-dimensional space using a frustrated efferocytosis assay [8]. In this assay, a glass coverslip is functionalized with an apoptotic cell-mimicking supported lipid bilayer (SLB) containing 80% phosphatidylcholine (PtdCho) and 20% PtdSer. Efferocytes are “parachuted” onto this substrate, and fluorescence microscopy is used to visualize the resulting efferocytic synapse (Fig. 1). By forcing synapse formation into a lateral plane, this technique allows for the high-resolution imaging of the synapse, while the planar

Fig. 1 The efferocytic synapse. A macrophage stained for MERTK (red) and β2 integrin (cyan) performing frustrated efferocytosis on apoptotic cell-mimicking SLBs. (a) Early synapse structure (t = 4 min). (b) Late synapse structure (t = 16 min), which demonstrates significant cell spreading. Macrophages will not form synapses on SLBs that lack phosphatidylserine (not shown)

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nature of the substrate allows the synapse to expand to a large size, thereby revealing key insights into synapse organization, receptor localization, and protein-protein interactions during efferocytosis. These analyses can be combined with additional readouts of cell activity (e.g., expansion rate of the synapse or focal exocytosis within the synapse), signaling (e.g., recruitment of signaling reporters to within the synapse), and single-molecule imaging of protein diffusion within the synapse to provide further information on synapse structure, function, and dynamics. This article will focus on the use of the frustrated efferocytic synapse model for studying cell spreading and protein diffusion within the synapse, but this approach can easily be combined with phagocytic signaling reporters such as those discussed in Chapter 6, to better understand the spatiotemporal regulation of signaling within the synapse. A key characteristic of transmembrane proteins, including efferocytic receptors, is the regulation of their diffusion on the plasma membrane. While membrane proteins diffuse 10–100 times slower than predicted by Brownian models [9], it is possible to classify their movement as free, confined, or directed. During free diffusion, the receptor undergoes random-walk motion characterized by brief pauses (due to trapping between lipids or in other short-lived membrane structures) followed by motion in a random direction [10]. In confined diffusion, long-lasting diffusion barriers such as actin-based corrals compartmentalize diffusion into discrete microdomains [11]. In directed diffusion, the receptor undergoes active transport along a linear path [12]. Analyzing receptor diffusion can provide unique insights into specific cellular processes. For instance, immunoreceptors such as FcεRI are known to undergo clustering and loss of free diffusion upon ligand recognition, which may enhance its downstream signaling [13], while diffusion barriers prevent FcR-inactivating phosphatases from entering sites of active Fc-receptor signaling [14]. As the frustrated efferocytosis assay stratifies the synapse across two-dimensional space, the lateral diffusion of receptors can be analyzed as a function of their movement in the x- and y-axes. In this chapter, we provide a protocol where the diffusion of fluorescently labeled efferocytic receptors are analyzed during frustrated efferocytosis by an established single particle tracking (SPT) algorithm (Fig. 2). This protocol requires high-speed timelapses (≤100 ms frame intervals), captured at regular intervals throughout synapse formation, thereby providing measures of diffusion mode (free, confined, directed), diffusion coefficient of each mode, and confinement areas for each receptor at each timepoint. Thus, the overall diffusion of efferocytic receptors can be studied as a function of time, with changes in their diffusive behavior providing insight into the changes in receptor clustering, actin structuring, and other processes as the efferocytic synapse matures.

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Fig. 2 Frustrated efferocytosis assay. (a) General schematic of a frustrated efferocytosis assay combined with high-speed single-molecule tracking microscopy. (b) Simplified workflow for a single particle tracking experiment. (Diagram produced with BioRender)

This approach has both advantages and disadvantages. The most inherent drawback of frustrated efferocytosis is that it does not fully recapitulate efferocytosis in vivo, which involves the engulfment of a finite 3D particulate, rather than an “infinite” 2D plane [15]. Thus, the circular synapse that is observed during frustrated efferocytosis does not recapitulate the events that occur as the efferosome seals, nor can it account for any processes that may occur to allow for the engulfment of complex threedimensional shapes [16]. Despite these limitations, frustrated

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efferocytosis provides a high-resolution view of the efferocytic cup in-plane with the lateral axis of a microscope – thereby providing the maximum possible resolution of synapse formation. Moreover, the supported lipid bilayer is freely diffusive, thereby better replicating the lipid ligands present on apoptotic cells compared to the immobilized opsonins often used for these assays [17]. Thus, phenomena such as receptor immobilization can be trusted as being driven by efferocytic signaling, rather than an artifact caused by an immobile substrate. While this assay is demonstrated with MERTK on macrophages, any efferocytic receptor and cell type should be compatible with this experimental approach.

2

Materials

2.1 Cell Culture and THP-1 Differentiation

1. THP-1 human monocyte cell line. 2. Roswell Park Memorial Institute Medium (RPMI) 1640 cell culture media supplemented with 10% fetal bovine serum (FBS). 3. 100 μg/mL in DMSO.

phorbol

12-myristate

13-acetate

(PMA)

4. 12-well plate. 5. 18 mm diameter, #1.5 thickness circular coverslips. 6. 15 mL conical centrifuge tubes. 7. Cell culture incubator, 37 °C, 5% CO2. 8. Benchtop centrifuge with swinging buckets. 9. Cell counter. 2.2 Preparation of Supported Lipid Bilayers

1. 10 mg/mL 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine (PtdCho) in chloroform. 2. 10 mg/mL 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphatidylserine (PtdSer) in chloroform. 3. 10 mg/mL chloroform.

biotin-phosphatidylethanolamine

4. 25 μL glass Hamilton syringe. 5. Amber glass vials with plastic caps. 6. Chloroform. 7. Parafilm. 8. Nitrogen tank with low-pressure regulator. 9. Fridge (4 °C). 10. 1× phosphate buffered saline (PBS). 11. Vortex mixer.

(PE)

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12. Avanti mini extruder kit with 0.1 μm polycarbonate exclusion filters. 13. Hot plate. 14. 1.7 mL microcentrifuge tubes. 15. 12-well plate. 16. 18 mm diameter, #1.5 thickness circular coverslips. 17. Vacuum flask connected to pump. 18. Rubber stopper. 19. 0.75 mg/mL streptavidin in 50% glycerol (fluorophoreconjugated, e.g., Alexafluor-647-Streptavadin). 2.3

Immunostaining

1. Primary antibody against proteins of interest (e.g., antiMERTK), preferably primary-labeled with an appropriate fluorophore (see Notes 1 and 2). 2. Secondary Fab antibody labeled with appropriate fluorophore conjugated Fab. 3. 1× PBS. 4. Live-cell imaging medium: 25 mM HEPES, 23.8 mM NaHCO3, 150 mM NaCl, 5 mM KCl, 2 mM CaCl2•2H2O, pH 7.4 in ddH2O. 5. Fridge (10 °C).

2.4 Frustrated Efferocytosis and Image Acquisition

1. Fluorescence microscope with motorized stage and heating chamber. Single particle tracking works best with high-speed image capture by widefield microscopy. Confocal systems should be avoided as the pinhole aperture can distort the point-spread function (PSF) of the fluorophores [18]. Herein, we utilize a Leica DMI6000B equipped with a Sedat Quad Filterset 100×/1.4 NA oil immersion objective with an additional 1.6 magnification and a high-sensitivity CMOS and EM-CCD camera. 2. Accutase cell dissociation reagent. 3. Microcentrifuge. 4. 1.7 mL microcentrifuge tubes. 5. Pooled human serum. 6. Live-cell imaging chamber.

2.5 Image Analysis for Single Particle Tracking

1. ImageJ or Fiji [19, 20] downloaded from https://imagej.net/. 2. An appropriate single-molecule tracking software package. Many are available, but in this protocol, we use the algorithm by Jaqaman et al., which is run in MatLab [21]. 3. Microsoft Excel.

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Methods

3.1 Cell Culture and THP-1 Differentiation

1. Grow THP-1 monocytes to a density of 1 × 106 cells/mL in 5 mL of RPMI supplemented with 10% FBS in a cell culture incubator at 37 °C and 5% CO2. 2. To split THP-1 monocytes, spin the cells at 400 × g at 4 °C for 5 min. Resuspend in 5 mL of warmed RPMI + 10% FBS. Count the cells using a hemocytometer, and split to a density as low as 1 × 105 cells/mL. The cells should not be allowed to exceed a density of 2 × 106 cells/mL. 3. Harvest cells by centrifuging 5 mL of cells at 400 × g at 4 °C for 5 min in a 15 mL conical tube. Decant the supernatant and resuspend the cells in 5 mL of warmed RPMI containing 10% FBS. Count the cells using a hemocytometer. 4. During the centrifugation in step 3, aseptically place 18 mm circular coverslips into each well of a 12-well plate. 5. Add 2.5 × 105 cells into each well with a coverslip. Bring media volume up to 1 mL with warmed RPMI containing 10% FBS. 6. To differentiate THP-1 monocytes into macrophages, add 100 ng/mL PMA and incubate at 37 °C and 5% CO2 for 72 h.

3.2 Functionalization of Coverslips with Supported Lipid Bilayers

1. To create liposomes, use a 25 μL glass syringe, and transfer lipids to an amber glass vial. For apoptotic mimics, add 15 μL PtdCho, 4 μL PtdSer, and 3.75 μL Biotin-PE. For control mimics, add 19 μL PtdCho and 3.75 μL Biotin-PE (see Note 3). 2. Evaporate the chloroform from the vial using a gentle stream of N2 gas, slowly rotating the vial to ensure no residual chloroform remains. Evaporation is complete when a translucent streak appears at the bottom of the vial. 3. Flush the vial an additional 20 min with a slow flow of N2 gas to ensure evaporation of any residual chloroform. 4. Assemble the Avanti mini extruder apparatus with a 0.1 μm polycarbonate exclusion filter according to manufacturer’s instructions. Equilibrate the membrane with 1 mL of PBS with a gastight syringe. Let the apparatus warm up to 65 °C using a hotplate. 5. Add 400 μL of PBS to the glass vial containing the dried lipid mixture, and vortex at max speed for 60 s. Transfer this mixture to a clean microcentrifuge tube. 6. Draw up the mixture using the clean 1000 μL gastight syringe. To draw up residual solution, close the tube, invert, and then tap the tube on a flat surface to allow the solution to fall into the cap. Draw up remaining solution inside of the cap, avoiding air bubbles.

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7. Remove air bubbles by holding the syringe upright and tapping the sides of the syringe. Expel any air bubbles that rise to the top. 8. Insert both syringes into the mini extruder apparatus on the hotplate. Let the syringe equilibrate to 65 °C. 9. Pass the mixture through the membrane filter by slowly depressing the syringe containing the lipid. The mixture must be passed through the filter exactly 11 times (see Note 4). 10. Transfer the filtered liposome solution into a 1.7 mL microcentrifuge tube. Make a working solution in a new microcentrifuge tube by diluting 200 μL of the solution into 800 μL of PBS. Flush the tube with N2. Liposomes can be stored at 4 °C for up to 4 days. 11. On the day of the experiment, place 18 mm circular coverslips in a glass vacuum flask connected to a pump. Cap the flask with a silicone rubber stopper or glass plug, and evacuate the flask to 0.5 microns in diameter [1]. Phagocytosis is an important process that spans both the innate and adaptive immune responses to foreign particles, pathogens, and malignant cells, as well as the clearance of apoptotic cells and other debris during tissue homeostasis. Given their relative ubiquity across tissue types, macrophages require a high degree of morphological and cytoskeletal plasticity [2]. This plasticity manifests in the eponymous cellular function of macrophages: phagocytosis, as well as the process of macropinocytosis, allows macrophages to engulf particles and large gulps of fluid.

Roberto J. Botelho (ed.), Phagocytosis and Phagosomes: Methods and Protocols, Methods in Molecular Biology, vol. 2692, https://doi.org/10.1007/978-1-0716-3338-0_6, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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Adhesion and migration of monocytes and macrophages are regulated in part by specialized structures called podosomes [3]. Podosomes are actin-rich protrusive structures found on the ventral side of cells which facilitates the engagement of integrins with the extracellular matrix. Podosomes rely on many of the same proteins as focal adhesions including talin, vinculin, and Src family kinases; however, podosomes have specific requirements for the Arp2/3 complex activator WASP (Wiskott-Aldrich syndrome protein) and Tks5 (tyrosine kinase substrate with five SH3 domains) [4, 5]. Curiously, podosome-like structures have also been described in the advancing pseudopods during the formation of phagosomes which facilitates the activation of integrins to aid in robust and efficient phagocytosis [6]. Taken together, the results suggest that the ability to form actin-dependent adhesive protrusions is critical to many functions of monocytes, macrophages, and other myeloid cells. Imaging the actin cytoskeleton, actin-binding proteins, and other cytoskeletal and endocytic processes frequently involve capturing and characterizing transient and spatiotemporally discrete events. Depending on the motivation of a given experiment, different options are available to interrogate the biological system. Super-resolution techniques, such as stimulated emission depletion (STED), are excellent for achieving a high degree of spatial resolution, especially when used with highly specific antibodies and optimal fluorophores. Alternatively, the introduction of genetically encoded fluorescently tagged proteins and biosensors allows for live cell imaging to enable the temporal dissection of the given process. Spinning-disk confocal microscopy with high-sensitivity digital cameras is excellent for long-term or high framerate microscopy. In recent years, post-acquisition techniques such as superresolution radial fluctuations (SRRF) have been developed and can be used in combination with spinning-disk confocal [7, 8]. However, SRRF typically requires ~100 rapidly acquired raw images to generate a high-quality final image, and thus this can be a challenge with living cells and dynamic processes. Fortunately, a new platform termed super-resolution spinning-disk confocal microscopy using optical photon reassignment (SoRa) has been developed which generates a sub-diffraction limited image from a single image acquisition [9]. SoRa leverages changes in both hardware and software to achieve a lateral resolution of 120 nm. Specifically, SoRa uses the conventional spinning-disk head with micro-lens to allow multifocal excitation of the sample but also makes use of micro-lenses— instead of simple pin holes—to capture additional photons plus a magnifying lens to increase the magnification 2.8× or 4× depending on the objective used (Fig. 1). These changes to the light path improve the XY resolution by 1.4×, and further deconvolution results in a final resolution of roughly 2× the standard optical

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Fig. 1 Comparison of the light pathway of traditional spinning-disc confocal and the SoRa. (a) Schematic illustration of a conventional spinning-disk microscope demonstrating the path of the excitation light from the laser to the sample and then the detection of light by a camera. (b) Conventional spinning-disc microscopes have a lens disc and a pinhole disc. (c) The SoRa system has a lens disc, but the pinhole disc is also outfitted with lens to enhance light capture. The system also has a magnification lens to spread the signal over more pixels of the camera

limit. For reference, Fig. 2 highlights podosome structures stained with AlexaFluor-488 labeled phalloidin and immunostained with an anti-vinculin antibody detected with a Cy5-conjugated secondary antibody captured with a traditional spinning disk and the SoRa disk on the same microscope. We also describe the material and methods used to acquire this image as well as use spinning disk or SoRa for live cell imaging of RAW macrophages.

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Materials

2.1 RAW 264.7 Murine Macrophage Cell Preparation

1. RAW 264.7 murine macrophage cell line (ATCC: TIB-71) (see Note 1). 2. 37 °C incubator with 5% CO2. 3. RPMI 1640 medium supplemented with 10% heat-inactivated fetal bovine serum (FBS) (see Notes 2 and 3). 4. T-25 tissue culture flasks. 5. Phosphate-buffered saline magnesium. 6. 1.7 cm cell scrapers. 7. Hemocytometer.

(PBS)

without

calcium

and

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Fig. 2 Comparison of F-actin structure imaged with spinning disc and SoRa. Raw 264.7 murine macrophage podosome structures stained with AlexaFluor-488 labeled phalloidin and immunostained with an anti-vinculin antibody detected with a Cy5-conjugated secondary antibody captured with a traditional spinning disk and the SoRa disk on the same microscope. The SoRa has improved resolution and is able to detect filamentous structures not seen using the same laser power and exposure time as the traditional CSU-W1. Scale bars, 10 μm

8. 12-well tissue culture plate. 9. 18 mm round glass coverslip. 10. Tabletop centrifuge. 2.2 Transient Transfection of RAW Cells

1. DNA transfection reagent (such as jetOPTIMUS or FuGENE HD) and associated buffers. 2. Low serum culture media (such as OptiMEM) (see Note 4). 3. DNA construct(s) of interest to express GFP or mCherry chimeric proteins (see Note 5). 4. 12-well culture plate. 5. 18 mm round glass coverslip. 6. RPMI + 10% FBS.

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1. Neon Electroporation system (ThermoFisher Scientific) (see Note 6). 2. 0.4% Trypan blue solution in 0.81% sodium chloride and 0.06% potassium phosphate. 3. Hemocytometer. 4. DNA construct of interest. 5. 12-well culture plate. 6. 18 mm round glass coverslip. 7. RPMI + 10% FBS.

2.4 Opsonization of Polystyrene Beads

1. 4.19 μm polystyrene beads in suspension (~10% solids [w/w] aqueous suspension) with 2% (v/v) divinylbenzene (DVB) bead composition (see Note 7). 2. 50 mg/mL human IgG in PBS stock solution (store at -20 °C). 3. Fluorescently conjugated anti-human IgG antibody.

2.5 Phagocytosis Live Imaging and Microscopy

1. SoRa spinning-disk confocal microscope 60× (NA ~ 1.42) or 100× (NA ~ 1.35) objective lens, light source, filter set appropriate for the fluorescent protein of choice, objective heater, and heated stage. 2. Magnetic Leiden chamber for live cell imaging with 18 mm coverslips. Our lab uses the Chamlide CMB magnetic chamber (Quorum Technologies). 3. Hank’s balanced salt solution (HBSS).

2.6 Immunofluorescence and Phalloidin Staining

1. Alexa Fluor-488 Phalloidin (Thermo Fisher Scientific) (see Note 8). 2. Mouse anti-vinculin antibody (EMD Millipore; RRID AB_2304338) (see Note 9). 3. Donkey anti-mouse IgG—Cy5 conjugated (Jackson ImmunoResearch) (see Notes 8 and 9). 4. 1% BSA in PBS. 5. PBS with 0.1% Triton X-100. 6. PBS with 0.1% Tween 20. 7. Mounting media with a high refractive index and additives to prevent photobleaching such as ProLong Gold Antifade Mountant (Invitrogen).

2.7

Image Analysis

1. The system used is supplied by Nikon, and image acquisition and deconvolution was accomplished using NIS-Elements. 2. For image analysis and figure generation, we routinely use Image J acquisition (see Note 10).

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Methods

3.1 RAW 264.7 Murine Macrophage Cell Preparation

1. Culture RAW 264.7 macrophages in RPMI + 10% FBS. Incubate in a suitable incubator at 37 °C with 5% CO2. 2. Cells should be passaged or seeded from 80% confluence. To passage cells, first add 10 mL of culture medium to a new T25 flask, and place it in the incubator to equilibrate for approximately 30 min. 3. While media from step 2 equilibrate, take the original T25 containing the confluent culture. Aspirate the media. 4. Pipetting gently to ensure no bubbles are introduced, wash adherent culture with 10 mL sterile pre-warmed PBS, taking care to tilt the flask and ensure full coverage. 5. Aspirate PBS and gently detach cells with a cell scraper. 6. Add 10 mL pre-warmed RPMI + 10% FBS, and transfer cell suspension in a 15 mL conical tube for centrifugation. 7. Centrifuge cell suspension at 500 × g for 5 min. 8. Aspirate the supernatant from the pellet and resuspend with 3 mL of the equilibrated media. 9. Using a hemocytometer or automated cell counter, determine the number of cells in suspension. 10. Seed the new 1.0 × 106 cells.

flask with

pre-warmed

medium with

11. Top up the new flask with fresh media to ensure 10 mL volume, and incubate at 37 °C with 5% CO2 (see Notes 11 and 12). 12. Surplus cells from passaging can be used to seed plates for use in phagocytosis assays or transfection. When seeding a 12-well plate for transfection and live imaging, place a sterile 18 mm round glass coverslip at the bottom of each well in use (see Note 13). 13. Seed wells at a density of 0.1 × 106 cells per well (see Notes 11 and 12). 14. Add 1 mL culture medium (RPMI + 10% FBS) to each well in use. 3.2 Transient Transfection of RAW Cells (See Note 14)

1. Culture RAW 264.7 murine macrophages as per Subheading 3.1 to 60–80% confluency. 2. Six hours prior to transfection, change culture media for OptiMEM (see Note 4). 3. For each well intended for transfection, dilute 0.5 μg of a DNA construct in 100 μL of JetOPTIMUS buffer. Vortex and spin down.

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4. Add 0.5 μL of Jet Optimus reagent, vortex, spin down, and incubate for 10 min at room temperature. 5. Add combined buffer and reagent mixture to target wells and label accordingly. 6. Incubate the plate at 37 °C with 5% CO2 for 4 h before replacing the medium with fresh RPMI + 10% FBS. 7. Incubate plate at 37 °C with 5% CO2 for 24–48 h. 3.3 Electroporation of RAW Cells ( See Note 14)

1. Macrophage cell lines are refractory to transient transfections especially when larger plasmids (>7 kb) are used. Electroporation is an alternative approach. 2. Suspend adherent RAW cells as per methods in Subheading 3.1 for culture/subculture of adherent RAW cells in steps 3–7. 3. Count cells via hemocytometer. 4. Transfer 1 × 106 cells in suspension to a 15 mL tube, and centrifuge at 500 × g for 5 min. 5. Aspirate supernatant and resuspend the cell pellet in 100 μL resuspension buffer R, and add 5 μg DNA (see Note 15). 6. Add 3 mL electrolytic buffer into the Neon Tube, and place it into the Neon Pipette Station. 7. Set electroporation parameters (1680 V, 20 ms pulse width, 1 pulse). 8. Using the Neon Tip, draw 100 μL of resuspended cells taking care to avoid bubbles. 9. Activate electroporation system. 10. Dispense cell suspension (100 μL) into 1 well of a 12-well tissue culture plate (with 18 mm round glass coverslip) with RPMI + 10% FBS.

3.4 Opsonization of Polystyrene Beads

1. Mix opsonization targets at a ratio of 100 μL PBS, 10 μL bead slurry, and 10 μL human IgG solution (see Note 16). For 12-well plates, we recommend preparing 100 μL final working solution to afford for 5 μL of final bead suspension per well. 2. Incubate in a heated shaker at 37 °C with mild agitation for 1 h. 3. Wash opsonized beads two times with PBS, first pelleting beads at 15,000 × g for 30 s, aspirating the supernatant, and then resuspending in 100 μL PBS. 4. Prepared phagocytic targets can be stored at 4 °C until usage (ideally within 24 h of preparation).

3.5 Phagocytosis Live Imaging and Microscopy

1. Prior to removing cells from the incubator, ensure start-up process of spinning-disk confocal microscope is complete, with imaging software fully loaded, light source powered, lasers

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set to the correct wavelength for chosen conjugated fluorophores, and stage heater having reached a stable temperature of 37 °C. 2. Prepare a cooler with ice to maintain low temperature, and prolong macrophage viability during the imaging process. 3. Remove the plate of transfected macrophages from the incubator, and place the plate directly on ice, aspirating media and replacing it with 1 mL ice-cold HBSS (see Note 17). 4. Using forceps, carefully remove the coverslip from the 12-well plate, and transfer it to the chamber slide. Reattach the upper portion of the chamber slide taking care to modulate the speed and force of magnetic fixation with the thumb and forefinger to avoid coverslip breakage. Pipette sufficient HBSS to ensure saturation of coverslip (200 μL). 5. With minimal buffered solution on cells, mount the chamber slide on the stage at 60×/oil immersion lens, and find your ideal region of interest (ROI), preferably with multiple macrophages in focus without signs of damage or cell stress like blebbing. 6. Engage perfect focus (if available on your microscope software) to prevent focus drift during imaging or recording. Tune offset to focus immediately above the coverslip (Z = ~0) to capture lamellipodial protrusions. 7. When preparing to acquire time-lapse images and/or Z-stacks, aim for -1 to 16 μm with 0.4–0.8 μm steps DNA (see Notes 12 and 18). 8. Suspend 5 μL of previously prepared opsonized beads in 300 μL HBSS, and add this novel dilution to the coverslip. 9. Start recording time-lapse video/Z-stacks of ROI with parameters outlined in step 7. 3.6 Immunofluorescence of Podosomes in Resting Macrophages

1. Following protocols outlined in Subheading 3.1 or Subheading 3.2, grow cells to 80% confluence in a 12-well tissue culture plate with coverslips. 2. Aspirate media and replace with 4% PFA, and allow cells to fix for 30 min at room temperature. 3. Aspirate 4% PFA mixture and wash three times with Tris buffered saline or PBS containing 20 mM glycine to quench unreacted PFA. 4. Permeabilize fixed cells with PBS with 0.1% Triton X-100 for 30 min at room temperature. 5. Replace the solution with a blocking solution of 1% BSA/PBS solution, and incubate for 60 min.

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6. Add mouse anti-vinculin antibody in 1% BSA/PBS supplemented with 0.1% Tween 20, and incubate for 1 h at room temperature or overnight at 4 °C. 7. Wash three times for 10 min each wash with PBS/0.1% Tween 20. 8. Incubate goat anti-mouse secondary antibody in 1% BSA/PBS with 0.1% Tween 20 for 30–60 min at room temperature. 9. Wash three times for 10 min each wash with PBS/0.1% Tween 20. 10. Place a drop of ProLong Gold mountant on a glass slide, using forceps remove the coverslip and touch to a Kimwipe to remove excess media, and place cells facing down on the ProLong Gold mounting medium. 11. Place samples in the dark and allow 24 h for curing of the mounting medium. 12. Image as in Subheading 3.5. 3.7

Image Analysis

1. Open images using chosen image analysis suite (such as ImageJ or Nikon NIS-Elements) (see Note 10). 2. Determine the region of interest (ROI) for quantification (e.g., phalloidin IF tagged actin or fluorophore-conjugated F-actin reporter at the phagocytic cup). Measure the mean fluorescence per pixel in the region of interest. 3. Select a sample of background without fluorescence, and subtract this value from the experimental mean fluorescence in the ROI.

4

Notes 1. THP-1 cells or J774 cells are suitable alternative cell lines to Raw 264.7 cells. The described protocols also work with isolated peripheral blood mononuclear cells, bone marrowderived macrophages, or similarly isolated macrophages. Given the opsonization process uses IgG, methods are optimized for examining FcγR-mediated phagocytosis. 2. On occasion, we culture RAW cells in 5% serum-containing medium which results in a longer doubling time. 3. Many of the commonly used and readily available media have high levels of sodium bicarbonate (3700 mg/L) and are better suited to 10% CO2 incubators. The RPMI used in our lab contains 1500 mg/L. 4. JetOPTIMUS meditated transfection performs better in low serum media. Transfection of smaller plasmids (~5 kb) without

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replacing the RPMI + 10% FBS is successful. However, for larger more difficult-to-transfect plasmids (7–8 kb) is more efficient if the RPMI + 10% FBS is replaced with OptiMEM Minimal Essential Medium 4–6 h prior to transfection. 5. Many labs use the traditional EGFP- and mCherry-tagged constructs. However, newer fluorescent proteins have become available in the last 5–10 years. Many of these proteins have desirable features including better stability, less photobleaching, and superior brightness. Options to replace EGFP include mClover, mEmerald, and mNeonGreen, while mScarlet and mApple are brighter alternatives to mCherry and a better spectral match to the commonly used 561 nm excitation laser. 6. Protocols are optimized for use with Thermo Fisher’s Neon Electroporation system and Raw 264.7 macrophages. Other systems and cell lines may require adjustment for optimization. 7. Bangs Beads also sells latex beads with or without 2% DVB. In our experience, the antibodies do not stick to polystyrene beads lacking 2% DVB. 8. Phalloidin conjugates can be obtained across the full spectral range; primarily, the focus is on ensuring minimal overlap between F-actin staining (phalloidin) and fluorescent secondary antibody selected for staining vinculin. 9. Protocol and blocking optimized for use with listed primary and secondary antibodies. Manufacturers usually provide optimal dilution with the occasional need for adjustment. 10. Many confocal microscopes include optimized software packages (NIS-Elements), multiplatform commercial alternatives exist (e.g., ImageJ), as well as open-source alternatives (μManager). 11. The optimal seeding density of RAW 264.7 cells for ~80% confluence in 3 days is approximately 0.03 × 106 cells/cm2 surface area. Scale accordingly to chosen culture flask or well sizes. 12. If the cell count noted lower percent viability, lower viability can sometimes be remediated by allowing 4–6 h incubation for cells to settle to the flask surface and then aspirating nonadherent cells and supplying fresh media. 13. When placing coverslips in a 12-well plate, it can help to create a basic suction tool to reduce the handling of glass coverslips, ease the separation of stacked coverslips, and reduce the potential for surface scratching or breakage. To do so, take a 200 μL sterile pipette tip, and with a sterile cutting implement remove the distal 0.5 cm of the tip attempting to ensure a flat and uniform cut without distortion, with the plane of the cut parallel to the opening at the broader end of the tip. This tip

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can then be loosely fitted into the aspirator, allowing the use of suction to transiently adhere glass coverslips. Hold from the suction tool should be sufficient to remove one glass coverslip, and should issues arise releasing the coverslip, gently press the edge of the coverslip against the edge of the well until the suction is broken. 14. Survivability and efficiency of transfection are broadly impacted by the method used to introduce DNA and the size of the plasmid you are attempting to deliver. For smaller constructs and when high efficiency is not required such as single-cell microscopy, our lab uses transfection reagents (JetOPTIMUS). However, when larger plasmids are required or populationbased experiments we use electroporation. 15. The optimal cell suspension density for Neon Electroporation is 6 × 107 cells/mL with a total of 0.9–1.2 × 106 cells in suspension. 16. Polystyrene beads come in varying sizes from 0.5 to greater than 25 μm. Bead size partially determines the rate of attachment of opsonized targets and the duration of particle engulfment. We typically use beads with a diameter of 3–5 μm. Selecting phagocytic targets in this range affords a greater potential of capturing multiple internalization events when examining selected ROI in live imaging but may represent a trade-off, providing a smaller area to observe spatiotemporally discrete aspects of engulfment. To this end, we select 3.87 μm beads to try to find a compromise between increased odds of capturing multiple internalization events in live imaging while also allowing a slightly larger viewing area for examining actin dynamics during internalization. 17. This method is optimized for parachuting in beads with a greater volume to ensure a more uniform dispersal of beads. Preference is for parachuting by virtue of more temporally staggered internalization events to capture, and less potential for clumping, or overdispersal of beads to the periphery of the chamber slide during handling. Alternatively, beads can be introduced at the concentration prepared in Subheading 3.4 and plates centrifuged at 300 × g for 1 min to force the beads to the bottom of wells. 18. Optimal acquisition parameters include reducing exposure, intensity, and power of the laser to minimize photobleaching and phototoxicity. Ideal Z-sectioning should record 1 stack per 15 s, and total acquisition time should be between 10 and 20 min. Anything beyond that risks photobleaching and phototoxicity and likely misses the window of particle internalization.

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Acknowledgments Work on macrophage and phagocytosis is supported by a Canadian Institutes of Health Research Project Grant (PJT1655968) to G.D.F. G.D.F is also supported by a Tier 1 Canada Research Chair in Multiomics of Lipids and Innate Immunity. Figures were generated using BioRender and Adobe Illustrator. References 1. Uribe-Querol E, Rosales C (2020) Phagocytosis: our current understanding of a universal biological process. Front Immunol 11:1066 2. Davidson AJ, Wood W (2020) Macrophages use distinct actin regulators to switch engulfment strategies and ensure phagocytic plasticity in vivo. Cell Rep 31(8):107692 3. Wiesner C, Le-Cabec V, El Azzouzi K, Maridonneau-Parini I, Linder S (2014) Podosomes in space: macrophage migration and matrix degradation in 2D and 3D settings. Cell Adhes Migr 8(3):179–191 4. Murphy DA, Courtneidge SA (2011) The “ins” and “outs” of podosomes and invadopodia: characteristics, formation and function. Nat Rev Mol Cell Biol 12(7):413–426 5. Burger KL, Davis AL, Isom S, Mishra N, Seals DF (2011) The podosome marker protein Tks5

regulates macrophage invasive behavior. Cytoskeleton 68(12):694–711 6. Ostrowski PP, Freeman SA, Fairn G, Grinstein S (2019) Dynamic podosome-like structures in nascent phagosomes are coordinated by phosphoinositides. Dev Cell 50(4):397–410 7. Culley S, Tosheva KL, Pereira PM, Henriques R (2018) SRRF: universal live-cell super-resolution microscopy. Int J Biochem Cell Biol 101: 74–79 8. Gustafsson N, Culley S, Ashdown G, Owen DM, Pereira PM, Henriques R (2016) Fast live-cell conventional fluorophore nanoscopy with ImageJ through super-resolution radial fluctuations. Nat Commun 7(1):1–9 9. Azuma T, Kei T (2015) Super-resolution spinning-disk confocal microscopy using optical photon reassignment. Opt Express 23(11): 15003–15011

Chapter 7 Filamentous Bacteria as Targets to Study Phagocytosis Akriti Prashar, Maria Cecilia Gimenez, Serene Moussaoui, Iram Sobia Khan, and Mauricio R. Terebiznik Abstract Filamentous targets are internalized via phagocytic cups that last for several minutes before closing to form a phagosome. This characteristic offers the possibility to study key events in phagocytosis with greater spatial and temporal resolution than is possible to achieve using spherical particles, for which the transition from a phagocytic cup to an enclosed phagosome occurs within a few seconds after particle attachment. In this chapter, we provide methodologies to prepare filamentous bacteria and describe how they can be used as targets to study different aspects of phagocytosis. Key words Phagocytosis, Target morphology, Filamentous bacteria, Phagosome maturation, Phagocytic cup formation

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Introduction The canonical model of phagocytosis has been delineated using spheroidal targets [1]. However, the uptake of these targets occurs within seconds making the study of the initial events of phagocytosis such as the formation of a phagocytic cup, its remodeling and closing, and the scission of the nascent phagosome from the plasma membrane technically challenging. This limitation can be overcome by different techniques that hinder the ability of macrophages to enclose targets including parachuting phagocytes onto a defined planar surface coated with IgG (also known as frustrated phagocytosis), using pharmacological treatments, dominant negative mutants, or gene silencing approach [2, 3]. Nonetheless, these methodologies arrest phagocytosis, presenting limitations for the study of phagocytic cup remodeling, phagosome formation, and maturation. In contrast to spheroidal targets, filamentous bacteria are internalized via long-lasting phagocytic cups that extend along the length of the targets, which can be longer than 50 μM.

Roberto J. Botelho (ed.), Phagocytosis and Phagosomes: Methods and Protocols, Methods in Molecular Biology, vol. 2692, https://doi.org/10.1007/978-1-0716-3338-0_7, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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Consequently, the resulting phagocytic cups can easily persist for several minutes, depending on the length of the filament. This allows for the study of phagocytic cup formation and remodeling and phagosome formation and maturation in unprecedented spatiotemporal detail. Also, since these phagocytic cups fuse with endosomal compartments, as we described elsewhere in [4], these targets could also be utilized for studying events that correspond to phagosome maturation for spheroidal targets. Furthermore, by using biological targets that phagocytes may potentially encounter in vivo, additional parameters such as target surfaces and opsonins can be modified to examine different signaling pathways and their roles in the process of phagocytosis. These filamentous bacterial targets can be easily modified by cross-linking to fluorescent probes and can be used in real-time microscopy-based assays. Using this model, we recently demonstrated that filamentous target morphology affects phagosome maturation. This has important consequences for the outcome of phagocytosis as longer filaments of Legionella pneumophila survived and replicated more frequently than the bacillary forms in macrophages [4]. The methods described here can be used to study other filamentous pathogens like fungi or specialized host cells like neutrophils to assess the role of target morphology in host-pathogen interactions in addition to studying phagosome morphogenesis. This chapter provides details regarding the use of filamentous bacteria as phagocytic targets to study the progression of the various stages involved in phagocytosis [4].

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Materials 1. Phosphate-buffered saline (PBS): 0.144 g/L KH2PO4, 9.00 g/L NaCl, 0.795 g/L Na2HPO4. 2. Primary antibodies: anti-Legionella pneumophila IgG monoclonal antibody, anti-Escherichia coli IgG polyclonal antibody, anti-Salmonella IgG polyclonal antibody (obtained from EMD Millipore, BIORAD, and Thermo Fisher Scientific, respectively. See Note 1). 3. Purified human immunoglobulin G (IgG). 4. PBS with 5% (w/v) skim milk. 5. Fluorescently labeled secondary antibodies. 6. PBS with 4% (v/v) paraformaldehyde (PFA). 7. Dulbecco’s Modified Eagle Medium (DMEM) supplemented with 10% heat-inactivated fetal bovine serum (FBS). 8. PBS with 0.1% (v/v) Triton X-100.

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9. Liquid buffered yeast extract (BYE): 54.9 mM ACES buffer [(N –(2-Acetamido)-2 aminoethanesulfonic acid)], 10 g/L yeast extract, 6.84 mM α-ketoglutarate, 2.28 mM L-cysteine HCl, and 33.6 μM ferric pyrophosphate. 10. Buffered charcoal yeast extract agar (BCYE): 54.9 mM ACES, 10 g/L yeast extract, 6.84 mM α-ketoglutarate, 2.28 mM Lcysteine HCl, 33.6 μM ferric pyrophosphate, 11.4 g/L bacteriological agar, 2 g/L acid-washed activated charcoal with HCl, 100 μg/mL thymidine. 11. Luria broth Lennox (LB Lennox). 12. LB Lennox agar: 20 g/L Luria broth Lennox, 15 g/L agar bacteriological grade. 13. 10 mg/mL gentamicin in water, stored at room temperature. 14. Calcium chloride, dihydrate, molecular biology grade, min 99.5%. 15. Glycerol sterile solution. 16. pUltra plasmids (pUltra-RFP-GM, pUltra-GFP-KM) [5]. 17. 50 mg/mL kanamycin monosulfate, resuspended in water, stored at 20  C. 18. DQ-BSA resuspended in 1 PBS to stock concentration of 1 mg/mL and stored at 4  C for 2–4 weeks (Thermo Fisher Scientific, Burlington, ON, Canada). 19. 1-Ethyl-3-(3-dimethylaminopropyl)carbodiimide (or carbodiimide) resuspended in water, prepared fresh before use. 20. 25 mg/mL ciprofloxacin in water; stored at

20  C.

21. Bacteria for generating targets: Escherichia coli, Escherichia coli strain PAT84 (PMID:6991482 [6, 7]), Salmonella typhimurium, or Legionella pneumophila. 22. RAW 264.7 macrophages (ATCC TIB-71™; see Note 2). 23. 1.0 N potassium hydroxide. 24. 0.1 M sodium borate, pH 8.0 prepared the day of the experiment. 25. 1.0 M glycine in water prepared fresh before being diluted for use in experiments. 26. #1.5 thickness circular cover glasses (12 and 18 mm). 27. Microscope slides. 28. Fluorescent mounting media; store at 4  C. 29. Confocal microscope for fixed and/or live-cell imaging.

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Methods All reagents and media were prepared using MiliQ water. All bacterial culturing procedures were performed using standard aseptic techniques and according to biosafety regulations.

3.1 Obtaining Filamentous Bacterial Targets

Elongated polystyrene particles have been utilized as targets for phagocytosis [8]. However, filamentous bacteria offer an advantage over these particles, as they are longer but more flexible and therefore easily accommodated inside the macrophages. This property allows macrophages to internalize filaments >100 μM. Filamentous bacterial targets can be opsonized, or utilized without opsonization, and can be generated from bacteria expressing fluorescent proteins and conjugated to bioactive, fluorogenic, and fluorescent probes for microscopy-based, real-time functional assays [9, 10].

3.1.1 Generating Filamentous Bacterial Targets of Escherichia coli or Salmonella typhimurium Using Antibiotics

Filamentous growth in bacteria can be induced by using antibiotics that inhibit bacterial cell division as described below. 1. Prepare Luria broth (LB) media, autoclave to sterilize, and store as per the manufacturer’s directions. Generally, sterile LB media can be stored at 4  C for several months. 2. Take room temperature LB media, and transfer 3 mL to a sterile 10 mL culture tube with dual position cap. 3. Using a sterile inoculating loop, transfer bacteria to this culture tube from stocks maintained at 80  C in sterile LB media containing 25% glycerol (v/v). 4. Place the culture tubes in a shaking incubator, and shake at 200 rpm at 37  C overnight. 5. Make a 1:10 dilution of the overnight culture in LB, and measure the optical density at 600 nm (OD600). Prepare a subculture in a bacterial culture tube using LB to obtain a final volume of 3 mL and a starting OD600 of 0.05. Add 0.5 μg/mL of ciprofloxacin (see Note 3) to the culture, and place in a shaking incubator to shake at 200 rpm at 37  C for 4 hours (h). 6. Pellet the bacteria by centrifugation at 1000  g for 10 min at 4  C. 7. Wash the pellet twice with 1 PBS and resuspend in 4% PFA in PBS to kill and fix bacteria. These PFA-treated bacteria can be stored at 4  C for several months (see Note 4).

3.1.2 Generating Filamentous Bacterial Targets of a Thermosensitive ftsZ Escherichia coli Mutant Strain

Bacterial filamentation can be induced by mutations in genes controlling cell division [11, 12]. The filamenting mutant Escherichia coli strain PAT84 (E. coli PAT84) is a thermosensitive ftsZ mutant defective in cell division that grows as multinucleated filaments when the temperature is shifted from 30 to 42  C [6, 7]. Resultant filaments are relatively homogeneous in size (see

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Fig. 1 Filamentous bacterial targets. (a) PFA-fixed E. coli PAT84 filaments. Bacterial filaments were generated as outlined in Subheading 3.1.2. (b) PFA-fixed L. pneumophila filaments. Bacterial filaments were generated as outlined in Subheadings 3.1.6 and 3.1.7

Fig. 1a), and their length can be manipulated by incubating the bacteria at 42  C for different periods of time as described below. 1. Prepare LB Lennox media and autoclave to sterilize. Allow the media to cool down until media reaches room temperature. Media can be stored at 4  C for several months. 2. Use a sterile loop or pipette tip to scrape frozen bacteria from the top of the glycerol stock, and inoculate it into a culture tube with dual position cap filled with 5–7 mL LB Lennox media. 3. Place the tube containing bacteria in a shaking incubator set at 30  C. Shake the cultures at 200 rpm for 12 h. 4. Transfer 5 mL of bacterial culture to a sterile 125 mL Erlenmeyer containing 50 mL of LB Lennox. 5. Transfer the Erlenmeyer to a shaking incubator set to 30  C and 200 rpm until an OD600 of 0.2 is reached, which usually takes 2–3 h. 6. Dilute the bacteria culture 1:4 into a sterile 125 mL Erlenmeyer (i.e., 10 mL of bacteria culture +40 mL of LB Lennox media), and incubate as in step 3. 7. Repeat step 4 one more time (see Note 5). 8. Dilute the bacterial culture as described in step 4, and switch the temperature of the shaking incubator to 42  C to induce filamentation. To reduce the entanglement of bacteria filaments, reduce the agitation velocity to 150 rpm. Incubate for 3.5 h.

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9. Add 20 mL of fresh LB Lennox media to the bacterial culture (70 mL total volume), and incubate for an additional 1.5 h at 42  C. 10. Add 10 mL of fresh LB Lennox to the bacterial culture (80 mL total volume), and incubate for an additional 1 h at 42  C (see Note 6). 3.1.3 Engineering E. coli PAT84 to Express Fluorescent Proteins

1. Prepare chemically competent bacteria following established protocols such as the Hanahan method (also called calcium chloride method) [13]. 2. Take one aliquot of 100 μL of competent bacteria, and divide it into two aliquots of 50 μL. Use one aliquot as a control of heat shock by adding sterile water (volume equal to plasmid added to the second aliquot). 3. Use the second aliquot for plasmid transformation. Add 100–200 ng of pUltra-GFP-KM or pUltra-RFP-GM plasmid. Keep on ice for 30 min. 4. Place the tube in a water bath set to 42  C for 60 s. Place the tubes back on ice immediately afterward, and incubate for 5 min. 5. Add 800 μL of LB Lennox media (pre-warmed at 37  C) per aliquot, and place it in a shaking incubator set to 30  C and 200 rpm for 1 h. 6. Spread 100 μL of bacteria coming from the control aliquot on LB Lennox agar plates without antibiotic (should see full growth if heat shock procedure was performed correctly). Spread 100 μL of bacteria transformed with pUltra-GFP-KM or pUltra-RFP-GM on LB Lennox agar supplemented with 50 μg/mL of kanamycin or with 30 μg/mL of gentamicin, respectively. 7. Invert the plate and incubate it at 30  C for 24–48 h. 8. When bacterial colonies are detected, transfer a single colony from the LB Lennox agar containing antibiotic into a culture tube with dual position cap filled with 5–7 mL LB Lennox media and supplemented with either 50 μg/mL of kanamycin or 30 μg/mL of gentamicin. 9. Place the tube containing bacteria in a shaking incubator set to 30  C. Shake the cultures at 200 rpm for 12 h. 10. To make filaments, proceed as described in Subheading 3.1.2.

3.1.4 Preparing Fixed Filaments of E. coli PAT84

1. Transfer 50 mL of bacterial culture from Subheading 3.1.3, step 8, to a 50 mL conical tube. Centrifuge at 1000  g for 10 min to pellet the bacteria. Wash twice with 50 mL of sterile PBS.

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2. Resuspend the pellet in 15 mL 4% PFA in PBS. Using a laboratory rotator, incubate for 15 min at room temperature. Fixed bacteria can be stored in 4% PFA at 4  C for several months (see Notes 4 and 7). 3.1.5 Generating Filamentous Bacterial Targets of L. pneumophila in BCYE Agar

Filamentous Legionella pneumophila (L. pneumophila) targets can be obtained by growing bacteria in broth cultures or by resuspending colonies from cultures on agar as described below. Bacteria grown using these procedures yield cultures that are heterogeneous and contain bacteria of different lengths [4] (see Fig. 1b). However, they are enriched in long bacterial filaments [14]. For studies requiring homogeneous populations of filamentous bacterial targets, the cultures can be filtered as described in [15]. 1. Prepare BCYE media and autoclave to sterilize. Cool the media to 56  C and add L-cysteine, ferric pyrophosphate, and thymidine supplements that have been filter-sterilized (see Note 8). 2. Adjust pH to 6.9 using 1 N KOH that has been filter-sterilized. Keep stirring the media to ensure that the charcoal remains well suspended. 3. Pour 25 mL of media into sterile bacteriological plates, and allow them to cool overnight at room temperature. Store the plates at 4  C (see Note 9). 4. Using a sterile inoculating loop, streak L. pneumophila from frozen glycerol stocks maintained at 80  C in sterile LB media containing 25% glycerol (v/v). Invert the plate and incubate it at 37  C and 5% CO2 for 3–4 days to obtain bacterial colonies. 5. Bacteria growing on these plates are used as an inoculum to obtain filamentous targets to study phagocytosis (see Note 10).

3.1.6 Generating Filamentous L. pneumophila in BYE Media

1. Prepare BYE broth media and sterilize by autoclaving. Cool the media to 56  C, and add L-cysteine and ferric pyrophosphate that have been filter-sterilized (see Note 8). 2. Adjust pH to 6.9 using 1 N KOH that has been filter-sterilized. Store media at 4  C (see Note 11). 3. Transfer 25 mL of media to a sterile 125 mL Erlenmeyer flask. Remove a single bacterial colony from a 3–4-day-old BCYE agar plate using a sterile inoculating loop, and resuspend the bacteria well in the media. 4. Place the flask containing the bacteria in a shaking incubator set to 37  C. Shake the cultures at 100 rpm until they reach an OD600 of 3.5–4.0, which usually takes 24 h. By using these slow agitation conditions, the cultures obtained will be enriched in filamentous bacteria.

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3.1.7 Preparing Fixed Filaments of L. pneumophila

Transfer 25 mL of the OD600 3.5–4.0 L. pneumophila culture to a 50 mL conical tube, and process it as described above for E. coli PAT84, in Subheading 3.1.5.

3.2 Phagocytosis Assay Using Filamentous Bacterial Targets

Although the following methodology mentions RAW 264.7 macrophage cell line, it can be used to perform phagocytosis on any cultured macrophage cell line, primary macrophages, or monocytederived macrophages. 1. Grow RAW 264.7 cells in DMEM supplemented with 10% FBS (hereafter referred to as tissue culture medium) at 37  C and 5% CO2. Use 25 cm3 cell culture-treated flasks to provide cells with a consistent growth surface. 2. Once cells are ~80–85% confluent, remove the medium from the flask containing macrophages, and replace with 5 mL of fresh tissue culture medium. Gently scrape cells using a cell scraper, and dislodge cells by gently pipetting up and down. 3. Count cells using a hemocytometer and plate macrophages to achieve a final density of 5  105 cells per well (18 mm diameter) at the time of performing the assay. 4. Wash glass coverslips in sterile ultrapure water (see Note 12). Add 1 coverslip to each well of a 12-well tissue culture-treated plate. To each well, add the calculated volume of cell suspension from step 3, and distribute contents by swirling. 5. Incubate cells at 37  C with 5% CO2 to allow them to adhere. 6. Prepare the fixed bacterial targets by diluting the stock to yield 1 mL of bacteria at OD 2.0 in 1 PBS. Wash targets three times by centrifugation for 2 min at 10,000  g, removing the supernatant and resuspending in PBS. 7. Opsonize targets by incubating with anti-bacteria antibody (see Subheading 2 for details) for 1 h at room temperature (see Notes 13 and 14). Wash targets three times as described above, and resuspend pellet in 1 mL of PBS. 8. Place the macrophages plated on tissue culture plates in a precooled centrifuged to cool cells at 15  C for 5 min (see Note 15). Add 50 μL of the opsonized fixed bacterial targets to each well, yielding a 150:1 target-to-cell ratio (see Note 16). 9. Centrifuge the plate at 15  C and 300  g for 5 min to synchronize target binding. Incubate the cells at 37  C and 5% CO2 for 5 min to enhance the attachment of bacterial targets to the macrophages (see Note 17). 10. Wash the cells as follows: remove media from the wells and replace with 1 mL of PBS warmed to 37  C. Swirl contents and remove PBS. Repeat for a total of three washes. Add 1 mL of fresh tissue culture medium equilibrated at 37  C and 5% CO2

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Fig. 2 Internalization of filamentous bacterial targets through a long-lasting phagocytic cup. (a) RAW 264.7 macrophage expressing PM-GFP internalizing a PFA-fixed E. coli PAT84 filament. (b) RAW 264.7 macrophage

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to each well, and incubate cells for desired periods. As illustrated in Fig. 2a, b, two different stages of phagocytosis can be captured by fixing cells at different time points. In Fig. 2a, a long E. coli PAT84 filament was partly internalized by a macrophage at 15 min post phagocytosis. In Fig. 2b, a fully internalized E. coli PAT84 filament enclosed within a large phagosome at 2 h post phagocytosis. 11. The cells can be visualized by live- or fixed-cell microscopy. If cells are processed for fixed-cell microscopy, remove the medium and wash cells three times with PBS as described. Remove PBS and add 1 mL of 4% PFA in PBS to each well. Incubate at room temperature for 20 min and wash three times with PBS. Store coverslips in PBS for further processing. 3.3 Measuring Phagocytic Internalization Using Filamentous Bacteria

A phagocytic index assessing the uptake of spheroidal targets can be obtained by distinguishing between external and intracellular targets by immunofluorescence (also referred to as differential immunofluorescence) [16]. However, the prolonged phagocytic uptake stage observed with filamentous bacteria provides an interval where targets are partially internalized in a phagocytic cup. Therefore, by using differential immunolabeling as outlined below, it is not only possible to determine phagocytic index but also to measure the length of segments of the targets inside the phagocytic cup over time and determine the kinetic parameters of target internalization. This is illustrated in Fig. 2c where PFA-fixed RFP-expressing L. pneumophila is being internalized through a long-lasting phagocytic cup formed in a RAW 264.7 macrophage 10 min after the binding was synchronized. The external segment of the filament that was differentially immunolabeled as described in Subheading

ä Fig. 2 (continued) expressing PM-GFP that has fully internalized a PFA-fixed E. coli PAT84 filament. (a, b) Cells were transiently transfected with PM-GFP overnight using Lipofectamine LTX following the manufacturer’s protocol. PFA-fixed E. coli PAT84 filaments were generated as outlined in Subheading 3.1.2 and used to perform a phagocytosis assay as described in Subheading 3.2. Cells were washed and fixed (a) 15 min or (b) 120 min after binding was synchronized. (c) RAW 264.7 macrophage expressing PM-GFP internalizing a PFA-fixed L. pneumophila filament. Cells were transiently transfected with PM-GFP overnight using Fugene HD following the manufacturer’s protocol. PFA-fixed L. pneumophila filaments were generated as outlined in Subheading 3.1.6 and used to perform a phagocytosis assay as described above in Subheading 3.2. Cells were washed and fixed 10 min after the binding was synchronized. External segments of the filament were differentially immunolabeled as described in Subheading 3.3, step 1. Images to the right are higher magnifications of the framed region from the main panel. Phagocytic cup containing a filamentous bacterial target undergoing maturation by fusion with lysosomes. (d) RAW 264.7 cells were incubated with DQ-BSA for 1 h to label the late endosomes and lysosomes as outlined in Subheading 3.4, step 3. Phagocytosis was performed by adding PFA-fixed, L. pneumophila filaments as described in Subheading 3.2. Cells were fixed 20 min after the target attachment was synchronized. External segments of the filament were differentially immunolabeled as outlined in Subheading 3.3, step 1. Images to the right show a higher magnification of the phagocytic cup labeled with DQ-BSA delivered to this compartment via endosomal fusion

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3.3, step 1, is shown in blue. Images to the right are higher magnifications of the framed region from the main panel. All steps outlined can be carried out at room temperature for 1 h unless otherwise noted. 1. After performing the phagocytosis assay as described in Subheading 3.2, incubate PFA-fixed cells with 1 mL of 5% skim milk for blocking. 2. Wash cells three times with PBS, and incubate with a primary antibody against your bacteria of interest diluted in blocking solution to label the filamentous bacterial targets. 3. Wash cells three times with PBS, and incubate with the desired fluorescent dye-conjugated secondary antibody diluted in the blocking solution. This procedure will immunolabel the segments of the filamentous bacteria not yet internalized by the macrophage. Refer to Note 18 if fluorescent bacterial targets are used for phagocytosis assays. 4. Wash cells three times with PBS, and incubate with 1 mL of 0.1% Triton X-100 for 20 min to permeabilize cell membranes. 5. Block and incubate again with the primary antibody as described in steps 2 and 3. 6. Following three washes with PBS, incubate cells with a different fluorescent dye-conjugated secondary antibody to allow for the distinction between external and internalized bacterial targets. This procedure will immunolabel the entire length of the bacterial filament, including the section internalized by the macrophage. 7. Wash cells three times with PBS, and mount the coverslip onto glass slides using fluorescent mounting media. 8. Acquire confocal z-stacks series to observe the entire filament. 9. Using imaging analysis software such as Image J (NIH) or an equivalent, measure the length of the filamentous targets labeled in the two rounds of immunolabeling. The difference in the total length stained after the second round of immunolabeling and the length of the filament stained after the first round of immunolabeling can be used to indicate the length of the filament internalized by the macrophage. 10. To complement the approach described above in steps 1–9, the phagocytosis assay can be performed in cells expressing recombinant probes for plasma membrane markers such as PM-GFP (a chimeric GFP fused to a myristoylation/palmitoylation sequence from Lyn kinase) or GPI-GFP (glycosyl phosphatidylinositol-GFP) to delineate the phagocytic cups [17, 18].

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3.4 Remodeling and Maturation of Phagocytic Cups and Phagosomes

The transition from phagocytic cups to phagosomes requires the remodeling of the lipids and the associated proteins of the plasma membrane, rearrangement of the actin cytoskeleton, as well as the focal exocytosis of endomembranes at the phagocytic cup [19]. The study of these processes using spheroidal targets is challenging due to the fast pace at which these events take place. Given the long duration of the phagocytic cup stadia for filamentous targets, greater spatial and temporal resolution can be achieved without the requirement of genetic and/or pharmacological approaches to stall the process in order to study these remodeling events.

3.4.1 Phagocytic Cup Remodeling

Using filamentous bacterial targets and the protocol described below, we studied the remodeling of actin as well as phosphoinositides (PtdIns) at the phagocytic cup using fixed- and live-cell imaging as described in [4]. 1. Perform a phagocytosis assay as described in Subheading 3.2. 2. Fix cells using 4% PFA in PBS following the desired incubation periods, and differentially immunolabel the filamentous targets as described in Subheading 3.3 to distinguish between the external and intracellular bacterial segments. 3. Stain cells using fluorophore-conjugated phalloidin to label F-actin and to visualize the phagocytic cups. 4. Mount the coverslips onto glass slides using fluorescent mounting media. 5. Acquire confocal z-stacks to analyze target internalization over time and actin distribution around the particle to assess actin remodeling. 6. To complement the fixed-cell imaging approach described in steps 1–4 with live-cell imaging, perform a phagocytosis assay as outlined in Subheading 3.2 using fluorescent bacterial targets in cells stably or transiently expressing fluorescent actin probes (see Note 19). Similarly, to examine PtdIns remodeling, cells can be transfected with recombinant probes to determine changes in PtdIns levels (e.g., PLCδ-PH-GFP to visualize phosphatidylinositol 4,5-bisphosphate levels) [20].

3.4.2 Phagocytic Cup Maturation

The canonical model of phagocytosis requires the nascent phagosome to separate from the plasma membrane for maturation to occur. However, similar to phagosome maturation, the phagocytic cups formed for filamentous bacteria fuse with endosomes and lysosomes. The large surface presented by the elongated phagocytic cups can be used for mechanistic studies of highly dynamic membrane fusion and fission events, protein complex formation and disassembly, and membrane lipid remodeling processes associated with phagosome maturation. Using antibodies, GFP chimeric constructs for early and late endosomal proteins, or fluorogenic

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endosomal probes, and following the protocol in Subheading 3.2, we assessed the maturation of phagocytic cups though fixed- and live-cell imaging as described in [4]. Fluid phase uptake of DQ-BSA can be used to label endosome and lysosomes, and their delivery to the phagocytic cup can be followed by fixed- or live-cell microscopy (see Note 20) (Fig. 1d) where a cell with lysosomes labeled with DQ-BSA is internalizing a PFA-fixed, RFP-expressing L. pneumophila filament. Cells were fixed 20 min after attachment was synchronized, and the external segment of the bacterial filament was immunolabeled (blue). Images to the right show a higher magnification of the phagocytic cup labeled with DQ-BSA delivered to this compartment via endosomal fusion. The steps described below are for labeling endosomal compartments with DQ-BSA but can be used for other fluid phase markers including fluorescent-labeled dextrans. 1. Dilute DQ-BSA in tissue culture media according to the manufacturer’s instructions. 2. Replace the media used to culture cells with media containing DQ-BSA. 3. Incubate cells with DQ-BSA solution for 1 h at 37  C and 5% CO2. 4. Remove the media and wash cells twice with PBS warmed to 37  C. 5. Incubate cells in fresh tissue culture media that is pre-warmed to 37  C for 1 h at 37  C and 5% CO2 for the desired period to chase the probes into early or late endosomes or lysosomes. 6. Perform a phagocytosis assay as described in Subheading 3.2. 7. Fix cells in 4% PFA in PBS for 20 min at room temperature for fixed-cell imaging or process for live-cell imaging. 3.4.3 Assessing the Luminal Environment of Phagocytic Cups and Phagosomes During Maturation

To assess the luminal environment of a phagocytic cup and its acidification and acquisition of hydrolytic properties during the transition to a phagosome, fluorescent probes can be cross-linked to filamentous bacterial targets for real-time microscopy-based assays. We successfully used this approach to analyze the acquisition of hydrolytic properties by phagocytic cups containing filamentous bacterial targets during their transition to phagosomes using livecell microscopy [4]. Although the following method describes the cross-linking of DQ-BSA to filamentous targets, this technique can be modified to measure acidification [21], generation of ROS, etc. 1. Resuspend 1  109 PFA-fixed filaments in PBS with 25 mg/ mL of carbodiimide cross-linker.

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2. Mix by agitation at room temperature for 15 min. 3. Wash cells three times with 1 mL of 0.1 M sodium borate coupling buffer (pH 8.0) to remove excess cross-linker. 4. Use 500 μL of coupling buffer with 1.0 mg DQ-BSA (or another fluorogenic probe) and 4 mg/mL IgG to opsonize the targets for 12–16 h with gentle rotation at room temperature. 5. Spin the targets at 3000  g and wash three times with 250 mM glycine in PBS (pH 7.2) to quench the probe. Vortex in between wash steps for 5 min. 6. Wash the substrate-bound targets three times with PBS. 7. Cool cells for 5 min at 15  C by placing in a precooled centrifuge. 8. Add 50 μL of the opsonized targets to each well, and synchronize binding by centrifugation at 15  C and 300  g for 5 min. 9. Incubate the cells at 37  C and 5% CO2 for an additional 5 min. 10. Remove media from the wells and wash three times with PBS pre-warmed to 37  C. 11. Replace with fresh tissue culture media pre-warmed to 37  C, and incubate cells at 37  C and 5% CO2 for desired periods. 12. The cells can be visualized by live- or fixed-cell microscopy.

4

Notes 1. Other antibodies against the bacteria of interest exist and could be employed. 2. RAW 264.7 macrophages must be handled using Biosafety Level 2 (BSL-2) practices and procedures. 3. Ciprofloxacin is an antibiotic that belongs to the fluoroquinolone family and prevents bacterial replication by inhibiting DNA gyrase. Usage at sublethal doses activates the bacterial SOS stress response, inducing filamentation without causing cell death [4, 12]. 4. We did not observe any difference with respect to the stability of the filaments between storing the fixed bacteria at 4  C resuspended in 4% PFA or PBS. If storing in PFA, wash the bacteria twice with PBS or wash and incubate with 125 mM glycine in PBS (pH 7.2) for 15 min at room temperature to quench any residual PFA before using for an experiment. 5. Steps 4 and 5 usually take 4 h to complete. 6. Longer incubation periods at 42  C are not advised because bacterial viability will be compromised beyond this time period. An incubation period of 6 h is ideal if one wants to obtain

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filaments longer than 50 μM. If shorter filaments are desired (15–20 μM), this incubation time should be reduced to 3.5 h. 7. Fixing with PFA preserves the filamentous bacteria, allowing for the study of effects of target morphology on phagocytosis, excluding any interference from potential bacterial effectors. Live bacteria can be used to investigate the impact of filamentous morphology on pathogenicity. 8. Warming helps to dissolve the ferric pyrophosphate for making the BYE and BCYE media. 9. Plates stored at 4  C can be used for up to 4 months. In our experience, using plates older than 4 months reduces the proportion of longer filaments. 10. To ensure that the cultures are enriched in filamentous bacteria, use freshly streaked plates. Using older plates or plates stored at 4  C to start cultures yields cultures enriched in bacillary forms of the bacteria. 11. BYE media stored at 4  C can be used for up to 4 months. In our experience, using media older than 4 months reduces the proportion of longer filaments obtained in cultures. 12. Coverslips are stored in 70% ethanol for sterilization. Prior to use for plating cells, they are washed thoroughly in sterilized ultrapure water to remove the ethanol. Alternatively, coverslips can be sterilized by autoclaving prior to use. 13. The phagocytosis assays can be performed without opsonization of the targets. However, opsonization with IgG antibodies increases binding and internalization efficiencies. 14. Although an anti-Lp1 antibody is used to opsonize targets in this chapter, nonspecific IgG, such as donkey or human IgG, can be used as well. 15. Unlike the phagocytosis of spherical targets, where target binding can be synchronized by cooling cells to 4  C while inhibiting target uptake, the attachment of longer filamentous targets is inhibited at 4  C. This could be due to actin-dependent reorientation of the targets needed to gain access to their short axes as described in [22] in order for phagocytosis to occur or the lack of the receptors needed for target attachment with high avidity. Therefore, the attachment is synchronized at 15  C. 16. A late-exponential culture of bacterial filaments (OD600 2.0) contains approximately 1.5  109 bacteria/mL. Using this approximation, cells are presented with fixed bacterial targets with a target-to-cell ratio of 150:1. 17. Cells are incubated with filamentous targets for 5 min following synchronization of binding to enhance attachment. If

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washed immediately following centrifugation, the efficiency with which longer filaments bind is greatly reduced. 18. If the fixed filamentous bacterial targets express a fluorescent protein such as GFP or RFP, cell membrane permeabilization and immunolabeling of internalized targets are not required. 19. If desired, external segments of the filamentous bacteria can be immunolabeled prior to performing live-cell imaging. Place cells on ice, and replace media with cold media containing the fluorescent dye-conjugated secondary antibody of choice against the primary antibody used to opsonize targets, for 5 min. Wash the unbound antibody and proceed with livecell imaging. As labeling in the cold is only done for 5 min, we found that using a higher concentration (two to three times greater than what is used for fixed-cell labeling) of secondary antibody provided better labeling. 20. DQ-BSA is a self-quenched protease substrate that is converted into fluorescent peptides once hydrolyzed by lysosomal proteases.

Acknowledgments We thank Dr. Ana Isabel Rico and Dr. Miguel Vicente, CNB-CSIC Spain, for sharing the E. coli PAT84 strain and for their keen input on the generation of filaments. This work was funded by the Natural Sciences and Engineering Council of Canada (RGPIN2018-05734 and RGPAS2018-522692) to M.R. Terebiznik. References 1. Champion JA, Mitragotri S (2006) Role of target geometry in phagocytosis. Proc Natl Acad Sci U S A 103(13):4930–4934. https:// doi.org/10.1073/PNAS.0600997103 2. Swanson JA (2008) Shaping cups into phagosomes and macropinosomes. Nat Rev Mol Cell Biol 9(8):639–649. https://doi.org/10. 1038/NRM2447 3. Wright SD, Silverstein SC (1984) Phagocytosing macrophages exclude proteins from the zones of contact with opsonized targets. Nature 309(5966):359–361. https://doi. org/10.1038/309359A0 4. Prashar A, Bhatia S, Gigliozzi D, Martin T, Duncan C, Guyard C, Terebiznik MR (2013) Filamentous morphology of bacteria delays the timing of phagosome morphogenesis in macrophages. J Cell Biol 203(6):1081. https://doi.org/10.1083/JCB.201304095

5. Mavridou DAI, Gonzalez D, Clements A, Foster KR (2016) The pUltra plasmid series: a robust and flexible tool for fluorescent labeling of Enterobacteria. Plasmid 87–88:65–71. https://doi.org/10.1016/J.PLASMID.2016. 09.005 6. Ricard M, Hirota Y (1973) Process of cellular division in Escherichia coli: physiological study on thermosensitive mutants defective in cell division. J Bacteriol 116(1):314–322. https:// doi.org/10.1128/JB.116.1.314-322.1973 7. Lutkenhaus JF, Wu HC (1980) Determination of transcriptional units and gene products from the ftsA region of Escherichia coli. J Bacteriol 143(3):1281. https://doi.org/10.1128/JB. 143.3.1281-1288.1980 8. Champion JA, Mitragotri S (2009) Shape induced inhibition of phagocytosis of polymer particles. Pharm Res 26(1):244–249. https:// doi.org/10.1007/S11095-008-9626-Z

Filamentous Targets 9. Russell DG, Vanderven BC, Glennie S, Mwandumba H, Heyderman RS (2009) The macrophage marches on its phagosome: dynamic assays of phagosome function. Nat Rev Immunol 9(8):594–600. https://doi. org/10.1038/NRI2591 10. Yates RM, Hermetter A, Russell DG (2005) The kinetics of phagosome maturation as a function of phagosome/lysosome fusion and acquisition of hydrolytic activity. Traffic 6(5): 413–420. https://doi.org/10.1111/J. 1600-0854.2005.00284.X 11. Feucht A, Errington J (2005) ftsZ mutations affecting cell division frequency, placement and morphology in Bacillus subtilis. Microbiology (Reading) 151(Pt 6):2053–2064. https://doi. org/10.1099/MIC.0.27899-0 12. Harry E, Monahan L, Thompson L (2006) Bacterial cell division: the mechanism and its precison. Int Rev Cytol 253:27–94. https:// doi.org/10.1016/S0074-7696(06)53002-5 13. Green MR, Sambrook J (2018) The Hanahan method for preparation and transformation of competent Escherichia coli: high-efficiency transformation. Cold Spring Harb Protoc 2018(3):183–190. https://doi.org/10.1101/ PDB.PROT101188 14. Prashar A, Bhatia S, Tabatabaeiyazdi Z, ˜ o RA, Tang P, Low DE, Duncan C, Gardun Guyard C, Terebiznik MR (2012) Mechanism of invasion of lung epithelial cells by filamentous Legionella pneumophila. Cell Microbiol 14(10):1632–1655. https://doi.org/10. 1111/J.1462-5822.2012.01828.X 15. Bos J, Zhang Q, Vyawahare S, Rogers E, Rosenberg SM, Austin RH (2015) Emergence of antibiotic resistance from multinucleated bacterial filaments. Proc Natl Acad Sci U S A 112(1):178–183. https://doi.org/10.1073/ PNAS.1420702111 16. Steinberg BE, Grinstein S (2007) Assessment of phagosome formation and maturation by

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fluorescence microscopy. Methods Mol Biol 412:289–300. https://doi.org/10.1007/ 978-1-59745-467-4_19 17. Teruel MN, Blanpied TA, Shen K, Augustine GJ, Meyer T (1999) A versatile microporation technique for the transfection of cultured CNS neurons. J Neurosci Methods 93(1):37–48. https://doi.org/10.1016/S0165-0270(99) 00112-0 18. Nichols BJ, Kenworthy AK, Polishchuk RS, Lodge R, Roberts TH, Hirschberg K, Phair RD, Lippincott-Schwartz J (2001) Rapid cycling of lipid raft markers between the cell surface and Golgi complex. J Cell Biol 153(3): 529–541. https://doi.org/10.1083/JCB. 153.3.529 19. Lee WL, Mason D, Schreiber AD, Grinstein S (2007) Quantitative analysis of membrane remodeling at the phagocytic cup. Mol Biol Cell 18(8):2883–2892. https://doi.org/10. 1091/MBC.E06-05-0450 20. Stauffer TP, Ahn S, Meyer T (1998) Receptorinduced transient reduction in plasma membrane PtdIns(4,5)P2 concentration monitored in living cells. Curr Biol 8(6):343–346. https://doi.org/10.1016/S0960-9822(98) 70135-6 21. Naufer A, Hipolito VEB, Ganesan S, Prashar A, Zaremberg V, Botelho RJ, Terebiznik MR (2018) pH of endophagosomes controls association of their membranes with Vps34 and PtdIns(3)P levels. J Cell Biol 217(1): 329–346. https://doi.org/10.1083/JCB. 201702179 22. Mo¨ller J, Luehmann T, Hall H, Vogel V (2012) The race to the pole: how high-aspect ratio shape and heterogeneous environments limit phagocytosis of filamentous Escherichia coli bacteria by macrophages. Nano Lett 12(6): 2901–2905. https://doi.org/10.1021/ NL3004896

Chapter 8 The Derivation and Use of HoxB8-Driven Conditionally Immortalized Macrophages Shranjit S. Lail, Neil McKenna, and Robin M. Yates Abstract The use of Hox-driven conditionally immortalized immune cells has significantly increased in biomedical research over the past 15 years. HoxB8-driven conditionally immortalized myeloid progenitor cells maintain their ability to differentiate into functional macrophages. There are multiple benefits to this conditional immortalization strategy including the ability for unlimited propagation, genetic mutability, primary-like immune cells (macrophages, dendritic cells, and granulocytes) on demand, derivation from variety of mouse strains, and simple cryopreservation and reconstitution. In this chapter, we will discuss how to derive and use these HoxB8-conditionally immortalized myeloid progenitor cells. Key words HoxB8, Conditional immortalization, Differentiation, Macrophages, Phagosomes

1

Introduction Furthering our understanding of the immune system is vital to deciphering the mechanisms of immune response and dysregulation which underpins most common diseases and illnesses. Ex vivo models of innate and adaptive immune cells have facilitated countless advances in our understanding of immunology. Protocols for obtaining and maintaining primary immune cells ex vivo have been well-described and are the primary method used to study the immune system [1]. Research using primary immune cells is limited because these primary immune cells are short-lived, require constant sacrifice of mice, and are very difficult to genetically manipulate. With the use of a sophisticatedly simplistic method developed by Mark Kamps’ group in the mid-2000s, these limitations can be overcome [2, 3]. This method uses overexpression of the HoxB8 transcription factor to immortalize myeloid progenitors that have demonstrated the ability to differentiate into functional macrophages. In Wang et al. (2006), when these conditionally immortalized cells

Roberto J. Botelho (ed.), Phagocytosis and Phagosomes: Methods and Protocols, Methods in Molecular Biology, vol. 2692, https://doi.org/10.1007/978-1-0716-3338-0_8, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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were maintained in granulocyte-macrophage colony-stimulating factor (GM-CSF), removal of estrogen induced the majority of the cells (greater than 99%) to differentiate into macrophages [2]. The methods used in this chapter have been utilized in Lail et al. [4].

2

Materials

2.1 ER-HoxB8 DNA Isolation

1. E. coli DH5α. 2. Luria-Bertani broth, in ddH2O; store at room temperature. 3. 100 μg/mL ampicillin; aliquot and store at -20 °C. 4. Plasmid DNA Midiprep Kit. 5. Nanophotometer.

2.2 ER-HoxB8 Lentivirus Production

1. 10 cm tissue culture (TC)-treated plates. 2. 10 μg/mL poly-L-lysine, in PBS; store at 4 °C. 3. Phosphate buffered saline (PBS); store at room temperature. 4. Base medium: Dulbecco’s Modified Eagle Medium (DMEM), 10% bovine growth serum, 1 mM sodium pyruvate, 100 U/ mL penicillin/streptomycin, 2 mM L-glutamine. Store at 4 °C. 5. Lentivirus production is done using Human Embryonic Kidney 293T (HEK293T) cells. 6. Transfection medium: DMEM, 10% bovine growth serum, 1 mM sodium pyruvate, 2 mM L-glutamine. Store at 4 °C. 7. Transfection reagent. The authors routinely use Lipofectamine 2000. Store at 4 °C. 8. Opti-MEM. Store at 4 °C. 9. 0.45 μm polyethersulfone (PES) syringe filter. 10. 38 mL PPCO ultracentrifuge tubes. 11. 20% sucrose, in PBS; store at 4 °C. 12. DMEM. Store at 4 °C. 13. Ultracentrifuge. 14. Virus storage medium: DMEM, 30% fetal bovine serum, 1 mM sodium pyruvate, 100 U/mL penicillin/streptomycin, 2 mM L-glutamine. Store at 4 °C.

2.3 Bone Marrow Isolation

1. C57BL/6J mice. 2. DMEM. Store at 4 °C. 3. 25g needle. 4. Erythrocyte lysis buffer: 0.8% NH4Cl, in ddH2O; store at 4 °C. 5. 70 μm pore cell strainer.

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6. 10 ng/mL recombinant murine IL-3; aliquot and store at 20 °C. 7. 10 ng/mL recombinant murine IL-6; aliquot and store at 20 °C. 8. 10 ng/mL recombinant murine SCF; aliquot and store at 20 °C. 9. 6-well suspension non-TC-treated plates. 2.4 ER-HoxB8 Lentiviral Transduction

1. 12-well suspension non-TC-treated plates. 2. 10 μg/mL human Plasma Fibronectin Purified Protein, in PBS; store at 4 °C. 3. PBS; store at room temperature. 4. Virus storage medium: DMEM, 30% fetal bovine serum, 1 mM sodium pyruvate, 100 U/mL penicillin/streptomycin, 2 mM L-glutamine. Store at 4 °C. 5. Polybrene transduction enhancer; aliquot and store at -20 °C. 6. Base medium: DMEM, 10% bovine growth serum, 1 mM sodium pyruvate, 100 U/mL penicillin/streptomycin, 2 mM L-glutamine. Store at 4 °C. 7. GM-CSF-conditioned medium; aliquot and store at -20 °C (see Note 1). 8. 20 mM β-estradiol in ethanol; aliquot and store at -20 °C.

2.5 Macrophage Differentiation

1. Base medium: DMEM, 10% bovine growth serum, 1 mM sodium pyruvate, 100 U/mL penicillin/streptomycin, 2 mM L-glutamine. Store at 4 °C. 2. L929-conditioned medium; aliquot and store at -20 °C. 3. 10 mM fulvestrant, in DMSO; aliquot and store at -20 °C. 4. 10 cm sterile polystyrene Petri dishes. 5. PBS; store at room temperature.

3

Methods

3.1 ER-HoxB8 and Lentiviral Packaging Constructs DNA Isolation

1. A synthetic construct designed to express an ER-HoxB8 fusion protein is synthesized and cloned into the second-generation lentiviral transfer vector pCDH-MSCV-MCS-EF1α-Puro (see Note 2). 2. psPAX2, pCMV-VSV-G, and pCDH-ER-HoxB8 are transformed and selected in E. coli. 3. The E. coli are grown in Luria-Bertani broth with 100 μg/mL ampicillin overnight at 37 °C.

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Fig. 1 Schematic for the protocol for ER-HoxB8 and lentiviral packaging constructs DNA isolation. (Created with BioRender.com)

4. The cells are collected by centrifugation at 3500 × g for 10 min. 5. A Plasmid DNA Midiprep Kit is used to isolate DNA, following the manufacturer’s instructions. 6. The DNA concentrations nanophotometer.

are

determined

with

a

7. Schematic for the protocol for ER-HoxB8 and lentiviral packaging constructs DNA isolation (see Fig. 1). 3.2 Generating ERHoxB8 Lentivirus 3.2.1 Plating HEK293T Cells

1. Five 10 cm TC-treated plates produce sufficient quantities of virus to generate these HoxB8-driven conditionally immortalized cells. 2. The 10 cm TC-treated plates are coated with 2 mL of poly-Llysine diluted in PBS (to a final concentration of 10 μg/mL) and incubated for at least 15 min at 37 °C in order to encourage stronger cell attachment to the plate surface (see Note 3). 3. The poly-L-lysine is discarded and replaced with 10 mL of base medium (see Note 4). 4. HEK293T cells are then plated at a density of 3.5 × 106 cells per plate and incubated overnight at 5% CO2, 37 °C (see Note 5). 5. Schematic for the protocol for Day 1 of generating ER-HoxB8 lentivirus (see Fig. 2).

3.2.2

Transfection

1. The medium on the HEK293T plates is replaced with 10 mL of transfection medium (see Note 6). 2. Cells are transfected with 3 μg of the pCDH-ER-HoxB8 expression plasmid, 1.05 μg of the pantropic pseudotyping glycoprotein construct pCMV-VSV-G, and 1.95 μg of the

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Fig. 2 Schematic for the protocol for Day 1 of generating ER-HoxB8 lentivirus. (Created with BioRender.com)

Fig. 3 Schematic for the protocol for Day 2 of generating ER-HoxB8 lentivirus. (Created with BioRender.com)

second-generation lentiviral packaging vector psPAX2, per plate, respectively, using 2 μL of Lipofectamine 2000 per μg of DNA, as specified in the manufacturer’s instructions. 3. Briefly, DNA and Lipofectamine 2000 are each diluted in 0.5 mL of Opti-MEM per plate. 4. These solutions are mixed, incubated for 10 min at room temperature, and then added to the HEK293T cells. 5. Cells are incubated at 37 °C for 6–7 h, at which time the culture medium is replaced with 5 mL base medium for overnight incubation and virus production, followed by virus harvesting (see Note 7). 6. Schematic for the protocol for Day 2 of generating ER-HoxB8 lentivirus (see Fig. 3).

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Fig. 4 Schematic for the protocol for Day 3 of generating ER-HoxB8 lentivirus. (Created with BioRender.com) 3.2.3

Virus Harvesting

1. The virus-conditioned medium from each plate is collected at approximately 1 day following transfection and filtered through a 0.45 μm syringe filter to remove cellular debris and any potential contaminants. The medium is kept at 4 °C until the second harvest (step c). 2. An additional 5 mL of base medium is added to each plate, and the plates are incubated at 37 °C for 6–7 h. 3. The virus-conditioned medium from each plate is collected and filtered through a 0.45 μm syringe filter and kept overnight at 4 °C. 4. Schematic for the protocol for Day 3 of generating ER-HoxB8 lentivirus (see Fig. 4).

3.2.4

Ultracentrifugation

1. 25 mL of virus-conditioned medium is added to 38 mL polypropylene copolymer (PPCO) ultracentrifuge tubes. 2. 6 mL of 20% sucrose dissolved in PBS is underlaid (see Note 8). 3. The weight of the ultracentrifuge tubes are balanced using DMEM. 4. The tubes are centrifuged at 26,000 rpm for 2 h at 4 °C. 5. Following ultracentrifugation, the top layer is removed to the interphase, and then the remaining medium is carefully decanted (see Note 9). 6. The tubes are dried for approximately 15–30 min on a paper towel. 7. The virus is then resuspended in 200 μL of virus storage medium, then divided to 10 μL working aliquots, and kept at -80 °C. 8. Schematic for the protocol for Day 4 of generating ER-HoxB8 lentivirus (see Fig. 5).

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Fig. 5 Schematic for the protocol for Day 4 of generating ER-HoxB8 lentivirus. (Created with BioRender.com) 3.3 Isolating Bone Marrow

1. The tibiae, femora, and ilia are isolated from C57BL/6J mice. The bone marrow is removed by flushing the bones with a DMEM-filled syringe. 2. The bone marrow is centrifuged at 300 × g for 5 min. 3. 4 mL of erythrocyte lysis buffer is added to the bone marrow pellet and incubated at room temperature for approximately 4 min. 25 mL of DMEM is then added to stop the lysis process. 4. The bone marrow is strained through a 70 μm pore cell strainer to remove cell clumps for a uniform cell suspension, and centrifugation at 300 × g for 5 min. 5. The isolated bone marrow is then resuspended in medium containing 10 ng/mL recombinant murine IL-3, IL-6, and SCF to increase proliferation and improve the transduction efficiency of hematopoietic stem cells (HSC). 6. The cells are transferred to a 6-well suspension plate and incubated for 2 days at 5% CO2, 37 °C. 7. Schematic for the protocol for isolating bone marrow (see Fig. 6).

3.4 Transduction with ER-HoxB8 Lentivirus

1. 12-well suspension plates are coated with 500 μL of fibronectin (10 μg/mL in PBS) per well as a transduction efficiency enhancer and incubated at 37 °C overnight the day before transduction. 2. The fibronectin is removed and replaced with 1 mL of virus storage medium per well before lentiviral transduction. 3. The 10 μL virus aliquots are diluted to a final volume of 100 μL with virus storage medium and added to the appropriate wells.

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Fig. 6 Schematic for the protocol for isolating bone marrow. (Created with BioRender.com)

4. The HSC are collected by centrifugation and diluted to 500,000 cells/mL. 5. Polybrene is added to the cells as a transduction enhancer at concentration of 24 μg/mL. 6. 500 μL of cells is then added to each well. 7. The plate lids are sealed with parafilm to prevent aerosolization and escape of virus and centrifuged at 1500 × g for 90 min at 22 °C. 8. 3 mL of medium containing 5% GM-CSF-conditioned medium with 2 μM β-estradiol is added to each well, after centrifugation. In the presence of GM-CSF, these immortalized cells become monocyte-committed progenitor-like cells (“GB8s”). 9. The plates are then incubated overnight at 5% CO2, 37 °C. 10. The following day, a half medium exchange is performed in order to dilute the polybrene concentration in each well. 11. Cells are then passaged every couple of days (2–3 days), where non-transduced cells differentiate and adhere to the bottom of the plate, and only the supernatant is passaged, until the conditionally immortalized monocyte-committed progenitor-like cell line is created. 12. Schematic for the protocol for transduction with ER-HoxB8 lentivirus (see Fig. 7). 3.5 Macrophage Differentiation

1. Collect GB8 cells and centrifuge at 200 × g for 10 min (see Note 10). 2. Resuspend in 20 mL of base medium containing 20% L929conditioned medium with 10 μM fulvestrant (estrogen receptor antagonist) to allow for more efficient differentiation.

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Fig. 7 Schematic for the protocol for transduction with ER-HoxB8 lentivirus. (Created with BioRender.com)

3. Plate the cells into two 10 cm polystyrene Petri dishes. 4. On Day 2, add 10 mL base medium containing 20% L929conditioned medium to each Petri dish. 5. On Day 6, subculture the cells and plate on fresh 10 cm Petri dishes in 10 mL base medium containing 20% L929conditioned medium (see Note 11) as follows: remove medium from cells and replace with cold PBS. Incubate at 4 °C for 10 min. Gently dislodge cells with a cell scraper and centrifuge at 200 × g for 10 min. Resuspend cells in base medium containing 20% L929-conditioned medium and plate onto 10 cm polystyrene Petri dishes. 6. On Day 8, add 10 mL base medium containing 20% L929conditioned medium to each Petri dish. 7. The cells are fully differentiated 10 days following initial differentiation (see Note 12). These differentiated cells express the common murine monocyte/macrophage markers CD11b and F4/80 (see Fig. 8). The cell morphology of both the GB8 cells and macrophages derived from GB8 cells was imaged (see Figs. 9 and 10). 8. Schematic for the protocol for macrophage differentiation (see Fig. 11).

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Fig. 8 Macrophages derived from GB8 cells express the common murine monocyte/macrophage markers CD11b and F4/80. These macrophages derived from GB8 cells were screened for surface expression of CD11b and F4/80 by flow cytometry. The representative dot plot of FITC-conjugated anti-CD11b and PerCPconjugated anti-F4/80

4

Notes 1. GM-CSF-conditioned medium is generated by conditioning supernatant from stably selected CHO cells transfected with a construct directing expression of GM-CSF in base medium and collecting supernatant. 2. The ER-HoxB8 fusion protein was synthesized by fusing the estrogen-binding domain of the human estrogen receptor to the N-terminus of the human HoxB8 protein. This was cloned into a lentiviral transfer vector pCDH-MSCV-MCS-EF1α-Puro (System Biosciences, Palo Alto, CA). The mouse stem cell virus (MSCV) promoter allows for higher expression of the ER-HoxB8 fusion protein in mouse hematopoietic cells [5]. 3. The longer the plates are coated with polylysine, the better the HEK293 cell attachment. 4. Warm the base medium in a 37 °C water bath before using.

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Fig. 9 Brightfield images of macrophages derived from GB8 cells after staining with Diff-Quik. Macrophages derived from GB8 cells were plated on coverslips and stained using Diff-Quik. Coverslips were then mounted onto slides and observed on an optical microscope with a 40× objective

Fig. 10 Brightfield images of GB8 cells and macrophages derived from GB8 cells. Cells were plated on a μ-slide 8-well plate overnight in a 37 °C culture incubator. Cells were then observed on a Leica TCS SP confocal microscope with a 63× objective. (a) GB8. (b) GB8-MØ

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Fig. 11 Schematic for the protocol for macrophage differentiation. (Created with BioRender.com)

5. HEK293T cells produce far less virus when they have been passaged 12 times or more from frozen; in order to optimize viral yield, use as low a passage of HEK293T cells as practicable. 6. Warm the transfection medium in a 37 °C water bath before using. 7. All media, cells, and culture plates, following transfection, will contain virus; therefore, treat it as biohazardous as it has the potential to infect humans. 8. The sucrose should be added gently, and a clear interface should appear. 9. Virus is found in the pellet of this centrifugation. 10. The best conditions to differentiate conditionally immortalized monocyte-committed progenitor-like cells are to culture them to 70% confluency in a T25 flask with medium containing GM-CSF-conditioned medium, respectively, and 2 μM β-estradiol prior to differentiating. 11. Subculture the cells depending on the confluency of the cells. Ideal subculturing can range between 1:2 and 1:10. 12. Cells are ready for use 10–14 days following initial differentiation. References 1. Kondo M (2010) Lymphoid and myeloid lineage commitment in multipotent hematopoietic progenitors. Immunol Rev 238:37–46 2. Wang GG, Calvo KR, Pasillas MP, Sykes DB, H€acker H, Kamps MP (2006) Quantitative production of macrophages or neutrophils ex vivo using conditional Hoxb8. Nat Methods 3:287– 293 3. Odegaard JI, Vats D, Zhang L, RicardoGonzalez R, Smith KL, Sykes DB, Kamps MP, Chawla A (2007) Quantitative expansion of ES cell-derived myeloid progenitors capable of

differentiating into macrophages. J Leukoc Biol 81:711–719 4. Lail SS, Arnold CR, de Almeida LG, McKenna N, Chiriboga JA, Dufour A, Warren AL, Yates RM (2022) Hox-driven conditional immortalization of myeloid and lymphoid progenitors: uses, advantages, and future potential. Traffic 23:538–553 5. Ramezani A, Hawley TS, Hawley RG (2000) Lentiviral vectors for enhanced gene expression in human hematopoietic cells. Mol Ther 2:458– 469

Chapter 9 Quantitative Immunofluorescence to Study Phagosome Maturation and Resolution Me´lanie Mansat, Roya M. Dayam, and Roberto J. Botelho Abstract Cells such as macrophages and neutrophils can internalize a diverse set of particulate matter, illustrated by bacteria and apoptotic bodies through the process of phagocytosis. These particles are sequestered into phagosomes, which then fuse with early and late endosomes and ultimately with lysosomes to mature into phagolysosomes, through a process known as phagosome maturation. Ultimately, after particle degradation, phagosomes then fragment to reform lysosomes through phagosome resolution. As phagosomes change, they acquire and divest proteins that are associated with the various stages of phagosome maturation and resolution. These changes can be assessed at the single-phagosome level by using immunofluorescence methods. Typically, we use indirect immunofluorescence methods that rely on primary antibodies against specific molecular markers that track phagosome maturation. Commonly, progression of phagosomes into phagolysosomes can be determined by staining cells for Lysosomal-Associated Membrane Protein I (LAMP1) and measuring the fluorescence intensity of LAMP1 around each phagosome by microscopy or flow cytometry. However, this method can be used to detect any molecular marker for which there are compatible antibodies for immunofluorescence. Key words Immunofluorescence, Antibodies, Lysosomes, LAMP1, Organelles, Macrophages, Phagosome maturation

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Introduction During phagocytosis, particulate matter such as bacteria and apoptotic bodies are sequestered into phagosomes [1, 2]. Phagosomes then undergo a striking alteration in their properties through phagosome maturation. This process is typically thought to follow a sequence, whereby phagosomes first fuse with early and late endosomes and ultimately with lysosomes, transforming into phagolysosomes [1–3]. Interestingly, as particles are degraded, phagosomes then undergo a recycling process whereby they fragment into smaller vesicles in a manner dependent on membrane contact sites, lipid

Roberto J. Botelho (ed.), Phagocytosis and Phagosomes: Methods and Protocols, Methods in Molecular Biology, vol. 2692, https://doi.org/10.1007/978-1-0716-3338-0_9, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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signaling, the cytoskeleton, and clathrin [2, 4, 5]. Thus, maturation converts phagosomes from inert to highly acidic and degradative organelles that catalyze the decomposition of the internalized particle [1, 2]. Through this process, phagosomes acquire and divest various proteins and lipids associated with each stage of phagosome maturation: from a plasma membrane-like to a lysosome-like stage and to its eventually fragmentation, assuming the particles are degradable [1, 2, 6]. For example, phagosomes lose phosphatidylinositol4,5-bisphosphate and actin filaments associated with the plasma membrane [7–9], then acquire and rapidly eliminate EEA1 and phosphatidylinositol-3-phosphate associated with early endosomes [10–12], and ultimately acquire endo-lysosomal proteins such as LAMP1, V-ATPase complex, and the Rab7 GTPase and effectors like RILP [10, 13–17]. While this is the canonical phagosome maturation, many infectious agents have evolved strategies that circumvent phagosome maturation. For example, Mycobacterium tuberculosis prevents fusion of phagosomes with lysosomes after they are engulfed by macrophages, thereby protecting themselves from lysosomal degradation [18–22]. To understand phagosome maturation and how pathogens usurp this machinery, we need methods to track maturation progress. Immunofluorescence techniques are commonly used to determine and visualize phagosome maturation by staining for well-characterized markers of phagosome maturation, in particular EEA1 and Hrs for early phagosomes and LAMP1 for phagolysosomes [6]. Typically, we employ indirect immunofluorescence, whereby fixed cells are exposed to primary antibodies against a specific marker, followed by a fluorescently labeled secondary antibody that binds to the primary antibody [23]. Cells can then be semiquantitatively or quantitatively analyzed by confocal or epifluorescence microscopy [24] or by flow cytometry. Flow cytometry can measure the total intensity of the fluorescent tag per particle (could be a cell or isolated phagosomes) for thousands of particles, but does not provide any information about the localization of the target protein [25]. In contrast, microscopy can provide us with the location and relative fluorescence signal but is typically low-throughput [24]. Here, we detail methodology to quantify phagolysosome maturation in macrophage cell lines using immunofluorescence against LAMP1 and clathrin, which is important for phagosome resolution [4], of whole cells or isolated phagosomes. This method can be adopted to stain other markers, including for double-staining.

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Materials 1. RAW 264.7 macrophages (ATCC) (or other cell type). 2. Cell culture medium (complete DMEM medium): Dulbecco’s Modified Eagle Medium (DMEM) supplemented with 10% fetal bovine serum (FBS). 3. Fresh 1× phosphate buffered saline (PBS). 4. Fixation solution: freshly made 4% paraformaldehyde (PFA) in 1× PBS. 5. 100 mM glycine in 1× PBS; store at 4 °C. 6. Permeabilization solution: 100% ice-cold methanol or 0.1% Triton X-100. 7. Primary antibodies: for example, rat anti-mouse LAMP1 monoclonal antibodies (Developmental Hybridoma; see Note 1); rabbit anti-clathrin heavy chain monoclonal antibodies (Cell Signaling D3C6; see Note 1). 8. Secondary antibodies: fluorescently conjugated secondary antibodies (Donkey anti-rat or anti-rabbit). 9. Mounting medium. 10. Blocking solution: 0.5–1% bovine serum albumin (BSA) in 1× PBS. 11. Polystyrene plain (hydrophobic) microspheres beads with diameter of 3.87 μm. 12. 10 mg/mL immunoglobulin G (IgG) from human serum, reconstituted in 150 mM NaCl. 13. Bacteria expressing fluorescent protein such as pBADmCherry-Escherichia coli (E. coli) (Addgene: Plasmid #54630). 14. 100 mg/mL ampicillin stock solution. Store at -20 °C. 15. 1 M L-arabinose stock solution in milli-Q H2O. Store at -20 °C. 16. LB broth: add 10 g tryptone, 5 g yeast extract, 10 g NaCl to 1 L ddH2O. 17. Microscope slides. 18. Glass cover slips. 19. Homogenization buffer for phagosome isolation (see Note 2): 20 mM Tris (pH 7.4), 2.5 μL/mL protease inhibitor cocktail, 1 mM AEBSF, 1 mM MgCl2, 1 mM CaCl2, 1 mg RNase, 1 mg DNase. 20. Sucrose gradient for phagosome isolation: 60% sucrose in 1× PBS. 21. Microscope (e.g., spinning disk confocal).

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22. Ultracentrifuge. 23. Ultracentrifuge tubes. 24. Centrifuge with adaptor for culture plates. 25. Nutator. 26. Flow cytometer. 27. Spectrophotometer. 28. Cuvette for spectrophotometer.

3 3.1

Methods Cell Culture

1. Seed RAW 264.7 macrophages onto T25 flask in complete DMEM medium, and grow at 37 °C and 5% CO2 until cells reach 70–80% confluency (see Note 3). 2. Next day, wash cells with 5 mL PBS. 3. Then add 10 mL new complete DMEM medium and scrape cells gently. 4. Take 1 mL of cell suspension and seed cells onto 12-well plate (see Note 4) containing glass coverslips to obtain 50–60% confluency using complete DMEM (see Note 5). 5. Grow cells at 37 °C and 5% CO2.

3.2 Particle Preparation 3.2.1 mCherryEscherichia coli (E. coli) Production and Fluorescent Protein Expression

This protocol applies to bacteria carrying a plasmid for L-arabinoseinducible expression of a fluorescent protein. Bacteria without label or constitutively expression of a fluorescent protein can also be used. If using bacteria without a fluorescent protein, antibodies against the bacteria may need to be used to identify the bacteria. 1. Start an overnight preculture of the bacteria in 5 mL LB media containing 100 μg/mL ampicillin at 37 °C, shaking 200 rpm. 2. The day after, add 5 mL of the overnight bacteria culture into a 50 mL LB media (with antibiotics) at 37 °C on shaking until the OD600 is between 0.6 and 0.8. 3. Induce protein expression by adding 13 mM L-arabinose for 3 h at 37 °C, shaking at 200 rpm. From this step, keep the bacteria in the dark by wrapping tubes in foil. 4. Centrifuge the bacteria at 3000 × g for 5 min, and wash three times with PBS. Before the last wash, measure the OD600 (see Note 6). 5. Fix the bacteria with 4% PFA for 15 min. 6. Replace PFA with 100 mM glycine and incubate for another 15 min to quench the remaining PFA. 7. Wash again three times with PBS and add PBS to achieve a final suspension of bacteria of 1 OD600/mL. Store at 4 °C in the dark until use.

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1. Add 50 μL of PBS, 10 μL of polymer beads, and 10 mg/mL of human IgG into a microcentrifuge tube and use a nutator to mix it for 30 min at room temperature (see Note 7). 2. Wash excess IgG with 1× PBS and centrifuge at 1200 × g for 1 min. Remove the supernatant and repeat this step three times (see Note 8). 3. Resuspend beads in PBS (see Note 9).

3.3 Phagocytosis, Phagosome Maturation, and Phagosome Resolution

1. Add opsonized beads to the media containing cells and incubate at 37 °C and 5% CO2 for 20 min (see Note 10).

3.3.1 Phagocytosis of IgG-Coated Beads

3. Replace the PBS with pre-warmed media and incubate the cells (chase) at 37 °C and 5% CO2 for the desired time (see Note 11).

2. After 20 min pulse, remove the unbound beads from the media by washing the cells with PBS three times.

4. After the desired chase time, fix cells with 4% PFA for 15 min at room temperature. 5. Replace PFA with 100 mM glycine, and incubate for another 15 min at room temperature to quench the remaining PFA. 6. Remove glycine, add 100% ice-cold methanol, and incubate for 3–5 min on ice to permeabilize the cells (see Note 12). 7. Wash the cells three times with 1% BSA. Antibodies can now be used (see below). 3.3.2 Phagocytosis of mCherry-E. coli and Fixation

1. Resuspend the mCherry-E. coli for a final suspension of 0.0025 OD600/mL in pre-warmed cell media on a 15 mL tube (this will allow to have around five bacteria phagocytized by cells). The number of bacteria may need to be optimized depending on experiment. 2. Replace the cell medium with the one containing mCherryE. coli, and centrifuge the plate at 400 × g for 5 min. Then, incubate at 37 °C. This is the phagocytosis pulse. 3. After 30 min of phagocytosis, remove the unbound bacteria from the media by washing the cells with PBS three times (see Note 13). 4. Replace the PBS with pre-warmed media, and incubate the cells at 37 °C and 5% CO2 for the desired time (see Note 11). This is the chase period. 5. After the desired chase time, fix cells with 4% PFA for 15 min at room temperature. 6. Replace PFA with 100 mM glycine, and incubate for another 15 min at room temperature to quench the remaining PFA.

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7. Remove glycine, and wash three times with PBS. If the use of antibodies is needed, replace with a permeability solution such as 0.1% Triton X-100, and incubate for 10 min to permeabilize the cells (see Note 12). 8. Wash the cells three times with PBS and incubate 1 h with 1% BSA (blocking step before using antibodies). 3.4 Antibody Staining of Whole Phagocytes

1. Add desired primary antibodies according to manufacturer’s specifications or after optimization. For example, we use rat anti-LAMP1 antibody with a dilution of 1:200 in PBS, and incubate for 1 h at room temperature (see Note 14). Another example, we use rabbit anti-clathrin antibody with a dilution of 1:200 in PBS, and incubate for 1 h at room temperature. 2. Wash the cells three times with 0.5% BSA every 5 min for 15 min to remove the unbound primary antibodies. 3. Add secondary antibodies with a dilution of 1:1000 in 1× PBS, and incubate for 1 h at room temperature in the dark (see Note 15). 4. Wash the excess secondary antibodies three times with 0.5% BSA every 5 min for 15 min. 5. Mount the cover slips on a microscope slide using mounting medium, and store the slide in a microscope slide box at 4 °C until microscopy.

3.5 Microscopy and Analysis of Whole Phagocytes 3.5.1 Maturation of Intact Phagosomes

There are different ways to quantify phagosome maturation using microscopy, from semiquantitative to quantitative. Below we describe both. 1. Acquire images (see Note 16) of 10–15 fields of view per condition using a confocal microscope (see Note 17). 2. Open the images using Image J. If image is an RGB stack, split the channels into DIC and fluorescence (Image J: Image → color → split channels). 3. Quantification (a) For semiquantitative phagosome maturation using visual inspection, convert the image of interest to 8-bit. Then use the pseudo-color processing (see Note 18) to help determine the intensity of LAMP1 (or other signal) around the phagosome (Image J: Image → lookup tables → fire). Count the total number of phagosomes in each field-of-view, and assign each phagosome as being LAMP1 positive, partial, or negative based on the intensity of LAMP1 around each phagosome (Fig. 1a, c) (see Note 19). (b) For quantitative phagosome maturation, draw regions of interest around all phagosomes present per cell. Measure

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Fig. 1 LAMP1 acquisition by phagosomes by whole-cell immunofluorescence. (a) Phagosomal acquisition of LAMP1 in control and PIKfyve inhibited (apilimod, a PIKfyve inhibitor) RAW macrophages. Cells were fixed and stained with LAMP1 antibodies, arrows point to LAMP1-positive phagosomes, and arrow heads point to LAMP1 negative phagosomes. (b) Quantitative analysis of phagosome maturation in whole cell: mean intensity of LAMP1 was measured around each phagosome in control and apilimod-treated cells. Images can be 16-bit (preferred) or 8-bit (as shown here). (c) Semiquantitative analysis of phagosome maturation in RAW macrophages: applying pseudo-color to the images and assigning each phagosome as LAMP1 positive, partial, or negative. Using 8-bit images, white-yellow color indicates high intensity of LAMP1 with grayscale intensities of 225-180 and was assigned LAMP1 positive, orange to red indicates the partial intensity of LAMP1 (grayscale intensities of 180-80), and purple to blue indicates the absence of LAMP1 (grayscale intensities of 80-0) around the phagosomes, which was assigned LAMP1 negative

average fluorescence intensity for each region of interest (phagosome). Include a background area outside of cells. Import values to a spreadsheet for processing such as background subtraction and averaging per cell. 3.5.2 Imaging and Quantification of Fragmented Phagosomes

1. Acquire images (see Note 16) of 10–15 fields of view per condition using a confocal microscope (see Note 17). 2. Open the images using Image J. If necessary, split the channels to have independent images regarding the fluorescence (Image J: Image → color → split channels) (Fig. 2a, b).

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Fig. 2 Whole-cell immunofluorescence to study phagosome maturation and resolution using bacteria expressing a fluorescent protein. (a) RAW macrophages engulfed mCherry-E. coli, which were then chased for 1 h, 5 h, or overnight to elicit maturation and resolution. Cells were then fixed and stained with LAMP1 antibodies (in green). Scale bar = 10 μm. (b) Phagosome maturation was followed as in (a). Cells were then fixed and stained with clathrin antibodies (in green). Scale bar = 10 μm. Magnification of the boxed area shown on the bottom. Scale bar = 5 μm. (c) Quantitative analysis of the phagosome fragmentation during the time: fragmented phagosomes were separated in small (below 1.54 μm2) and large (above 1.55 μm2) particles. (d) Ratio between the small and the large particles during the phagocytosis essay

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3. First, a threshold is set to exclude background and to select only the fluorescent signal (Image J: Image → Adjust → Threshold) (see Note 20). 4. This threshold is converted into a mask (Image J: Process → Binary → Convert to mask). 5. A watershed is applied to segment the particles (Image J: Process → Binary → Watershed) (see Note 21). 6. Then, particles below 1.54 μm2 and particles above 1.55 μm2 are counted (Image J: Analyze → Analyze particles). Those numbers are divided by the number of cells in the image (Fig. 2c, d) (see Note 22). 3.6 Immunofluorescence and Flow Cytometry Analysis of Isolated Phagosomes

1. Grow RAW macrophages to 80% confluency in T-25 culture flasks with complete DMEM (one flask for each condition, e.g., control, drug treatment, etc.).

3.6.1 Cell Culture for Phagosome Isolation

3. Add 3 mL of new DMEM medium into each flask.

3.6.2 Phagosome Isolation

1. Sucrose gradient preparation: pipette 1 mL of 60% sucrose into 1 mL ultracentrifuge tube and centrifuge at 50,000 × g for 1 h at 4 °C (see Note 23). After 1 h, place the tubes on ice without disturbing.

2. Next day, wash the cells with 5 mL of 1× PBS.

2. To opsonize beads, add 60 μL of beads and 120 μL of 10 mg/ mL of human IgG in 300 μL of PBS, and rotate for 30 min at room temperature. Wash beads 3× in 1 mL of PBS by centrifuging. 3. Add 300 μL of opsonized beads into each flask containing cells, and incubate for 30 min at 37 °C and 5% CO2 (see Note 24). 4. Wash cells three times with cold PBS to remove the unbound beads. 5. Replace the PBS with pre-warmed media, and incubate the cells at 37 °C and 5% CO2 for 60 min (chase) (see Note 25). 6. After 60 min chase, remove the media and add 10 mL of cold homogenization buffer to the T-25 flask, and gently scrape the cells. 7. Transfer the buffer containing the scraped cells into a 15 mL conical tube, and centrifuge at 500 × g, 4 °C, for 5 min. 8. To disrupt cells, after centrifugation, the pellet is resuspended in 1 mL of homogenization buffer and passed through a syringe with 22-gauge needle, five to ten times (see Note 26), and centrifuge for 5 min, 4 °C, at 1000 × g (see Note 27).

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9. The pellet is then resuspended in 200 μL of PBS and transferred into a 1 mL ultracentrifuge tube containing the sucrose gradient (see Note 28). 10. Beads are separated from the sample by centrifuging the sucrose gradient at 21,000 × g for 10 min at 4 °C. 11. Beads are withdrawn from the sucrose gradient using a syringe with 22-gauge needle, transferred into a fresh microcentrifuge tube, and washed with ice-cold PBS (see Note 29). 12. Fix the beads with 4% PFA for 20 min at room temperature. 13. Replace PFA with 100 mM glycine to quench the remaining PFA, and incubate for 20 min at room temperature (see Note 30). 3.6.3 LAMP1 and Clathrin Staining of Isolated Phagosomes

1. Add primary antibodies (anti-LAMP1 and anti-clathrin heavy chain antibodies [4] with a dilution of 1:200 in PBS), and incubate for 1 h at room temperature (see Note 31). 2. To remove the unbound primary antibodies, spin down the isolated phagosomes at 2000 × g for 1 min and remove the supernatant (excess primary antibodies). 3. Wash phagosomes with 500 μL of 0.5% BSA and centrifuge at 2000 × g for 1 min; repeat this step three times every 5 min for 15 min. 4. Add secondary antibodies (with a dilution of 1:1000 in PBS), and incubate for 1 h at room temperature in the dark. 5. Wash the excess secondary antibodies three times with 0.5% BSA every 5 min for 15 min by spinning down the phagosomes after each wash using a centrifuge at 2000 × g for 1 min. 6. Either resuspend the phagosomes in 20 μL of 1× PBS and mount on a microscope slide using a cover slip and mounting media and use a microscope to visualize the isolated phagosomes, or resuspend them in 500 μL of 1× PBS for flow cytometry (see Note 32).

3.6.4 Microscopy and Analysis of Isolated Phagosomes

1. Acquire z-stack images of 10–15 fields of view per condition using a confocal microscope (see Note 33). 2. Open the images using Image J and make a single image out of the z-stack (Image J: Stacks → Z-project → assign the star and stop slice → in the projection type select sum slices and press OK; Fig. 3a). 3. For semiquantitative analysis, use the pseudo-color processing (see Note 18) to help determine the intensity of LAMP1 around the phagosome (Image J: Image → lookup tables → fire). 4. For quantitative analysis, select a phagosome (ROI), and measure the mean intensity of LAMP1 (Fig. 3b).

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Fig. 3 Immunofluorescence of isolated phagosomes. (a) Matured phagosomes isolated from control and PIKfyve inhibited (apilimod) RAW macrophages and stained with LAMP1 antibodies. Arrow points to the LAMP1-positive phagosome, and arrow head points to the LAMP1 negative phagosome. (b) Quantification of the intensity of LAMP1 in isolated phagosomes in control and apilimod-treated RAW macrophages; mean intensity of LAMP1 was measured in each phagosome 3.6.5 Flow Cytometry and Analysis of Isolated Phagosomes

1. Resuspend the phagosome prep by agitating and then inject into the flow cytometer. 2. Run the flow cytometer to measure the total intensity of LAMP1 per phagosome (see Note 34). 3. While running the flow cytometer (see Note 35), select the population of the isolated phagosomes by drawing a gate, and measure the intensity of the fluorescent probe of the selected region (see Note 36). 4. Using the available flow cytometry software, obtain either a histogram or density plot, followed by statistical values such as mean and median intensity of the fluorescent probe (see Note 37).

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Notes 1. Other antibodies to LAMP1 and clathrin exist. In addition, antibodies to other markers (e.g., EEA1, LAMP2, CD63, or other protein of interest) could be employed. Lastly, if the antibodies are raised in different species, one could employ double immunofluorescence staining. In addition, always check the compatibility of an antibody for immunofluorescence or verify if unknown. 2. Homogenization buffer has to be prepared fresh. However, stock solution of Tris can be made and stored at 4 °C for a few weeks, which can be used to make the homogenization buffer. It is very important to adjust the pH of Tris to 7.4 with HCl.

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3. Other cells such as primary macrophages, neutrophils, and dendritic cells can also be used as professional phagocytes to perform this experiment. 4. For a 6-well plate, add 2 mL of cell suspension into each well containing a glass coverslip. 5. If cells are to be transfected the next day, then the confluency of the cells should be adjusted to 30–40% while seeding onto a glass cover slip. 6. The OD600 number will be useful to know the number of bacteria. For the final step, the bacteria will be stored to a concentration of 1 OD600/mL and will be ready to use. Bacteria can be used for up to a month if stored at 4 °C. 7. This is for a 12-well plate; if using 6-well, simply double the volume of all the reagents. 8. After opsonization of the beads, it is very important to wash the beads with 1× PBS, ideally three times to remove the unbound/excess IgG. The excess IgG will occupy the Fcγ receptors and may reduce the number of internalized beads. 9. There are multiple types of polymers and synthetic materials like silica and glass. Other particles like IgG-coated red blood cells or IgG-coated bacteria can be considered. 10. Phagocytosis can be synchronized by placing the plate containing cells and beads (or bacteria) on ice for 10 min or centrifuging the plate at 400 × g for 5 min at room temperature. Wash cells three times with 1× PBS, and add complete DMEM and incubate at 37 °C and 5% CO2. 11. Chase time is highly dependent on the purpose of the experiment. For example, if one is interested to look at the initial events of phagosome maturation and stain for early phagosomal markers such as EEA1, then the chase time may be 5–15 min. Late phagosome maturation should be 30–60 min. And for phagosome fragmentation or resolution, the time can vary from 3 h (the fragmentation is starting) to overnight (the fragmentation is complete) (Fig. 2a, b). 12. Triton is a typical permeabilization reagent. However, permeabilization depends on antibodies and on the structures or organelles studied. For example, other permeabilization reagents can be used like the digitonin or the saponin (transitory permeabilization) that are described to be gentler than Triton. For this specific LAMP1 antibody, it is recommended that 100% ice-cold methanol be used to permeabilize cells. Therefore, it is important to look at antibody specifications before proceeding with this proposed method.

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13. It is important to wash well to remove the remaining bacteria. Lift the coverslip during the washes to help remove trapped bacteria. Also, the phagocytic pulse time should be optimized to your own requirements. 14. It is very important to use the right dilution of the antibodies. Thus, check the data sheet provided by the antibody supplier, or optimize by trying different dilution factors. 15. Secondary antibodies need to be matched to the species in which the primary antibody was raised. For double labeling experiments, the secondary antibodies must be cross-absorbed against IgGs from the other species to prevent cross-reactivity with other secondary antibodies. Always check the data sheet provided by the supplier to ensure cross-absorption was performed. 16. Acquiring single images in a single XY plane is often sufficient for quantifying phagosome maturation (e.g., rings of LAMP1 around beads) or fragmentation as long as only particles in focus with that plane are counted. Alternatively, one can obtain and collapse a full z-stack set to quantify all phagosomes within a field-of-view, but this is not always necessary. Additionally, the number of images will also depend on number of cells per field and objective being used. A minimum of 100 cells per condition per experiment should be quantified for more robust analysis. 17. Epi-fluorescent microscope reflect light, so using polymer beads will results in a high background noise and will make it difficult to determine whether a phagosome is positive for LAMP1 or not. Therefore, confocal microscopy is ideal when using polymer beads. Epifluorescence microscopy can be used when using red blood cells or bacteria for phagocytosis though. 18. Using pseudo-coloring is a semiquantitative method to determine whether a phagosome is matured or not. Data can be quantitatively analyzed by selecting a region of interest around each phagosome, and measure the mean intensity of the fluorescent probe, in this case LAMP1, and subtract the background (Fig. 1b). 19. White-yellow color indicates high intensity of LAMP1 with grayscale intensities of 255-180 and is designated as LAMP1 positive, orange to red indicates the partial intensity of LAMP1 (grayscale intensities of 180-80), and purple to blue indicates the absence of LAMP1 (grayscale intensities of 80-1) around the phagosomes and can be designated as LAMP1 negative (Fig. 1a, c). 20. The aim of a threshold is to only consider the intensity corresponding of the particles and exclude the background. Before anything, make sure to have the dark background

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check. Click on “auto” to have an idea of the threshold you will need. Adjust it to have only the intensity corresponding of the desired particles. Keep the same threshold number for all the images of the experiment. It could be useful to test different threshold on different control images to make the right choice before using the same threshold on all the images. This step is somewhat subjective and thus the user needs to be careful. Blind quantification is recommended to reduce bias. 21. After the mask is set, two or more particles can become merged due to the binary mask. The watershed tool helps separate these objects into individual particles. This is not perfect, but assuming the same error rate per condition, it should have a neutral effect. 22. If preferred, save each cell as a separate image and apply the analysis on a cell-by-cell basis. In addition, the number of smaller fragments is likely underscored in this approach since often they are less bright and may be lost during thresholding. 23. The centrifuge should be balanced. So, after pipetting 1 mL of sucrose into each tube, use a balance to measure the weight of each tube, and adjust by adding or removing sucrose solution to ensure the tubes are within 0.5 g from each other. 24. Pulse time can be different based on the purpose of the experiment. For synchronization, place the flask on ice for 10 min (bead binding time), then wash off the unbound beads three times with cold PBS and add pre-warmed DMEM medium into each flask, and incubate for a specific chase time. Alternatively, spin beads onto cells if a plate-rotor adaptor is available. 25. Chase time is dependent on the purpose of the experiment. If looking at late maturation, then 1 h chase is enough for the phagosomes to mature. However, if one is interested to look at the early stages of maturation, then cells can be incubated for a shorter (5–10 min) period of time. 26. As alternatively to syringe-based lysis, a French press can also be used to disrupt the cells. But the orifice must be adjusted to avoid breaking phagosomes. 27. All the solutions need to be ice-cold, and the tubes containing the cells need to be placed on ice throughout the syringing process. 28. It is very important not to disturb the sucrose gradient. Therefore, add the sample dropwise onto the gradient. 29. Immunostaining of isolated phagosomes requires multiple steps of washing and centrifugation. It is very important to pay attention when removing the supernatant and not to disturb the pellet after each centrifugation. If the pellet is disrupted, sample needs to be centrifuged again.

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30. Since this is a long experiment, this is a perfect step in which one can pause and leave the beads on glycine at 4 °C until the next day to perform immunofluorescent. 31. There is no need to add 1 mL of primary antibody dilution into each tube—200 μL is sufficient. After adding the antibody solution, resuspend the pellet to ensure homogeneous staining of beads. 32. The volume of PBS being added to the phagosomes for flow cytometry is highly dependent on the amount of phagosomes obtained at the end of the experiment (200–500 μL of PBS is ideal). 33. The thickness of each slice is dependent on the size of the beads being used in the experiment. Since beads are very small (2–4 μm), we typically use 0.3 μm thickness for each slice. Number of images depends on the number of phagosomes per field. It should have >100 phagosomes total per condition per experiment. 34. Before running the sample, it is very important to resuspend isolated phagosomes by gentle pipetting in PBS to obtain a homogenous suspension. 35. Run your sample at a low speed; however, if your solution containing the isolated phagosomes is dilute and the counterwindow shows a count of less than 100 phagosomes/s, then switch to medium speed or high speed. 36. While running the sample, one might see two populations parted from each other. It is very important to select the right population (singlet phagosomes). Sometimes phagosomes can be associated and show up as a doublet (might look bigger in size and appears brighter than single phagosomes). It is easy to distinguish phagosome aggregates from singlets, since they constitute a very small population and appear bigger in the forward scatter axis. 37. There are several flow cytometry analysis software available. Older flow cytometers may not provide median intensity of the fluorescent probe. However, software such as FCS Express can use the raw data and provide the median values. References 1. Levin R, Grinstein S, Canton J (2016) The life cycle of phagosomes: formation, maturation, and resolution. Immunol Rev 273:156–179. https://doi.org/10.1111/imr.12439 2. Fountain A, Inpanathan S, Alves P, Verdawala MB, Botelho RJ (2021) Phagosome maturation in macrophages: eat, digest, adapt, and

repeat. Adv Biol Regul 82:100832. https:// doi.org/10.1016/j.jbior.2021.100832 3. Uribe-Querol E, Rosales C (2020) Phagocytosis: our current understanding of a universal biological process. Front Immunol 11:1066. https://doi.org/10.3389/fimmu.2020. 01066

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4. Lancaster CE, Fountain A, Dayam RM, Somerville E, Sheth J, Jacobelli V, Somerville A, Terebiznik MR, Botelho RJ (2021) Phagosome resolution regenerates lysosomes and maintains the degradative capacity in phagocytes. J Cell Biol 220:e202005072. https://doi.org/10.1083/jcb.202005072 ˜ o-Rendo´n F, 5. Levin-Konigsberg R, Montan Keren-Kaplan T, Li R, Ego B, Mylvaganam S, DiCiccio JE, Trimble WS, Bassik MC, Bonifacino JS, Fairn GD, Grinstein S (2019) Phagolysosome resolution requires contacts with the endoplasmic reticulum and phosphatidylinositol-4-phosphate signalling. Nat Cell Biol 21: 1234–1247. https://doi.org/10.1038/ s41556-019-0394-2 6. Levin-Konigsberg R, Mantegazza AR (2021) A guide to measuring phagosomal dynamics. FEBS J 288:1412–1433. https://doi.org/10. 1111/febs.15506 7. Levin R, Grinstein S, Schlam D (2015) Phosphoinositides in phagocytosis and macropinocytosis. Biochim Biophys Acta 1851:805–823. https://doi.org/10.1016/j.bbalip.2014. 09.005 8. Scott CC, Dobson W, Botelho RJ, CoadyOsberg N, Chavrier P, Knecht DA, Heath C, Stahl P, Grinstein S (2005) Phosphatidylinositol-4,5-bisphosphate hydrolysis directs actin remodeling during phagocytosis. J Cell Biol 169:139–149. https://doi.org/10.1083/jcb. 200412162 9. Botelho RJ, Teruel M, Dierckman R, Anderson R, Wells A, York JD, Meyer T, Grinstein S (2000) Localized biphasic changes in phosphatidylinositol-4,5-bisphosphate at sites of phagocytosis. J Cell Biol 151:1353–1367. https://doi.org/10.1083/jcb.151.7.1353 10. Vieira OV, Botelho RJ, Rameh L, Brachmann SM, Matsuo T, Davidson HW, Schreiber A, Backer JM, Cantley LC, Grinstein S (2001) Distinct roles of class I and class III phosphatidylinositol 3-kinases in phagosome formation and maturation. J Cell Biol 155:19–25. https://doi.org/10.1083/jcb.200107069 11. Fratti RA, Backer JM, Gruenberg J, Corvera S, Deretic V (2001) Role of phosphatidylinositol 3-kinase and Rab5 effectors in phagosomal biogenesis and mycobacterial phagosome maturation arrest. J Cell Biol 154:631–644. https://doi.org/10.1083/jcb.200106049 12. Vieira OV, Harrison RE, Scott CC, Stenmark H, Alexander D, Liu J, Gruenberg J, Schreiber AD, Grinstein S (2004) Acquisition of Hrs, an essential component of phagosomal maturation, is impaired by mycobacteria. Mol Cell Biol 24:4593–4604.

https://doi.org/10.1128/MCB.24.10. 4593-4604.2004 13. Kim GHE, Dayam RM, Prashar A, Terebiznik M, Botelho RJ (2014) PIKfyve inhibition interferes with phagosome and endosome maturation in macrophages. Traffic 15:1143–1163. https://doi.org/10.1111/ tra.12199 14. Sun-Wada G-H, Tabata H, Kawamura N, Aoyama M, Wada Y (2009) Direct recruitment of H+-ATPase from lysosomes for phagosomal acidification. J Cell Sci 122:2504–2513. https://doi.org/10.1242/jcs.050443 15. Harrison RE, Bucci C, Vieira OV, Schroer TA, Grinstein S (2003) Phagosomes fuse with late endosomes and/or lysosomes by extension of membrane protrusions along microtubules: role of Rab7 and RILP. Mol Cell Biol 23: 6494–6506. https://doi.org/10.1128/MCB. 23.18.6494-6506.2003 16. Pitt A, Mayorga LS, Stahl PD, Schwartz AL (1992) Alterations in the protein composition of maturing phagosomes. J Clin Invest 90: 1978–1983. https://doi.org/10.1172/ JCI116077 17. Nguyen JA, Yates RM (2021) Better together: current insights into phagosome-lysosome fusion. Front Immunol 12:636078. https:// doi.org/10.3389/fimmu.2021.636078 18. Vergne I, Fratti RA, Hill PJ, Chua J, Belisle J, Deretic V (2004) Mycobacterium tuberculosis phagosome maturation arrest: mycobacterial phosphatidylinositol analog phosphatidylinositol mannoside stimulates early endosomal fusion. Mol Biol Cell 15:751–760. https:// doi.org/10.1091/mbc.e03-05-0307 19. Chua J, Deretic V (2004) Mycobacterium tuberculosis reprograms waves of phosphatidylinositol 3-phosphate on phagosomal organelles. J Biol Chem 279:36982–36992. https://doi.org/10.1074/jbc.M405082200 20. Sachdeva K, Goel M, Sudhakar M, Mehta M, Raju R, Raman K, Singh A, Sundaramurthy V (2020) Mycobacterium tuberculosis (Mtb) lipid mediated lysosomal rewiring in infected macrophages modulates intracellular Mtb trafficking and survival. J Biol Chem 295:9192– 9210. https://doi.org/10.1074/jbc.RA120. 012809 21. Gutierrez MG (2013) Functional role(s) of phagosomal Rab GTPases. Small GTPases 4: 148–158. https://doi.org/10.4161/sgtp. 25604 22. Santucci P, Aylan B, Botella L, Bernard EM, Bussi C, Pellegrino E, Athanasiadi N, Gutierrez MG (2022) Visualizing pyrazinamide action by live single-cell imaging of phagosome

Immunofluorescence and Phagosome Maturation acidification and mycobacterium tuberculosis pH homeostasis. mBio 13:e0011722. https:// doi.org/10.1128/mbio.00117-22 23. Aoki V, Sousa JX, Fukumori LMI, Pe´rigo AM, Freitas EL, Oliveira ZNP (2010) Direct and indirect immunofluorescence. An Bras Dermatol 85:490–500. https://doi.org/10.1590/ s0365-05962010000400010

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24. St Croix CM, Shand SH, Watkins SC (2005) Confocal microscopy: comparisons, applications, and problems. BioTechniques 39:S2– S5. https://doi.org/10.2144/000112089 25. Brown M, Wittwer C (2000) Flow cytometry: principles and clinical applications in hematology. Clin Chem 46:1221–1229

Chapter 10 Approaches to Measuring Reductive and Oxidative Events in Phagosomes Shranjit S. Lail, Dale R. Balce, Johnathan Canton, and Robin M. Yates Abstract The phagosome is a redox-active organelle. Numerous reductive and oxidative systems play both direct and indirect roles in phagosomal function. With the advent of newer methodologies to study these redox events in live cells, the details of how redox conditions change within the maturing phagosome, how they are regulated, and how they influence other phagosomal functions can be investigated. In this chapter, we detail phagosome-specific, fluorescence-based assays that measure disulfide reduction and the production of reactive oxygen species in live phagocytes such as macrophages and dendritic cells, in real time. Key words Phagosome, Lysosome, Oxidation, Disulfide reduction, Redox

1

Introduction Many cellular functions and signaling pathways are modulated by reductive and oxidative (redox) chemistries. The redox environment and its influence on the function of endosomes, lysosomes, and phagosomes have recently garnered much attention [1]. Reductive mechanisms that influence disulfide reduction of protein-based antigens are required for efficient processing of disulfide-containing antigens within the phagosomal lumen [2]. In addition, reductases such as gamma-interferon inducible lysosomal thiol reductase (GILT) influence the proteolytic processing of antigens through modulation of the activities of thiol-based proteases such as the cysteine cathepsins [3, 4]. On the oxidative side, the NADPH oxidase complex (NOX2), which is responsible for the production of reactive oxygen species (ROS) in these cellular compartments, is also critical to many immune functions of phagocytes. In addition to microbial killing, NOX2-mediated ROS production can also control the activity of redox-sensitive

Shranjit S. Lail and Dale R. Balce are co-authorship. Roberto J. Botelho (ed.), Phagocytosis and Phagosomes: Methods and Protocols, Methods in Molecular Biology, vol. 2692, https://doi.org/10.1007/978-1-0716-3338-0_10, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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proteases in these organelles, resulting in differential processing of antigens [5]. Numerous cellular assays exist to monitor redox events; however, not all are suitable for specific evaluation in endosomes/lysosomes and phagosomes [6]. Phagosome-specific real-time evaluation of a variety of phagosomal chemistries such as proteolysis, acidification, and phagosome-lysosomal fusion has allowed characterization of the phagosomal lumen under various activation states of different phagocytes [7]. The adaptation of these technologies to monitor phagosome-specific redox events has led to the discovery of the characteristics and functions of phagosomal reductive and oxidative mechanisms [4, 8–10]. In this chapter, we describe four phagosome-specific fluorescence-based redox assays that measure phagosomal disulfide reduction and ROS production in real time in live macrophages. These assays evaluate the reductive or oxidative capacity of the phagosome in a population of phagocytes upon internalization of experimental particles bearing various redox reporters. We describe detailed protocols of the generation of phagosome-specific redox reporter-linked beads and the use of these beads to assess phagosome-specific redox measurements in adherent phagocytes in a multi-well format. Compared to other commonly used redox probes, the reagents used in these assays provide high signal to noise ratios and are stable in the presence of other phagosomal chemistries (e.g., pH) [6]. Furthermore, as these experimental protocols require minimal manipulation, they are easily adapted for high-throughput analysis [9, 11].

2

Materials

2.1 Preparation of Redox Reporter Beads

1. 3.0 μm carboxylate-modified silica particles/beads 5% suspension (Si-COOH, Kisker Biotech, Steinfurt, Germany) (see Note 1). 2. 1.5 mL polypropylene tubes with silicone O-rings. 3. Cyanamide. Store as desiccate at 4 °C. Protect from moisture. 4. Phosphate-buffered saline (PBS). Store at room temperature. 5. Dextran, amino-modified, 70,000 MW. Store as desiccate. 6. Coupling buffer: ddH2O, 0.1 M sodium borate, pH 8.0. Store at room temperature. 7. Quenching buffer: PBS pH 7.2 containing 250 mM glycine. Filter-sterilize through 0.22 μm filter. Store at 4 °C. 8. 5 mg/mL BODIPY FL L-cystine, in DMSO; aliquot and store at -20 °C. Protect from light. 9. Sodium azide 2% aqueous solution. Store at room temperature.

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10. MES buffer: ddH2O, 0.1 M 2-[N-Morpholino]ethanesulfonic acid (MES), 0.5 M NaCl, pH 6.0. Store at room temperature. 11. N-(3-dimethylaminopropyl)-N′-ethylcarbodiimide hydrochloride (EDC). Store as desiccate. Protect from moisture. 12. N-hydroxysulfosuccinimide sodium salt (Sulfo-NHS). Store as desiccate. Protect from moisture. 13. 5 mg/mL (+)-Biotin N-hydroxysuccinimide ester (NHS-Biotin), in DMSO; aliquot and store at -20 °C. 14. 5 mg/mL Alexa Fluor 594 carboxylic acid, succinimidyl ester (AF594-SE), in DMSO; aliquot and store at -20 °C. Protect from light. 15. 1 mg/600 μL OxyBURST Green H2HFF BSA, in coupling buffer; aliquot and store at -20 °C. Protect from light. Thaw on ice prior to use. 2.2 Cell Preparation and Handling

1. Macrophage cells such as primary bone marrow-derived macrophages (BMMØs), dendritic cells (BMDCs), Hoxb8derived macrophages (GB8-MØs), and/or macrophage-like cells (J774A.1). 2. BMMØ growth media: Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% fetal bovine serum (FBS), 2 mM l-glutamine, 1 mM sodium pyruvate, 100 U/mL penicillin-streptomycin, 20% conditioned media derived from the supernatant of M-CSF producing L929 cells. Store at 4 °C. 3. BMDC growth media: Roswell Park Memorial Institute medium (RPMI) supplemented with 5% FBS, 2 mM l-glutamine, 10 mM HEPES, 0.5 mM β-mercaptoethanol, 100 U/mL penicillin-streptomycin, 20% conditioned media derived from the supernatant of Ag8653 melanoma cells transfected with murine GM-CSF cDNA. Store at 4 °C. 4. J774 Medium: DMEM supplemented with 10% bovine growth serum (BGS), 2 mM l-glutamine, 1 mM sodium pyruvate, 100 U/mL penicillin-streptomycin. Store at 4 °C. 5. GB8 Medium: RPMI supplemented with 10% bovine growth serum (BGS), 2 mM l-glutamine, 1 mM sodium pyruvate, 100 U/mL penicillin-streptomycin, 5% conditioned media derived from the supernatant of CHO cells transfected with murine GM-CSF, 2 μM β-estradiol. Store at 4 °C. 6. 10 mM fulvestrant, in DMSO; aliquot and store at -20 °C. 7. 96-well assay plates: 96-well μClear black clear bottom. 8. T25 TC-treated flask. 9. 10 cm tissue culture (TC)-treated plate. 10. Trypsin-Versene mixture; aliquot and store at -20 °C.

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2.3 Performing Redox Measurements on Live Cells

1. Assay buffer: tissue culture grade PBS supplemented with 1 mM CaCl2, 2.7 mM KCl, 0.5 mM MgCl2, 5 mM dextrose, and 0.25% gelatin. Filter-sterilize through a 0.22 μm filter. Store at 4 °C. Warm to 37 °C prior to use. 2. 10 mg/mL anti-biotin IgG: mouse monoclonal (clone BN-34) as ascitic fluid, in PBS; aliquot and store at -20 °C. 3. 10 mg/mL anti-bovine serum albumin IgG, in PBS; aliquot and store at -20 °C. 4. 2 mg/mL IgG from human serum, in sterile water; aliquot and store at -20 °C. 5. 1 mg/mL zymosan particles, in bovine growth serum; aliquot and store at -20 °C. 6. 10 mM Amplex UltraRed, in DMSO; aliquot and store at 20 °C. Protect from light. 7. 1 U/μL horseradish peroxidase (HRP), in PBS; aliquot and store at -20 °C. 8. Fluorescence microplate reader with temperature control. Plate readers may be filter- or monochromator-based. The authors routinely use an Envision microplate reader (Perkin Elmer) to monitor phagosomal oxidative capacity and a Fluostar Optima (BMG Labtech) to monitor phagosomal disulfide reduction. 9. Clear μ-slide 8-well plate. 10. 10 mg/mL nitroblue tetrazolium, in ddH2O, made fresh each time. Aliquot and store at room temperature. 11. 4% paraformaldehyde, in PBS; aliquot and store at 22 °C. 12. 50 mM ammonium chloride, in PBS; store at 4 °C. 13. Confocal microscope. The authors routinely use a Leica TCS SP confocal microscope to visualize cells.

3

Methods

3.1 Preparation of Redox Reporter Beads 3.1.1 Preparation of Dextran-Linked ReductaseReporter Beads

1. 50 mg of carboxylate-modified silica beads (1 mL of manufacturer stock solution) is transferred to a low binding 1.5 mL screw-capped polypropylene tube. Beads are pelleted and washed twice with PBS by centrifugation at 6000 × g at room temperature for 30 s using a benchtop microtube centrifuge and vortex. 2. 30 mg of the heterobifunctional cross-linking reagent cyanamide is freshly dissolved in 1 mL PBS and incubated with the beads at room temperature with agitation for 15 min. 3. During the incubation period, 10 mg of amine-modified dextran is dissolved in 1 mL coupling buffer.

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4. The beads are washed twice with ice-cold coupling buffer to remove excess cyanamide. The cyanamide-activated beads are then resuspended in the dextran/coupling buffer solution and incubated at room temperature with agitation for 2 h (see Note 2). 5. The beads are washed twice with 1 mL PBS to remove unconjugated amine-modified dextran, and the remaining aminereactive groups are quenched by incubation with quenching buffer for 5 min at room temperature with agitation. The dextran-coupled beads can be now conjugated with BODIPY FL L-cystine (steps 6–8) or can be stored at 4 °C for later use in 1 mL quenching buffer with 10 μL of 2% sodium azide. 6. Dextran-coupled beads are washed twice with 1 mL MES buffer and resuspended in 100 μL MES buffer in a 1.5 mL screw-capped polypropylene tube. 7. 20 μL of 16 mg/mL EDC in MES (make fresh and use immediately), 27.5 μL of 32 mg/mL sulfo-NHS in MES (make fresh and use immediately), and 20 μL of 5 mg/mL BODIPY-FL L-cystine in DMSO are added (in order as listed with vortexing between additions) to the dextran-coupled bead suspension and incubated with agitation for 2–4 h at room temperature in the dark. 8. The beads are washed twice with 1 mL coupling buffer and resuspended in 500 μL coupling buffer (see Note 3). 9. 5 μL of 5 mg/mL NHS-Biotin and 2 μL of 5 mg/mL AF594SE are added to the resuspended beads and incubated with agitation for 1–2 h at room temperature in the dark. 10. Unreacted NHS-Biotin and AF594-SE are quenched and removed by washing beads twice with quenching buffer. 11. The now-completed dextran-linked reductase-reporter beads are resuspended in 1 mL quenching buffer with 10 μL of 2% sodium azide as a preservative, enumerated using a hemocytometer and stored at 4 °C in the dark. 3.1.2 Preparation of ROS-Reactive OxyBURST Beads

1. 5 mg of carboxylate-modified silica beads (0.1 mL of manufacturer stock solution) is transferred to a low binding 1.5 mL screw-capped polypropylene tube. Beads are pelleted and washed twice with PBS by centrifugation at 6000 × g at room temperature (RT) for 30 s using a benchtop microtube centrifuge and vortex (see Notes 4 and 5). 2. Beads are resuspended in a solution containing 25 mg of the heterobifunctional cross-linker cyanamide in 1 mL PBS and incubated with agitation for 15 min at room temperature. 3. The cyanamide-activated beads are then washed twice with ice-cold coupling buffer (to remove excess cyanamide) and

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resuspended in 100 μL of the OxyBURST/coupling buffer solution (see Note 2). The bead/OxyBURST/coupling buffer suspension is then incubated with agitation in the dark at room temperature for 1–3 h (see Note 6). 4. The beads are washed twice with 1 mL coupling buffer and then resuspended in 200 μL coupling buffer containing 1 μL of 5 mg/mL AF594-SE and incubated with agitation in the dark at room temperature for an additional 15 min. 5. The now-completed OxyBURST-linked reporter beads are washed twice and resuspended in 1 mL PBS, enumerated using a hemocytometer and stored at 4 °C in the dark. For best results, the beads should be used within 5–7 days. 3.2 Cell Preparation and Handling 3.2.1 BMMØs and BMDCs

1. Bone marrow is flushed from the tibias, femurs, and ilia of mice with DMEM (for BMMØs) or RPMI (for BMDCs) and centrifuged at 230 × g at 4 °C for 10 min. 2. To obtain BMMØs, freshly isolated marrow is resuspended in BMMØ growth media and plated onto non-treated petri dishes (approximately eight to ten dishes per mouse). Seven days after initial plating, BMMØs are diluted 1:2 and replated onto new petri dishes in BMMØ growth media. Approximately 10–14 days after initial plating, BMMØs are ready for use. 3. To obtain BMDCs, freshly isolated marrow is resuspended in RPMI containing 10% fetal bovine serum, 2 mM L-glutamine, 1 mM sodium pyruvate, and 100 U/mL penicillinstreptomycin and cultured in a 75 cm2 tissue culture flask overnight. The next day, nonadherent cells are harvested from the flask, resuspended in BMDC growth media, and plated onto 100 mm treated tissue culture dishes (approximately six to eight dishes per mouse). Every 2 days for 10 days, half of the growth media is removed from the culture dish and replenished with new growth media. Approximately 10 days after initial plating, BMDCs are ready for use. 4. To harvest BMMØs, growth media are removed from confluent, fully differentiated BMMØs and replaced with cold PBS. The cells are incubated at 4 °C for 10 min to facilitate BMMØ detachment from the culture dish. BMMØs are gently dislodged with a cell scraper and centrifuged at 230 × g at 4 °C for 10 min. 5. To harvest BMDCs, growth media are removed and 5 mL pre-warmed trypsin-EDTA is added to the culture dish. Following a 5 min incubation at 37 °C, cells are gently dislodged with a cell scraper and centrifuged at 230 × g at 4 °C for 10 min. 6. BMMØs and BMDCs are plated onto 96-well assay plates to 100% confluency. Approximately 1.2 × 105 cells are required

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per well in a volume of 100 μL of complete medium to establish a confluent monolayer. Plates are kept at room temperature for 15 min before incubating at 37 °C overnight to ensure an even monolayer will be established (see Note 7). 3.2.2

GB8-MØs

1. Thaw a vial of frozen HoxB8-conditionally immortalized myeloid progenitor cells (GB8s) in a 37 °C water bath (see Note 8). Add cells to 10 mL of pre-warmed GB8 medium and centrifuge at 200 × g for 10 min. 2. Resuspend GB8 cells in 10 mL of pre-warmed GB8 medium, and plate into a T25 flask. Incubate at 5% CO2, 37 °C. 3. Passage GB8 cells when confluent into a new T25 with pre-warmed GB8 medium at a dilution of 1:10. 4. Differentiate GB8 cells into GB8 macrophages (GB8-MØs) using BMMØ growth media with 10 μM fulvestrant (see Note 8). Approximately 10–14 days after initial plating, GB8-MØs are ready for use. 5. GB8-MØs are plated onto 96-well assay plates to 100% confluency. Approximately 1.2 × 105 cells are required per well in a volume of 100 μL of complete medium to establish a confluent monolayer. Plates are kept at room temperature for 15 min before incubating at 37 °C overnight to ensure an even monolayer will be established (see Note 7).

3.2.3

J774s

1. Thaw a vial of frozen J774 cells in a 37 °C water bath. Add cells to 10 mL of pre-warmed J774 medium and centrifuge at 200 × g for 10 min. 2. Resuspend in 10 mL of pre-warmed J774 medium and plate onto 10 cm TC-treated plate. Incubate at 5% CO2, 37 °C. 3. Passage cells when confluent into new 10 cm TC-treated plate at dilution of 1:10. Dispose of old media, rinse with 2 mL PBS, and add 2 mL of trypsin. Incubate for 4 min at 37 °C. Add 10 mL of pre-warmed J774 medium and remove adherent cells using a cell scraper. Collect cells in 50 mL conical tube and pellet cells at 200 × g for 10 min, followed by resuspension in 10 mL pre-warmed J774 medium. Add 1 mL of cells onto new 10 cm TC-treated plate that contains 9 mL of pre-warmed J774 medium. 4. J774 are plated onto 96-well assay plates to 100% confluency. Approximately 1.2 × 105 cells are required per well in a volume of 100 μL of complete medium to establish a confluent monolayer. Plates are kept at room temperature for 15 min before incubating at 37 °C overnight to ensure an even monolayer will be established (see Note 7).

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3.3 Performing Redox Measurements in Live Cells 3.3.1 Assessment of Phagosome-Specific Disulfide Reduction

1. Shortly prior to phagosomal assessment, the well medium is removed, and the adherent cell monolayer is washed twice with 100 μL of pre-warmed assay buffer (37 °C). 2. 50 μL of pre-warmed assay buffer is added to each well prior to the addition of experimental beads. In this volume, cells can be treated with water soluble inhibitors/compounds if desired. In addition, background readings of cells without experimental beads can be obtained at this stage. 3. 100 μL of the reductase bead stock is washed twice with 1 mL PBS (gentle vortex, or by pipetting up and down) and resuspended in 200 μL PBS containing 5 μL anti-biotin IgG (10 mg/mL). Beads are incubated with agitation in the dark at room temperature for 15 min (see Note 9). 4. Beads are washed once (gentle vortex, or by pipetting up and down) with 1 mL PBS and resuspended in 1 mL PBS. 5. The appropriate concentration of beads required per well is then determined. Approximately two to four beads per cell are optimal. As a starting point, 20 μL of the opsonized reductase bead stock solution is resuspended in 1 mL assay buffer. 50 μL is added to a test well containing a confluent monolayer of cells. After a 5–10 min incubation period, the number of beads per cell is visually inspected under a light microscope. The bead dilution is then adjusted accordingly to achieve approximately two to four beads per cell. 6. 50 μL of the reductase beads at the appropriate working dilution in assay buffer is added to each experimental well (see Note 10). 7. The assay plate is inserted into the plate reader, and substrate fluorescence (SF) 488/520 nm and calibration fluorescence (CF) 594/620 nm (excitation λ/emission λ) are recorded. Disulfide reduction is indicated by an increase in fluorescence as the disulfide linker is cleaved resulting in the de-quenching of the disulfide-linked BODIPY fluorophores (see Note 11). 8. Substrate/calibration fluorescence is recorded every 2–5 min for 1–2 h. 9. Data is exported into a spreadsheet application such as Microsoft Excel. RFU (relative fluorescence unit) = (SF - SFbackground)/(CF - CFbackground) is calculated for each time point and plotted against time. An example of analyzed data is shown in Fig. 1.

3.3.2 Assessment of Phagosome-Specific ROS Production

1. Shortly prior to phagosomal assessment, the well medium is removed, and the adherent cell monolayer is washed twice with 100 μL of pre-warmed assay buffer (37 °C). 2. 50 μL of pre-warmed assay buffer is added to each well prior to the addition of experimental beads. In this volume, cells can be

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Fig. 1 Assessment of phagosome-specific disulfide reduction in (a) BMMØs and (b) BMDCs derived from C57Bl6 (WT) or GILT-deficient (GILT-/-) mice. Phagosomal disulfide reduction is significantly decreased in the absence of GILT. As disulfide reduction in the phagosome is sensitive to the presence of ROS, disulfide reduction is increased in the presence of the NOX2 inhibitor diphenyleneiodonium

treated with water soluble inhibitors/compounds if desired. In addition, background readings of cells without experimental beads can be obtained at this stage. 3. OxyBURST bead stock is washed once with 1 mL PBS (gentle vortex, or by pipetting up and down) and resuspended in 200 μL PBS containing 5 μL anti-BSA IgG (10 mg/mL). Beads are incubated with agitation in the dark at room temperature for 30 min (see Note 9). 4. Beads are washed once (gentle vortex, or by pipetting up and down) with 1 mL PBS and resuspended in 1 mL PBS. 5. The appropriate concentration of beads required per well is then determined. Approximately two to four beads per cell are optimal. As a starting point, 20 μL of the opsonized OxyBURST bead stock solution is resuspended in 1 mL assay buffer. 50 μL is added to a test well containing a confluent monolayer of cells. After a 5–10 min incubation period, the number of beads per cell is visually inspected under a light microscope. The bead dilution is then adjusted accordingly to achieve approximately two to four beads per cell. 6. 50 μL of the OxyBURST bead working solution in assay buffer is then added to each well (see Note 10). 7. The assay plate is inserted into the plate reader, and substrate fluorescence (SF) 488/520 nm and calibration fluorescence (CF) 594/620 nm (excitation λ/emission λ) are recorded. Oxidation of the OxyBURST substrate by phagosomal ROS leads to increased fluorescence of the fluorophore (see Note 12).

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Fig. 2 Assessment of phagosome-specific ROS production in (a) BMMØs and (b) BMDCs derived from C57Bl6 (WT) or NOX2 complex-deficient (Cybb-/- and Ncf-/-) mice. Phagosomal ROS is effectively absent in gp91-deficient (Cybb-/-) and p47-deficient (Ncf-/-) phagocytes

8. Data is exported into a spreadsheet application such as Microsoft Excel. RFU = (SF - SFbackground)/(CF - CFbackground) is calculated for each data point and plotted against time. An example of analyzed data is shown in Fig. 2. 3.3.3 Measurement of Extracellular H2O2

To complement the OxyBURST bead assay described above, a simple assay to measure extracellular release of H2O2 in response to the phagocytosis of opsonized zymosan particles can be used. Amplex UltraRed reacts with H2O2 in a stoichiometry of 1:1 to produce the fluorescent resorufin; thus, this assay can be used to quantify extracellular concentrations of H2O2 when regressed to a standard curve. 1. Shortly prior to phagosomal assessment, the well medium is removed, and the adherent cell monolayer is washed twice with 50 μL of pre-warmed assay buffer (37 °C). 2. Serum-opsonized zymosan particles (100 mg zymosan/mL fetal bovine serum) are resuspended in assay buffer to a final concentration of 500 μg/mL. 50 μL is added to each well. 50 μL of assay buffer without zymosan are added as background controls. 3. The assay plate is incubated for 2 h in a 37 °C culture incubator. 4. The well supernatant is transferred to a fresh 96-well assay plate. 5. In a clean 15 mL conical tube, Amplex UltraRed and HRP are added to freshly degassed PBS at a concentration of 20 μM and 40 U/mL, respectively. 50 μL of the Amplex UltraRed/HRP/ PBS solution is immediately added to each well supernatant. The assay plate is incubated for 30 min in the dark at room temperature.

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Fig. 3 Assessment of extracellular H2O2 produced by BMMØs, GB8-MØs, and J774 cells upon internalization of serum-opsonized zymosan particles. Data is presented as relative to BMMØs. Error bars denote SEM. *P < 0.05

6. Amplex Red fluorescence ~571/585 nm (excitation λ/emission λ) is then measured using a microplate reader. 3–5 measurements from each well and from each well replicate are collected and averaged. 7. Data is exported into a spreadsheet application such as Microsoft Excel. Background values from no zymosan control samples are deducted from each data set. An example of analyzed data is shown in Fig. 3. 3.3.4 Measurement of Reactive Oxygen Species Production

To highlight the phagosomal reactive oxygen species production, a simple assay to measure reactive oxygen species production in response to activation of the phagocyte by an opsonized phagocytic target, zymosan particles, can be used [12, 13]. The production of reactive oxygen species is measured by the production of formazan, an insoluble deposit that becomes evident in phagosomes where reactive oxygen species are formed. 1. Plate 300 μL of phagocytes on a μ-slide 8-well plate (250,000 cells per well), and incubate overnight in a 37 °C culture incubator. 2. For serum opsonization of the zymosan, pellet 10 mg of zymosan at 10,000 × g for 5 min. Resuspend the pellet in 1 mL of bovine growth serum. Incubate for 1 h at 37 °C. Wash the zymosan with PBS three times after the incubation. 3. Discard the media from the μ-slide 8-well plate and gently wash with PBS three times. 4. Add 50 μL of serum-opsonized zymosan particles (10 mg/mL) to the wells.

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Fig. 4 Assessment of reactive oxygen species produced by C57Bl6 (WT) and NOX2 complex-deficient (Cybb-/-) GB8-MØs cells upon internalization of serum-opsonized zymosan particles. Data is presented as relative to C57Bl6 (WT) GB8-MØs. Error bars denote SEM. *P < 0.05

5. Add 50 μL of NBT to each of the wells and incubate at 37 °C, 7% CO2, for 2 h. 6. The cells are then gently washed with PBS three times. The cells are fixed using 200 μL 4% PFA and incubated at 22 °C for 20 min. 7. The cells are gently washed with PBS three times again. 200 μL of ammonium chloride is added to the cells which are incubated at 22 °C for 5 min. The cells are washed gently with PBS three times before imaging. 8. The cells are then visualized on a confocal microscope. Brightfield images were taken with a 63× objective. The NBT reagent reacts with reactive oxygen species to produce a black compound, formazan. The relative reactive oxygen species production was calculated by measuring the average number of formazan positive phagosomes per phagocyte. An example of analyzed data is shown in Figs. 4 and 5.

4

Notes 1. 1.5–5.0 μM silica or latex beads can also be used. 2. Washing at this step should be done quickly and dextran/ OxyBURST solution added immediately after washing. 3. Beads at this stage should be a deep orange in color, indicating successful conjugation of the BODIPY FL L-cystine substrate. 4. As OxyBURST beads must be made fresh, quantities have been adjusted to yield a small batch of beads (approximately enough for 100 wells of a 96-well assay plate).

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Fig. 5 Brightfield images of reactive oxygen species production by C57Bl6 (WT) and NOX2 complex-deficient (Cybb-/-) GB8-MØs cells upon internalization of serum-opsonized zymosan particles. Cells were then observed on a Leica TCS SP confocal microscope with a 63× objective. Arrow highlights presence of formazan. (a) C57Bl6 (WT) GB8-MØ. (b) NOX2 complex-deficient (Cybb-/-) GB8-MØ

5. PBS and coupling buffers are first degassed in a water bath sonicator prior to use. 6. Depending on the objectives of the experiment, the surface of the beads can be covalently coupled to IgG in lieu of (noncovalent) opsonization with anti-BSA IgG prior to the addition of the beads to cells (Subheading 3.3.2, step 3). Hence, if desired, 1 μL of human IgG (2 mg/mL) is added to the bead/OxyBURST solution during the 1–3 h incubation. 7. The authors usually plate a minimum of three wells for each experimental condition and additional “test” wells for empirically titrating bead numbers. 8. To generate and differentiate HoxB8-conditionally immortalized myeloid progenitor cells (GB8s), follow detailed protocol in The Derivation and Use of HoxB8-Driven Conditionally Immortalized Macrophages chapter or Lail et al. [14]. 9. IgG opsonization of the experimental beads is most efficient when performed immediately prior to use. 10. Vortex working bead solution vigorously before applying to cells to avoid clumping of beads. 11. A variety of fluorescence plate reader platforms can be used to monitor monolayer phagosomal activity; however, a few specific functionalities are required. Fluorescence plate readers

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with bottom read and height optimization are required as BMMØs and BMDCs establish adherent monolayers. Temperature control is required for kinetic measurements in live cells. 12. The same instrument protocol used to monitor BODIPY FL L-cystine de-quenching can be used to monitor OxyBURST substrate oxidation. References 1. Yates RM (2013) Redox considerations in the phagosome: current concepts, controversies, and future challenges. Antioxid Redox Signal 18:628–629 2. Collins DS, Unanue ER, Harding CV (1991) Reduction of disulfide bonds within lysosomes is a key step in antigen processing. J Immunol 147:4054–4059 3. Phan UT, Arunachalam B, Cresswell P (2000) Gamma-interferon-inducible lysosomal thiol reductase (GILT): maturation, activity, and mechanism of action. J Biol Chem 275: 25907–25914 4. Balce DR, Allan ER, McKenna N, Yates RM (2014) γ-Interferon-inducible lysosomal thiol reductase (GILT) maintains phagosomal proteolysis in alternatively activated macrophages. J Biol Chem 289:31891–31904 5. Allan ER, Tailor P, Balce DR, Pirzadeh P, McKenna N, Renaux B, Warren AL, Jirik FR, Yates RM (2014) NADPH oxidase modifies patterns of MHC class II–restricted epitopic repertoires through redox control of antigen processing. J Immunol 192:4989–5001 6. Balce DR, Yates RM (2013) Redox-sensitive probes for the measurement of redox chemistries within phagosomes of macrophages and dendritic cells. Redox Biol 1:467–474 7. Yates RM, Russell DG (2008) Real-time spectrofluorometric assays for the lumenal environment of the maturing phagosome. Methods Mol Biol 445:311–325 8. Balce DR, Li B, Allan ER, Rybicka JM, Krohn RM, Yates RM (2011) Alternative activation of

macrophages by IL-4 enhances the proteolytic capacity of their phagosomes through synergistic mechanisms. Blood 118:4199–4208 9. Balce DR, Greene CJ, Tailor P, Yates RM (2016) Endogenous and exogenous pathways maintain the reductive capacity of the phagosome. J Leukoc Biol 100:17–26 10. VanderVen BC, Yates RM, Russell DG (2009) Intraphagosomal measurement of the magnitude and duration of the oxidative burst. Traffic 10:372–378 11. Rybicka JM, Balce DR, Khan MF, Krohn RM, Yates RM (2010) NADPH oxidase activity controls phagosomal proteolysis in macrophages through modulation of the lumenal redox environment of phagosomes. Proc Natl Acad Sci 107:10496–10501 12. Canton J, Blees H, Henry CM, Buck MD, Schulz O, Rogers NC, Childs E, Zelenay S, Rhys H, Domart MC, Collinson L (2021) The receptor DNGR-1 signals for phagosomal rupture to promote cross-presentation of deadcell-associated antigens. Nat Immunol 22: 140–153 13. Canton J, Khezri R, Glogauer M, Grinstein S (2014) Contrasting phagosome pH regulation and maturation in human M1 and M2 macrophages. Mol Biol Cell 25:3330–3341 14. Lail SS, Arnold CR, de Almeida LG, McKenna N, Chiriboga JA, Dufour A, Warren AL, Yates RM (2022) Hox-driven conditional immortalization of myeloid and lymphoid progenitors: uses, advantages, and future potential. Traffic 23:538–553

Chapter 11 Measuring Phagosomal pH by Fluorescence Microscopy Gerone A. Gonzales and Johnathan Canton Abstract Dual-wavelength and dual-fluorophore ratiometric imaging has become a powerful tool for the study of pH in intracellular compartments. It allows for the dynamic imaging of live cells while accounting for changes in the focal plane, differential loading of the fluorescent probe, and photobleaching caused by repeated image acquisitions. Ratiometric microscopic imaging has the added advantage over whole-population methods of being able to resolve individual cells and even individual organelles. In this chapter, we provide a detailed discussion of the basic principles of ratiometric imaging and its application to the measurement of phagosomal pH, including probe selection, the necessary instrumentation, and calibration methods. Key words Phagosome, Dual-wavelength ratio imaging, Dual-fluorophore ratio imaging, Fluorescence microscopy, pH, Fluorescein, Oregon Green, SNARF1, CypHer5e, pHrodo

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Introduction Phagocytosis denotes a complex biological process in which relatively large particles (>0.5 μm) can be bound and engulfed by a eukaryotic cell. The process itself is evolutionarily ancient and represents the primary means by which single-celled phagotrophic eukaryotes, such as those belonging to the Dictyostelium genus, acquire nutrients [1]. In metazoans, phagocytosis is central to the maintenance of homeostasis, serving to scavenge and clear dead cells and particulate debris. In our bodies, for example, approximately 200–300 billion cells undergo apoptosis or some other form of cell death each day [2]. It is through phagocytosis (referred to as efferocytosis when targeting dead cells) that these cells are cleared and their components recycled [2]. Other homeostatic by-products cleared by phagocytosis include fibrillar β-amyloid in the brain, the shed outer segments of photoreceptor cells in the eye, and the nuclei extruded by erythroblasts at sites of erythroid maturation [3–5]. Phagocytosis also plays a central role in both the execution of innate immune defense and the initiation of the adaptive phase of immunity. Invading pathogens can be recognized by specialized

Roberto J. Botelho (ed.), Phagocytosis and Phagosomes: Methods and Protocols, Methods in Molecular Biology, vol. 2692, https://doi.org/10.1007/978-1-0716-3338-0_11, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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receptors on the surface of phagocytic cells; these specific receptors not only immobilize the microorganism but also transduce signals that result in its engulfment and elimination of the associated threat. Enclosure of the pathogen in a membrane-bound intracellular compartment, termed a phagosome, serves two functions: (i) it allows for the compartmentalized killing of the invading organism, thereby limiting the release of inflammatory agents, and (ii) it targets antigenic material to a compartment where it can be loaded onto major histocompatibility molecules for presentation to cells of the adaptive immune system [6]. The abovementioned functions of phagocytosis rely on one salient feature—the acquisition of an acidic phagosomal lumen (≤pH 5) (Figs. 4b and 5b). Various degradative enzymes are sorted into phagosomes as they mature; however, some are zymogens and are only processed into their mature, active forms at low pH. Such is the case for most of the cathepsin family [7]. Other phagosomal enzymes have acidic pH optima and therefore only function effectively in an acidic environment [6]. Only fully acidified phagosomes possess the potent hydrolytic capacity required for the degradation of internalized particles. Moreover, resultant degradation products such as nucleosides, iron, and amino acids are translocated into the cytosol by proton-coupled transporters [8, 9]. It is not surprising then that dissipating phagosomal pH with ionophores or weak bases, such as chloroquine, has been used historically to inhibit functions associated with phagosomal processing [10–12]. It is through the acquisition of the vacuolar-type ATPase (V-ATPase) that luminal acidification is achieved. Shortly after sealing, a series of fusion and fission interactions with organelles of the endocytic pathway result in the accumulation of V-ATPases on the phagosomal membrane [13]. The V-ATPase is a multimeric pump that transports protons from the cytosol to the lumen of the phagosome using energy derived from the hydrolysis of ATP [14]. Since the movement of protons across the phagosomal membrane is an electrogenic process, continued activity of the V-ATPase, without some form of charge compensation, would result in the generation of a self-inhibitory electrical potential [15–20]. Although there is a dearth of information on how the necessary charge compensation is achieved on phagosomes, it can be inferred from work done in lysosomes that either the inward movement of anions (such as Cl-) or the outward movement of cations (such as Na+ and K+) can provide a counterion current, thereby limiting the buildup of an inhibitory potential [20]. Attempts to measure phagosomal pH have been made from the very dawn of macrophage biology. Indeed, shortly after discovering phagocytes, E´lie Metchnikoff made the first approximations of phagosomal pH. By allowing phagocytic protists to engulf bits of litmus paper, he observed a color change in the sealed phagosome indicative of an acidic lumen [21]. Later, investigators began to

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adsorb indicator dyes such as neutral red, bromophenol blue, and bromocresol green to microbes that could be ingested by mammalian phagocytes [22–25]. The resultant color changes of the indicator dyes gave rough estimates of changes in pH over time. These measurements, however, were purely qualitative and not amenable to precise calibrations. Around this time, Ohkuma and Poole pioneered a more accurate, sensitive, and quantitative method to estimate the pH of a different intracellular organelle, the lysosome [26]. Their technique involved targeting a pH-sensitive fluorophore, in this case fluorescein, to the lysosomal compartment. Fluorescein has a peak fluorescence emission at 520 nm when excited at 490 nm (Fig. 1a). That emission is exquisitely sensitive

Fig. 1 Fluorescent pH sensors are most sensitive near their pKa. (a) Fluorescein (10 μg/mL) was dissolved in calibration buffers at the indicated pH values. An excitation wavelength spectral sweep was performed from 400 to 500 nm at 10 nm intervals, recording emission at 520 nm using the SpectraMax Gemini EM fluorescence plate reader. (b) Oregon Green (10 μg/mL) was dissolved in calibration buffers at the indicated pH. An excitation wavelength spectral sweep was performed from 400 to 500 nm at 10 nm intervals, recording emission at 520 nm, as above. (c) Background-subtracted fluorescence values at 490 and 440 nm excitation from (a) and (b) were used to generate a ratio of 490/440 vs. pH curve. (d) SNARF1 (5 μg/mL) was dissolved in calibration buffers at the indicated pH. An emission wavelength spectral sweep between 560 and 700 nm was performed with excitation at 488 nm, as above. Background-subtracted fluorescence values at the 640 and 580 nm emission wavelengths were used to generate a ratio of 640/580 vs. pH curve (inset)

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to changes in pH (Fig. 1a). However, other factors, such as photobleaching of the probe, can also affect the overall emission intensity (Fig. 3b). Therefore, normalizing to a second, relatively pH-insensitive, excitation wavelength (440 nm) (Figs. 1a, 3a, and 4a) allows for the measurement of changes in pH while accounting for other concerns such as photobleaching (Fig. 3b), changes in focal plane (Fig. 3c), and the amount of fluorophore. This technique is referred to as dual-excitation ratio fluorescence imaging. The ratio of the emission intensities recorded with excitation at 490 and 440 nm (ratio 490/440) can then be converted into pH by using an in situ calibration procedure. This entails clamping the pH of the intracellular compartment using ionophore-containing calibration solutions (Figs. 3a and 4a, b) [27]. This technique was promptly adapted to measure phagosomal pH by covalently coupling fluorescein to phagocytic targets [10]. Fluorescence ratio imaging remains the most sensitive method for real-time measurement of the phagosomal pH. Several variations of this technique have emerged, including the measurement of whole populations of cells using a spectrofluorometer [28], individual cells using a flow cytometer [29], and even individual phagosomes using fluorescence microscopy [30]. Each of these approaches requires careful consideration. For example, population-based methods like spectrofluorometry and flow cytometry, although useful for the rapid measurement of large populations of cells, do not distinguish between phagocytic targets that have been internalized and those that are merely bound to the cell surface; therefore, this data represents a composite of the pH in the immediate environment of both internalized and uninternalized particles. Also, recent work has shown that there is a great degree of heterogeneity between cells and even individual phagosomes [31]. Only fluorescence microscopy can resolve the idiosyncrasies of individual phagosomes. In this chapter, we will focus on the application of ratio fluorescence microscopy using several pH-sensitive fluorophores for the dynamic measurement of phagosomal pH. We will discuss the advantages and disadvantages of dual-wavelength fluorescence imaging using a single fluorophore and dual-fluorophore fluorescence imaging whereby a pH-sensitive and a pH-insensitive fluorophore are coupled to phagocytic targets. 1.1

Instrumentation

A comprehensive schematic of the required hardware for ratio fluorescence imaging can be found in a recent chapter on the measurement of lysosomal pH (see ref. 27). In short, a highintensity arc lamp is required as a light source. The type of lamp used should be matched to the desired excitation wavelengths. The two most commonly used light sources are xenon and mercury arc lamps. Xenon lamps provide a relatively even output spectrum, whereas mercury lamps are characterized by distinct, sharp emission peaks [32]. If the discrete emission peaks of the mercury lamp do

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not match the desired excitation wavelengths for the pH probe, suboptimal excitation of the probe may occur. Xenon lamps also have the added benefit of more stable emission over time, rendering them better suited for real-time or quantitative fluorescence imaging. From the arc lamp, light is carried through a fiber-optic cable to a computer-controlled, shuttered excitation filter wheel. The filter wheel should allow for a very rapid (on the order of milliseconds) transition between wavelengths. The filtered light then passes through a filter cube containing a dichroic mirror that reflects the light up through the objective lens to the biological sample. The light emitted from the fluorophore is then collected by the objective lens, passes through the dichroic mirror, and is filtered through a second computer-controlled, shuttered filter wheel. This emission filter wheel should allow for equally rapid transitions between filters, particularly if performing dual emission wavelength fluorescence imaging. The emitted light can then be collected by a chargecoupled device (CCD) camera. Ideally, the detector should have a small pixel size and a large imaging area, which allow for a large dynamic range of gray-scale intensity values as well as the capacity for shorter exposure times. These features facilitate the simultaneous measurement of individual cells or phagosomes that may be heterogeneously labeled with the fluorescent probe. Pixel binning can be applied when resolution must be sacrificed to increase sensitivity. Lastly, the appropriate software capable of integrating the exposure times, filter transitions, acquisition intervals, and other parameters is required. 1.2

Probe Selection

An initial consideration when selecting a probe to measure organelle pH is appropriate subcellular targeting. Rough estimates of phagosomal pH can be obtained by using fluorescent weak bases that partition into acidic compartments. This is the basis of several commercially available probes such as acridine orange and the LysoTracker/LysoSensor variants. The relatively low membrane permeability of the protonated form of these probes, combined with the much larger permeability of the uncharged species, results in their accumulation in acidic organelles. This is often used to determine whether or not a phagosome has acidified; however, these are purely qualitative probes and cannot be used to estimate pH values accurately. Moreover, weak base probes also label lysosomes and other acidic compartments and are therefore not phagosome-specific [27]. The accumulation of the protonated form of these acidotropic dyes can also result in alkalizing or osmotic effects in the organelle(s) where they accumulate. Also, any dynamic experiments involving the dissipation of organelle pH with ionophores or V-ATPase inhibitors will result in the irreversible loss of these dyes from the compartment being monitored.

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Lastly, some LysoTracker probes can photo-convert to species emitting at alternate wavelengths and are therefore not suitable for two-color imaging [33]. A far more specific way of targeting the pH probe to the phagosome is to covalently link it to the phagocytic target. A number of the pH-sensitive probes come as either an isothiocyanate or a succinimidyl ester. When incubated under slightly basic conditions, these amine-reactive groups allow the probes to attach covalently to primary amines, to yield a thiourea or stable amide bond, respectively. This approach has the advantage of specific phagosomal targeting, no loss of probe upon pH dissipation, and minimal alkalinizing or osmotic effects as only a finite amount of the probe is covalently bound to the phagocytic target. Upon binding protons, a number of fluorescent molecules undergo either a change in emission intensity or shifts in the emission spectrum. This feature allows for these molecules to be used as sensors of the proton concentration in cellular compartments. After correction for any fluorescence background, the emission intensities at two excitation (dual excitation) or emission (dual emission) wavelengths can be used to generate a ratio that reports the pH [34–36]. This is referred to as dual-wavelength fluorescence microscopy, and, as discussed briefly earlier, this is the basis of fluorescein pH sensitivity but also of other probes such as Oregon Green and SNARF1 (Fig. 1). Covalent coupling of these probes to phagocytic targets allows for ratio fluorescence determinations of phagosomal pH (Figs. 3a, 4a, and 5a). In some instances, the appropriate equipment for dual-wavelength fluorescence microscopy may not be readily available. A compromise is to couple two separate fluorophores to the phagocytic target—one which changes emission intensity upon binding protons (Figs. 2a, b, and 5a) and another that is insensitive to pH (Figs. 2c and 5a). This is referred to as dual-fluorophore fluorescence microscopy, and, in addition to fluorescein, Oregon Green, and SNARF1 (all of which are excited at ~490 nm), several new pH-sensitive fluorophores are amenable to this approach including pHrodo (excited at ~560 nm) and CypHer5e (excited at ~640 nm) (Figs. 2 and 5). For these pH-sensitive probes, an important consideration is their dissociation constant (pKa). Since the pKa represents the pH at which half of the probe is protonated, it also represents the pH value at which it is most sensitive to changes in proton concentration. Phagosomal pH spans a very large range of values, starting at near neutral shortly after sealing and rapidly acidifying to near pH 5.0 [37]. In addition, some cell types, such as neutrophils and human M1 macrophages, can exhibit significantly alkaline phagosomal pH values [37, 38]. This represents a challenge when selecting an appropriate pH sensor. As can be noted in Figs. 1 and 2, fluorescein (pKa = 6.4), pHrodo (pKa = 6.5), and CypHer5e (pKa = 6.1) are useful for changes in pH near their pKa

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Fig. 2 pH-sensitive and pH-insensitive fluorophores can be used in dual-fluorophore ratio imaging. (a) CypHer5e (10 μg/mL) was dissolved in calibration buffers at the indicated pH values. An emission wavelength spectral sweep between 655 and 695 was performed at 10 nm intervals with an excitation at 640 nm. (b) pHrodo (10 μg/mL) was dissolved in calibration buffers at the indicated pH values. An emission wavelength spectral sweep between 570 and 620 nm with 10 nm intervals was performed with an excitation at 560 nm. (c) AF488 (2 μg/mL) was dissolved in calibration buffers at the indicated pH values. An emission wavelength spectral sweep between 515 and 545 nm with 10 nm intervals was performed with an excitation at 488 nm. (d) Background-subtracted fluorescence values at 666 and 520 nm emission from (a) and (c) were used to generate a ratio of 666/520 vs. pH curve (blue curve). Background-subtracted fluorescence values at 590 and 520 nm emission wavelengths from (b) and (c) were used to generate a ratio of 590/520 vs. pH curve (black curve)

(Figs. 1 and 2), but they become relatively useless at more basic or acidic values. Oregon Green (pKa = 4.7) is, therefore, a more useful probe for measuring subtle changes in an acidified phagosome while being relatively useless during the early phases of phagosomal acidification (Fig. 1b, c). Neither fluorescein nor Oregon Green are ideal for measurements of the slightly alkaline phagosomal pH values obtained in neutrophils and M1 macrophages; for this another probe, SNARF1 (pKa = 7.5) is a more suitable choice (Fig. 1d) [37].

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In addition, particular care must be taken when employing dual-fluorophore fluorescence imaging for measuring phagosomal pH. In this approach, a ratio is generated from the emissions of a pH-sensitive probe (e.g., fluorescein, pHrodo, or CypHer5e) and a second pH-insensitive probe (e.g., rhodamine or Alexa Fluor 488). In many cases, differential bleaching of the two probes is unavoidable and can result in illumination- dependent changes of the ratio that are not indicative of actual pH changes (Fig. 3d). In view of the above caveats, the stage of phagosome maturation and the desired experimental manipulations should be considered carefully when selecting the appropriate pH sensor(s). The method detailed below will employ dual-wavelength fluorescence microscopy, although a very similar approach can be employed for dual-fluorophore fluorescence microscopy. 1.3

Calibration

The ratios obtained by dual-wavelength or dual-fluorophore fluorescence imaging are proportional to the pH; however, to make quantitative determinations, they must be converted to pH by calibrating in buffers of known pH. Several approaches to calibration have been used over the years. A relatively simple method involves exposing the particles to be used as phagocytic targets that have been labeled with the pH sensor to buffers of known pH (Fig. 3a). The ratios can then be plotted against pH and the experimental values interpolated. Although this method is quick and comparatively simple, it works on the assumption that the dye behaves similarly inside a phagosome as it does in vitro, and this may not be the case. A preferred method is to collect experimental data and then calibrate the same particles in situ [39]. This method involves bathing the cells in a series of high potassium buffers (approximately the same concentration as potassium in the cytosol) of various known pH values and adding the K+/H+ antiporter nigericin. This manipulation equilibrates the extracellular pH with the cytosolic pH and likely also that of intracellular compartments, including the phagosomal lumen. The result is a “clamping” of the pH of intracellular compartments at (approximately) the pH of the bathing solution. Dual-wavelength or dual-fluorophore ratios can then be obtained at the imposed pH values, and this can be used to generate an in situ calibration curve (Figs. 4b and 5b). Although this technique makes no assumptions about the behavior of the dye in different physiological settings, it does assume that the concentration of K+ in the phagosome is the same or similar to the concentration of K+ in the cytosol, which may not be the case. Moreover, not all cells are equally sensitive to nigericin [40]. Nevertheless, this method has been used with relative success over the years and will be outlined below.

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Fig. 3 Basic principles of dual-wavelength ratio imaging. (a) Fluorescein-labeled zymosan particles were attached to an 18 mm coverslip and imaged in a series of calibration buffers at the indicated pH values. 490/440 ratios were plotted against time. Images of individual fluorescein-labeled zymosan particles acquired in each calibration buffer with both 490 and 440 nm excitation were used to generate ratio images using the RatioPlus macro in ImageJ. The resulting pseudo-colored ratio images are depicted (insets). (b) Fluoresceinlabeled zymosan particles attached to a coverslip were imaged for 15 min, alternating between 490 and 440 nm excitation wavelengths every minute. The background-subtracted and normalized fluorescence for the 490 and 440 nm wavelengths alone and the ratio of 490/440 are shown. The traces illustrate the effect of progressive photobleaching on the fluorescence recorded at the individual wavelengths and on the ratio of 490/440. (c) Fluorescein-labeled zymosan particles were imaged as in (b), but the plane of focus was changed during the acquisition at the three points indicated by red arrows to illustrate the effect of changes in the focal plane on the fluorescence recorded at the individual wavelengths and on the ratio of 490/440. (d) Zymosan particles co-labeled with fluorescein and Alexa 555, a red dye used as reference, were imaged as in (b). The differential photobleaching over time of fluorescein and Alexa 555 is shown, and the resulting effect on the ratio FITC/Alexa 555 is shown. Note that application of the ratio does not adequately compensate for the bleaching induced loss of fluorescence, which occurs at different rates for the two dyes

2 2.1

Materials Cell Lines

1. RAW264.7 murine macrophage-like cells (see Note 1). 2. Complete RPMI1640 medium: RPMI1640 supplemented with 5% v/v heat-inactivated fetal bovine serum (FBS). Store at 4 °C.

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Fig. 4 Live-cell imaging of phagosomal pH using dual-wavelength ratio imaging. (a) Primary human macrophages were allowed to bind fluorescein-labeled zymosan and initiate phagocytosis. A zymosan particle in a phagocytic cup is denoted showing fluorescence with 490 nm excitation. The same particle was observed for 15 min, capturing a picture every 1 min. The background-corrected pseudo-colored 490/440 ratio image is shown at the start of phagocytosis and at 8 and 15 min after engulfment. Scale bar = 10 μm. (b) The background-corrected 490/440 ratio for the phagosome depicted in (a) was plotted against time (blue points). An in situ calibration was then carried out on the same cell (red points)

Fig. 5 Live-cell imaging of phagosomal pH using dual-fluorophore ratio imaging. (a) Raw264.7 murine macrophage-like cells were seeded with CypHer5e-AF488-conjugated silica beads to initiate phagocytosis. The same particle was observed for 20 min, capturing a picture every 2 min. The background-corrected pseudo-colored 666/520 ratio image is shown at the start of phagocytosis and at 5, 7, and 14 min after engulfment. (b) The background-corrected 666/520 ratio for the phagosome depicted in (a) was plotted against time (blue points). An in situ calibration of pH 4.5, 5.5, and 7.5 was then carried out (red points)

2.2

Reagents

1. 25 mg/mL fluorescein isothiocyanate (FITC) in anhydrous DMSO. Store at -20 °C. 2. 25 mg/mL Oregon Green 488 succinimidyl ester in anhydrous DMSO. Store at -20 °C. 3. 25 mg/mL SNARF1 succinimidyl ester in anhydrous DMSO. Store at -20 °C. 4. 2 mg/mL zymosan A bioparticles in PBS (see Note 2). Store at -20 °C.

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5. 1 mg/mL human IgG in PBS. Store at 4 °C. 6. Hank’s balanced salt solution (HBSS). Store at 4 °C. 7. Phosphate-buffered saline (PBS). Store at 4 °C. 8. 10 mg/mL nigericin, free acid in ethanol. Store at -20 °C. 9. Potassium-rich calibration buffer: 140 mM KCl, 1 mM MgCl2, 1 mM CaCl2, 5 mM glucose, and a buffer suitable for the desired pH (see Note 3). The solutions are adjusted to desired pH values using 1 M KOH or 1 M HCl for alkalization and acidification, respectively. 2.3 Microscopy System Requirements

1. External high-intensity arc lamp (such as X-Cite 120, EXFO Photonic Solutions, Inc.). 2. Inverted light microscope (such as the Leica DM4). 3. Magnetic Chamlide chamber. 4. Computer-controlled excitation and emission filter wheels (such as Lambda 10-2 optical filter changer, Sutter Instrument Company). 5. Appropriate dichroic filter cubes. 6. Electron-multiplied charge-coupled device camera (such as the Cascade II EMCCD camera, Photometrics). 7. Image analysis software (such as MetaFluor fluorescence ratio imaging software, Molecular Devices).

3

Methods

3.1 Labeling of Phagocytic Targets with pH Sensors

1. Centrifuge 500 μL of the 2 mg/mL stock solution of zymosan particles at 6000 × g for 15 s (see Notes 4 and 5). 2. Aspirate the supernatant, and resuspend the zymosan particles in 1 mL of PBS to wash. 3. Centrifuge the particles once more at 6000 × g for 15 s, and repeat the wash a total of three times. 4. Resuspend the zymosan in either 500 μL of 0.5 mg/mL FITC, 0.5 mg/mL Oregon Green succinimidyl ester, or 0.5 mg/mL SNARF1 succinimidyl ester in PBS adjusted to pH 8.3 (see Note 6). 5. Incubate for 1 h at room temperature, under constant agitation.

3.2 IgG Opsonization of Zymosan

1. Dilute 100 μL of the fluorophore-labeled zymosan in 400 μL of PBS. 2. Centrifuge at 6000 × g for 15 s and resuspend the particles in 200 μL of PBS.

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3. Add 4 μL of human IgG from the 1 mg/mL stock, and rotate at room temperature for 1 h (see Note 7). 4. Wash five times with PBS and resuspend the pellet after the final wash in 100 μL of PBS. 5. Opsonized zymosan should be used immediately and not stored long term. 3.3 Phagocytosis Assay

1. One day before performing the phagocytosis assay, seed RAW264.7 cells onto 18 mm coverslips in a 12-well plate containing complete RPMI1640. 2. On the day of the experiment, remove one coverslip from the 12-well plate, and place it into a magnetic Chamlide chamber (see Note 8). 3. Fill the chamber with 500 μL of HBSS pre-warmed to 37 °C. 4. Pipette 5 μL of the opsonized fluorophore-labeled zymosan into the magnetic chamber with the cells. 5. Centrifuge at 500 × g for 1 min, to bring the particles into contact with the RAW264.7 cells. 6. Place the magnetic chamber onto the heated stage (37 °C) of the microscope. 7. Turn the lights off in the imaging room to minimize the contribution of stray light to the fluorescence recordings. 8. Using a 40× objective, bring the cells into focus and scan the coverslip for cells that have zymosan particles attached (see Note 9). 9. Launch the imaging protocol in the software, and configure the acquisition parameters, making sure that there are no saturated pixels in the field. 10. Select the regions of interest (individual phagosomes) and a background region (see Note 10). 11. Acquire a series of images, alternating between either the desired excitation or emission wavelengths (see Note 11). Keep the number of acquired images to the minimum required for the desired time interval to prevent excessive photobleaching of the probe and phototoxicity to the cells.

3.4 In Situ Calibration

1. Pre-warm all the calibration buffers to 37 °C. 2. Wash the cells 2× with the first calibration buffer (see Note 12). 3. In a 1.5 mL microcentrifuge tube, combine 500 μL of the first calibration buffer with nigericin at a final concentration of 10 μg/mL. Vortex the solution for 5 s, and immediately add it to the cells in the Chamlide chamber.

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4. Wait for 5 min for the pH of the phagosome to equilibrate to the pH of the calibration buffer. Acquire three images 1–2 min apart. If the ratios obtained from each image are very different, wait for an additional 3–5 min to allow the pH to equilibrate. 5. Once equilibrated, acquire five images to determine the wavelength ratio that corresponds to the pH of the calibration solution. 6. Repeat steps 2–5 for each calibration solution. 7. Obtain a pH calibration curve by plotting the backgroundcorrected wavelength ratios against their corresponding pH values. 8. The experimentally observed phagosomal pH can then be estimated by interpolating the pH values from the calibration curve generated (Figs. 4b and 5b).

4

Notes 1. Raw264.7 cells are a macrophage-like cell line and were selected on the basis of them being professional phagocytes. They express several phagocytic receptors including Fc receptors and complement receptors. Other cell lines, such as HeLa cells, are not professional phagocytes, do not express phagocytic receptors and are therefore not ideal for studying phagocytic events. 2. Zymosan was chosen here due to its relative stability. Once dissolved in PBS, zymosan can be stored for several years at 20 °C. Other particles, such as sheep red blood cells, are commonly used for phagocytosis assays and are amenable to labeling with isothiocyanate or succinimidyl ester conjugates, but are not as stable for long-term storage. Moreover, sheep red blood cells must be opsonized, as they do not bear any ligands for mammalian phagocytic receptors. Zymosan, on the other hand, is readily able to bind phagocytic dectin receptors without any form of opsonization. 3. The buffer should be selected based on optimal buffering capacity near the desired pH. Example buffers (shown in parentheses) for select pH values are: pH 4.0 (acetate-acetic acid), pH 4.5 (acetate-acetic acid), pH 5.0 (acetate-acetic acid), pH 5.5 (2-[N-morpholino]ethanesulfonic acid [MES]), pH 6.0 (MES), pH 6.5 (MES), pH 7.0 (N-[2-hydroxyethyl]piperazine-N-[2-ethanesulfoinc acid] [HEPES]). 4. The phagocytic target chosen requires consideration. Here we describe the use of zymosan—a particle composed of protein– carbohydrate complexes from yeast cell walls. The presence of

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β-1,3-glycosidic linkages allows this particle to be recognized by dectin receptors and internalized by phagocytic cells. 5. Not all cells are equally phagocytic, nor are all targets equally stimulatory. The appropriate selection of a phagocytic target requires careful consideration. For example, RAW264.7 cells only poorly engulf unopsonized zymosan due to a paucity of dectin receptors on their surface. Therefore, unopsonized zymosan would not be suitable as a phagocytic target in this experiment. Secondly, not all forms of phagocytosis are identical. It should be noted that opsonizing targets with different opsonins triggers different phagocytic receptors and hence different internalization and maturation pathways. Therefore, the particle/opsonin should be carefully chosen to match the desired experimental question. 6. As discussed above, this protocol takes advantage of the commercial availability of succinimidyl ester and isothiocyanate forms of various pH-sensitive fluorophores. Other reagents, such as antibodies directly conjugated to pH sensors, can also be employed. Antibodies directly conjugated to pH sensors, such as fluorescein, are commercially available. Using this approach typically requires a primary antibody directed against the phagocytic target itself, followed by labeling with a secondary anti- body conjugated to a pH sensor. Of course, this will effectively render the target particle immunoglobulinopsonized. Therefore, if other forms of phagocytosis—such as complement-mediated phagocytosis or dectin-mediated phagocytosis—are to be studied, direct labeling of the target with an isothiocyanate or succinimidyl ester conjugate is more suitable. 7. Although primary human and mouse macrophages readily internalize unopsonized zymosan through dectin receptors, the RAW264.7 macrophage-like cells used in this protocol do not internalize unopsonized zymosan efficiently, due to a paucity of dectin receptor expression. For this reason, the zymosan must be opsonized with IgG prior to the assay to foster phagocytosis through Fc receptors, which are abundantly expressed on RAW264.7 cells. 8. A Chamlide chamber is a coverslip holder consisting of two interlocking parts and an O-ring. A coverslip is placed in the chamber and a seal is achieved by the magnetic attraction of the two interlocking parts. The coverslip can then be bathed in complete RPMI 1640. The Chamlide fits into a slot on the heated stage of the microscope allowing for continuous, livecell imaging. Finding a cell that is in the process of engulfing a particle can take practice. It is not sufficient to select a particle that is merely in the vicinity of a phagocytic cell. It is helpful to look for cells that have already extended pseudopodia around

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the phagocytic target to increase the likelihood of capturing a phagocytic event. 9. A background region of interest should be selected in the extracellular space near to the region of interest surrounding the phagosome, and its fluorescence determined at the wavelengths of choice. The background values for both wavelengths should be subtracted separately from the values recorded at the corresponding wavelength in the phagosome region of interest, prior to the generation of the ratio. This allows for the calculation of a properly background-corrected ratio. 10. Bear in mind that some ratiometric probes are dual-excitation probes (fluorescein and Oregon Green) and some are dual emission probes (SNARF1). This means that either the excitation filters (490 and 440 nm for fluorescein and Oregon Green) or the emission filters (640 and 580 nm for SNARF1) will have to be alternated during acquisition. 11. The order of addition of the calibration buffers is theoretically unimportant. However, further deviation of the calibration buffer from neutral pH values does seem to affect the ability of the cell to stay attached to the coverslip. In general, a good practice is to start with the neutral pH buffer and to proceed sequentially to more acidic or alkaline calibration buffers. 12. All in all, the technique of fluorescence ratio imaging represents a powerful, reliable, and relatively simple method for assessing phagosomal pH. Measurements can be made in individual cells and even individual phagosomes. This allows for the observation of cell/phagosome heterogeneity that is inevitably masked by whole-population techniques. By carefully considering the pH sensor, phagocytic target, and calibration method chosen, this technique will continue to provide valuable information about the maturation and luminal biochemistry of phagosomes.

Acknowledgments We would like to thank members of the Canton, Yates, and Grinstein labs for helpful discussions. References 1. Bloomfield G, Traynor D, Sander SP et al (2015) Neurofibromin controls macropinocytosis and phagocytosis in Dictyostelium. Elife 4:e04940

2. Arandjelovic S, Ravichandran KS (2015) Phagocytosis of apoptotic cells in homeostasis. Nat Immunol 16:907–917 3. Koenigsknecht J, Landreth G (2004) Microglial phagocytosis of fibrillar β-amyloid

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through a β1 integrin-dependent mechanism. J Neurosci 24:9838–9846 4. Kevany BM, Palczewski K (2010) Phagocytosis of retinal rod and cone photoreceptors. Physiology (Bethesda) 25:8–15 5. Manwani D, Bieker JJ (2008) Chapter 2: The erythroblastic island. In: Current topics in developmental biology. Academic Press, pp 23–53 6. Blum JS, Wearsch PA, Cresswell P (2013) Pathways of antigen processing. Annu Rev Immunol 31:443–473 7. Turk V, Stoka V, Vasiljeva O et al (2012) Cysteine cathepsins: from structure, function and regulation to new frontiers. Biochim Biophys Acta 1824:68–88 8. Canton J (2014) Phagosome maturation in polarized macrophages. J Leukoc Biol 96: 729–738 9. Wreden CC, Johnson J, Tran C et al (2003) The H+-coupled electrogenic lysosomal amino acid transporter LYAAT1 localizes to the axon and plasma membrane of hippocampal neurons. J Neurosci 23:1265–1275 10. Geisow MJ, D’Arcy Hart P, Young MR (1981) Temporal changes of lysosome and phagosome pH during phagolysosome formation in macrophages: studies by fluorescence spectroscopy. J Cell Biol 89:645–652 11. Claus V, Jahraus A, Tjelle T et al (1998) Lysosomal enzyme trafficking between phagosomes, endosomes, and lysosomes in J774 macrophages: enrichment of cathepsin H in early endosomes*. J Biol Chem 273:9842– 9851 12. Hart PD, Young MR, Jordan MM et al (1983) Chemical inhibitors of phagosome-lysosome fusion in cultured macrophages also inhibit saltatory lysosomal movements. A combined microscopic and computer study. J Exp Med 158:477–492 13. Flannagan RS, Jaumouille´ V, Grinstein S (2012) The cell biology of phagocytosis. Annu Rev Pathol 7:61–98 14. Maxson ME, Grinstein S (2014) The vacuolartype H+-ATPase at a glance – more than a proton pump. J Cell Sci 127:4987–4993 15. Cuppoletti J, Aures-Fischer D, Sachs G (1987) The lysosomal H+ pump: 8-azido-ATP inhibition and the role of chloride in H+ transport. Biochim Biophys Acta 899:276–284 16. Dell’Antone P (1979) Evidence for an ATP-driven “proton pump” in rat liver lysosomes by basic dyes uptake. Biochem Biophys Res Commun 86:180–189 17. Graves AR, Curran PK, Smith CL et al (2008) The Cl-/H+ antiporter ClC-7 is the primary

chloride permeation pathway in lysosomes. Nature 453:788–792 18. Harikumar P, Reeves JP (1983) The lysosomal proton pump is electrogenic. J Biol Chem 258: 10403–10410 19. Ohkuma S, Moriyama Y, Takano T (1983) Electrogenic nature of lysosomal proton pump as revealed with a cyanine dye. J Biochem 94:1935–1943 20. Steinberg BE, Huynh KK, Brodovitch A et al (2010) A cation counterflux supports lysosomal acidification. J Cell Biol 189:1171–1186 21. Metchnikoff E (1893) Lectures on the comparative pathology of inflammation delivered at the Pasteur Institute in 1891. Рипол Классик 22. Sprick MG (1956) Phagocytosis of M. tuberculosis and M. smegmatis stained with indicator dyes. Am Rev Tuberc 74:552– 565 23. Pavlov EP, Solov’ev VN (1967) [pH changes of cytoplasm in phagocytosis of microbes stained with indicator dyes]. Biull Eksp Biol Med 63: 78–81 24. Mandell GL (1970) Intraphagosomal pH of human polymorphonuclear neutrophils. Proc Soc Exp Biol Med 134:447–449 25. Jensen MS, Bainton DF (1973) Temporal changes in PH within the phagocytic vacuole of the polymorphonuclear neutrophilic leukocyte. J Cell Biol 56:379–388 26. Ohkuma S, Poole B (1978) Fluorescence probe measurement of the intralysosomal pH in living cells and the perturbation of pH by various agents. Proc Natl Acad Sci U S A 75: 3327–3331 27. Canton J, Grinstein S (2015) Chapter 5: Measuring lysosomal pH by fluorescence microscopy. In: Platt F, Platt N (eds) Methods in cell biology. Academic Press, pp 85–99 28. Yates RM, Russell DG (2008) Real-time spectrofluorometric assays for the lumenal environment of the maturing phagosome. In: Deretic V (ed) Autophagosome and phagosome. Humana Press, Totowa, pp 311–325 29. Vergne I, Constant P, Lane´elle G (1998) Phagosomal pH determination by dual fluorescence flow cytometry. Anal Biochem 255: 127–132 30. Steinberg BE, Grinstein S (2007) Assessment of phagosome formation and maturation by fluorescence microscopy. In: Quinn MT, DeLeo FR, Bokoch GM (eds) Neutrophil methods and protocols. Humana Press, Totowa, pp 289–300 31. Schlam D, Bohdanowicz M, Chatilialoglu A et al (2013) Diacylglycerol kinases terminate diacylglycerol signaling during the respiratory

Measuring Phagosomal pH by Fluorescence Microscopy burst leading to heterogeneous phagosomal NADPH oxidase activation*. J Biol Chem 288:23090–23104 32. Webb DJ, Brown CM (2013) Epi-fluorescence microscopy. In: Taatjes DJ, Roth J (eds) Cell imaging techniques: methods and protocols. Humana Press, Totowa, pp 29–59 33. Freundt EC, Czapiga M, Lenardo MJ (2007) Photoconversion of Lysotracker Red to a green fluorescent molecule. Cell Res 17:956–958 34. Grynkiewicz G, Poenie M, Tsien RY (1985) A new generation of Ca2+ indicators with greatly improved fluorescence properties. J Biol Chem 260:3440–3450 35. Tsien RY (1989) Chapter 5: Fluorescent indicators of ion concentrations. In: Taylor DL, Wang Y-L (eds) Methods in cell biology. Academic Press, pp 127–156 36. Tsien RY, Rink TJ, Poenie M (1985) Measurement of cytosolic free Ca2+ in individual small

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cells using fluorescence microscopy with dual excitation wavelengths. Cell Calcium 6:145– 157 37. Canton J, Khezri R, Glogauer M et al (2014) Contrasting phagosome pH regulation and maturation in human M1 and M2 macrophages. Mol Biol Cell 25:3330–3341 38. Jankowski A, Scott CC, Grinstein S (2002) Determinants of the phagosomal pH in neutrophils*. J Biol Chem 277:6059–6066 39. Thomas JA, Buchsbaum RN, Zimniak A et al (1979) Intracellular pH measurements in Ehrlich ascites tumor cells utilizing spectroscopic probes generated in situ. Biochemistry 18: 2210–2218 40. Chow S, Hedley D, Tannock I (1996) Flow cytometric calibration of intracellular pH measurements in viable cells using mixtures of weak acids and bases. Cytometry 24:360–367

Chapter 12 Multiplexed Phagosomal Assays for the Detection and Quantification of Bidirectional Exchange Between the Phagolysosomal Lumen and Extracellular Space Jenny A. Nguyen, Catherine J. Greene, Samuel Cheung, and Robin M. Yates Abstract The phagolysosome is an antimicrobial and degradative organelle that plays a key role in macrophagemediated inflammation and homeostasis. Before being presented to the adaptive immune system, phagocytosed proteins must first be processed into immunostimulatory antigens. Until recently, little attention has been given to how other processed PAMPs and DAMPs can stimulate an immune response if they are sequestered in the phagolysosome. Eructophagy is a newly described process in macrophages that releases partially digested immunostimulatory PAMPs and DAMPs extracellularly from the mature phagolysosome to activate vicinal leukocytes. This chapter outlines approaches to observe and quantify eructophagy by simultaneously measuring several phagosomal parameters of individual phagosomes. These methods use specifically designed experimental particles capable of conjugating to multiple reporter/reference fluors in combination with real-time automated fluorescent microscopy. Through the use of high-content image analysis software, each phagosomal parameter can be evaluated quantitatively or semiquantitatively during post-analysis. Key words Phagosome, Phagolysosome, Macrophage, Eructophagy, Proteolysis, Acidification, Secretion

1

Introduction Recently we described a novel secretory process whereby macrophages can release partially digested phagolysosomal contents (including newly generated PAMPs) into the extracellular space through discrete, controlled events—a process we termed eructophagy [1, 2]. Eructophagy is currently known to be induced by pro-inflammatory stimuli, inhibited by IL-4, and regulated by mTOR and is dependent on key autophagy proteins, including fusion machinery of degradative and secretory autophagy. Considering the potential for phagolysosomes to release a wide variety of

Roberto J. Botelho (ed.), Phagocytosis and Phagosomes: Methods and Protocols, Methods in Molecular Biology, vol. 2692, https://doi.org/10.1007/978-1-0716-3338-0_12, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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bioactive molecules through this process, including hydrolases, antimicrobial agents, exogenous and endogenous protein antigens, exosomes, and potentially pathogenic factors, toxins, and infectious virions or prions, the impact of eructophagy on health and disease may be widespread. Eructophagy involves several microenvironmental and biochemical changes that occur within the lumen of the mature phagolysosome. Brief neutralization of the phagolysosome can be observed during which soluble products are lost from the lumen [1, 2]. Upon neutralization, the phagolysosomes quickly reacidify and recommence proteolysis within minutes. Thus, eructophagy provides a unique opportunity to study phagosome dynamics as it progresses through phagocytosis to maturation to secretion. As eructophagy is a transient process, cellular fixation is not practical for observing and quantifying this phenomenon—it can only be observed live. Due to the heterogeneity of phagosomes in terms of maturation and function, some phagolysosomes undergo eructophagy repeatedly, while others do not. Therefore, single assessment of individual phagosomes is favored over population-based approaches. To study eructophagy, we have adapted previously developed reporter bead-based fluorometric techniques to simultaneously measure multiple lumenal parameters of individual phagosomes in a population of live macrophages [3–5]. Using automated highcontent microscopic imaging and image analysis, this approach identifies and tracks phagosomes while recording relative fluorescent intensities of reporter and reference fluors. While more laborious than population-based assays, assessing multiple phagosomal properties within individual phagosomes enables the investigation of phagosomal lumenal chemistry, phagosomal autonomy, and heterogeneity [5]. To ensure that each measurement accurately reflects the experimental objective and is not overly influenced by other phagosomal parameters or proximal fluorophores, it is important to carefully consider the limitations of multiple fluorophores or substrates when designing multiplexed phagosomal assays [5]. Ensure that fluor combinations that do not have overlapping emission spectra and possible resonance energy transfer between fluor combinations should be considered and controlled. Furthermore, each fluor must be considered for its suitability and stability in the phagosomes. In general, calibration or reference fluors should be stable at low pH, and their fluorescent quantum yields should be unaffected by changes in phagosomal pH (pH ~7.5–4.0). In certain cases, a combination of reporter fluors that can capture all the desired phagosomal parameters may not be available. Analyzing and evaluating results in such scenarios requires consideration of the limitations of each fluor.

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Fig. 1 Schematic of the cellulase-resorufin-cellobioside eructophagy reporter assay. Macrophages phagocytose experimental beads conjugated to the enzyme cellulase and a tracing fluor (AF647SE) into phagosomes. These phagosomes undergo rapid maturation into mature phagolysosomes via fusion with lysosomes and are allowed to fully mature for 1 h post-phagocytosis. Subsequently, the cell membrane-impermeable substrate of cellulase, resorufin-cellobioside, is added to the extracellular medium. During eructophagy, content-mixing occurs between the phagosomal lumen and extracellular milieu, allowing cellulase to access resorufin-cellobioside for hydrolysis. The liberated resorufin groups emit fluorescence when excited and are used to report phagolysosomes undergoing eructophagy. (Adapted from Greene et al. (2022) [1]. This figure was created with BioRender.com)

In this chapter, we describe the gold standard for measuring the frequency of eructophagy per phagolysosome—the cellulase-resorufin-cellobioside eructophagy reporter assay. The enzyme cellulase is conjugated onto experimental beads which are phagocytosed by macrophages into phagosomes (Fig. 1). Cells are exposed to a cell membrane-impermeable cellulase substrate called resorufin-cellobioside, which cannot interact with cellulase without eructophagy to allow for bidirectional exchange of material between the phagosome lumen and extracellular milieu. Upon cleavage by cellulase, this substrate fluoresces and provides a reliable indication of eructophagy. In this approach, high-content microscopic imaging and image analysis are used to identify and track phagosomes and record the relative intensities of reporter and

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Fig. 2 Schematic of the DQ Green BSA/pHrodo eructophagy reporter assay. Macrophages phagocytose experimental beads conjugated to the fluorescent protease substrate DQ Green BSA, the pH sensor pHrodo, and a tracing fluor (AF647SE) into phagosomes. Both DQ Green BSA and pHrodo are self-quenched in non-proteolytic environments with neutral pH. Upon fusion with lysosomes, the mature phagolysosome acquires an acidic lumen with proteolytic capacity. pHrodo fluoresces in the acidic environment, while proteases cleave bound peptides which dequench and fluoresce. During eructophagy, the liberated fluorescent peptides are lost from the phagosomal lumen to the extracellular space. Simultaneously, the loss of protons to the extracellular space results in neutralization of the phagolysosomal lumenal pH. (Adapted from Greene et al. (2022) [1]. This figure was created with BioRender.com)

reference fluorophores. Additionally, we will describe how this assay can be modified to evaluate changes in multiple lumenal parameters during eructophagy including phagosome pH and proteolytic abilities (Fig. 2).

2 Materials 2.1 Cells, Reagents, and Buffers

1. Eructophagy is measured in primary bone marrow-derived macrophages (BMM∅s). However, these approaches can be employed for all primary macrophages, bone marrow-derived dendritic cells, or macrophages derived from conditionally immortalized myeloid precursors expressing ER-HoxB8 (see Note 1 and Chapter 8).

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2. 96-well assay plates: 96-well μClear black clear bottom (Greiner Bio-One). See Note 2. 3. BMM∅ growth media: Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% fetal bovine serum (FBS), 2 mM L-glutamine, 1 mM sodium pyruvate, 100 U/mL penicillin-streptomycin, and 20% conditioned medium from the supernatant of macrophage colony-stimulating factor (M-CSF)-producing L929 cells. 4. Assay buffer: Dulbecco’s phosphate-buffered saline (DPBS) supplemented with 1 mM CaCl2, 2.7 mM KCl, 0.5 mM MgCl2, 5 mM glucose, 0.1% calf skin gelatin (see Note 2), and 10 mM HEPES. Filter sterilize through a 0.22 μm filter. Store at 4  C. Warm to 37  C before use. 5. Coupling buffer: 0.1 M sodium borate in ddH2O. Adjust pH to 8.0 with 10 M NaOH. Filter sterilize through 0.22 μm filter. Store at room temperature. 6. Quenching buffer: 250 mM glycine in DPBS pH 7.2. Filter sterilize through 0.22 μm filter. Store at 4  C. 7. 2% sodium azide, aqueous solution. Store at room temperature. 8. 3.0 μm carboxylate-modified silica particles/beads (Si-COOH) 5% suspension (Kisker Biotech, Steinfurt, Germany) (see Note 3). 9. Cyanamide. Store as desiccate at 4  C. Protect from moisture. Warm to room temperature before use. 10. Reporter enzyme: cellulase from Trichoderma viride (Cellulysin®; Millipore Sigma). Alternatively, α-amylase from Bacillus licheniformis (LD Carlson Co.) or β-glucosidase from almonds (Sigma Aldrich) can be used. Store at 4  C. 11. Reporter enzyme substrate: 5 mg/ml of resorufin-cellobioside (for cellulase), resorufin maltotriose (for α-amylase), or resorufin glucopyranoside (for β-glucosidase), dissolved in high quality anhydrous dimethylsulfoxide (DMSO). Aliquot and store at 20  C. Protect from light and moisture (Marker Gene Technologies). 12. Proteolysis sensor: 1 mg/mL DQ Green BSA, dissolved in coupling buffer. Store at 20  C. Protect from light and moisture (Thermo Fisher Scientific). 13. pH sensor: 5 mg/mL pHrodo Red succinimidyl ester (pHrodo-SE), dissolved in high quality anhydrous dimethylsulfoxide (DMSO). 14. 2 mg/mL IgG from human serum, dissolved in sterile ddH2O. Aliquot and store at 20  C.

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15. Reference fluor: 5 mg/mL Alexa Fluor 647 succinimidyl ester (AF647SE), dissolved in DMSO. Other pH and ROS stable fluorophores that are spectrally distinct to the reporter fluors can be used as reference fluors. Aliquot and store at 20  C. Protect from light and moisture (Thermo Fisher Scientific). 2.2 Imaging Instrument and Analysis Software

1. IN Cell Analyzer 2000 Live C TEMP/LH/EC High-Content Analysis system (GE Healthcare Life Sciences). The instrument has environmental control for CO2 and humidity and temperature control hardware installed for the imaging chamber. The instrument is equipped with polychroic QUAD band mirrors that have been optimized for the installed excitation filters (350  50, 490  20, 579  24, and 645  30 nm) and emission filters (455  50, 525  20, 624  40, and 705  72 nm). 2. IN Cell Analyzer 2000 Acquisition Software (Version 4.6) (GE Healthcare Life Sciences). 3. Quantification software: IN Cell Developer Toolbox 1.9.3 (x64) as a component of the IN Cell Investigator image analysis software package (Version 1.6.3) (GE Healthcare Life Sciences). 4. Analysis software: Spotfire DecisionSite® analytics software integrated with IN Cell Investigator image analysis software package (Version 1.6.3) (TIBCO Software, Inc.).

3

Methods

3.1 Preparation of Experimental Reporter Particles

1. 10 mg of 3.0 μm carboxylate-modified Si-COOH particles (200 μL of manufacturer stock) is added to a low binding polypropylene 1.5 mL screw-cap microtube. Pellet the beads and remove the manufacturer’s storage buffer by brief centrifugation at 6000  g at room temperature using a bench-top centrifuge and three DPBS washes. 2. 30 mg of the heterobifunctional cross-linker; cyanamide is freshly dissolved in 1 mL of coupling buffer and incubated with the Si-COOH particles at room temperature for 15 min with agitation. Proceed to Subheading 3.2 for cellulase reporter beads or Subheading 3.3 for DQ Green BSA and pHrodo reporter beads.

3.2 Preparation of Cellulase Reporter Beads

1. During the incubation period, 20 mg of cellulase from Trichoderma viride and 0.01 mg human IgG (2 μL of a 2 mg/mL stock; optional) are dissolved in 1 mL of coupling buffer. 2. Quickly after incubation, the Si-COOH particles are washed three times with ice-cold coupling buffer to remove excess

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cyanamide. The cyanimide-activated Si-COOH particles are resuspended in the cellulase/IgG solution and incubated at room temperature for 30 min with agitation. This allows for covalent coupling of cellulase and IgG to the Si-COOH particles through the cyanimide cross-linker. 3. The Si-COOH particles are washed twice with 500 μL of coupling buffer to remove unbound protein and incubated with 500 μL of quenching buffer for 10 min at room temperature with agitation to quench unreacted amine moieties. 4. The Si-COOH particles are washed twice with 500 μL of coupling buffer and resuspended in 1 ml of coupling buffer containing 1 μL of 5 mg/mL AF647SE. The particles are covered and incubated at room temperature for 15 min with agitation to allow the particle bound proteins to covalently bind to the amine-reactive fluors. 5. The particles are washed three times with 1 mL DPBS to remove excess fluor and incubated with 500 μL of quenching buffer for 10 min at room temperature with agitation. 6. The particles are washed three times with 1 mL DPBS and can be stored at 4  C for up to 10 days (see Note 4). 3.3 Preparation of DQ Green BSA and pHrodo Reporter Beads

1. During the incubation period, 1 mg of DQ Green BSA and 0.01 mg of 0.01 mg human IgG (optional) are dissolved in 1 mL of coupling buffer. 2. Quickly after incubation, the Si-COOH particles are washed three times with ice-cold coupling buffer to remove excess cyanamide. The cyanimide-activated Si-COOH particles are resuspended in the DQ Green BSA/IgG solution and incubated at room temperature for 3–6 h or overnight with agitation. This allows for covalent coupling of Dq Green BSA and IgG to the Si-COOH particles through the cyanimide crosslinker. 3. See Subheading 3.2, step 3. 4. The Si-COOH particles are washed twice with 500 μL of coupling buffer and resuspended in 1 ml of coupling buffer containing 1 μL of 5 mg/mL pHrodo-SE and 1 μL of 5 mg/ mL AF647SE. The particles are covered and incubated at room temperature for 30 min with agitation to allow the particle bound proteins to covalently bind to the amine-reactive fluors. 5. See Subheading 3.2, step 5. 6. The labeled particles are washed three times with 1 mL DPBS and can be stored at 4  C for later use in 1 mL quenching buffer with 10 uL of 2% sodium azide as a preservative.

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3.4 Cell Preparation and Handling

1. Bone marrow is flushed from the femurs, tibias, and ilia of mice with DMEM and centrifuged at 230  g at 4  C for 10 min. 2. To derive BMM∅s, freshly isolated bone marrow is resuspended in BMM∅ growth medium and plated onto non-treated 10 cm petri dishes (approximately eight to ten dishes per mouse). After 7 days of initial plating, BMM∅s are subcultured 1:2 in BMM∅ media and plated onto fresh 10 cm petri dishes. The cells are allowed to differentiate for 10–14 days after initial plating before use. 3. To seed the BMM∅s for assays, the growth medium is removed from the fully differentiated BMM∅s and replaced with cold DPBS. The cells are incubated at 4  C for 10 min to allow for cell detachment from the untreated 10 cm culture dish. The BMM∅s are gently lifted with a cell scraper and centrifuged at 230  g at 4  C for 10 min. 4. The BMM∅ cell pellet is resuspended in 1 mL of BMM∅ growth medium, and the cells are numerated using Trypan blue exclusion on a hemocytometer. 5. BMM∅s are plated onto 96-well plates at 25% confluency (approximately 30,000 cells in 100 μL of BMM∅ medium) per well in a maximum of 48 wells. The authors also plate additional “test” wells for titrating bead numbers. The plates are incubated at room temperature for 15 min to establish an even monolayer before incubating at 37  C overnight.

3.5 Imaging System Setup

1. The system is pre-warmed to 37  C for at least an hour prior to imaging (see Note 5). 2. The 40 magnification objective is selected. 3. The required channels and their excitation emissions are assigned as follows: Cy3 (Resorufin), Cy5 (AF647SE) and the brightfield channel will default to DAPI. 4. It is recommended that the exposure value is set to 0.02 s for all channels and at 10% lamp intensity to avoid phototoxicity. This can be adjusted as required. All images are acquired in 2D mode. 5. Adjust the offset for each channel, which is optimized based the experimental plate used. If the recommended 96-well plate is used, we have found an offset of 6 for Cy3 and 5 for Cy5 work best (see Note 6). 6. To representatively capture eructophagy, images are acquired over a 180 s time interval for 2 h. However, the time interval can be adjusted as desired. Under these experimental conditions, eructophagy can be observed for up to 10 h postmaturation of the phagolysosome. 7. Laser-guided autofocus is used to refocus at the start of each timepoint.

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8. For Image Processing, the High pass filter is set to a value of 10 with the Invert phase polarity disabled. The sharpest images can be obtained by setting the Contrast angle to 135 and the Combiner prism to 0 and disabling the Intensity modulation. 3.6 Acquisition of Images

1. Prior to imaging, growth medium is removed from the BMM∅ monolayer, and the adhered cells are washed twice with 100 μL of pre-warmed assay buffer (37  C). 2. 50 μL of pre-warmed assay buffer is added to each well before adding beads. 3. The appropriate concentration of beads is then determined. Approximately one to two beads per cell are optimal. Initially, 5 μL of the cellulase bead stock (prepared in Subheading 3.2) is diluted into 1 mL assay buffer. 50 uL of the working dilution of beads is added to a test well containing cells. After a 5–10-min incubation period, the number of beads per cell is visualized using a light microscope. The working dilution of beads is then adjusted accordingly to achieve one to two beads per cell, as a higher number becomes more difficult to track accurately. 4. 50 μL of the appropriate working dilution of beads is added to each well. The plate is incubated at room temperature for 10 min to allow the beads to evenly settle onto the cells (see Note 7). An additional 100 μL of pre-warmed assay buffer can be added per well for assays exceeding 2 h. 5. The assay plate is loaded into the pre-warmed imaging chamber of the IN Cell Analyzer 2000. The previously programmed protocol (Subheading 3.5) is loaded, the sample wells are selected, and the acquisition protocol is run (see Note 8). 6. After image acquisition is complete, the computer will automatically generate the image stack file in xdce format. Note that fluorescent tracking described in the next step using the IN Cell Investigator software cannot be performed without the xdce file.

3.7

Analysis

Analysis of the acquired image data is performed using the IN Cell Investigator software and Spotfire DescisionSite data visualization software. The IN Cell Investigator software requires an xdce file generated during image acquisition using the IN Cell Developer software. A custom algorithm is used to identify fluorescent beads in each field of view based on their dimensions and characteristics. The following section will describe how to identify, track, and record fluorescent intensities of the phagosomally localized resorufin “flashes” and the 3.0 μm reporter particles conjugated to the calibration fluor. 1. Load the xdce file into the IN Cell Investigator software by selecting View/Analyze Image Stack.

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2. To create a protocol, select Protocol Explorer. The individual target image sets (New Target Type) corresponding to the individual channels are selected (Source Image) and named (e.g., enter “resorufin” for the Cy3 channel and “calibration” for the Cy5 channel). 3. From the options under the channel name, select the appropriate representative color (e.g., red for Cy3 and magenta for Cy5). To increase the accuracy of the automated detection of the phagocytosed beads, under Targets, select Filled and Remove single pixel targets. 4. Under Segmentation and Change Segmentation Type, the predefined algorithm, Nuclear detection is selected to accurately detect 3.0 μm beads. 5. The Minimum target area is changed to match the bead size of 3.0 μm (mean diameter). The bead position and area that will be analyzed can be previewed with the Preview selected operation option. This verifies that all the beads have been correctly identified and should be performed on at least two fields of view. 6. The Sensitivity is adjusted depending on the fluorescent signal of the bead. We use a value between 90 and 95 to yield maximal coverage of bead fluorescence without interference from background noise. 7. Under Postprocessing, Border Object Removal is selected and adjusted to a ten-pixel distance from all borders to remove background signal from the edge of the images. Optional: Fill Holes is selected if the core of the particle is nonfluorescent (e.g., if using solid-core polystyrene rather than Si-COOH beads). 8. Repeat steps 3–7 for each fluorescent channel. 9. To obtain a numerical value for fluorescent intensity per bead at each time point, add a Dens-Level measure under the Measures option for each channel. For the Cy3 (resorufin) target set, the source image is from the Cy3 channel. For the Cy5 (calibration) target set, add two Dens-Level measures—one for the Cy3 channel and one for Cy5. 10. It is necessary to implement a cell (bead) tracking function to quantify the fluorescence associated with beads over time as cells and phagosomes move within the field of view. Under Dynamic Behaviour, select Cell Tracking. The calibration fluor, Alexa 647-SE, is selected as the Track Target Set (see Note 9). For the 3.0 μm beads, Use Intensity as extra tracking is enabled as an additional tracking parameter. By default, the Outlier detection has a value of 4 and the Tracking Method is preset as proximity.

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Fig. 3 Quantification of eructophagy events per phagolysosome in BMM∅s following a 1 h incubation with cellulase-conjugated experimental beads. Lipopolysaccharide from Escherichia coli (10 ng/μL) and supraphysiological levels of essential amino acids were added to the assay medium prior to measurement in order to demonstrate induction and inhibition of eructophagy. Each data point represents a well replicate after 2 h of imaging

11. Save the protocol once the parameters have been inputted. Start the analysis by selecting the New Analysis Wizard and the preprepared protocol. The data can be exported in Microsoft Excel format to allow the analysis program to generate a copy of all quantified parameters in a Microsoft Excel file. 3.8 Using Spotfire DecisionSite to Visualize Data in Graphical Format

Once the IN Cell Developer analysis is complete, the software automatically exports the data as a Microsoft Excel file in addition to IN Cell Analyzer-specific files. The frequency of eructophagyspecific period can be quantified (Fig. 3). 1. Once connected to the Spotfire DecisionSite software, the data file in .xml format is uploaded into the software. 2. The fluorescence change in each particle over time is viewed as a 3D plot. When plotting the 3D graph, the left axis is set to Ch2. Dens-Level or Ch3. Dens-Level to display the fluorescent channel of interest. The top axis is set to LinkedTrackID and the bottom axis is set to This Time Index. 3. Under Properties, the line connection was set to link the data points by LinkedTrackID and by the order of ThisTimeIndex. The size of the data points is minimized, and the 3D

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Fig. 4 Representative 3D plots from Spotfire DescisionSite and corresponding images from the IN Cell 2000 Developer software. Individual phagosomes were monitored for eructophagy in BMM∅s over a 2-h period following a 1-h incubation with cellulase-conjugated experimental beads. Increased fluorescence intensity or “spikes” in the resorufin channel are indicative of eructophagy events. The Spotfire DecisionSite software package was used to generate the graphs

perspective is decreased to a minimum. Each fluorescent bead can be cross-referenced on the 3D plot and on the original image by selecting any data point or area on an image. Representative Spotfire DecisionSite 3D plots and corresponding images from the IN Cell Developer software are shown in Figs. 4 and 5.

Fig. 5 Representative 3D plots from Spotfire DescisionSite and corresponding images from the IN Cell 2000 Developer software. Individual phagosomes were monitored for eructophagy in BMM∅s over a 2-h period following a 1-h incubation with DQ Green BSA- and pHrodo-conjugated experimental beads. Coinciding decreases in fluorescence intensity in both the DQ Green BSA and pHrodo channel are indicative of eructophagy events. The Spotfire DecisionSite software package was used to generate the graphs

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4. Analysis of eructophagy events is conducted per well, per field of view, and per “track” (corresponding to a bead-containing phagosome). Tracks tracked for the entirety of the imaging period (spanning the entire bottom axis) were quantified as phagolysosomes. Eructophagy events were quantified over time as well-separated elevations of fluorescence intensity in the Cy3 channel, with no elevations in the Cy5 channel.

4

Notes 1. Eructophagy was observed to occur at significantly lower frequencies in primary dendritic cells than macrophages [1]. Eructophagy cannot be observed in traditionally immortalized phagocyte cell lines due to a aberrant mTOR and PI3K pathways. 2. 96-well μClear black clear bottom (Greiner Bio-One) plates are used for live-cell imaging in this protocol. Alternatively, other IN Cell Analyzer 2000 compatible microplates could be used. 3. Carboxylate-modified 1–5.0 μm polystyrene or latex beads can also be used. 4. It is recommended that after 10 days, a fresh batch of reporter particles are made prior to conducting the assay. We have noted a decline in cellulase activity after this period. 5. The imaging chamber should be preheated to prevent unwanted plate movement caused by thermal changes during acquisition. 6. The fluorescent “flashes” corresponding to the cleavage of resorufin-cellobioside were observed only when eructophagy or cell death occurs. If no signal is observed in the Cy3 channel, it may be necessary to image for 10 min to observe eructophagy and adjust the exposure/offset as required. 7. A smaller volume may allow particles to settle faster. Particle size and density also affect the rate of settling. 8. In our standard assay, we acquire images for 2 h at intervals of 3 min. During eructophagy, bidirectional exchange occurs between the phagosome lumen and extracellular space for approximately 3 min. If necessary, the acquisition period and time interval can be adjusted. 9. Tracking calculations should be based on a constant fluorescent signal unaffected by environmental changes (i.e., the reporter fluor Alexa 647-SE).

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References 1. Greene CJ, Nguyen JA, Cheung SM, Arnold CR, Balce DR, Wang YT, Soderholm A, McKenna N, Aggarwal D, Campden RI, Ewanchuk BW, Virgin HW, Yates RM (2022) Macrophages disseminate pathogen associated molecular patterns through the direct extracellular release of the soluble content of their phagolysosomes. Nat Commun 13(1):3072. h ttps://d oi.org/10.1038/s414 67-02230654-4 2. Nguyen JA, Greene CJ, Yates RM (2022) Eructophagy: macrophages use autophagic machinery to burp out parts of their meal. Autophagy:1–3. https://doi.org/10.1080/ 15548627.2022.2138006 3. Yates RM, Hermetter A, Russell DG (2009) Recording phagosome maturation through the

real-time, spectrofluorometric measurement of hydrolytic activities. Methods Mol Biol 531: 157–171. https://doi.org/10.1007/978-159745-396-7_11 4. Yates RM, Russell DG (2008) Real-time spectrofluorometric assays for the lumenal environment of the maturing phagosome. Methods Mol Biol 445:311–325. https://doi.org/10.1007/ 978-1-59745-157-4_20 5. Cheung S, Greene C, Yates RM (2017) Simultaneous analysis of multiple Lumenal parameters of individual phagosomes using high-content imaging. In: Botelho R (ed) Phagocytosis and phagosomes: methods and protocols. Springer, New York, pp 227–239. https://doi.org/10. 1007/978-1-4939-6581-6_15

Chapter 13 Quantitative Spatio-temporal Analysis of Phagosome Maturation in Live Cells Patricia Rosell Are´valo, Beren Aylan, and Maximiliano G. Gutierrez Abstract Phagocytosis and phagosome maturation are central processes to the development of the innate and adaptive immune response. Phagosome maturation is a continuous and dynamic process that occurs rapidly. In this chapter we describe fluorescence-based live cell imaging methods for the quantitative and temporal analysis of phagosome maturation of beads and M. tuberculosis as two phagocytic targets. We also describe simple protocols for monitoring phagosome maturation: the use of the acidotropic probe LysoTracker and analyzing the recruitment of EGFP-tagged host proteins by phagosomes. Key words Phagosome, Macrophage, Live cell imaging, Lysosome, Mycobacterium

1

Introduction Phagocytosis is the process by which professional and non-professional phagocytes ingest particles whose size normally exceeds 1 μM [1]. The resulting intracellular organelles, termed phagosomes, progress through biochemical changes that modify the composition of both their limiting membrane and contents, by a sequence that resembles the progression of the endocytic pathway [2, 3]. This process is referred to as phagosome maturation and bestows the phagosome with degradative properties, which are central to the microbicidal function of phagosomes and the first line of defense against infection in multicellular organisms. During evolution, many intracellular pathogens developed strategies to survive within host cells and manipulate phagosome maturation. A good example is the intracellular pathogen Mycobacterium tuberculosis (Mtb) that manipulates the cellular trafficking machinery and disrupts phagosomal membrane integrity to survive within macrophages [4–10]. Phagosome maturation is a highly complex and dynamic pathway, being the result of multiple rounds of vesicular fusion but also

Roberto J. Botelho (ed.), Phagocytosis and Phagosomes: Methods and Protocols, Methods in Molecular Biology, vol. 2692, https://doi.org/10.1007/978-1-0716-3338-0_13, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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phagosomal membrane tubulation and fission (e.g., recycling) and other types of transient interactions such as “kiss and run” fusion or tubular contacts [2, 11, 12]. Thus, in addition to complete fusion with other organelles, transient and rapid contacts are also responsible for many aspects of the maturation process [11, 13]. As well as being dynamic, maturation of a phagosome to a phagolysosome can take different amount of time depending on its contents and the ligand that triggers internalization. Monitoring the process by specific late endocytic markers shows that formation of a phagolysosome can take between 15 and 60 min [14, 15]. Therefore, studying the association of specific endocytic markers with phagosomes by methods such as indirect immunofluorescence or immunogold labelling of ultrathin thawed cryosections in fixed cells has technical limitations. For example, rapid events (during the first 2–3 min of phagosome formation) or transient events such as “kiss and run” fusion or tubular structures cannot be analyzed in real time using fixed samples. When compared to techniques with low temporal resolution such as immunolabelling of fixed cells, Western blot of isolated phagosomes, proteomics, or flow cytometry [16–18], live cell imaging provides a complementary tool to investigate phagosome maturation progression continuously over long periods of time, providing important information about bacterial replication and host cell activation in real time [19]. Compared to “high content” techniques such as flow cytometry of phagosomes (phagoFACS), live cell imaging is time-consuming. However, when combined with automated or semi-automated quantitative analysis, live cell imaging is very powerful and provides critical, and sometimes unique, information about changes in composition of phagosomes in real time, as well as transient recruitment of membrane repair machinery and inter-organelle contacts, at the single cell and individual phagosome level [19, 20]. Therefore, live cell imaging provides complete spatiotemporal dynamics information of phagosome membrane composition and damage/repair which may not be captured by immunolabelling of fixed cells. Here, we describe simple methods to investigate the dynamic association of late endocytic markers with phagosomes using live cell imaging, as a measurement of phagosome maturation. We also explain the process required for semi-automated quantitative image analysis in a typical experiment.

2

Materials

2.1 Cells, Buffers, and Solutions 2.1.1

Cells

1. Bone marrow from C57BL/6 mice (see Note 1). 2. KOLF2 iPSC cells (HPSI0114i-kolf_2 iPSC, Public Health England Culture Collections, Cat #77650100).

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3. Leukocyte cones (NC24) (NHS Blood and Transplant service, UK) (see Note 2). 4. Mycobacterium tuberculosis expressing H37Rv E2-CRIMSON (see Note 3). 2.1.2

Culture Media

1. Bone marrow-derived macrophages (BMM) complete medium: Roswell Park Memorial Institute (RPMI) 1640 Medium supplemented with GlutaMAX, 10% fetal calf serum (FCS). During differentiation steps medium is further supplemented with 20% L929 fibroblast culture supernatant. Store at 4
C (see Note 4). 2. Human monocyte-derived macrophages (HMDM) complete medium: Roswell Park Memorial Institute (RPMI) 1640 Medium with GlutaMAX and HEPES, 10% heat-inactivated FCS, 100 ng/mL hM-CSF or 10 ng/mL of hGM-CSF. Store at 4
C. 3. Human-induced pluripotent stem cells (iPSC) culture medium: Essential 8™ Medium (Gibco, Thermo Scientific, USA) (see Note 5). Store at 4
C. 4. Embryonic body (EB) culture medium: Essential 8™ Medium (Gibco, Thermo Scientific, USA) supplemented with Y-27632 ROCK inhibitor (see Note 6), 50 ng/mL hBMP4 50 ng/mL hVEGF, 20 ng/mL hSCF. Store at 4
C. 5. Factory complete medium: X-VIVO15 (Lonza, Switzerland) supplemented with 1% (v/v) Glutamax, 0.1% (v/v) β-mercaptoethanol, 100 ng/mL hM-CSF, 25 ng/mL hIL-3. Store at 4
C. 6. Human-induced pluripotent stem cell-derived macrophages (iPSDM) differentiation medium: X-VIVO15 (Lonza, Switzerland) supplemented with 1% (v/v) Glutamax, 0.1% (v/v) β-mercaptoethanol, 100 ng/mL hM-CSF. Store at 4
C. 7. 7H9 medium for mycobacteria: Middlebrook 7H9 broth base (Fluka Analytical, Sigma-Aldrich, Germany) in distilled water, supplemented with BBL Middlebrook ADC Enrichment (BD Diagnostics, USA), 0.05% (v/v) Tween 80, and 0.2% glycerol. Store at 4
C.

2.1.3 Reagents for the Preparation of Phagocytic Targets

1. Polystyrene Beads. 1. 2.5% (w/v) 3 μM polystyrene beads (Krisker Biotech, Germany). 2. 1 M MES buffer, pH 6.7 (Sigma-Aldrich, Germany). 3. 10 mg/mL EDAC (1-ethyl-3-[3-dimethylaminopropyl] carbodiimide hydrochloride) (Sigma-Aldrich, Germany).

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4. Stop buffer pH 9.4 (1% Triton-X100 in 10 mM Tris–HCl (Sigma-Aldrich, Germany)). 5. Mouse IgG (Rockland, USA) or fluorophore-conjugated mouse IgG (Jackson ImmunoResearch laboratories, USA). 2. Silica Beads. 1. 50 mg/mL (w/v) 3 μM silica beads, Flou-Red, surface NH2 (Krisker Biotech, Germany, PSI-R3.0NH2). 2. 50 mg/mL (w/v) 3 μM silica beads, Flou-Green, surface NH2 (Krisker Biotech, Germany, PSI-G3.0NH2). 3. 10% (w/v) Bovine Serum Albumin. 2.1.4 Labelling of Intracellular Organelles

1. LysoTracker® Blue DND-22 (L7525), Red DND-99 (L7528), or Far Red (L12492) (Invitrogen, USA). 2. Neon™ Electroporation System (MPK5000), 100 μL Neon™ Electroporation Kit (MPK10096) (Thermofisher, USA). 3. DNA/plasmids of fluorescent genes.

2.1.5 Preparation of BMM

1. Phosphate-buffered saline pH 7.4 (PBS).

2.1.6 Preparation of HMDM

1. Ficoll-Paque Premium (GE Healthcare, USA).

2. 70% ethanol.

2. MACS rinsing solution (Miltenyi Biotec, UK). 3. RBC lysing buffer (Sigma-Aldrich, Germany). 4. 1% BSA. 5. Anti-CD14 magnetic beads (Miltenyi Biotec, UK).

2.1.7 Preparation of Human [21] and EB

1. Vitronectin XF (Stem Cell Technologies, Canada). 2. TrypLE™ (Gibco, Thermo Scientific, USA). 3. DPBS buffer. 4. Anti-Adherence Rinsing Solution (Stem Cell Technologies, Canada).

2.2

Equipment

2.2.1 Preparation of BMM

2.2.2 Preparation of HMDM

1. Two pairs of autoclaved scissors and forceps. 2. Petri dishes. 3. Syringe and 25G 5/8 (0.5  16 mM) needle (Terumo, Germany). 1. LS column (Miltenyi Biotec, UK). 2. QuadroMACS separator magnet (Miltenyi Biotec, UK). 3. Scrappers (Sarsted, Germany).

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1. 40 μM cell strainer (Corning, USA). 2. T225 EasyFlask (ThermoFisher, USA). 3. Scrappers.

2.2.4

Live Cell Dishes

2.2.5 Microscope and Environmental Chamber

1. 35 mM glass bottom dishes with 12 mM or 22 mM glass aperture, thickness #1.5 (WillCo Wells, Germany). 1. Inverted confocal microscope (see Note 7). 2. HC PL APO CS2 63.0/1.40 OIL objective. 3. Argon (488 nM), DPSS (561 nM), HeNe (633 nM) lasers. 4. Environmental control chamber (37
C, 5% CO2, 20–30% humidity) (see Note 8). 5. Type 37 immersion oil (Cargille Laboratories, USA). 6. Leica Applications Suite (LAS) software for image acquisition. 7. Fiji (Fiji is just ImageJ), available for download at http://fiji.sc (NIH, USA).

3

Methods

3.1 Preparation of Macrophages

In order to carry out phagocytosis experiments, it is possible to use either mouse macrophage cell lines such as RAW264.7 macrophages (not covered in this chapter) or primary mouse bone marrow-derived macrophages (BMM). The use of mouse primary cells is considered to be more physiologically relevant regarding macrophage function. In cases where a particular knockout mouse is available, they can be used to analyze the role of specific host factors in phagosome biology. Primary cells also avoid the problem of bias of clonal selection. However, BMM are more challenging to use for experiments requiring expression of fluorescently tagged markers since transfection efficiency is consistently very low. The use of lentiviral-based expression vectors significantly improves transfection yields, but sometimes with noticeable macrophage activation [22]. Nucleoporation is a good alternative, although macrophage functionality can be compromised. In cases where fundamental questions regarding intracellular trafficking are addressed, RAW264.7 macrophages represent a suitable model to study the localization of specific markers and their kinetics of association to phagosomes. In fact, many important discoveries in phagosome maturation studies in RAW264.7 macrophages were reproduced in BMM [23]. Primary human macrophage models are very limited and prone to donor to donor variability. We described here the use of the monocyte-derived macrophages (HMDM) and stem cell-derived

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macrophages (iPSDM). Both systems are more physiologically relevant in the context of human biology [24, 25]. 3.1.1 Preparation of BMM

1. Perform culling of the mice by cervical dislocation. 2. After culling, use a set of sterile/autoclaved scissors and forceps to dissect and remove the femur and tibia bones from the mouse hind legs (see Note 9). 3. Remove the mouse tissue from the bones, and collect them in PBS (see Note 10). 4. In a sterile Petri dish, rinse the bones with 70% ethanol, and then transfer to a new Petri dish, and repeat once more with 70% ethanol and once with PBS. Then transfer the bones to a new Petri dish containing culture medium. 5. Cut off the ends of the bones to make the bone marrow accessible (see Note 11). 6. Flush the medullary cavity with ice-cold culture media using a syringe attached to a 25G needle. Note: If the bones are flushed properly, they will appear translucent/white. 7. Centrifuge the cells at 350  g for 10 min at 4
C to obtain a pellet. The pellet is then resuspended in BMM complete medium and plated in sterile microbiology uncoated 9 cm Petri dishes. Incubate the cells at 37
C in 5% CO2 atmosphere. Note: Count using a hemocytometer or an automated cell counter, and plate approximately 4  106 cells in 10 mL BMM culture media per dish. 8. Every 48 h aspirate 70–80% of the medium from the Petri dishes, and replace with fresh complete medium for at least 5 days to differentiate into macrophages. 9. After differentiation, cells should be transferred into glass bottom dishes at the concentration required for the experiment. Wash BMM twice with PBS to remove non-adherent cells. Add 5 mL of ice-cold PBS to each dish, and leave them on ice at 4
C until all cells are completely detached. Cells should be detached after 10–15 min. 10. Carefully collect the cells by washing the dishes and centrifuge at 350  g for 10 min. 11. Resuspend the pellet in BMM complete cell culture medium. 12. Count and dilute BMM to the required density, and transfer the cells into the recess of the glass bottom live cell dishes. 13. For live cell imaging, 4  106 cells per 22 mM aperture dish in 1 mL medium are transferred and subsequently incubated at 37
C in 5% CO2 atmosphere.

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1. Human monocytes were prepared from Leucocyte cones (NC24) supplied by the NHS Blood and Transplant service [26] (see Note 12). 2. Isolate the white blood cells by centrifugation on Ficoll-Paque Premium for 60 min at 300  g. 3. Collect the mononuclear cells, and wash twice with MACS rinsing solution to remove platelets and red blood cells. 4. Incubate the remaining samples with 10 mL RBC lysing buffer per pellet for 10 min at room temperature. 5. Wash the cells with rinsing buffer, and resuspend in 80 μL MACS rinsing solution supplemented with 1% BSA and 20 μL anti-CD14 magnetic beads per 108 cells. 6. After 20 min on ice, wash the cells in MACS/BSA solution, and resuspend at a concentration of 108 cells/500 μL in MACS/BSA solution, and further pass through an LS column in the field of a QuadroMACS separator magnet. 7. Wash the LS column three times with MACS/BSA solution, then elute CD14 positive cells, centrifuge, and resuspend in HMDM complete medium. Note: Plate approximately 4–5  106 cells in 10 mL BMM culture media per dish. 8. Incubate the dishes in a humidified 37
C incubator with 5% CO2. 9. After 3 days, add an equal volume of fresh complete media including 10 ng/mL of hGM-CSF. 10. After differentiation, cells should be transferred into glass bottom dishes at the concentration required for the experiment. 11. Wash HMDM once with PBS to remove non-adherent cells. Add 3 mL of 0.5 mM EDTA in ice-cold PBS. 12. Leave them on ice at 4
C until all cells are completely detached. 13. Add 7 mL PBS and gently scrape them using cell scrapers. 14. Pellet the cells by centrifugation, and resuspend in RPMI medium containing 10% FBS.

3.1.3 Preparation of iPSDM (As Alternative to BMM or HMDM)

1. Maintain iPSC in Vitronectin XF-coated plates with Essential 8™ Medium. 2. Once the confluency reaches 70%, wash the cells one time with 1 DPBS, and add 3 mL of Versene solution. 3. Incubate the cells at 37
C for 5 min. 4. Remove the Versene solution gently, and resuspend the cells within Essential 8™ Medium.

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5. Plate the cells in 1:6 ratio until iPSCs reach the required quantity to proceed EBs preparation. 6. Prior to EBs seeding, equilibrate an AggreWellTM plate with anti-adherence rinsing solution. 7. Add 0.5 mL anti-adherence rinsing solution to the wells, and centrifuge at 3000  g for 3 min to remove any bubbles. 8. Aspirate and rinse the wells with 1 mL 1 DPBS. 9. Add 1 mL of EB culture medium. 10. Centrifuge plate at 3000  g for 3 min, and place the plate in CO2 incubator at 37
C. 11. Remove medium from iPSC-containing plate and rinse once with 1 DPBS. 12. Aspirate DPBS, and add 1 mL room temperature TrypLE, and incubate at 37
C for 5 min. 13. Dilute the TrypLE solution containing the cells with 1 DPBS (1:10), and collect the cells gently in a centrifuge tube. 14. Centrifuge at 400  g for 5 min. 15. After the centrifugation, remove the DPBS-TrypLE solution, and resuspend the cells in EB culture medium. 16. Ensure a final cell concentration of 4.0  106/mL. 17. Add 1 mL of the cell suspension to AggreWell (making a total volume of 2 mL). 18. Centrifuge the plate at 100  g for 3 min. 19. Examine the cell distribution after the centrifugation, and return the plate to CO2 incubator at 37
C. 20. Change the medium daily by removing twice 50% of EB culture medium. 3.1.4 Preparing iPSDM for Experiments

1. On day 4, harvest the EBs by gently flushing out of the well with pipetting. 2. Filter the collected EBs through an inverted 40 μm cell strainer. 3. Seed the EBs from each AggreWell into one T225 flask in 30 mL Factory complete medium. 4. Feed the monocyte factories once per week with 20 mL of Factory complete medium for 5 weeks until an adequate number of monocytes are observed in the supernatant. 5. Weekly harvest 20–40 mL of the supernatant, and following the harvest feed the factories with 20–40 mL Factory complete medium. 6. Centrifuge the supernatant at 300  g for 5 min. 7. Resuspend the monocytes in iPSDM differentiation medium.

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8. Plate 4–5  106 cells per 10 cm Petri dish or 10–12  106 cells per 15 cm petri dish to differentiate over 7 days. 9. On day 4 after monocyte plating, replace 50% of the media with fresh iPSDM differentiation medium. 10. After 7 days of differentiation, wash one time with 1 DPBS. 11. Add 3 mL of Versene solution to 10 cm Petri dish or 5 mL to 15 cm Petri dish. 12. Incubate at 37
C for 15 min. 13. Dilute the Versene solution with 1 DPBS (1:3), and gently scrape the macrophages. 14. Centrifuge the suspension at 300  g, and check the viability of the macrophages by resuspending in iPSDM complete medium. 3.2 Preparation of Phagocytic Targets

There are several particles that can be used to study phagosome maturation by live cell imaging. Here we will be describing two different methods: firstly, the use of inert particles as a model of phagocytosis for polysterene and silica beads and, secondly, mycobacteria (Mycobacterium tuberculosis H37Rv) as an intracellular pathogen that modulates normal phagosome maturation.

3.2.1 Polystyrene and Silica Beads

Polystyrene and silica beads are inert particles that can be used to study phagosome maturation. Fcγ-receptor-mediated internalization is one of the best characterized internalization mechanisms involved in phagocytosis and further phagosome maturation [2]. Opsonization of the polystyrene beads by coating them with bovine serum albumin (BSA) and/or the appropriate IgG is required to ensure that they are recognized and internalized by macrophages. Furthermore they can be coated with fluorescent dyes to visualize them.

Human IgG Coating 1. Mix 400 μL polystyrene beads (3 μm polystyrene beads (2.5% (w/v))) with 100 μL MES buffer, pH 6.7, and 50 μg of mouse IgG. 2. Mix the solution of beads for 15 min in a rotating wheel at room temperature. 3. Add 14 μL of EDAC (10 mg/mL in sterile water) to the beads, and mix the whole solution in a rotating wheel for 1 h at room temperature. Then add another 14 μL of EDAC and mix for a further 1 h. 4. The coupling reaction is stopped by washing with Stop buffer (1% Triton-X 100 in 10 mM Tris–HCl, pH 9.4) three times. During these washing steps, the bead solution is centrifuged,

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and the supernatant is removed before the beads are resuspendend in 1 mL Stop buffer. 5. The IgG-coated beads are stored in PBS at 4
C. PBS was added to the beads to give a final concentration of 1% (w/v). Bovine Serum Albumin (BSA) Opsonization Cells were seeded on the day prior to the experiment. 1. For initial washing, mix 100 μL of beads in 1 mL of 1 PBS (1: 10 dilution). Centrifuge the beads at 10,000  g for 5 min at room temperature, and remove PBS. 2. Opsonize the beads in 1 mL of 10% BSA in 1 PBS, and incubate for 1 h at room temperature or alternatively overnight at 4
C while gently rotating. 3. Upon opsonization, wash the beads in 1 PBS (as previously described). Resuspend the beads in 1 mL cell culture media at an approximated concentration of 3.6  109 beads/mL. 4. Further dilute the beads in cell culture before addition to the cells. For 50,000 cells, the ideal dilution would be 1:400. Add 100 μL of 1:400 dilution per well.

3.2.2 Preparation of Mycobacteria for Infection

5. Centrifuge plate at 100  g for 5 min. Incubate for 30 min at 37
C to allow internalization. After 30 min, wash the cells twice with 1 PBS, and replace with cell culture media. Mycobacterium tuberculosis H37Rv (Mtb) expressing E2-CRIMSON was used in the experiments shown in Figs. 1 and 2 to analyze the association of LysoTracker or various phagosomal markers to bacteria. 1. 1.Grow mycobacteria in rolling 50 mL falcon tubes to mid-exponential phase (OD600 0.5–1) in 7H9 medium for mycobacteria at 37
C (see Note 13). 2. Pellet bacteria in a 50 mL falcon tube by centrifuging at 2000  g for 5 min at room temperature. 3. Wash the pellet two times with PBS. 4. Add 2.5–3.5 mM sterile glass balls to mycobacterial pellets (see Note 14). 5. Break bacterial clumps by vigorously shaking the bacterial pellet together with the glass balls for 1 min (see Note 15). 6. Resuspend the bacteria in cell culture medium with respect to the cell background that is in use. 7. Spin down any remaining clumps by centrifugation at 320  g for 5 min. 8. The upper part of the supernatant after centrifugation contains a suspension of single bacteria (rather than clumps of bacteria).

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Fig. 1 Overview of the experimental setting in live cell imaging of phagosome maturation. (a) Mouse macrophage cell models used include bone marrow-derived macrophages (BMDMs), prepared from mouse legs, and RAW 264.7 cell line. Human macrophage cell models used include monocyte-derived macrophages (MDMs), extracted from blood samples, and induced pluripotent stem cell-derived macrophages (iPSDMs). (b) Overview of the sequential steps involved in the preparation of mycobacteria, here shown for Mtb, for the infection of macrophages. (c) Overview of experimental set-up. Macrophages prepared in (a) are plated in a glass bottom live cell dish. On the day of the experiment, LysoTracker is added to the cells for 30 min prior to infection with Mtb prepared in (b). The dish is placed in a holder and transferred to the microscope for imaging. (d) Image settings used for acquisition of time-lapse movies using a Leica TCS SP5 or TCS SP8 confocal microscope. (Figure created with BioRender.com)

9. Measure the OD600 (blank using cell culture medium), and calculate the volume needed to add to the macrophages assuming that an OD600 of 1 represents 1 108 bacteria. For live cell

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Fig. 2 Diagram of the image analysis flow and representative data. Overview of the different steps during the image analysis of association of LysoTracker to Mtb. Detailed description can be found in Subheading 3.5.2. (a) Image preparation for analysis. The image is opened in Fiji, and one focal plane is selected for the following analysis. In the images LysoTracker Red DND-99 is shown in red and EGFP-Mtb in green. (b) Selection of the bacteria. The channels with the different colors are split and bacteria are thresholded. (c) Image calculations. The thresholded image is dilated and eroded, and then the two images are subtracted from each other to result in the selection of only the regions surrounding the bacteria. (d) Analysis. The measurements are set and the association of LysoTracker to Mtb is analyzed. (e) Association of LysoTracker with EGFP-Mtb H37RV in BMM. Quantification shows mean  SEM from ten cells in two biological experiments

imaging in BMM with LysoTracker, we added Mtb at a multiplicity of infection [27] of 2 in 1 mL BMM medium and MOI of 1 in 1 mL for human macrophages (see Note 16). 3.3 Labelling of Intracellular Organelles

There are multiple methods to investigate live dynamics of phagosome maturation; here we will be focusing on two methods: one using a fluorescent probe that labels acidic organelles and another using expression of fluorescently tagged proteins in macrophages. Additionally, transfer of pre-loaded fluorescent dextran into phagosomes can also be used which has previously been described in detail [11, 28].

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LysoTracker is a fluorescent dye linked to a weak base and is used to label and track acidic compartments in live cells [29, 30]. The weak base causes it to be only partially protonated at neutral pH, allowing LysoTracker to freely cross cell membranes in live cells. The probe is then trapped in acidic compartments following protonation of the base. There are different commercially available LysoTracker probes: LysoTracker Red, Green, and Blue. In our experience the most reliable probe is the LysoTracker Red and Far Red, since it gives a bright fluorescent signal, without fast bleaching or high cytotoxicity. LysoTracker Blue might be required in cases where red and green channels are already used. However, LysoTracker Blue has the disadvantage of bleaching faster and being more cytotoxic, and therefore only short time periods can be imaged using this dye. We would recommend to use this probe for acquiring snapshots at spaced time points (e.g., every hour) rather than continuous image acquisition. It is very important to consider that LysoTracker association with beads or bacterial phagosomes could be both due to acidification of this compartment and therefore direct LysoTracker accumulation or due to delivery by fusion with LysoTracker positive compartments. In the second case, the dye is trapped in an acidic compartment by protonation of its base and then gets delivered into the phagosome [30]. 1. LysoTracker solution is diluted to 50 nM in complete medium (1:10,000 dilution) (see Note 17). 2. LysoTracker solution is added to macrophages 30 min before addition of beads/mycobacteria in 1 mL media. 3. LysoTracker containing medium is replaced, and cells are washed three times with 1 PBS before beads or mycobacteria are added as described in Subheading 3.2.

3.3.2

Cell Transfection

To monitor association of cellular markers to phagosomes, macrophages are transfected with plasmids expressing fluorescently tagged proteins. Here we describe one protocol to efficiently transfect macrophages using the Neon™ Electroporation System (refer to manufacturer’s protocol for further details) (see Note 18). 1. Harvest the macrophages (as previously described), count the cells, and wash once in 1 mL of 1 PBS. Centrifuge at 300  g for 5 min and remove PBS. 2. Per 100 μL electroporation reaction, resuspend 1  106– 1.1  106 cells and 10 μg of plasmid in 100 μL of Buffer R. Mix thoroughly. 3. Using the 100 μL pipette tip, aspirate the cells (see Note 19). Electroporate using the following set-up for iPSDMs: 1400 V, 30 ms, 1pulse in 3 mL of Buffer E.

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4. Immediately add cells to the culture media and mix gently before plating. 3.4

Live Cell Imaging

There are various suitable microscopes available to perform live cell imaging, including wide-field fluorescent, spinning-disk, and confocal laser scanning microscopes. There are both advantages and disadvantages between the different systems, and important parameters to consider are stability, photocytotoxicity, photobleaching, and resolution [28]. For the live cell imaging studies described here, we used a confocal laser scanning microscope since it provides high resolution with relatively low phototoxicity. In combination with a microscope environmental chamber, it provides a very stable and reliable system. An overview of the experimental design is shown in Fig. 1. 1. Transfer cells into glass bottom live cell dishes as described in Subheadings 3.1.1 and 3.1.2. 2. Prepare a single cell mycobacterial solution, and add bacteria at a MOI of 2 in 1 mL total volume to the BMM for live cell imaging as described in Subheading 3.2.2 (see Note 20). 3. For live cell imaging, an environmental control chamber providing 37
C, 5% CO2, and 20–30% humidity in a closed chamber is needed (see Notes 21 and 22). 4. Acquire images from the bright-field channel to check both for cell morphology and in the case of beads for their localization (see Note 23). 5. Always save the data in the native file format of the imaging system used.

3.5

Image Analysis

3.5.1 Analysis of the Association of Various Markers to IgGCoated Beads

The image analysis can be performed using various software packages; here we will describe the use of the freely available javabased Fiji platform. Fiji is an ImageJ distribution containing additional preloaded plugins. It is available for download at http:// fiji.sc. Macros and plugins used are mentioned in notes. An overview of the imaging analysis flow and a representative experiment are shown in Fig. 2. We have previously described in detail the image analysis of the association of various markers to bead-containing phagosomes [28]. Therefore, we will only briefly outline the steps required for this analysis and focus in the next section on bacteria-containing phagosome analysis that is more challenging. In general, many steps in the analysis of association of different markers to beads or mycobacteria are very similar [28]. 1. Open the image saved in the native file format in Fiji using the “LOCI” – “Bio-Formats Importer,” and split the different

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channels into separate windows. The details of these steps can be found in Subheading 3.5.2. 2. Set the measurement parameters (“Analyze” – “Set Measurements. . .”), and redirect to the channel containing the signal from the probe or marker. 3. Select the channel with the beads—bright field or fluorescence depending on experimental set-up—and select a circular region of interest (ROI) on the internalized bead using the “oval selection tool.” 4. Start the analysis one frame before complete internalization, similar to analysis with mycobacteria. Starting with that frame, select “Analyze” – “Measure.” A table with the results for this measurement should appear. 5. Repeat the process for the following image time frames. As the beads move inside the cell, the ROI might need to be adjusted for localization exactly on the bead. 6. Plot the measurements obtained in the results table against the time after internalization (see Note 24). 3.5.2 Analysis of the Association of Various Markers to Mycobacteria

1. Open the file to be analyzed in Fiji by either using the plugin “LOCI” – “Bio-Formats Importer” or dragging and dropping the file into Fiji which should automatically use this plugin (see Note 25). 2. Choose the following options for opening the file, “view as hyperstack,” and in the case of files larger than the available amount of random access memory (RAM) for the computer, select “virtual” (see Note 26). 3. Create a duplicate image containing only the one focal plane with the bacterium. If the bacterium analyzed changes focal plane throughout the time-lapse movie, split the movie into several shorter files using the “Make Substacks. . .” command (“Image – Stack – Tools – Make Substack”). Select only the channels, focal plane, and time frames containing the bacterium and marker to be analyzed. Start the time-lapse movie one or two frames before the bacterium is completely internalized (see Note 27). 4. To facilitate further analysis, the short time-lapse movies can be made into a single file again using the “Concatenate...” command (“Image – Stacks – Tools – Concatenate”). Select the files in the order they appear in the original time-lapse movie. 5. Split the different channels (“Image – Color – Split Channels”). 6. Select the channel containing the bacteria. Adjust the threshold to select only the bacterial region (“Image – Adjust – Threshold”). Select a thresholding setting that will result in

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thresholding of bacteria over the complete time-lapse movie. After selecting “ok” make sure that in the window that opens “Calculate threshold for each image” is not ticked (see Note 28). 7. Starting from the thresholded image, select “Fill Holes” (“Process – Binary – Fill Holes”), and process all images. 8. Duplicate the thresholded image selected (“Image – Duplicate”). Tick the box next to “Duplicate Hyperstack.” 9. Select the duplicated image, and reduce the size of the thresholded area by 1 pixel (“Process – Binary – Erode”). 10. Select the original thresholded image, and increase the size of the thresholded area by 1 pixel (“Process – Binary – Dilate”). 11. Subtract the duplicated eroded image from the original dilated one (“Process – Image Calculator...”). In the window that opens, select “Create new window” and then “Process all images.” 12. Select the newly generated image which should now show a thresholded area around the location of the bacteria. The size of the ring can be adjusted by multiple rounds of dilation or erosion before subtracting the two thresholded images from each other. 13. Go to “Analyze” – “Set Measurements. . .,” and select area, mean, gray value, standard deviation, integrated density, display label, and any other parameter that you wish to measure. 14. In the same window, click “redirect to,” and select the channel containing the signal from the probe or marker. In Fig. 2 we redirected to the channel containing the signal from LysoTracker. 15. Go to “Analyze” – “Measure particles. . . .” Select the size of the particles to be analyzed: “Size (pixel^2): 5 – Infinity,” “Circularity: 0.00–1.00,” “Show: Outlines,” “Display results.” The lower size can be adjusted depending on the size of the particle/bacteria measured. Selecting a value between 5 and 10 in most cases is suitable to stop measurement of thresholding artifacts. 16. A results table will appear with the values for the signal intensity in the channel with the marker in each thresholded area in every single image. In the case of bacteria, the area of the thresholded regions can change significantly depending on bacterial size and orientation. To compare between different results, use a measurement that is independent of area, e.g., mean intensity (see Note 29). 17. Check in the Results window showing the outlines that only one thresholded area corresponding to the location of the

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analyzed bacterium is measured. In some cases cells will internalize more than one bacterium, which can result in the measurement of the signal from two particles. In this case only select the measurements obtained from the same single bacterium. 18. Plot the measurement against time. In Fig. 2 we plotted the mean intensity against time after internalization.

4

Notes 1. Animal care and all the procedures, including training, described in this chapter follow the institutional and national guidelines. 2. Mycobacterium tuberculosis H37 is one of the most commonly used laboratory strains. The H37 strain is further distinguished as “virulent” (Rv) or “avirulent” (Ra). 3. Human samples and all the procedures described in this chapter follow the institutional and national guidelines. 4. Macrophage colony-stimulating factor (M-CSF) is secreted by L929 cells and is used in the form of L929-conditioned medium. Supernatants of L929 cells provide a cheap source of M-CSF but contain many other growth factors. A more expensive alternative is to use purified growth factors such as granulocyte-macrophage colony-stimulating factor (GM-CSF) or (M-CSF). 5. Essential 8™ Medium is sensitive to temperature and should be warmed at room temperature. 6. Y-27632 ROCK inhibitor should be used only the day of EB seeding. It should be removed from the EB culture medium in the following days. 7. For live cell imaging, Leica TCS SP5 AOBS laser scanning confocal microscope equipped with HyD detectors and Leica SP8 inverted confocal microscope equipped with HyD detectors (Leica Microsystems, Germany) were used due to availability. 8. Temperature stability is critical for long-term imaging since very subtle variations in temperature during image acquisition will cause focus drift. Alternatively, many companies offer autofocus correction. Different types of environmental boxes are available in the market; we use a customized box from EMBL technology transfer (Heidelberg, Germany). 9. The hind legs are longer and will give a higher BMM yield. 10. To reduce risk of contamination, perform all following steps in a class II biosafety cabinet.

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11. Use a different set of sterile/autoclaved scissors and forceps then during the removal of the bones from mice to avoid contamination. Macrophages are sensitive to activation by lipopolysaccharides, and glassware should be avoided. 12. Ethical approval to work with human macrophages needs to be in place. 13. M. tuberculosis is a biosafety level 3 (BSL3) pathogen and should be handled in a BSL3 laboratory obeying all appropriate regulations. 14. The amount of glass balls added depends on the size of the pellet; the volume of both pellet and balls should be similar. 15. M. tuberculosis is notorious for clumping; it is particularly important to have a single cell suspension for live cell imaging to image single events. 16. M. tuberculosis should be added at a low MOI where possible. Depending on the cell type used for infection, the MOI might have to be adjusted. If there are too many extracellular bacteria, reduce the MOI, and/or wash cells before the image acquisition. 17. Very high concentrations of LysoTracker can result in labelling of the bacteria. 18. In addition to Neon™ Electroporation System, Lonza Nucleofector™ (Lonza, Switzerland) is another example to efficiently transfect macrophages. 19. Ensure there are no bubbles inside the tip. 20. In the case of infection with Mtb, add parafilm around the dish, and place it in a live cell dish-clamping holder. 21. In the experiment shown in Fig. 2, we used the following settings which can also be seen in the panel on the right side of the figure: acquisition mode—xyzt, resolution 1024  1024 pixels, 400 Hz scanning speed, 3 line averaging, sequential mode acquisition, z-stack with four steps, interval between image acquisitions 5 min. 22. The imaging settings should be adjusted according to the experiment. If the acquisition of a very fast event is sought, it might be necessary to adjust both the resolution/line averaging and number of z-planes to decrease the interval between image acquisitions. For longer time-lapse movies with bacteria, it is best to acquire images as a z-stack choosing multiple focal planes as both cells and bacteria can move during the image acquisition and can otherwise not be followed over time. 23. In the case of transfected cells, bright-field channels can also be used for looking at the localization of surrounding non-transfected cells.

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24. In contrast to bacteria, the size and shape of inert particles will not change during the imaging process. Therefore, instead of plotting the “mean” value obtained, it is also possible to plot the intensities independent of the area (“IntDen” column in the results table). 25. The file should have been saved in the native file format of the imaging system used as mentioned in Subheading 3.4. In the case of Leica confocal microscope, it will be a .lif file. 26. Choosing to view the file as hyperstack will make it easier to follow the time-lapse movie. Furthermore, this step is essential for splitting the image for analysis of data from one focal plane. Viewing the file as “virtual” reduces the capacity needed to open and then view the file and therefore helps speeds up the analysis process for large files. 27. It is essential to analyze the association of markers to bacteria from only one focal plane to avoid the signal from another location in the cell from resulting in a false-positive bacterial association signal. 28. During the thresholding step, select a lower limit of a value n depending on the signal intensity in the images and the maximal value 255 as the upper limit. 29. The mean intensity refers to the total signal intensity measured for each particle divided by the area of the particle.

Acknowledgments We thank the host-pathogen interactions in tuberculosis laboratory for useful discussions and comments on the manuscript. This work was supported by the Francis Crick Institute (to MGG), which receives its core funding from Cancer Research UK (FC001092), the UK Medical Research Council (FC001092), and the Wellcome Trust (FC001092). This project has received funding from the European Research Council [31] under the European Union’s Horizon 2020 research and innovation program (grant agreement n
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Chapter 14 Measurement of Salmonella enterica Internalization and Vacuole Lysis in Epithelial Cells Jessica A. Klein, TuShun R. Powers, and Leigh A. Knodler Abstract Establishment of an intracellular niche within mammalian cells is key to the pathogenesis of the gastrointestinal bacterium, Salmonella enterica serovar Typhimurium (S. Typhimurium). Here we will describe how to study the internalization of S. Typhimurium into human epithelial cells using the gentamicin protection assay. The assay takes advantage of the relatively poor penetration of gentamicin into mammalian cells; internalized bacteria are effectively protected from its antibacterial actions. A second assay, the chloroquine (CHQ) resistance assay, can be used to determine the proportion of internalized bacteria that have lysed or damaged their Salmonella-containing vacuole and are therefore residing within the cytosol. Its application to the quantification of cytosolic S. Typhimurium in epithelial cells will also be presented. Together, these protocols provide an inexpensive, rapid, and sensitive quantitative measure of bacterial internalization and vacuole lysis by S. Typhimurium. Key words Salmonella enterica, Gentamicin protection assay, Chloroquine resistance assay, Salmonella-containing vacuole, Type III secretion system, Vacuole lysis, Epithelial cells, Colony-forming units

1

Introduction Many years ago, it was noted that intracellular bacteria are protected from the bactericidal action of antibiotics [1, 2]. This observation forms the basis for the gentamicin protection assay [3], a widely used technique to study the internalization of bacteria into mammalian cells. Gentamicin is an aminoglycoside antibiotic that has proven efficacy against many Gram-negative and Gram-positive bacteria. It is widely believed to be ineffective against intracellular bacteria because it does not penetrate mammalian cells. However, gentamicin can penetrate eukaryotic cells via endocytic and non-endocytic routes [4–6], albeit poorly, and accumulates in lysosomes, as well as the Golgi complex, cytoplasm, and nucleus [7, 8]. Rather than being membrane-impermeant, the lack of bactericidal activity of gentamicin against internalized bacteria is more

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likely due its relatively low cellular accumulation. The gentamicin protection assay requires no expensive equipment and generates quantitative, reproducible data in a relatively short time. While generally conducted in 24-well tissue culture plates, it can easily be adapted to 48-well or 96-well plates for the screening of bacterial mutant libraries or testing the effects of either chemical compounds or siRNA knockdown of mammalian genes on bacterial invasion, for example. Salmonella enterica serovar Typhimurium (S. Typhimurium) is a facultative, intracellular pathogen that causes self-limiting gastroenteritis in many animal species, including humans, after the ingestion of contaminated food and water. S. Typhimurium colonizes epithelial cells (enterocytes and goblet cells) and macrophage in the gastrointestinal tract of infected hosts. Bacterial entry into non-phagocytic epithelial cells is dependent upon a type III secretion system (T3SS1) encoded on Salmonella pathogenicity island-1 (SPI-1) [9]. T3SS1 translocates numerous type III effector proteins into host cells that alter host cytoskeletal architecture to generate large actin-rich plasma membrane protrusions, known as “ruffles,” that engulf the bacteria into a membrane-bound vacuole, the Salmonella-containing vacuole (SCV) (reviewed in [10]). Flagellabased motility is also required for S. Typhimurium entry into non-phagocytic cells [11–13], and SPI-1 and flagella are co-regulated under a variety of in vitro conditions [14–16]. Bacterial internalization is independent of T3SS2, a second T3SS encoded on SPI-2 [17, 18], which functions intracellularly to translocate type III effectors required for SCV maturation and positioning (reviewed in [19]). Here, we will describe the use of the gentamicin protection assay to quantify S. Typhimurium invasion into HeLa epithelial cells, a non-phagocytic human cell line that has been used extensively to decipher many aspects of the type III effector-mediated internalization event. While the majority of internalized S. Typhimurium remain confined within a membrane-bound vacuole in epithelial cells, a small but significant proportion lyse or damage their nascent phagosome to enter the host cell cytosol [20, 21]. Some of these cytosolic bacteria are eliminated by xenophagy [20, 22, 23], which is the autophagic degradation of bacteria, but some can escape recognition by selective autophagy receptors [24, 25] and ultimately hyper-replicate in the cytosol of epithelial cells [21, 25– 27]. This eventually leads to epithelial cell death by pyroptosis and lytic release of bacteria [26]. Chloroquine (CHQ) is a lysosomotropic agent that accumulates to high concentrations within endosomes, but does not access the cytosol [28]. The differential intracellular distribution of CHQ imparts its specificity against vacuolar but not cytosolic bacteria [29], although the mechanism of action of CHQ-dependent killing of vacuolar bacteria remains unknown. The CHQ resistance assay was initially used to identify

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Shigella flexneri mutants that did not lyse their internalization vacuole [30]. Recently, we adapted this assay to quantify cytosolic S. Typhimurium at different times post-infection (p.i.) in epithelial cells [21] and provide a detailed description of this protocol here. Like the gentamicin protection assay, the CHQ resistance assay is relatively inexpensive and readily scalable and generates quantitative data in a short timeframe.

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Materials 1. Use double-distilled or MilliQ water to make all reagents. 2. HeLa cervical epithelial cells (American Type Culture Collection (ATCC), CCL-2), low passage number (see Notes 1 and 2). 3. Growth medium: Eagle’s minimum essential medium with Earle’s balanced salts (EMEM), 2 mM L-glutamine, 1 mM sodium pyruvate, and 10% heat-inactivated fetal bovine serum (FBS) (see Notes 3, 4, and 5). 4. Trypsin EDTA: 0.25% trypsin, 2.21 mM EDTA in HBSS without sodium bicarbonate, calcium, and magnesium (see Note 3). 5. Phosphate-buffered saline without divalent cations (PBS--): 0.14 g/L KH2PO4, 9 g/L NaCl, 0.80 g/L Na2HPO4 (anhydrous), pH 7.4 (see Note 3). 6. Dulbecco’s PBS with divalent cations (DPBS++): 0.2 g/L KCl, 0.2 g/L KH2PO4, 8 g/L NaCl, 2.17 g/L Na2HPO4·7H2O, 0.1 g/L CaCl2 (anhydrous), 0.1 g/L MgCl2·6H2O, pH 7.4 (see Note 3). 7. Hanks’ balanced salt solution (HBSS): 0.14 g/L CaCl2 (anhydrous), 0.40 g/L KCl, 0.06 g/L KH2PO4, 0.098 g/L MgSO4 (anhydrous), 8 g/L NaCl, 0.048 g/L Na2HPO4 (anhydrous), 0.35 g/L NaHCO3, 1 g/L glucose, pH 7.25 (see Note 3). 8. 75 cm2 tissue culture flasks and 24-well tissue culture treated plates (see Note 3). 9. LB-Miller broth: 10 g/L tryptone, 5 g/L yeast extract, 10 g/ L NaCl. 10. LB-Miller agar plates: 10 g/L tryptone, 5 g/L yeast extract, 10 g/L NaCl, 15 g/L agar. 11. 100 mg/mL streptomycin in water (filter sterilize and store aliquots at -20 °C). 12. 50 mg/mL gentamicin in water (filter sterilize and store aliquots at -20 °C). 13. 52 mg/mL CHQ diphosphate salt (100 mM) in water (see Note 6).

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14. Lysis buffer: 0.2 g sodium deoxycholate per 100 mL water (filter sterilize and store at room temperature). 15. Glycerol stock of Salmonella enterica serovar Typhimurium SL1344. Dilute 0.75 mL of an overnight culture of S. Typhimurium (LB-Miller broth + 100 μg/mL streptomycin) with an equal volume of 30% (v/v) sterile glycerol in a 2 mL screw cap cryovial. Gently mix and freeze the tube at -80 °C (see Note 7). 16. Freshly streaked plate of S. Typhimurium. Using a sterile loop, pipette tip or toothpick, spread some bacteria from the top of the glycerol stock onto a LB-Miller agar plate containing 100 μg/mL streptomycin, and incubate at 37 °C overnight. Keep plate at 4 °C for 500-fold

10. Add washed bacteria to growth media in each well at a multiplicity of infection (MOI, the number of bacteria added per HeLa cell) of ~100. Start the lab timer (t0). Two to three technical replicates per bacterial culture are recommended. Allow infection to proceed for 10 min at 37 °C in a 5% CO2 incubator (see Note 11). 11. While the infection is underway, make serial dilutions of bacterial subcultures in DPBS++, and spot quintuple 10 μL aliquots on dry LB-Miller agar plates. Invert plates and incubate overnight at 37 °C. Count colony-forming units (CFU) the next day to calculate the “bacterial inoculum.” 12. Aspirate the media and wash monolayers thrice with 1 mL HBSS per well to remove most of the extracellular bacteria. Add fresh growth medium, and continue incubation for 20 min at 37 °C in a CO2 incubator (see Note 12). 13. At 30 min post-infection (p.i.), replace media with growth media containing 100 μg/mL gentamicin to kill any remaining non-internalized bacteria. Continue incubation for an additional 30 min (see Note 13).

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14. At 1 h p.i., wash monolayers once with DPBS++, and solubilize by pipetting up and down in 1 mL lysis buffer (see Note 14). 15. Collect lysis buffer and transfer to 1.5 mL Eppendorf tubes. Immediately make serial dilutions in DPBS++, and spot quintuple 10 μL aliquots of each dilution on dry LB-Miller agar plates. 16. Invert plates, incubate at 37 °C overnight, and count CFU for number of “internalized bacteria.” 17. Invasion efficiency is calculated as internalized bacteria/bacterial inoculum × 100% (see Fig. 1) (see Note 11). 3.2 Chloroquine Resistance Assay

1. Culture HeLa cells to confluency in growth medium in a 75 cm2 tissue culture flask at 37 °C in a 5% CO2 incubator. 2. Remove and discard the culture medium. Rinse the cell monolayer with 5 mL PBS-- and discard. 3. Add 1 mL trypsin-EDTA, and gently swirl the flask to cover the monolayer completely. Allow the cells to incubate for 2–5 min and dislodge by gently tapping the flask. 4. Add 4 mL growth medium and resuspend the cells by gentle pipetting. 5. Count the HeLa cells on a hemocytometer or cell counter. 6. Dilute the HeLa cells to 5 × 104/mL in growth medium, and add 1 mL per well in a 24-well plate. For this assay, you will need twice as many wells for each bacterial strain or time point compared to the gentamicin protection assay (see Fig. 2). Incubate at 37 °C in a CO2 incubator for 18–24 h prior to infection. 7. Inoculate 2 mL of LB-Miller broth containing 100 μg/mL streptomycin with a single colony of S. Typhimurium (from a freshly streaked agar plate) in a 10 mL snap cap polypropylene tube. Shake at 220 rpm for 16–18 h at 37 °C (see Note 8). 8. Inoculate 10 mL LB-Miller broth (no antibiotics) with 0.3 mL overnight culture in a 125 mL Erlenmeyer flask. Shake at 220 rpm for 3.5 h at 37 °C. The optical density (OD600) of the culture should be ~3.0, which corresponds to 3 × 109 CFU/mL (see Fig. 1) (see Notes 8 and 9). 9. Centrifuge 1 mL of bacterial subculture in a microcentrifuge tube at 6000 × g for 90 s. Remove the supernatant, and resuspend the bacterial pellet in 1 mL HBSS by gently pipetting up and down. Proceed directly to infect monolayers (see Note 10). 10. Add washed bacteria to growth media in each well at an MOI of ~100. Start the lab timer (t0). Two to three technical replicates per bacterial culture are recommended. Allow infection to proceed for 10 min at 37 °C in a 5% CO2 incubator (see Note 11).

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Fig. 2 CHQ resistance assay for determining cytosolic S. Typhimurium. (a) A typical 24-well tissue culture plate set up for the CHQ resistance assay. For each time point (1.5, 3, 5, and 7 h p.i.), set aside duplicate wells for total bacteria (gentamicin, black wells) and duplicate wells for cytosolic bacteria (gentamicin plus CHQ, gray wells). (b) HeLa cells were seeded in a 24-well plate as shown in (a) and infected with S. Typhimurium SL1344 wild type. One hour prior to each time point, two wells were treated with CHQ. At the indicated times, monolayers were solubilized, and CFU enumerated by plating on LB agar. Black circles, total bacteria; open squares, cytosolic bacteria. At 1.5 h and 7 h p.i., approximately 15% and 50% of bacteria are present in the cytosol, respectively [21]

11. Aspirate the media, and wash monolayers thrice with 1 mL HBSS per well to remove most of the extracellular bacteria. Add fresh growth medium, and continue incubation for 20 min at 37 °C in a CO2 incubator. 12. At 30 min p.i., replace media with growth media containing 100 μg/mL gentamicin to eliminate any remaining non-internalized bacteria. Continue incubation for an additional 1 h (see Notes 6 and 15). 13. Reduce gentamicin concentration to 10 μg/mL thereafter. 14. To determine the extent of SCV lysis, infected cells are incubated in the presence of gentamicin with or without CHQ for 1 h (see Fig. 2). For example, to quantify the percentage of bacteria in the cytosol at 90 min p.i., incubate half the designated wells with 100 μg/mL gentamicin and the other half with 100 μg/mL gentamicin plus 400 μM CHQ from 30–90 min p.i. For 5 h p.i., incubate half the designated wells with 10 μg/mL gentamicin and the other half with 10 μg/mL gentamicin plus 400 μM CHQ from 4–5 h p.i. 15. Wash monolayers once with DPBS++, and solubilize by pipetting up and down in 1 mL lysis buffer (see Note 14). 16. Collect lysis buffer and transfer to 1.5 mL Eppendorf tubes. Immediately make serial dilutions in DPBS++, and spot quintuple 10 μL aliquots of each dilution on dry LB-Miller agar plates.

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17. Invert and incubate overnight at 37 °C. 18. Cytosolic bacteria (CHQ-resistant bacteria) = CFU from HeLa cells treated with gentamicin and CHQ; total internalized bacteria = CFU from HeLa cells treated with gentamicin alone (see Fig. 2). The percentage of cytosolic S. Typhimurium at any time point is calculated as CHQ-resistant bacteria/total internalized bacteria × 100% (see Fig. 2).

4

Notes 1. Many journals are adopting requirements for cell line validation for publication. Obtain cell lines from a reputable source, such as American Type Culture Collection (ATCC). 2. Passage number affects numerous characteristics of cell lines, including morphology, proliferation, responses to stimuli, and genotype. Record cell line passage number, and use low passage number cells (40 reactions with phagolysosomes. For the preparation of early phagosomes (10′/0′, pulse/chase) or late phagosomes (10′/20′, pulse/ chase), we use 20 dishes (10 cm) because the phagosome yield is always lower than for phagolysosomes, due to the shorter pulse periods used for loading of latex beads into early or late phagosomes. 13. To purify early phagosomes, late phagosomes, or phagolysosomes, perform step 5, step 6, or step 7, and proceed with step 8. The time periods that latex beads take to reach early phagosomes, late phagosomes, or phagolysosomes may differ from one cell line to another, and pulse/chase protocols may need to be adjusted. Suitable pulse/chase protocols can be established by immunoblot analysis of phagosomes purified at different times after phagocytosis, as we have done in [15]. This also applies to the pulse/chase protocols used to label different maturation stage endosomes with ferrofluid [12]. 14. Proteins or agents to be tested in the fusion assay should be either dialyzed against or dissolved in HB. Reconstituted fusion is sensitive to PBS, possibly because the SNARE chaperone N-ethylmaleimide sensitive fusion factor (NSF), which is vital for all SNARE-mediated fusion events, is inhibited in phosphate buffers [18]. Moreover, cell-free fusion is sensitive to dimethyl sulfoxide (DMSO), an organic solvent that is often used to dissolve pharmacological inhibitors (our unpublished results): fusion is efficiently inhibited at concentrations above 0.3% (v:v) DMSO. If possible, dissolve inhibitors in different organic solvents. DMF, isopropanol, or ethanol can be applied at higher concentrations than DMSO. 15. Fix phagosome–early endosome fusion samples in HB containing 2.5% (v:v) glutardialdehyde and 2% (v:v) formaldehyde. This way, calcein (not directly fixable) is more efficiently entrapped in LBPs than with fixation with formaldehyde alone. Mount coverslips for phagosome–early endosome fusion in Mowiol containing 2.5% (w/v) Dabco (1,4-Diazabicyclo[2.2.2]octan) as an antifading reagent because calcein fluorescence bleaches quickly. Analyze samples containing early endosomes not later than the day after the experiment because calcein is not fixable and will leak from LBPs. 16. Absolute colocalization between phagosomes and endosome/ lysosome tracers in standard fusion reactions is typically approximately 15% [16]. We disregard experiments in which absolute colocalization is below 3%. Fusion efficiencies can significantly vary depending on the cytosol preparations.

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Acknowledgments This work was supported by a grant from the Deutsche Forschungsgemeinschaft (HA 1929/13-1) to Albert Haas. References 1. Fountain A, Inpanathan S, Alves P et al (2021) Phagosome maturation in macrophages: eat, digest, adapt, and repeat. Adv Biol Regul 82: 100832. https://doi.org/10.1016/j.jbior. 2021.100832 2. Haas A (2007) The phagosome: compartment with a license to kill. Traffic 8:311–330. https://doi.org/10.1111/j.1600-0854.2006. 00531.x 3. Vieira OV, Botelho RJ, Grinstein S (2002) Phagosome maturation: aging gracefully. Biochem J 366:689–704. https://doi.org/10. 1042/BJ20020691 4. Gotthardt D, Warnatz HJ, Henschel O et al (2002) High-resolution dissection of phagosome maturation reveals distinct membrane trafficking phases. Mol Biol Cell 13:3508– 3520. https://doi.org/10.1091/mbc.e0204-0206 5. Fairn GD, Grinstein S (2012) How nascent phagosomes mature to become phagolysosomes. Trends Immunol 33:397–405. https://doi.org/10.1016/j.it.2012.03.003 6. Omotade TO, Roy CR (2019) Manipulation of host cell organelles by intracellular pathogens. Microbiol Spectr 7. https://doi.org/10. 1128/microbiolspec.BAI-0022-2019 7. Guo M, H€artlova A, Dill BD et al (2015) High-resolution quantitative proteome analysis reveals substantial differences between phagosomes of RAW 264.7 and bone marrow derived macrophages. Proteomics 15:3169– 3174. https://doi.org/10.1002/pmic. 201400431 8. Herweg J-A, Hansmeier N, Otto A et al (2015) Purification and proteomics of pathogenmodified vacuoles and membranes. Front Cell Infect Microbiol 5:48. https://doi.org/10. 3389/fcimb.2015.00048 9. Levin-Konigsberg R, Mantegazza AR (2021) A guide to measuring phagosomal dynamics. FEBS J 288:1412–1433. https://doi.org/10. 1111/febs.15506 10. Mehendale N, Mallik R, Kamat SS (2021) Mapping sphingolipid metabolism pathways

during phagosomal maturation. ACS Chem Biol 16:2757–2765. https://doi.org/10. 1021/acschembio.1c00393. PMID: 34647453 11. Lu¨hrmann A, Haas A (2000) A method to purify bacteria-containing phagosomes from infected macrophages. Methods Cell Sci 22: 3 2 9 – 3 4 1 . h t t p s : // d o i . o r g / 1 0 . 1 0 2 3 / a:1017963401560 12. Becken U, Jeschke A, Veltman K et al (2010) Cell-free fusion of bacteria-containing phagosomes with endocytic compartments. Proc Natl Acad Sci U S A 107:20726–20731. https:// doi.org/10.1073/pnas.1007295107 13. Nguyen JA, Yates RM (2021) Better together: current insights into phagosome-lysosome fusion. Front Immunol 12:636078. https:// doi.org/10.3389/fimmu.2021.636078 14. Jeschke A, Haas A (2016) Deciphering the roles of phosphoinositide lipids in phagolysosome biogenesis. Commun Integr Biol 9: e1174798. https://doi.org/10.1080/ 19420889.2016.1174798 15. Jeschke A, Zehethofer N, Lindner B et al (2015) Phosphatidylinositol 4-phosphate and phosphatidylinositol 3-phosphate regulate phagolysosome biogenesis. Proc Natl Acad Sci U S A 112:4636–4641. https://doi.org/10. 1073/pnas.1423456112 16. Jeschke A, Haas A (2018) Sequential actions of phosphatidylinositol phosphates regulate phagosome-lysosome fusion. Mol Biol Cell 29:452–465. https://doi.org/10.1091/mbc. E17-07-0464 17. Fiani ML, Beitz J, Turvy D et al (1998) Regulation of mannose receptor synthesis and turnover in mouse J774 macrophages. J Leukoc Biol 64:85–91. https://doi.org/10.1002/jlb. 64.1.85 18. Block MR, Rothman JE (1992) Purification of N-ethylmaleimide-sensitive fusion protein. Methods Enzymol 219:300–309. https://doi. org/10.1016/0076-6879(92)19030-a

Chapter 18 An In Vitro System to Analyze Generation and Degradation of Phagosomal Phosphatidylinositol Phosphates Andreas Jeschke Abstract Phagosomes are formed when phagocytic cells take up large particles, and they develop into phagolysosomes where the particles are degraded. The transformation of nascent phagosomes into phagolysosomes is a complex multi-step process, and the precise timing of these steps depends at least in part on phosphatidylinositol phosphates (PIPs). Some such-called “intracellular pathogens” are not delivered to microbicidal phagolysosomes and manipulate the PIP composition of the phagosomes they reside in. Studying the dynamic changes of the PIP composition of inert-particle phagosomes will help to understand why the pathogens’ manipulations reprogram phagosome maturation. We here describe a method to detect and to follow generation and degradation of PIPs on purified phagosomes. To this end, phagosomes formed around inert latex beads are purified from J774E macrophages and incubated in vitro with PIP-binding protein domains or PIP-binding antibodies. Binding of such PIP sensors to phagosomes indicates presence of the cognate PIP and is quantified by immunofluorescence microscopy. When phagosomes are incubated with PIP sensors and ATP at a physiological temperature, the generation and degradation of PIPs can be followed, and PIP-metabolizing enzymes can be identified using specific inhibitory agents. Key words Phagocytosis, Phagocytes, Phosphatidylinositol phosphates, Phosphoinositide kinases, Lipid phosphatases, Phagosome, FYVE domain, Phosphoinositide detection, Phosphoinositide metabolism

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Introduction Phagocytosis is the uptake and degradation of large particles (>0.4 μm) by cells and can be subdivided into the processes of phagosome formation, maturation, and resolution [1]. Phagosomes are formed as phagocytes engulf large foreign or endogenous particles such as invading microorganisms or senescent red blood cells and eventually mature into phagolysosomes in which the ingested cargoes are killed and degraded [2]. Phagosome maturation involves acidification of the phagosome lumen and sequential fusion of phagosomes with early endosomes, late endosomes, and

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lysosomes [3]. Phagosome-with-lysosome fusion generates hybrid phagolysosomes from which lysosomes are reformed and acid hydrolases are recycled in a process termed “phagosome resolution” [4, 5]. The various sub-reactions of phagocytosis are hierarchically ordered and critically depend on phosphatidylinositol phosphates (PIPs). Therefore, it is not surprising that PIPs have evolved as the molecular targets of “intracellular pathogens” that evade being delivered to microbicidal phagolysosomes by manipulating the PIP composition of the phagosomes they reside in through secreted PIP-modifying effector proteins [6, 7]. PIPs are phosphorylated derivatives of the glycerophospholipid phosphatidylinositol (PI) with phosphate groups attached to the 3-, 4-, and/or 5-hydroxyl functions of the inositol head of PI. PIPs are either mono-phosphorylated [PI(3)P, PI(4)P, and PI(5)P], bis-phosphorylated [PI(3,4)P2, PI(3,5)P2, and PI(4,5)P2], or tris-phosphorylated [PI(3,4,5)P3] and are interconverted by kinases that attach phosphate groups to PI or a PIP or by phosphatases that dephosphorylate PIPs [8]. Most often, cells possess more than one enzyme for every type of PIP conversion. For example, PI (4)P can be generated by phosphorylation of PI through either of four different phosphatidylinositol 4-kinase (PI4K) isoforms: PI4K2A, PI4K2B, PI4K3A, or PI4K3B [9]. PIP-metabolizing enzymes are asymmetrically distributed within cells so that each subcellular compartment has a different set of PIPs [10]. PIPs regulate cellular processes by recruiting effector proteins to membranes. Most effector proteins bind to PIPs through conserved PIP-binding domains, such as PX (Phox homology), PH (Pleckstrin homology), or FYVE (Fab1, YOTB, Vac1, EEA1) domains [8, 11, 12]. Effector-derived PIP-binding domains that bind to a single PIP isomer have been fused to fluorescent protein tags and been widely used to analyze the PIP composition of subcellular compartments [8], including phagosomes. This approach has provided a detailed picture of the PIP metabolism especially during particle uptake at the plasma membrane and the early stages of phagosome maturation [13]. The analysis of PIP metabolism during late stages of phagosome maturation is difficult because ectopically expressed PIP sensors can inhibit PIP-dependent processes by sequestering PIPs. For example, expression of PI(3)P-binding protein domains inhibits phagolysosome formation [14] so that phagolysosomes cannot be readily analyzed for PI(3)P by expressed PI(3)P sensors. Therefore, it is advisable to analyze in the same samples the localization of both, a PIP sensor and a marker of phagolysosomes [5]. This way, one can directly test whether the lipid probe localizes to phagolysosomes or whether it binds to a pre-phagolysosome compartment that does not develop into a phagolysosome because the PIP to be visualized is sequestered.

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Alternatively, the PIP composition of late stage compartments of the phagocytic pathway has been analyzed with biochemically purified phagosomes [15, 16] by either assessing the binding of lipid probes or by extracting and quantifying PIPs through highperformance liquid chromatography- and/or mass spectrometrybased methods [15, 16]. These approaches have revealed that phagolysosomes contain PI(3)P, PI(4)P, and PI(4,5)P2. In this chapter, we provide a step-by-step protocol to follow the generation and degradation of PIPs on purified phagosomes. In this protocol, latex bead phagosomes (LBP) are purified and incubated with PIP-binding protein domains or antibodies. Binding of such PIP sensors to phagosomes indicates presence of the corresponding PIP and is quantified by immunofluorescence microscopy. This method can be used to determine which PIPs are present in phagosomes of different maturation stages. Moreover, when isolated phagosomes are incubated with ATP at a physiological temperature, generation and degradation of PIPs can be followed, and phagosome-associated PIP-modifying enzymes can be identified by specific (pharmacological) inhibitors (see Fig. 1).

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Materials

2.1 Purification of Glutathione-STransferase (GST) or GST-fused PIP-Binding Proteins from E. coli BL21(DE3)

1. Bacterial expression plasmids encoding GST from Schistosoma japonicum (i.e., pGEX4T1) fused to either two successive copies of mouse hepatocyte growth factor-regulated tyrosine kinase substrate (Hrs) FYVE domain (i.e., pGEX-5X-32xFYVE [17]) or to the Pleckstrin homology (PH) domain of human phospholipase C δ1 (i.e., pGEX4T1-PLCδ1-PH [18]). 2. LB (lysogeny broth)/amp: 5 g/L yeast extract, 10 g/L tryptone, 5 g/L NaCl, 100 μg/mL ampicillin sodium salt. 3. 50× protease inhibitor cocktail (PIC): 12.5 μM leupeptin, 25 mM 1,10-phenantroline, 35 μM pepstatin A, 5 mM pefabloc in deionized H2O. 4. Lysis buffer: 140 mM NaCl, 10 mM Na2HPO4, 1.8 mM KH2PO4, pH 7.5. 5. Glutathione sepharose (see Note 1). 6. 10 mL polypropylene chromatography columns. 7. Elution buffer: pH 7.5 (HCl).

50

mM

Tris,

10

mM

glutathione,

8. Reagent for protein determination (see Note 2). 9. Nitrocellulose dialysis membrane with 3 kDa molecular weight cutoff.

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Fig. 1 Detection of the generation of PIPs on latex bead phagosomes. (a) Workflow of the detection of PIPs on purified phagosomes. LBPs are purified and incubated in the presence of soluble cytosolic proteins, an ATP-regenerating system, and a PIP-binding probe in a suitable buffer at physiological temperature (1). A PIP is formed by action of a PI(P) kinase (2) and is bound by the PIP-binding probe (3). Phagosomes are reisolated from reaction mixtures, spun and fixed onto coverslips, and (4) stained for associated lipid-binding probes using suitable antibodies. Fluorescence microscopy is used to determine the amount of lipid-binding probe on phagosomes. (5) Representative fluorescence micrographs for the detection of PI(3)P on LBPs using the PI(3) P-binding 2xFYVE domain after incubation in the presence or absence of ATP. Bars, 5 μM. (b, c) Quantification of the binding of the 2xFYVE domain to phagosomes that had been incubated at 37 °C in the absence of ATP or in the presence of ATP and 4 mM PI3K inhibitor 3-MA, a Vps34-inhibiting antibody, or 15 μM RabGDI which extracts Rab GTPases from membranes and inhibits Rab GTPase-dependent processes. Data are means + SD from three independent experiments. (d, e) Quantification of the binding of a PI(4)P-binding antibody or a PI (4,5)P2-binding PH domain to phagosomes that had been incubated at 37 °C in the absence of ATP or with ATP

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2.2 Detection of PIPs on Purified LBPs by Immunofluorescence Microscopy

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1. J774E macrophage-like cells (see Note 3). 2. Phosphate-buffered saline (PBS): 137 mM NaCl, 2.7 mM KCl, 3.8 mM Na2HPO4, 1.45 mM KH2PO4, pH 7.4. 3. MES buffer: 50 mM 2-[N-morpholino]ethanesulfonic acid, pH 6.8 (NaOH). 4. EDAC (N-[3-dimethyl-aminopropyl]-N′-ethylcarbodiimide hydrochloride). 5. Tris buffer: 1.5 M Tris, pH 8.8 (HCl). 6. Storage buffer: PBS, 10 mg/mL bovine serum albumin (BSA), 1× PIC, 0.01% NaN3. 7. 1 μM carboxylate-modified latex beads. 8. Internalization medium (INT): Dulbecco’s modified Eagle Medium (DMEM). 9. DMEM/FCS: DMEM, 5% (v:v) fetal calf serum (FCS), 1% (v: v) glutamax, 1% (v:v) penicillin–streptomycin. 10. PBS/5 mM EDTA: PBS, 5 mM EDTA. 11. Homogenization buffer (HB): 250 mM sucrose, 20 mM HEPES, 0.5 mM EGTA, pH 7.2 (KOH). 12. Dounce homogenizer (Dura-Grind Stainless Steel Dounce Tissue Grinder (Wheaton) or similar). 13. 25% sucrose solution: 25% (w:v) sucrose, 20 mM HEPES, 0.5 mM EGTA, pH 7.2 (KOH). 14. 62% sucrose solution: 62% (w:v) sucrose, 20 mM HEPES, 0.5 mM EGTA, pH 7.2 (KOH). 15. Ultracentrifuge with swing rotor and centrifugation tubes suitable for the centrifugation of volumes of ≥12 mL (see Note 4). 16. Cytosol from J774E cells. 17. 10× ATP-regenerating system: 6 mg/mL ATP, 26 mg/mL creatine phosphate, 1 mg creatine kinase, 10 mM MgCl2, pH 7.2 (see Note 5). 18. DTT: HB, 100 mM dithiotreitol (DTT). 19. 10× salts: HB, 1 M KCl, 15 mM MgCl2. 20. IF (immunofluorescence) blocking buffer: PBS, 4% (w:v) BSA. 21. Antibody suitable for the detection of PI(4)P by immunofluorescence microscopy (see Note 6).

ä Fig. 1 (continued) and PI(4)P phosphatase Sac1, PI4K2 inhibitor adenosine, or PI4K2-inhibiting antibody 4C5G. Data are means + SD from three independent experiments. (f) Data summary. Phagolysosomes contain PI(3)P generated by Vps34 and Vps34 activity which is regulated by Rab GTPases, likely Rab7a. Phagolysosomes contain PI(4)P which is generated by PI4K2 and constantly converted to PI(4,5)P2. Converting enzymes are indicated. (Data are taken from Jeschke et al. [15])

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22. Antibody suitable for the detection of GST by immunofluorescence microscopy (see Note 7). 23. Inhibitory antibodies against Vps34 or class II PI4K (see Note 8). 24. 4% (v:v) formaldehyde in HB. 25. 50 mM NH4Cl in HB. 26. Mowiol.

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Methods

3.1 Purification of Glutathione-STransferase (GST) or GST-fused PIP-Binding Proteins from E. coli BL21(DE3), Genotype E. coli B dcm ompT hsdS(rB-mB-) gal

1. Inoculate 25 mL of LB/amp with E. coli BL21(DE3) transformed with pGEX4T1, pGEX-5X-3-2xFYVE, or pGEX4T1PLCδ1-PH, and incubate on a shaker for 16 h at 200 rpm and 37 °C. 2. Use 20 mL of the resulting culture to inoculate 1 L of LB/amp. Incubate culture for 3 h at 200 rpm and 37 °C. 3. Add isopropyl-β-D-galactopyranoside (IPTG) to a final concentration of 0.25 mM, and incubate for 16 h at 200 rpm and 26 °C. 4. All of the following steps are performed at 4 °C unless stated otherwise. 5. Harvest bacteria by centrifugation for 10 min at 6000 g and 4 ° C. 6. Resuspend bacteria in lysis buffer containing 1× PIC. 7. Homogenize bacteria by tip-sonication. 8. Centrifuge homogenate for 30 min at 16100 g and 4 °C. 9. Wash 0.3 mL glutathione sepharose twice with each 1 mL lysis buffer. 10. Mix homogenate with glutathione sepharose (0.3 mL per 10 mL homogenate), and incubate for at least 30 min on an end-over-end shaker. 11. Sediment glutathione sepharose by spinning for 2 min at 1800 g and 4 °C, discard supernatant, and resuspend in 10 mL lysis buffer. 12. Repeat step 11 twice. 13. Transfer glutathione sepharose to a 10 mL chromatography column. Discard flow-through. 14. Elute bound proteins by washing the column once with 0.5 mL and four times with 1 mL elution buffer. Collect elution fractions in separate tubes. Determine protein concentrations in elution fractions, combine fractions with highest

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protein concentration, and dialyze against 2000 volumes of HB for 16 h at 4 °C using a nitrocellulose dialysis membrane with 3 kDa molecular weight cut-off. 15. Determine protein concentration of dialyzed proteins. 16. Aliquot and snap-freeze proteins in liquid nitrogen and store at -80 °C. 3.2 Coating Latex Beads with BSA

1. Sediment 1 μM carboxylate-modified latex beads by spinning for 5 min at 16100 g. Discard supernatant and resuspend in 320 μL MES buffer. 2. Repeat step 1. 3. Discard supernatant, and resuspend beads in MES buffer containing 0.3 mg/mL BSA, and incubate for 15 min on a rocker. 4. Add EDAC to a final concentration of 0.1 mM, and incubate for 60 min on a rocker. 5. Add another 0.1 mM EDAC, and incubate bead suspension for 60 min on a rocker. 6. Add Tris buffer to a final concentration of 10 mM Tris to stop the conjugation reaction. 7. Sediment beads by spinning for 5 min at 16100 g, discard supernatant, and resuspend in 1 mL PBS. 8. Repeat step 7 three times. 9. Discard supernatant, resuspend beads in storage buffer, and store at 4 °C.

3.3 Preparation of Cytosol from J774E Macrophages

1. Sub-cultivate cells in a 1:3 ratio 3 days before the preparation of cytosol. 2. Discard medium. 3. All of the following steps are performed at 4 °C unless stated otherwise. Add 2 mL PBS per dish and detach cells using a rubber cell scraper. 4. Sediment cells by spinning at 160 g for 5 min at 4 °C, discard supernatant, and resuspend cells in 20 mL PBS/EDTA. 5. Sediment cells by spinning at 160 g for 5 min at 4 °C, discard supernatant, and resuspend cells in 20 mL HB. 6. Repeat step 5. 7. Resuspend cells in 2 mL of HB containing 1× PIC, and homogenize cells using a Dounce homogenizer. 8. Centrifuge homogenate for 60 min at 186000 g and 4 °C, collect supernatant, and centrifuge again for 5 min at 16100 g and 4 °C.

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9. Collect supernatant, determine protein concentration, and adjust to 15 mg/mL (protein) by adding HB. 10. Aliquot and snap-freeze cytosol in liquid nitrogen. Store cytosol at -80 °C until use. 3.4 Preparation of Mowiol

1. Dissolve 6 g glycerol and 2.4 g mowiol in 6 mL deionized H2O. 2. Incubate for 60 min at ambient temperature. 3. Add 12 mL 0.2 M Tris, pH 8.8 (HCl). 4. Incubate mixture for 120 min at ambient temperature and for 10 min at 50 °C. 5. Centrifuge mixture for 15 min at 2000 g and 4 °C. 6. Collect supernatants and store at -20 °C.

3.5 Purification of Latex Bead Phagolysosomes from J774E Macrophages

1. Sub-cultivate J774E macrophage-like cells at a 1:3 ratio 2 days before the experiment. Use ten confluent (10 cm diameter) cell culture dishes for phagosome preparation. 2. After 48 h, discard medium, and wash cells once with 5 mL PBS per dish. 3. Add 2.5 mL INT containing 1 μm BSA-coated latex beads per dish (“pulse” of latex beads into the cells). 4. Incubate for 30 min at 37 °C. Wash cells three times with 5 mL PBS (per dish), add 3 mL DMEM/FCS per dish, and incubate for 60 min at 37 °C (“chase” of already ingested beads to their final destination) (see Note 9). 5. From here on, all steps are performed at 4 °C unless stated otherwise. Discard medium and add 2 mL of PBS per dish. 6. Scrape off cells using a rubber cell scraper, and transfer cell suspension into a 50 mL tube. 7. Sediment cells by spinning for 5 min at 160 g and 4 °C. 8. Resuspend cells in 20 mL PBS/EDTA, and centrifuge for 5 min at 160 g and 4 °C. 9. Discard supernatant, resuspend cells in 20 mL HB, and centrifuge for 5 min at 160 g and 4 °C. 10. Discard supernatant, and resuspend cells in 2 mL HB containing 1× PIC. 11. Homogenize cells in a Dounce homogenizer. 12. Centrifuge homogenate for 5 min at 800 g and 4 °C to prepare a postnuclear supernatant (PNS). Collect PNS, resuspend the pellet in 1 mL of HB, and centrifuge for 1 min at 800 g and 4 °C. Combine the resulting supernatants.

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13. Mix 3 mL of the combined supernatants with 3 mL of 62% sucrose solution in an ultracentrifugation tube by gently pipetting up and down. 14. Carefully overlay with 3 mL 25% sucrose solution. 15. Carefully overlay with 3 mL of HB. 16. Centrifuge step gradients for 30 min at 18300 rpm and 4 °C. 17. Harvest phagosomes from the HB/25% sucrose solution interface from the top using a wide-bore pipette tip on a 200 μL micropipette. 3.6 Incubation of Phagosomes with Lipid-Binding Probes

1. The following steps refer to the preparation of a single PIP labeling reaction and are performed on ice unless otherwise stated. 2. Add 8 μL of HB or of HB containing the agent to be tested for inhibition of PIP-metabolizing enzymes to a reaction tube, and add 8.7 μL of purified LBPs. 3. Mix in a separate tube 4 μL of J774E cytosol (adjusted to 15 mg/mL protein), 3 μL of a 10× ATP-regenerating system, 3 μL of 10× salts, and 0.3 μL of 100 mM DTT, and fill to 11.3 μL by addition of HB (see Note 10). 4. Add 11.3 μL of the mixture to each reaction. 5. Add GST, GST-fused PIP sensor, or lipid-binding antibody (see Note 11) to reactions. 6. Fill reactions to a total volume of 30 μL by addition of HB. 7. Incubate reactions at 37 °C for 60 min in the presence or absence of PI(P) kinase (or phosphatase) inhibitors. 8. Set samples on ice, and mix with 30 μL of 62% sucrose solution by carefully pipetting up and down. Overlay with 1 mL of 25% sucrose solution and with 200 μL of HB, and centrifuge resulting density gradients for 30 min at 1800 g and 4 °C in a swingout rotor. 9. Harvest LBPs from the interface between HB and 25% sucrose solution, add 300 μL of HB containing 5 mg/mL BSA, and spin onto glass coverslips in 24-well plates for 15 min at 1800 g and 4 °C. 10. Discard supernatant and add 300 μL of 4% (v:v) formaldehyde in HB. Incubate at 4 °C for 16 h or at ambient temperature for at least 30 min. 11. Discard fixative, and incubate samples for 30 min at ambient temperature in 300 μL HB containing 50 mM NH4Cl to inactivate residual formaldehyde.

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12. Set coverslips with LBPs facing downward into 30 μL drops of IF blocking buffer, and incubate for 30 min at ambient temperature. 13. For the detection of GST or GST-tagged lipid probes, stain samples with an anti-GST antibody (see Notes 7 and 12) and a secondary antibody conjugated with a suitable fluor. For the detection of PI(4)P, reactions are incubated in the presence of an anti-PI(4)P antibody which is detected with a suitable fluorescently labelled secondary antibody diluted in IF blocking buffer. Immunostaining is performed at ambient temperature and by placing each coverslip with LBPs facing downward into 30 μL drops of antibody solution. 14. Incubate coverslips with primary antibodies for 60 min, rinse 5 times in PBS, and incubate with secondary antibodies for 30 min. 15. Rinse coverslips 5 times in PBS, twice in deionized H2O, and mount in 3 μL of Mowiol (see Notes 13 and 14). 3.7 Fluorescence Microscopy Analysis and Quantification of the Amounts of LipidBinding Probes on Phagosomes

To assess the relative amounts of PIPs in phagosome membranes, the fluorescence signals on these compartments, resulting from immunofluorescence labeling of the PIP-binding probes, are quantified. This is done using the image analysis software ImageJ (National Institutes of Health). 1. Export single channel micrographs as tiff-files. Open brightfield image in ImageJ (File>open). 2. Use the “Oval selection tool” and draw selection of the size of a LBP. 3. Move selection tool to mark the outlines of a LBP, and press “t” to add this selection to the “region of interest (ROI) manager.” 4. Left-click (and hold) onto the center of the selection, and move it to mark outlines of another LBP. Press “t.” 5. Repeat step 4 until all LBPs in the microscopic field have been defined as ROIs and added to the ROI manager. Move oval selection tool to an area devoid of phagosomes, and press “t.” This ROI defines the background fluorescence (see Note 15). 6. Open micrograph showing fluorescence signals corresponding to IF-marked LBP-associated lipid probes (File>open), and transform to 8-bit (Image>Type>8-bit). 7. Define parameters to be measured in ROIs. To measure the mean fluorescence in ROIs, choose “mean gray value” in the “Set measurements” menu (Analyze>Set measurements>tick “mean gray value”).

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8. Mark all ROIs in the ROI manager (Analyze>tools>ROI manager), and click “Measure.” This way, a table containing the measured mean fluorescence intensities in ROIs is generated. Values can be exported to a spreadsheet software for further processing. 9. Subtract background fluorescence (see step 5) from the fluorescence intensities measured in ROIs. 10. Determine the mean fluorescence intensities for at least 50 phagosomes.

4

Notes 1. We use Glutathione Sepharose 4B (GE Healthcare). 2. We use Protein Assay Dye Reagent Concentrate (Bio-Rad). 3. High mannose receptor cell line J774E [19] from P. D. Stahl (Washington University in St. Louis, St. Louis, MO) is cultivated at 37 °C in a humid atmosphere of 7% CO2 in DMEM containing 5% (v:v) fetal calf serum (FCS), 1% (v:v) glutamax, and 1% (v:v) penicillin–streptomycin. The here presented protocol can also be used with phagosomes purified from other phagocyte cell lines, such as RAW264.7 (our unpublished results) or THP1. Note that the times that latex beads take to reach early phagosomes, late phagosomes, or phagolysosomes may vary between different cell lines. 4. We use Optima L-80 XP ultracentrifuge with SW40 Ti rotor and Ultra-Clear centrifuge tubes (14 × 95 mm) from Beckman Coulter. 5. Adjust pH to 7 by adding KOH before addition of creatine kinase which is inactivated by exposure to acidic pH. ATP-regenerating system can be stored at -20 °C for several months. 6. We use mouse anti-PI(4)P IgM from Echelon Biosciences. 7. We use rabbit anti-GST Z-5 or mouse anti-GST B-14 from Santa Cruz Biotechnology. 8. We use rabbit anti-Vps34 [20], courtesy of Jonathan M Backer (Molecular Pharmacology, Belfer Institute for Advanced Biomedical Sciences, Albert Einstein College of Medicine, Bronx, NY 10461). This antibody is not commercially available. However, as for the inhibition of Vps34, specific pharmacological inhibitors, e.g., SAR405 [21] and Vps34-IN1 [22], can be used. For the inhibition of class II PI4Ks, we use mouse antiPI4K2 4C5G [23]. This antibody is not commercially available but can be purified from mouse hybridoma cell clone CRL-2538. These hybridoma cells can be purchased from ATCC.

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9. Within the pulse/chase protocol used here, latex beads are delivered to phagolysosomes in J774E macrophages. By varying the pulse/chase periods, latex beads can be used to also label early or late phagosomes. For J774E macrophages, early or late phagosomes can be purified after 10 min/0 min and 10 min/20 min (pulse/chase), respectively [15]. Early and late phagosomes can be tested for PIPs using the same procedures as described here for phagolysosomes. 10. Prepare a master mix sufficient for all planned PIP labeling reactions. Thaw aliquots of the ATP-regenerating system and of J774E cytosol, and prepare master mix just before addition to reactions. To test whether a PIP is generated by a kinase during the incubation, omit ATP from the incubations. Samples incubated on ice can indicate the amount of a PIP present on LBPs at the beginning of the incubation. Moreover, to test whether a PIP-metabolizing enzyme is cytosolic and recruited during the incubation or already phagosome-bound, cytosol can be omitted from the incubations. In fact, omission of cytosol does not reduce generation of PI(3)P on phagolysosomes, suggesting that the corresponding enzyme is already present on and copurified with phagolysosomes (our unpublished results). 11. As lipid sensors, GST-2xFYVE or GST-PLCδ1 PH is added at a final concentration of 2 μM. Similarly, GST (as a negative control) is added at a final concentration of 2 μM. For the detection of PI(4)P, mouse anti-PI(4)P IgM is added at a final concentration of 1:200 (v:v). 12. Dilute anti-GST antibodies specified in Note 7 in IF blocking buffer 1:100 (v:v). 13. The approach presented here relies on the binding of PIP-binding proteins to their cognate lipids and therefore only detects PIPs which are not already effector-complexed on the purified membranes (“free PIPs”). Moreover, the method is limited to the detection of PIPs for which reliable reporters have been developed [8]. 14. As done here, the PIP labeling approach can be used to detect the activities of PIP-metabolizing enzymes on phagosomes. To this end, purified phagosomes are incubated with cytosol, ATP, and lipid-binding probes, and PIPs present on phagosomes after the incubation are detected indirectly by association with lipid sensors. To detect free PIPs that had been generated in phagosome membranes within cells before phagosome purification, homogenates (PNS) prepared from cells after phagocytosis of latex beads are incubated with lipid-binding probes on ice (without

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addition of cytosol or ATP), and phagosomes are purified and assayed for bound lipid probes by immunofluorescence microscopy. This way, we have observed that latex bead phagolysosomes contain PI(3)P [15]. 15. Definition of ROIs can be automatized. To this end, fluorescently labeled latex beads need to be used. Fluorescent labeling should be uniform, i.e., all phagosomes should fluoresce with similar intensities. Open micrograph displaying fluorescence of labeled latex beads (File>open), and convert to 8-bit (Image>Type>8-bit). Transform micrograph into a binary (i.e., black and white) image (Process>Binary>Make Binary), and use “Analyze particles” function to identify LBPs as ROIs (Analyze>Analyze particles>tick “add to Manager”). By pressing “Ok,” ROIs will be identified and added to the “ROI manager.” Proceed with of Subheading 3.7, step 6.

Acknowledgments This work was supported by a grant from the Deutsche Forschungsgemeinschaft (HA 1929/13-1) to Albert Haas, Institute for Cell Biology, University of Bonn, Bonn, Germany. References 1. Fountain A, Inpanathan S, Alves P et al (2021) Phagosome maturation in macrophages: eat, digest, adapt, and repeat. Adv Biol Regul 82: 100832. https://doi.org/10.1016/j.jbior. 2021.100832 2. Haas A (2007) The phagosome: compartment with a license to kill. Traffic 8:311–330. https://doi.org/10.1111/j.1600-0854.2006. 00531.x 3. Desjardins M, Huber LA, Parton RG et al (1994) Biogenesis of phagolysosomes proceeds through a sequential series of interactions with the endocytic apparatus. J Cell Biol 124:677– 688. https://doi.org/10.1083/jcb.124.5.677 4. Lancaster CE, Fountain A, Dayam RM et al (2021) Phagosome resolution regenerates lysosomes and maintains the degradative capacity in phagocytes. J Cell Biol 220. https://doi. org/10.1083/jcb.202005072 ˜ o-Rendo´n F, 5. Levin-Konigsberg R, Montan Keren-Kaplan T et al (2019) Phagolysosome resolution requires contacts with the endoplasmic reticulum and phosphatidylinositol-4phosphate signalling. Nat Cell Biol 21:1234– 1247. https://doi.org/10.1038/s41556019-0394-2

6. Pizarro-Cerda´ J, Cossart P (2004) Subversion of phosphoinositide metabolism by intracellular bacterial pathogens. Nat Cell Biol 6:1026– 1 0 3 3 . h t t p s : // d o i . o r g / 1 0 . 1 0 3 8 / ncb1104-1026 7. Weber SS, Ragaz C, Hilbi H (2009) Pathogen trafficking pathways and host phosphoinositide metabolism. Mol Microbiol 71:1341–1352. https://doi.org/10.1111/j.1365-2958.2009. 06608.x 8. Posor Y, Jang W, Haucke V (2022) Phosphoinositides as membrane organizers. Nat Rev Mol Cell Biol:1–20. https://doi.org/10. 1038/s41580-022-00490-x 9. Balla A, Balla T (2006) Phosphatidylinositol 4-kinases: old enzymes with emerging functions. Trends Cell Biol 16:351–361. https:// doi.org/10.1016/j.tcb.2006.05.003 10. de Matteis MA, Godi A (2004) PI-loting membrane traffic. Nat Cell Biol 6:487–492. https:// doi.org/10.1038/ncb0604-487 11. Rusten TE, Stenmark H (2006) Analyzing phosphoinositides and their interacting proteins. Nat Methods 3:251–258. https://doi. org/10.1038/nmeth867

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12. Kutateladze TG (2010) Translation of the phosphoinositide code by PI effectors. Nat Chem Biol 6:507–513. https://doi.org/10. 1038/nchembio.390 13. Levin R, Grinstein S, Schlam D (2015) Phosphoinositides in phagocytosis and macropinocytosis. Biochim Biophys Acta 1851:805–823. https://doi.org/10.1016/j.bbalip.2014. 09.005 14. Vieira OV, Harrison RE, Scott CC et al (2004) Acquisition of Hrs, an essential component of phagosomal maturation, is impaired by mycobacteria. Mol Cell Biol 24:4593–4604. https://doi.org/10.1128/MCB.24.10. 4593-4604.2004 15. Jeschke A, Zehethofer N, Lindner B et al (2015) Phosphatidylinositol 4-phosphate and phosphatidylinositol 3-phosphate regulate phagolysosome biogenesis. Proc Natl Acad Sci U S A 112:4636–4641. https://doi.org/10. 1073/pnas.1423456112 16. Defacque H, Bos E, Garvalov B et al (2002) Phosphoinositides regulate membranedependent actin assembly by latex bead phagosomes. Mol Biol Cell 13:1190–1202. https:// doi.org/10.1091/mbc.01-06-0314 17. Gillooly DJ, Morrow IC, Lindsay M et al (2000) Localization of phosphatidylinositol 3-phosphate in yeast and mammalian cells. EMBO J 19:4577–4588. https://doi.org/10. 1093/emboj/19.17.4577 18. Dowler S, Currie RA, Campbell DG et al (2000) Identification of pleckstrin-homology-

domain-containing proteins with novel phosphoinositide-binding specificities. Biochem J 351:19–31. https://doi.org/10. 1042/0264-6021:3510019 19. Fiani ML, Beitz J, Turvy D et al (1998) Regulation of mannose receptor synthesis and turnover in mouse J774 macrophages. J Leukoc Biol 64:85–91. https://doi.org/10.1002/jlb. 64.1.85 20. Siddhanta U, McIlroy J, Shah A et al (1998) Distinct roles for the p110alpha and hVPS34 phosphatidylinositol 3′-kinases in vesicular trafficking, regulation of the actin cytoskeleton, and mitogenesis. J Cell Biol 143:1647–1659. https://doi.org/10.1083/jcb.143.6.1647 21. Ronan B, Flamand O, Vescovi L et al (2014) A highly potent and selective Vps34 inhibitor alters vesicle trafficking and autophagy. Nat Chem Biol 10:1013–1019. https://doi.org/ 10.1038/nchembio.1681 22. Bago R, Malik N, Munson MJ et al (2014) Characterization of VPS34-IN1, a selective inhibitor of Vps34, reveals that the phosphatidylinositol 3-phosphate-binding SGK3 protein kinase is a downstream target of class III phosphoinositide 3-kinase. Biochem J 463:413– 427. https://doi.org/10.1042/BJ20140889 23. Endemann GC, Graziani A, Cantley LC (1991) A monoclonal antibody distinguishes two types of phosphatidylinositol 4-kinase. Biochem J 273(Pt 1):63–66. https://doi.org/ 10.1042/bj2730063

Chapter 19 Dissecting Phagosomal Pattern Recognition Receptor-Dependent Signaling and Antigen MHC-II Presentation from Phagosomes in Murine Dendritic Cells Emilia Scharrig and Adriana R. Mantegazza Abstract Phagosomal pattern recognition receptor signaling promotes phagosome maturation and additional immune pathways such as proinflammatory cytokine secretion and antigen MHC-II presentation in antigen-presenting cells. In the present chapter, we describe procedures to assess these pathways in murine dendritic cells, professional phagocytes positioned at the interface between innate and adaptive immune responses. The assays described herein follow proinflammatory signaling by biochemical and immunological assays as well as antigen presentation of the model antigen Eα by immunofluorescence followed by flow cytometry. Key words Antigen presentation, Dendritic cells, Eα, Phagocytosis, MHC-II

1

Introduction Dendritic cells (DCs) are both professional phagocytes and antigen (Ag)-presenting cells that sense and signal extracellular cues via pattern-recognition receptors (PRRs) and integrate those signals while processing phagocytosed material, bridging innate to adaptive immunity [1]. PRR triggering (such as Toll-like receptor— TLR—stimulation) on DC phagosomes formed upon particle capture stimulates phagosome maturation to the production of proinflammatory cytokines, the degradation of phagosomal content, and Ag processing and presentation to T cells [2, 3]. The main downstream signaling pathways that promote phagosome maturation consist of a phosphorylation cascade that includes TLR-associated kinases such as interleukin 1 receptor-associated kinases 1/4 (IRAK1/4) and members of the mitogen-activated protein kinase (MAPK) family such as p38 and ultimately lead to the activation of transcription factors NF-κB and AP-1, favoring proinflammatory responses [4]. A later inflammatory response leads to the activation

Roberto J. Botelho (ed.), Phagocytosis and Phagosomes: Methods and Protocols, Methods in Molecular Biology, vol. 2692, https://doi.org/10.1007/978-1-0716-3338-0_19, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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of interferon regulatory factors (IRFs), promoting type-I interferon production [5]. PRR stimulation also optimizes the presentation of exogenous Ags on Major Histocompatibility Complex Class II (MHC-II) molecules [6, 7]. These consist of heterodimers of non-covalently coupled α and β chains initially associated with an invariant chain (Ii), which occludes the peptide binding site and prevents premature peptide binding [8]. Phagosomal PRR signaling also favors MHC-II transport to the endosomal system, which interacts with phagosomes along their maturation process [9, 10]. MHC-II molecules are then loaded with peptides generated during phagosomal content degradation and transported to the cell surface, leading to the presentation of phagocytosed antigen [6, 11]. While phagocytosed cargo may also be presented on MHC-I molecules in the process of Ag cross-presentation in DCs [7, 12], the present chapter will focus on the presentation of phagocytosed Ag on MHC-II molecules, using the model antigen Eα [13, 14]. Mouse MHC molecules are known as histocompatibility 2 (H2) molecules, and their nomenclature varies depending on the mouse strain. In the case of our model mouse strain C57BL/6J, the designation of MHC-II molecules H2-I-A is shortened to I-A followed by a subscript indicating the haplotype [15]. The formation of the Eα52–68 peptide:I-Ab complex is detected using the YAe antibody, either on the cell surface or intracellularly [16, 17]. In the following sections, we describe procedures to (i) assess PRR-dependent proinflammatory signaling by immunoblotting and the quantification of cytokine production after phagocytosis and (ii) evaluate MHC-II:peptide loading and presentation of phagocytosed Eα Ag by flow cytometry (see Fig. 1) in bone marrow-derived DCs. These procedures may also be applied to tissue-resident DCs [18, 19].

2 2.1

Materials Equipment

1. Tissue culture incubator (37 °C, 5% CO2). 2. Tissue culture centrifuge with plate adapters. 3. Microcentrifuge. 4. Biosafety cabinet. 5. Water bath (37 °C). 6. Acrylamide gel electrophoresis system. 7. Flow cytometer. 8. Plate reader.

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Fig. 1 Representation of phagosomal signaling and antigen presentation from DC phagosomes. After Eα-YFP/ LPS or PRR ligand-coated bead phagocytosis, phagosome maturation promotes Ag processing and proinflammatory signaling. In the signaling pathway, PRR triggering induces a phosphorylation cascade that includes p38 and p65 (1) and leads to proinflammatory cytokine production (2). In the Ag presentation pathway, the processed Eα52–68 peptide is loaded on MHC-II molecules in the maturing phagosome (3) and presented on the cell surface (4)

2.2

Mice

2.3 Reagents and Supplies 2.3.1

Cell Culture

1. C57BL/6J mice (see Note 1). 1. Fetal bovine serum (FBS; very low endotoxin for BMDC culture). 2. DC media: RPMI 1640 cell supplemented with 10% FBS, 50 μM 2-mercaptoethanol, 2 mM glutamine, 100 U/mL penicillin, 100 μg/mL streptomycin. 3. DC media containing 30% granulocyte-macrophage colonystimulating factor (GM-CSF) conditioned media from J558 plasmacytoma [17] or 20 ng/mL commercially available mouse recombinant GM-CSF [20] (see Note 2). 4. Non-tissue culture-treated 10 cm dishes for BMDC culture. 5. 96-well round-bottom plates.

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6. 3.5 cm tissue culture-treated dishes (p35). 7. 6 cm tissue culture-treated dishes (p60). 2.3.2

Assays

Bead Preparation and Phagocytosis

1. 3 μM polystyrene beads or polystyrene amino beads (see Note 3). 2. 8% glutaraldehyde. 3. CO2-independent medium or RPMI media with 25 mM HEPES. 4. Phosphate-buffered saline (PBS), pH 7.4.

PRR Signaling

1. PRR ligands such as bacterial lipopolysaccharide (LPS). 2. ELISA immunoplates. 3. IL-6/IL-12/TNF-α ELISA kits. 4. Antibodies: anti-mouse p38 MAPK, anti-mouse phospo-p38 MAPK (Thr180/Tyr182), anti-mouse p65, anti-mouse phospho-p65 (Ser 536). 5. Lysis buffer: 10 mM Na pyrophosphate, 10 mM Na-β-glycerophosphate, 40 mM HEPES, 4 mM EDTA, 1% Triton X-100, protease and phosphatase inhibitors cocktail. Adjust pH to 7.4 before adding the detergent (see Note 4). 6. Laemmli sample buffer.

Eα Ag Presentation

1. Eα–YFP (Gift from Drs Marion Pepper, University of Washington and Mark Jenkins, University of Minnesota). 2. Eα (52–68) peptide (AbßEp). 3. Bovine serum albumin (BSA). 4. Flow cytometry buffer: 0.5% BSA, 2 mM EDTA in PBS. 5. 2% paraformaldehyde (PFA) solution. 6. Perm-wash buffer (PBS 0.1% saponin, 0.5% BSA). 7. Antibodies: biotinylated anti-mouse Eα52–68 peptide bound to I-Ab, anti-mouse Eα52–68 peptide bound to I-Ab, anti-mouse CD11c APC, anti-mouse CD11c PE, streptavidin APC, antimouse CD11b APC, anti-mouse MHC-II APC.

3

Methods

3.1 Cell Isolation or Differentiation 3.2

Signaling

3.2.1 Preparation of LPSCoated Beads

Perform BM isolation and differentiation to DCs as indicated in Chapter 20. 1. The day before the experiment, take the required amount of polystyrene beads (according to manufacturer’s instructions) for a 10:1 ratio (beads:DCs) (see Notes 5 and 6).

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2. Wash by adding 1 mL of sterile PBS. 3. Centrifuge at 14,000 g for 5 min. 4. Repeats step 2 and 3. 5. Remove PBS and resuspend with LPS to a final concentration of 100 μg/mL in PBS. 6. Rotate overnight at 4 °C. 7. Centrifuge at 14,000 g for 5 min. 8. Save supernatant and freeze it (see Note 7). Resuspend beads in 1 mL PBS. 9. Centrifuge at 14,000 g for 5 min. 10. Add 1 mL of PBS and centrifuge at 14,000 g for 5 min. 11. Resuspend beads in initial bead volume. 3.2.2

DC Harvesting

1. To collect BMDCs from 10 cm dishes, take cells in suspension, and transfer them to a conical tube on ice. 2. Wash the remaining adherent cells with 5–10 mL PBS to remove the remaining media. Transfer the cell wash to the tube containing the cell suspension. 3. Add 10 mL of cold PBS to the dish, and incubate for 20–30 min on ice to allow the cells to detach. 4. Collect the cells by pipetting several times using a P1000 micropipette to detach the adherent cells (see Note 8), and pool with the initial cell suspension.

3.2.3

DC Stimulation

1. 12–18 × 106 immature BMDCs are required for a kinetic experiment of 6 time points (e.g., 0, 15, 30, 60, 120, 180 min). Seed 2 × 106 in 2 mL DC media + GM-CSF/ p35 plate. If a higher cell number is desired, seed 3–5 × 106/3– 5 mL DC media + GM-CSF/p60 plate. Plate the cells the day before the experiment (see Note 9). 2. Remove the suspension cells, and wash the plate once with DC media to remove loosely attached cells. 3. Place the plates over ice for 1 min, remove the media, and add media containing ligand-coated beads or bacteria. 4. Incubate the plates over ice for 15 min to allow the beads to settle down. 5. Transfer the plates to the incubator at 37 °C for 15 min (“pulse”). 6. Wash twice with warm DC media to remove the remaining suspension beads.

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7. Visualize the cells under the microscope to corroborate the efficiency of the washes. Wash again if unbound beads are present. 8. Add 2 mL of warm media per p35 or 3–5 mL per p60 plate. 9. Transfer the plates to the incubator and “chase” for the desired time points. 10. At each time point, collect and save the supernatant at -20 °C to measure cytokines by ELISA, according to manufacturer’s recommendations. (See Note 10). 11. Harvest the attached cells to assess protein phosphorylation by immunoblotting by adding 100 μL of lysis buffer per 1 × 106 cells. Lift cells using a cell lifter, transfer to a microcentrifuge tube, add the required volume of Laemmli sample buffer [21], and store until use at -20 °C. 3.3 Eα Antigen Presentation Assay 3.3.1 Eα-YFP-Coated Bead Preparation

This procedure has been adapted from ref. 22. 1. The day before the experiment, take the required volume of polystyrene amino beads for a 15:1 ratio of beads to cells (see Note 6). 2. Wash by adding 1 mL of sterile PBS. 3. Centrifuge at 14,000 g for 5 min and discard the supernatant. 4. Resuspend the pellet in 0.5 mL 8% glutaraldehyde per 1.7 × 108 beads. 5. Rotate for 4–6 h at room temperature (RT). 6. Centrifuge at 14,000 g for 5 min and discard the supernatant. 7. Wash by adding 1 mL of sterile PBS. 8. Centrifuge at 14,000 g for 5 min and discard the supernatant. 9. Resuspend the beads in PBS with Eα-YFP to a final concentration of 5 mg/mL in 1 mL per 1.7 × 108 beads. 10. Rotate overnight at 4 °C, protected from light. 11. Centrifuge the beads at 14,000 g for 5 min. Save the supernatant and store it at -20 °C (see Note 7). 12. Wash by adding 1 mL of sterile PBS. 13. Centrifuge at 14,000 g for 5 min and discard the supernatant. 14. Repeat steps 12 and 13 once. 15. Resuspend in PBS in the original volume of beads.

3.3.2 Eα-YFP-Coated Bead Phagocytosis, Antigen Loading, and Presentation

After Eα-YFP-coated bead phagocytosis by DCs, Eα-YFP is degraded during phagosome maturation, the resulting Ags are loaded on MHC-II molecules and peptide: MHC-II complexes are finally presented on the cell surface. The following procedure allows the detection of intracellular and cell surface Eα52–68

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peptide:I-Ab complexes using the YAe antibody. In parallel incubation of DCs with soluble Eα52–68 peptide allows to control for MHC-II surface expression and serves as a positive control for Eα52–68 peptide:I-Ab detection. A phagocytosis control is also required to ensure that all cell types tested have a comparable ability to capture the beads. 1. A total of 12 × 106 DCs are required for a kinetic experiment over 18 h (four times points). Harvest BMDCs on the seventh day of culture on ice to prevent activation, as in Subheading 3.2.2 (see Note 9). 2. Centrifuge at 200 g for 7–10 min. Remove supernatant, and resuspend cells at a final concentration of 1 × 106 cells/mL in DC media. 3. Separate 3 × 106 total cells in DC media for six controls without beads, and leave them on ice until performing the staining. Controls: (1) Surface peptide binding to MHC-II molecules; (2) CD11c-PE staining; (3) CD11c-APC staining; (4) Unstained cells; (5 and 6) BMDC phenotype (see Note 11) (see Fig. 2). 4. Prepare 7 × 106 cells to pulse with beads in 700 μL of CO2independent media. From these, two controls will be performed: (7) phagocytosis control and (8) Avidin binding control (see Note 11) (see Fig. 2).

Fig. 2 Example of experimental layout for the Eα Ag presentation assay. Controls 1–8 are described in Note 11. Controls without beads: (1) Surface peptide binding to MHC-II molecules, (2) CD11c-PE staining, (3) CD11c-APC staining, (4) Unstained cells, (5, 6) BMDC phenotype. Controls with beads: (7) Phagocytosis control and (8) Avidin binding control

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5. Place the cells in CO2-independent media in a water bath at 37 °C for 2 min, and add the required number of beads. Resuspend and incubate for 20 min (“pulse”) (see Note 12). 6. Stop bead uptake with 10 mL of ice-cold PBS. 7. Centrifuge at 200 g for 5 min. 8. Resuspend in 7 mL of DC media to have 1 × 105 cells / 100 μL. Separate 500,000 cells for control 1 (phagocytosis control) and 500,000 cells for control 2 (avidin labeling control) (see Note 11). 9. Seed cells in triplicates in a round-bottom 96-well plate to perform the “chase” and subsequent surface and intracellular staining (see Note 13) (see Fig. 2). 10. Place the plate in the incubator (37 °C, 5% CO2), and incubate for the desired time points (“chase”). Times 0, 3, 6, and 18 h are recommended for initial screening (use one plate per time point). 11. For the surface peptide binding control (see Note 11), peptide serial dilutions should be performed. Recommended peptide concentrations, 0.1–10 μM. 12. After the chase, place the plate on ice to stop the Ag processing/presentation process. 3.3.3 Immunofluorescence

1. At the end of each chase time, centrifuge the plate at 300 g for 5 min, and remove the supernatant. 2. Incubate the cells with biotinylated anti-Eα (biotinylated YAe; dil. 1/50) for the non-permeabilized conditions and surface peptide binding control (control 1, see Note 11) or unconjugated anti-Eα (YAe; dil 1/50) for the permeabilized conditions (to block the Eα Ag presented at the surface), in flow cytometry buffer (see Fig. 3). 3. Add corresponding antibodies to control cells (controls 2, 3, 5, 6, 8). Keep unstained cells on ice (controls 4 and 7). See Note 11. 4. Incubate for 1 h at 4 °C. 5. Centrifuge at 300 g for 5 min. 6. Remove supernatant and wash with flow cytometry buffer. 7. Centrifuge at 300 g for 5 min. 8. Repeat steps 6 and 7 once. For Eα52–68 peptide: I-Ab cell surface staining (non-permeabilized conditions), continue with steps 22–26. For intracellular Eα52–68 peptide: I-Ab staining (permeabilized conditions), continue with steps 9–21. For the control cells, proceed to step 26. 9. Fix on ice with PFA 2% for 15 min.

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Fig. 3 Representation of the immunofluorescence steps to be performed on the 96-well plate after pulse and chase. The plate is divided in two, to represent the non-permeabilized (left side) and permeabilized (right side) conditions. In the non-permeabilized condition, MHC-II:Eα52–68 peptide complexes are detected on the cell surface by YAe-biotin. The permeabilized condition—after surface MHCII:Eα52–68 peptide blocking with unlabeled YAe and fixation—detects intracellular MHC-II:Eα52–68 peptide complexes using YAe-biotin in permeabilization buffer. YAe-biotin is finally detected by avidin in both conditions. DCs are labeled with anti-CD11c antibody. Note that while permeabilized cells are incubated with YAe-biotin, non-permeabilized cells remain in flow cytometry buffer

10. Wash with perm-wash buffer. 11. Centrifuge at 300 g for 5 min. 12. Remove supernatant and incubate with biotinylated-YAe in perm-wash buffer for 1 h at RT. 13. Wash with perm-wash buffer. 14. Centrifuge at 300 g for 5 min. 15. Repeat steps 13 and 14 once. 16. Incubate with avidin-APC (dil 1/200) and anti-CD11c (dil 1/200) in perm-wash buffer for 45 min–1 h at RT. 17. Wash with perm-wash buffer. 18. Centrifuge at 300 g for 5 min. 19. Wash with flow cytometry buffer. 20. Centrifuge at 300 g for 5 min. 21. Resuspend in 300 μL of flow cytometry buffer. Cells are ready to process by flow cytometry. 22. Incubate with avidin-APC (dil 1/200) and anti-CD11c (dil 1/200) in flow cytometry buffer for 45 min–1 h at 4 °C. 23. Wash with flow cytometry buffer.

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24. Centrifuge at 300 g for 5 min. 25. Optional: Fix with PFA 2% for 15 min on ice, wash and centrifuge as in steps 19 and 20. 26. Resuspend in 300 μL of flow cytometry buffer. Cells are ready to process by flow cytometry. 3.3.4

4

Flow Cytometry

After performing compensation with the corresponding controls, run samples and controls. For data visualization using FlowJo or similar software, gate on the cell population CD11chi (DCs)/ Eα52–68 peptide: I-Ab (YAe)+. Determine percentage of CD11chi/ YAe+ DCs over time, compared to time 0 of the chase, as shown in ref. 23.

Notes 1. This strain is commercially available, e.g., from The Jackson Laboratories (strain #000664). All mouse studies should be performed in compliance with veterinary regulations on the humane care and use of laboratory animals. 2. The preparation of GM-CSF conditioned media from J559 plasmacytoma allows for the availability of large amounts of GM-CSF at relatively lower costs than recombinant GM-CSF (rGM-CSF). However, the use of rGM-CSF is less timeconsuming and might be preferred for small-scale experiments. 3. For reference: Polybeads, Polysciences Inc. (cat#17134-15), or Polybeads amino, Polysciences Inc. (cat#17145-5). 4. Lysis buffer can be stored at -20 °C without Triton X-100 and protease/phosphatase inhibitors. Prepare 90 mL buffer at a 1.11× concentration. Freeze in 9 mL aliquots. On the day of the experiment, add 1 mL Triton X-100 10% to the 9 mL solution (final concentration 1%) and protease/phosphatase inhibitors for a 1× solution. 5. Alternative PRR stimuli should be considered, such as ligands for TLR2 (e.g., peptidoglycan), TLR5 (e.g., flagellin), or TLR3 (e.g., polyI:C). Recommended concentration for beadcoating is 100 μg/mL. Additionally, bacteria such as E. coli, S. aureus, or Salmonella may be chosen to probe anti-bacterial responses testing different multiplicities of infection (MOI). For BMDCs, we found that a 5:1 bacteria:DC ratio is sufficient to trigger proinflammatory responses in a timely manner. Increased MOI (10:1) is required for splenic DCs. 6. The ratio between beads and cells should be titrated depending on the cell population, particularly if working with knock-out, knock-down, or chemically treated DCs, in which phagocytic capacity may be reduced.

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7. The supernatant may be reused once, as LPS or Eα-YFP is in significant excess to ensure sufficient binding to beads. 8. Collect cells carefully—bubbles may cause cell activation. 9. If performing the experiment on tissue-resident DCs, cells are used on the same day of isolation and seeded at a concentration of 1 × 106 cells/mL in DC media. 10. Once the supernatants are collected and centrifuged to eliminate possible remaining cells, they can be stored at -20 °C until the ELISA is performed. 11. Summary of controls: Without beads 1. Surface peptide binding to MHC-II molecules: cells + Eα peptide—to control for MHC-II surface expression; it is expected to be similar between tested cell types if MHC-II surface levels are comparable. 2. CD11c-PE staining—flow cytometer compensation control. 3. CD11c-APC staining—flow cytometer compensation control. 4. Unstained cells—flow cytometer compensation control and for cell size gating 5. BMDC phenotype: CD11c + CD11b—cell phenotype control; BMDCs are CD11chi/CD11b+. 6. BMDC phenotype: MHC-II + CD11c—cell phenotype control; control for MHC-II surface expression. With beads 7. Phagocytosis control: DCs + beads, no staining—to evaluate the percent of cells that phagocytose beads. 8. Avidin binding control: Avidin-APC + CD11c-PE—immunofluorescence negative control for Eα52–68 peptide: I-Ab staining. 12. Pulse time may be varied depending on the cell type and whether DCs have been knocked out, knocked down, or chemically treated, which can affect their phagocytic activity. For initial screening, we recommend trying pulses of 20 min to 1 h. For pulses longer than 20 min, add complete DC media, and place cells in the CO2 incubator to continue the pulse. 13. The surface staining (non-permeabilized condition) will indicate the amount of MHC-II:Eα52–68 peptide presented on the cell surface. After blocking surface MHC-II:Eα52–68 peptide complexes, the intracellular staining (permeabilized condition) will allow the detection of intracellular MHC-II:Eα52–68 peptide complexes.

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Acknowledgments This work was supported by NIH grant R01 AI137173. The summary figure was created using Biorender.com through Thomas Jefferson University library portal. References 1. Steinman RM, Hemmi H (2006) Dendritic cells: translating innate to adaptive immunity. Curr Top Microbiol Immunol 311:17–58 2. Janeway CA Jr, Medzhitov R (2002) Innate immune recognition. Annu Rev Immunol 20: 197–216. https://doi.org/10.1146/annurev. immunol.20.083001.084359 3. Kagan JC, Iwasaki A (2012) Phagosome as the organelle linking innate and adaptive immunity. Traffic 13(8):1053–1061. https://doi. org/10.1111/j.1600-0854.2012.01377.x 4. Kawai T, Akira S (2010) The role of patternrecognition receptors in innate immunity: update on Toll-like receptors. Nat Immunol 11(5):373–384 5. Kumar H, Kawai T, Akira S (2011) Pathogen recognition by the innate immune system. Int Rev Immunol 30(1):16–34 6. Villadangos JA, Schnorrer P, Wilson NS (2005) Control of MHC class II antigen presentation in dendritic cells: a balance between creative and destructive forces. Immunol Rev 207:191–205. https://doi.org/10.1111/j. 0105-2896.2005.00317.x 7. Mantegazza AR, Magalhaes JG, Amigorena S, Marks MS (2013) Presentation of phagocytosed antigens by MHC class I and II. Traffic 14(2):135–152. https://doi.org/10.1111/ tra.12026 8. Cresswell P (1996) Invariant chain structure and MHC class II function. Cell 84(4): 505–507. https://doi.org/10.1016/s00928674(00)81025-9 9. Niedergang F, Grinstein S (2018) How to build a phagosome: new concepts for an old process. Curr Opin Cell Biol 50:57–63. https://doi.org/10.1016/j.ceb.2018.01.009 10. Savina A, Amigorena S (2007) Phagocytosis and antigen presentation in dendritic cells. Immunol Rev 219:143–156 11. Blander JM, Medzhitov R (2006) On regulation of phagosome maturation and antigen presentation. Nat Immunol 7(10): 1029–1035. https://doi.org/10.1038/ ni1006-1029 12. Guermonprez P, Saveanu L, Kleijmeer M, Davoust J, Van Endert P, Amigorena S (2003)

ER-phagosome fusion defines an MHC class I cross-presentation compartment in dendritic cells. Nature 425(6956):397–402. https:// doi.org/10.1038/nature01911 13. Rudensky A, Rath S, Preston-Hurlburt P, Murphy DB, Janeway CA Jr (1991) On the complexity of self. Nature 353(6345):660–662. https://doi.org/10.1038/353660a0 14. Rudensky Y, Preston-Hurlburt P, Hong SC, Barlow A, Janeway CA Jr (1991) Sequence analysis of peptides bound to MHC class II molecules. Nature 353(6345):622–627. https://doi.org/10.1038/353622a0 15. Bishop GA (1998) H2 class II. Encyclopedia of immunology, 2nd edn, pp 1040–1045. https://doi.org/10.1006/rwei.1999.0270 16. Lee HK, Mattei LM, Steinberg BE, Alberts P, Lee YH, Chervonsky A, Mizushima N, Grinstein S, Iwasaki A (2010) In vivo requirement for Atg5 in antigen presentation by dendritic cells. Immunity 32(2):227–239 17. Mantegazza AR, Guttentag SH, El-Benna J, Sasai M, Iwasaki A, Shen H, Laufer TM, Marks MS (2012) Adaptor protein-3 in dendritic cells facilitates phagosomal toll-like receptor signaling and antigen presentation to CD4(+) T cells. Immunity 36(5):782–794 18. Savina A, Peres A, Cebrian I, Carmo N, Moita C, Hacohen N, Moita LF, Amigorena S (2009) The small GTPase Rac2 controls phagosomal alkalinization and antigen crosspresentation selectively in CD8(+) dendritic cells. Immunity 30(4):544–555 19. Geem D, Medina-Contreras O, Kim W, Huang CS, Denning TL (2012) Isolation and characterization of dendritic cells and macrophages from the mouse intestine. J Vis Exp 63: e4040. https://doi.org/10.3791/4040 20. Winzler C, Rovere P, Rescigno M, Granucci F, Penna G, Adorini L, Zimmermann VS, Davoust J, Ricciardi-Castagnoli P (1997) Maturation stages of mouse dendritic cells in growth factor-dependent long-term cultures. J Exp Med 185(2):317–328 21. Laemmli UK (1970) Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227(5259):

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Chapter 20 Examining the Kinetics of Phagocytosis-Coupled Inflammasome Activation in Murine Bone Marrow-Derived Dendritic Cells Daniel J. Netting and Adriana R. Mantegazza Abstract In the present chapter, we describe procedures to assess NLRP3 and NLRC4 inflammasome assembly by immunofluorescence microscopy or live cell imaging, together with inflammasome activation by biochemical and immunological techniques upon phagocytosis. We also include a step-by-step guide to automating the counting of inflammasome “specks” after imaging. While our focus resides on murine bone marrowderived dendritic cells differentiated in the presence of granulocyte-macrophage colony-stimulating factor, which results in a cell population that resembles inflammatory dendritic cells, the strategies described herein may apply to other phagocytes as well. Key words ASC speck, Dendritic cells, Imaging, Inflammasome, NLRC4, NLRP3, Pattern recognition receptors, Phagosome, Pyroptosis

1

Introduction Dendritic cells (DCs) are professional phagocytes that continuously sample their microenvironment and respond to extracellular cues such as pathogen- or damage-associated molecular patterns (PAMPs or DAMPs) via pattern recognition receptors (PRRs) [1, 2]. DCs engulf bacteria and particles in the process of phagocytosis, resulting in particle enclosure into de novo formed phagosomes [3]. While phagosomes initially bear plasma membrane components such as PRRs, they subsequently mature through fusion and fission events with the endolysosomal system and other organelles [4, 5], thereby acquiring cargo and membrane components, such as endosomal PRRs [6, 7]. Engagement of PRRs or similar receptors triggers immune responses according to the encountered threat [8]. Some phagocytosed cargo, such as commensal or non-pathogenic bacteria, elicits mild to moderate types of immune responses, mainly characterized by the secretion of

Roberto J. Botelho (ed.), Phagocytosis and Phagosomes: Methods and Protocols, Methods in Molecular Biology, vol. 2692, https://doi.org/10.1007/978-1-0716-3338-0_20, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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pro-inflammatory cytokines. In contrast, pathogenic bacteria, crystals, or alum present in vaccine formulations may cause damage to the phagosomal membrane, resulting in leakage of phagosomal contents into the cytosol [8–10]. Breaching of phagosomal membrane integrity sets off an alarm for the activation of a “high-threat” type of immune response: the assembly of the inflammasome. Inflammasomes are cytosolic multiprotein complexes formed in response to PAMPs, DAMPs, or membrane damage. Inflammasomes are composed of a cytosolic PRR serving as a sensor protein, pro-caspase-1, and typically contain the adaptor protein apoptosisassociated speck-like protein containing a caspase activation and recruitment domain (CARD) [ASC], which bridges the sensor to pro-caspase-1 [9, 11, 12]. Pro-caspase-1 undergoes self-cleavage, achieved after inflammasome assembly, becoming an active enzyme. Upon activation, caspase-1 cleaves its target substrates, such as the pro-cytokines pro-IL-1β and pro-IL-18, into their active forms [13]. Caspase-1 also cleaves the pore-forming protein gasdermin-D (GSDMD), enabling the release of N-terminal fragments, which oligomerize and form plasma membrane pores. This may result in a type of cell death known as pyroptosis, accompanied by the release of the highly inflammatory cytokines IL-1β and IL-18 [14]. However, cytokine release may also occur from living cells [15, 16]. Five canonical inflammasomes have been described to date, each of which possesses a different sensor protein, three of which are nucleotide-binding domain leucine-rich repeat-containing receptors (NLRs) [12, 17, 18]. The NLR family pyrin domain containing 3 (NLRP3) inflammasome comprises NLRP3, ASC, and caspase-1. NLRP3 is the most studied inflammasome in terms of the broad array of stimuli that can trigger its activation, such as K+ and Cl- efflux, cathepsin leakage, bacteria, and microbial virulence factors [17, 19]. Following the triggering of the sensor protein, the sensor may associate with ASC via their respective pyrin domains (PYD), enabling nucleation of ASC into punctate structures known as “ASC specks” [20]. ASC recruits caspase-1 by interactions with the CARD domain in each respective protein. The association of multiple pro-caspase-1 proteins with the ASC speck brings the proteins in close enough proximity to undergo auto-cleavage, thereby activating caspase-1 [21]. ASC specks are considered the sites of inflammasome activity and therefore may be assessed to probe inflammasome activation [20] (Fig. 1). The assembly of most inflammasomes, such as NLRP3, absent in melanoma 2 (AIM2), and pyrin, requires a “priming” step, which consists of the transcription of pro-IL-1β and sensor proteins [22–25]. Priming is mediated by PRRs such as Toll-like receptors (TLRs) initial sensing at the plasma membrane or on phagosomes [5, 6]. In contrast, in the case of the NLR family CARD containing 4 (NLRC4) inflammasome, the sensors are constitutively expressed

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Fig. 1 Representation of the biological steps leading to canonical inflammasome activation in mouse BMDCs derived from ASC-citrine mice. Note that step 1, Priming, is required for most NLRs, except NLRC4. Bottom panels show fluorescence microscopy (a) and merge with bright-field (BF; b) images of activated NLRC4 inflammasomes (ASC “specks”) after STm stimulation. Nuclei are stained with DAPI

and do not require a priming step for their assembly, which occurs rapidly in response to certain pathogenic factors [26] (Fig. 1). NLR family apoptosis inhibitory proteins (NAIPs) serve as the sensors for the NLRC4 inflammasome, detecting bacterial flagella and type III secretion system components (rod and needle proteins) [26]. In response to detecting these ligands, NAIPs can interact with NLRC4, coordinating inflammasome assembly. Interestingly, due to NLRC4 containing a CARD, assembly of the NLRC4 inflammasome does not require the adaptor ASC, though assembly is facilitated by the presence of ASC [18]. In addition to canonical inflammasomes, non-canonical inflammasomes have also been described and serve as cytosolic sensors of lipopolysaccharide (LPS), a component of the outer membrane of gram-negative bacteria, and modified self-lipids [15]. The non-canonical inflammasome is composed of caspase-11 in mice and caspase-4 or caspase-5 in humans. These proteins serve as both LPS sensors, as they oligomerize by using LPS as a platform, and inflammatory caspases [27]. Following oligomerization around LPS, pro-caspases-4, 5, or 11 undergo auto-cleavage into their active forms [28]. These caspases are not known to cleave pro-IL1β or pro-IL-18 but rather cleave GSDMD into its active form

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[29]. However, non-canonical inflammasomes cooperate with NLRP3 inflammasomes to potentiate its activation [30]. Pro-caspase-11 is not expressed constitutively, unlike pro-caspase-1. Therefore, activation of the non-canonical inflammasome requires priming by PRRs and the production of type-I interferon [30, 31]. The following sections describe procedures to assess mainly the activation of the canonical NLRP3 and NLRC4 inflammasomes and the non-canonical caspase-11-dependent inflammasome over time in mouse bone marrow-derived DCs (BMDCs), using biochemical, immunological, and imaging techniques. BMDCs resemble monocyte-derived DCs or “inflammatory” DCs in vivo [32– 35]. Concordantly, BMDCs respond rapidly to PRR stimulation, are highly responsive to inflammasome activators, and secrete abundant amounts of IL-1β, as we and others have shown [15, 36, 37]. The bacterium Salmonella enterica serovar Typhimurium (STm) is used as a model for the activation of the NLRC4 inflammasome in its flagellated state and for the activation of the NLRP3 inflammasome and the caspase-11 non-canonical inflammasome in its non-flagellated state [31, 38, 39]. Aluminum hydroxide (alum), monosodium urate (MSU) crystals, and toxin-coated beads are additionally used as models for the activation of the NLRP3 inflammasome [17]. Similar methodologies may be applied to probe AIM2 inflammasomes (triggered by microbial DNA) [23, 40] or pyrin inflammasomes (triggered by certain Rho GTPase-modifying bacterial toxins) [25] as indicated in cited references. Because NLRP1 inflammasomes are activated by anthrax lethal factor in only certain mouse strains [41] and by viral proteases [42], these will not be discussed in the present chapter.

2 2.1

Materials Mice

1. C57BL/6J wild-type mice. 2. ASC-citrine mice (see Note 1).

2.2

DNA Constructs

1. Fluorescently tagged ASC, NLRC4, AIM2 retroviral constructs for BMDC transduction (e.g., plasmids #60199 (Addgene), a gift from Dr. R. Vance; #51667 (Addgene) and MSCV ASC-GFP [37], gifts from Dr. E. Alnemri). These will allow the visualization of ASC or NLR “specks,” as a readout for inflammasome activation.

2.3

Cell Culture

1. Filter sterilized DC media: RPMI 1640, 10% heat-inactivated fetal bovine serum (FBS; very low endotoxin for BMDC culture; see Note 2), 2 mM L-glutamine, 55 μM betamercaptoethanol, and 100 Units/mL penicillin–100 μg/mL streptomycin solution (except for experiments requiring bacterial stimulation).

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2. DC media containing 30% granulocyte-macrophage colonystimulating factor (GM-CSF) conditioned media from J558 plasmacytoma [43] or 20 ng/mL commercially available mouse recombinant GM-CSF [44] (see Note 3). 3. Filter sterilized imaging media: Leibovitz’s L15 media without phenol red, with 10% low endotoxin FBS. 4. Platinum E (Plat-E) retroviral packaging cell line [45] (see Note 4). 5. Filter sterilized Plat-E culture media: DMEM, 10% heatinactivated FBS, 2 mM L-glutamine, 100 Units/mL penicillin–100 μg/mL streptomycin solution, 1 μg/mL puromycin, 10 μg/mL blasticidin, and 1 mM sodium pyruvate. 6. Filter sterilized Plat-E transfection media: DMEM, 10% heatinactivated FBS, 2 mM L-glutamine. 7. Phosphate-buffered saline Ca2+/Mg2+-free (PBS). 8. PBS with Ca2+/Mg2+. 9. 0.25% Trypsin-EDTA. 10. Filter sterilized red blood cell (RBC) lysis buffer: 0.1 mM EDTA, 155 mM NH4Cl, 10 mM KHCO3. 11. Non-tissue culture-treated 10 cm Petri dishes. 12. Non-tissue culture-treated 6-well plates. 13. Poly-D-lysine-coated glass bottom p35 dishes (see Note 5). 14. Cell-bind dishes for transfections. 2.4 NLRC4/NLRP3 Inflammasome Stimuli

Salmonella Typhimurium Culture 1. LB broth: 10 g/L tryptone; 10 g/L NaCl; 5 g/L yeast extract in H2O. 2. LB agar plates: 10 g/L tryptone; 10 g/L NaCl; 5 g/L yeast extract; 15 g/L agar in H2O. 3. Streptomycin: 50 mg/mL stock solution. 4. 5 M NaCl.

2.5 NLRP3 Inflammasome Stimuli

1. Ultrapure LPS stock solution. 2. Alum stock solution. 3. MSU stock solution. 4. Lysteriolysin (LLO) or desired bacterial toxin. 5. 3 μM polystyrene beads.

2.6 Other Reagents and Materials

1. 100% w/v trichloroacetic acid (TCA) stock solution. 2. 2% paraformaldehyde (PFA). 3. 8 mg/mL polybrene stock solution.

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4. Lipofectamine or similar transfection reagent. 5. Laemmli sample buffer. 6. Anti-mouse CD11c and CD11b antibodies for flow cytometry. 7. Sterile scalpels and scissors. 8. 1 mL syringes. 9. 25G needles. 10. ELISA immunoplates. 2.7 Instruments and Software

1. Spectrophotometer. 2. Bacterial and tissue culture incubators. 3. Bacterial shaker. 4. Microcentrifuge. 5. Biosafety level 2 cabinet. 6. Tissue culture centrifuge (with a swinging rotor, multiwell plate adapters and temperature range 4–35 °C). 7. Automated cell counter or hemocytometer. 8. Polyacrylamide gel electrophoresis and immunoblotting equipment. 9. Flow cytometer. 10. Fluorescence microscope. A confocal microscope is not required unless examining protein colocalization. 11. Plate reader. 12. NIH ImageJ software (freely available at https://imagej.nih. gov).

3

Methods

3.1 Preparation of Reagents

1. LB broth: Combine 10 g tryptone, 10 g NaCl, and 5 g yeast extract in 1 L H2O. Sterilize by autoclaving. 2. LB agar: Combine 5 g tryptone, 5 g NaCl, 2.5 g yeast extract, and 7.5 g agar in 500 mL H2O. Autoclave the solution to sterilize and melt the agar. Once ~50 °C, add appropriate antibiotics, and pour 15 mL into 10 cm Petri dishes. 3. 5 M NaCl: Dissolve 29.2 g NaCl in 100 mL H2O. Sterilize by autoclaving or by filtering through a 0.22 μM filter. 4. 2× LPS: Final concentration is 60 ng/mL LPS for a 2× solution. From a 5 mg/mL ultrapure LPS stock, dilute 1:100 in DC media. Then, dilute the 1:100 dilution to 1:800 in DC media.

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5. 5× alum: Final concentration is 1000 μg/mL for a 5× solution. From a 40 mg/mL stock, dilute 1:40 in DC media. 6. 3× alum: Final concentration is 600 μg/mL for a 3× solution. From a 40 mg/mL stock, dilute 1:66 in DC media. 7. 5× MSU: Final concentration is 1000 μg/mL for a 5× solution. From a 5 mg/mL stock, dilute 1:5 in DC media. 8. 3× MSU: Final concentration is 600 μg/mL for a 3× solution. From a 5 mg/mL stock, dilute 1:8 in DC media. 3.1.1 Preparation of Toxin-Coated Beads

3.2 BM Isolation and BMDC Differentiation

1. Recombinant LLO is commercially available or may be purified from E. coli strain DP-3570 expressing six histidine-tagged LLO (kindly provided by Daniel Portnoy, University of California, Berkeley) and coated onto 3 micron polystyrene beads as described in refs. 46, 47 and detailed in Chapter 19 (see Note 6). This procedure was adapted from ref. 44. 1. Sacrifice 6–12-week-old mouse according to veterinary guidelines on euthanasia. If analyzing ASC speck formation is desired, BMDCs from a reporter mouse model expressing fluorescently tagged ASC may be used (e.g., ASC-citrine mouse). 2. Remove skin and hair from hind legs. Cut away as much muscle as possible from the femur and tibia and around the hip. 3. Cut the hind leg at the hip to avoid damaging the femur (see Note 7). Cut off the feet of the removed legs, and transfer cleaned legs to a dish on ice containing 5 mL DC media (see Note 8). 4. Take dishes to biosafety cabinet, and transfer to a new dish containing 5 mL DC media to rinse the legs. 5. Place one leg in a dry dish (or dish lid), and use a scalpel to scrape away any remaining muscle. Keep the other leg in a dish containing media on ice. 6. Gently separate femur and tibia at the knee joint using the scalpel, but avoid breaking the bone. 7. Place the tibia back into the dish on ice containing DC media while harvesting the femur. 8. Using the scalpel, cut each end of the femur. Try to cut off the smallest amount of bone possible that will give an opening to the BM. 9. Fill a 1 mL syringe with a 25G needle with DC media, and insert the needle into each end of the cut femur.

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10. Flush the BM from the femur into a 15 mL conical tube (see Note 9). Flush the bone six times in total, three times from each end. Place the tube containing BM on ice. 11. Gently separate the remainder of the foot of the mouse from the tibia using the scalpel, but avoid breaking the bone. 12. Using the scalpel, cut each end of the tibia. 13. Fill a 1 mL syringe with a 25G needle with DC media, and insert into each end of the cut tibia. 14. Flush the BM from the tibia into a 15 mL conical tube (see Note 9). Flush the bone six times in total, three times from each end. Place the tube containing BM on ice. 15. Repeat steps 5–14 with the remaining leg. 16. Centrifuge the BM cells at 200 g for 10 min at 4 °C and discard supernatant. 17. Resuspend cells in 1 mL room temperature RBC lysis buffer per leg. Incubate at room temperature for 2 min. 18. Add 10 mL DC media to cells to halt lysis. 19. Centrifuge at 200 g for 10 min at 4 °C. 20. Resuspend in 20 mL DC media + 30% GM-CSF per leg. 21. Count cells using an automated cell counter or a hemocytometer, and add media as necessary to get the concentration to 1 × 106 cells/mL. The expected yield is ~2 × 107 BM cells per leg. 22. Plate 1 × 107 cells in a non-tissue culture-treated 10 cm dish. If transfecting cells, plate 3 × 106 cells in each well of a non-tissue culture-treated 6-well plate. 3.3 Seeding and Splitting BM Cells

1. Three days after cell isolation, split cells by collecting the supernatant containing suspension cells in a conical tube. Wash the remaining loosely adherent cells with 10 mL PBS, and transfer the wash to the same conical tube. 2. Add 1 mL trypsin to the dish and incubate at 37 °C for 7–10 min. 3. Add 10 mL DC media to the cells and collect the detached cells. Transfer to the same conical tube. 4. Centrifuge at 200 g for 10 min at 4 °C. Remove supernatant with a sterile non-pyrogenic pipette, and resuspend cells in 20 mL DC media (see Note 10). 5. Plate 10 mL of resuspended cells in each of two non-tissue culture-treated 10 cm dishes. Incubate at 37 °C for 3 more days.

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6. Repeat steps 2–6. Cells are ready to use after 2–4 days. 7. Assess cell differentiation to BMDCs by immunofluorescence followed by flow cytometry (CD11chi/CD11b+) [43] as described in Chapter 19. 3.4 Transfecting Plat-E Cells and Seeding BM Cells for Retroviral Transduction

1. The afternoon before transfecting cells, harvest Plat-E cells. Wash a flask of Plat-E cells with 10 mL PBS, and add 1.5 mL trypsin to the flask. Incubate at 37 °C for 1–2 min. 2. Add 15 mL Plat-E media to the flask and harvest cells into a 50 mL conical tube. 3. Count cells using an hemocytometer.

automated cell counter or a

4. Centrifuge cells at 200 g for 10 min at 4 °C. 5. Discard supernatant, and resuspend in Plat-E transfection media to a concentration of ~8.5 × 105 cells/mL. 6. Plate ~8.5 × 106 cells in a 10 cm cell-bind dish. 7. The next morning, transfect with desired fluorescently tagged retroviral constructs using a transfection reagent such as lipofectamine 2000 (Invitrogen) following the manufacturer’s instructions (see Note 11). 8. Isolate BM on the same day of Plat-E transfection. Plate 3 × 106 BM cells in each well of non-tissue culture-treated 6-well plates. 9. Two days after isolating BM cells, collect suspension cells in a conical tube. Add 0.5 mL DC media to each well to prevent adherent cells from drying. Centrifuge collected cells for 7 min at 200 g at 4 °C. 10. Collect retrovirus-containing supernatant from transfected Plat-E cells in conical tubes. Centrifuge at 1000 g for 10 min at 4 °C. 11. Resuspend BM cells in retrovirus-containing supernatant, and transfer cells to the corresponding wells containing adherent BM cells. Add polybrene to each well to a final concentration of 8 μg/mL. 12. Centrifuge at 600 g for 2 h at 35 °C (retroviral transduction). Incubate the cells at 37 °C with CO2 (in a cell culture incubator) for 10–20 min. 13. Collect suspension cells in a conical tube. Add 0.5 mL DC media (with GM-CSF) to each well to prevent adherent cells from drying out. Centrifuge collected cells for 7 min at 200 g. 14. Resuspend BM cells in 3.5 mL DC media (with GM-CSF), and transfer cells to the corresponding wells containing adherent BM cells.

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15. Incubate for 3 days. If retroviral vectors contain antibiotic selection genes, change media, add corresponding antibiotics, and continue culture for 4–6 more days, adding new media with antibiotics every 3 days (see Note 12). 3.5 Salmonella Typhimurium Culture

1. Two days before the experiment, streak out Salmonella enterica serovar Typhimurium (hereafter referred to as STm) from a frozen glycerol stock onto an LB agar plate containing 100 μg/ ml streptomycin. Incubate at 37 °C overnight (see Note 13). 2. One day before the experiment, inoculate 3 mL LB broth with a single colony from the STm plate. Incubate at 37 °C overnight, shaking at 200 rpm with 100 μg/ml streptomycin (see Note 14). 3. If stimulating the NLRC4 inflammasome, a flagellated culture of STm must be generated. On the morning of the experiment, combine 75 μL of the overnight culture and 78 μL of 5 M NaCl in 3 mL LB broth. Incubate the culture without shaking at 37 ° C for 3 h to induce flagella expression (see Note 15). 4. During the LPS priming step (see Subheading 3.7), measure the OD600 of the culture, and calculate the concentration of bacteria with the assumption that an OD of 1.00 equals 5 × 108 bacteria (see Note 16). 5. Collect 1 × 108 bacteria, and centrifuge at 8000 g for 3 min in a microcentrifuge at room temperature (see Note 17). 6. Carefully pipette off the media, and resuspend the pellet in 5 mL DC media without antibiotics for a concentration of 2 × 107 bacteria/mL (see Note 18).

3.6 Harvesting and Plating BMDCs

1. The day before the experiment or on the same day (see Note 19), collect BMDC supernatants containing suspension cells, and transfer them to a conical tube on ice. 2. Wash the remaining adherent cells with 2 or 5 mL PBS, depending on whether cells are collected from 6-well plates or 10 cm dishes, respectively. Transfer the wash to the tube containing the suspension cells. 3. Add 2 or 5 mL of PBS to the adherent BMDCs, and incubate the plate or dish on ice for 30 min. Following the incubation, tilt the plate/dish, and, using a P1000 micropipette, pipette the PBS onto the cells to detach them from the surface of the plate/dish (see Note 20). Repeat this until most of the cells have been detached (approximately 10 times), and transfer the collected adherent cells to the tube from the previous step (see Note 21). 4. Count the cells using an automated cell counter or hemocytometer, and centrifuge them at 200 g for 7–10 min. Remove supernatant and resuspend cells in DC media (without

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antibiotics if stimulating with STm) to a final concentration of 2 × 106 cells/mL if plating in a 96-well plate or 3.5 × 105 cells/ mL if plating in a p35 dish (2 mL). If stimulating caspase-11dependent non-canonical inflammasome is desired, resuspend in DC media containing 100 ng/mL LPS (without antibiotics) to allow for overnight priming (see Note 22). 5. If assessing cytokine production, cell death, protein levels, and cleavage, seed each well of a 96-well plate with 100 μL resuspended cells (2 × 105 cells). Treatments should be done in triplicate, including LPS and non-treated controls. 6. If visualizing inflammasome activation by fluorescent microscopy, seed a poly-D-lysine-coated glass bottom p35 dish with 2 mL resuspended cells (7 × 105–1 × 106 cells). 3.7 Cell Priming and Stimulation in 96-Well Plate

This procedure has been adapted from ref. 36 for NLRP3 stimulation. 1. The morning after plating the cells (see Note 19), prime the BMDCs that will be stimulated with NLRP3 stimuli and the LPS control cells. Add 100 μL 2× LPS to each well, giving a final LPS concentration of 30 ng/mL. Incubate the plate at 37 ° C for 2.5 h. Caspase-11 priming is done overnight (see Subheading 3.6). 2. NLRC4 activation does not require priming [37]. Add 100 μL DC media without antibiotics to the wells where STm will be added. 3. After priming, perform inflammasome stimulation. Into the primed wells, add 50 μL 5× inflammasome stimuli. Incubate cells at 37 °C according to the following (see Note 23): (a) Alum or MSU—4–16 h; 100–300 μg/mL. (b) STm without flagella (NLRP3)—2–4 h; MOI = 5:1 (STm:BMDC) (see Note 24). (c) STm without flagella (caspase-11)—4–24 h; MOI = 5:1 (STm:BMDC). (d) STm with flagella—0.5–1 h; MOI = 5:1 (STm:BMDC). (e) Toxin-coated beads (ratio between 10 and 20 beads/ BMDC). 4. If stimulating with STm, add gentamicin to the appropriate wells at the end of the inflammasome stimulation period, to kill extracellular bacteria and halt the stimulation. Dilute 50 mg/ mL gentamicin stock to 1.25 mg/mL in DC media. Add 20 μL gentamicin to a final concentration of 100 μg/mL. 5. Centrifuge plates at 200 g for 5 min. Transfer 200 μL of the supernatant from each well to a separate 96-well plate, being careful not to touch the cells at the bottom of the plate (see Note 25). Store at -20 °C (see Note 26). Discard the remaining 50 μL of supernatant.

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6. The remaining cells on the bottom of the wells may be stored at -80 °C in 20 μL Laemmli sample buffer [48] and subsequently assessed for caspase-1 and gasdermin-D expression and cleavage by immunoblotting [39]. Alternatively, cell pellets may be stored for RNA isolation in recommended solutions for nucleic acid preservation at -80 °C. 3.8 Caspase-11 Detection by TCA Precipitation

1. For caspase-11 detection, precipitate cell supernatants adding a final concentration of 10% TCA (v/v) for 1 h on ice [31, 49]. Cell pellets are stored in Laemmli sample buffer in a concentration of 100 μL/1 × 106 cells. 2. Centrifuge precipitated supernatants at 20,000 g for 30 min at 4 °C. 3. Discard supernatants and air-dry pellets. 4. Resuspend protein pellets in Laemmli sample buffer 2× with 2-mercaptoethanol, and heat at 95 °C for 5 min [31, 49]. Precipitated supernatants from 3 × 106 BMDCs are recommended to be loaded per well on polyacrylamide gels.

3.9 Cell Priming and Stimulation in GlassBottom p35 Dishes for ASC Speck Visualization by Fluorescence Microscopy or Live Cell Imaging

1. The morning after plating the cells (see Note 19), remove unattached cells, and add 1 mL DC media. If stimulating BMDCs with STm, use media without antibiotics. If stimulating the NLRP3 inflammasome, add LPS to a final concentration of 30 ng/mL, and incubate at 37 °C for 2.5 h. If stimulating the NLRC4 inflammasome or the caspase-11 non-canonical inflammasome, proceed to the next step. 2. Stimulate the cells by adding 500 μL 3× inflammasome stimuli. Incubate the cells at 37 °C according to the following: (a) Alum or MSU—4–16 h; 100–400 μg/mL (see Note 27). (b) STm without flagella (NLRP3)—2–4 h; MOI = 5:1 (STm:BMDC). (c) STm without flagella (caspase-11)—4–24 h; MOI = 5:1 (STm:BMDC). (d) STm with flagella—15 min–1 h; MOI = 5:1 (STm: BMDC). 3. If stimulating with STm, add gentamicin to the appropriate dishes after stimulation. Dilute 50 mg/mL gentamicin stock to 1.5 mg/mL in DC media. Add 100 μL gentamicin, to a final concentration of 100 μg/mL. 4. Aspirate supernatant and wash twice in 1.5 mL PBS with Ca2+/ Mg2+. Add imaging media if the cells are being imaged while alive to detect fluorescent ASC or NLR speck formation over time. Otherwise, proceed to the next step. 5. Aspirate PBS and add 1.5 mL 2% PFA to the cells. Fix on ice for 20 min, protected from light.

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6. Aspirate 2% PFA and wash once with 1.5 mL PBS. 7. Aspirate PBS, and add 1.5 mL permeabilization buffer (e.g., 0.2% saponin, 2% bovine serum albumin optionally containing DAPI at a concentration of 1 μg/mL to stain nuclei and count activated cells) (see Note 28). Incubate protected from light at room temperature for 20 min. 8. Aspirate permeabilization buffer and wash once with 1.5 mL PBS. Store in 1.5 mL PBS protected from light. 3.10 ASC Speck Counting

This section describes how to count activated BMDCs differentiated from ASC-citrine mice using an ImageJ macro (see Note 29). 1. Open the file containing images of stimulated ASC-citrine BMDCs in ImageJ (see Note 30). 2. Duplicate the image [Image -> Duplicate]. run("Duplicate...", "title=[Speck Counts]");

3. Open “Window/Level” [Image -> Adjust -> Window/ Level]. Set “Window Center” to 250. Set “Window Width” to 10. Hit apply (see Note 31). run("Window/Level..."); setMinAndMax(250, 10); run("Apply LUT");

4. Open “Threshold” [Image -> Adjust -> Threshold]. Make sure “Dark background” is checked (see Note 32). Hit apply. setAutoThreshold("Default dark"); //run("Threshold..."); setOption("BlackBackground", false); run("Convert to Mask");

5. Open “Binary Options” [Process -> Binary -> Options]. Set “Iterations” to 3. Set “Count” to 3. Set “Do:)” to “Dilate.” Leave all other options as they are. run("Options...", "iterations=3 count=3 do=Dilate");

6. Run “Watershed” [Process -> Binary -> Watershed]. run("Watershed");

7. Open “Analyze Particles” [Analyze -> Analyze Particles]. Set “Size” to “3-Infinity” [pixel units]. Set “Show:” to “Outline.”

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Check “Display results” and “Summarize.” The “Count” tab in the “Summary” window will show the number of specks counted. run("Analyze Particles...", "size=3-Infinity pixel show=Outlines display summarize");

3.11

Count Nuclei

1. Open the file containing images of the DAPI-stained nuclei of stimulated BMDCs in ImageJ (see Note 29). 2. Duplicate the image [Image -> Duplicate]. run("Duplicate...", "title=[Nuclei counts]");

3. Open “Window/Level” [Image -> Adjust -> Window/ Level]. Set “Window Center” to 500. Set “Window Width” to 10. Hit apply (see Note 31). run("Window/Level..."); setMinAndMax(500, 10); run("Apply LUT");

4. Open “Remove Outliers” [Process -> Noise -> Remove Outliers]. Set “Radius” to 5.0, “Threshold” to 50, and “Which outliers” to “Bright” (see Note 33). run("Remove Outliers...", "radius=5 threshold=50 which=Bright");

5. Open “Threshold” [Image -> Adjust -> Threshold]. Make sure “Dark background” is checked. Hit apply (see Note 32). setAutoThreshold("Default dark"); //run("Threshold..."); setOption("BlackBackground", false); run("Convert to Mask");

6. Open “Binary Options” [Process -> Binary -> Options]. Set “Iterations” to 3. Set “Count” to 3. Set “Do:)” to “Close.” Leave all other options as they are (see Note 34). run("Options...", "iterations=3 count=3 do=Close");

7. Run “Watershed” [Process -> Binary -> Watershed]. run("Watershed");

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8. Open “Analyze Particles” [Analyze -> Analyze Particles]. Set “Size” to “5-Infinity.” Set “Show:” to “Outline.” Check “Display results” and “Summarize.” The “Count” tab in the “Summary” window will show the number of nuclei counted. run("Analyze Particles...", "size=5-Infinity pixel show=Outlines display summarize");

4

Notes 1. These strains are commercially available from The Jackson Laboratories (strains #000664 and #030744). All mouse studies should be performed in compliance with veterinary regulations on the humane care and use of laboratory animals. 2. Due to the sensitivity of the BMDC culture to endotoxins that may drive BMDC maturation, very low endotoxin FBS should be used for BMDC culture (e.g., Defined FBS, Hyclone SH30070.03). 3. The preparation of GM-CSF conditioned media from J559 plasmacytoma allows for the availability of large amounts of GM-CSF at relatively lower costs than recombinant GM-CSF (rGM-CSF). However, the use of rGM-CSF is less timeconsuming and might be preferred for small-scale experiments. 4. Plat-E cells are derived from the 293T cell line and are commercially available. 5. For reference, Mattek #P35GC-1.5-14-C. 6. Other bacterial toxins and virulence factors, such as components of the T3SS system, may be coated to polystyrene beads to probe phagocytosis-coupled inflammasome activation. 7. A considerable amount of BM is in the head of the femur. The leg is removed at the hip to ensure this portion of the femur remains intact. 8. Up to this step may be performed on the lab bench. 9. If media sprays out of the top, this means the opening is not big enough. Cut a little bit more off of the end of the bone and try again, until media is able to flow out of the bone. 10. Due to the sensitivity of the BMDC culture to endotoxins that may drive BMDC maturation, the use of an aspirator is not recommended for this procedure. Using a sterile, non-pyrogenic serological pipette ensures that BMDCs do not come in contact with any unwanted stimuli.

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11. Lipofectamine 2000 is used in our lab, but other transfection reagents may be used instead. If other reagents are used, they may need to be optimized for use in this procedure. 12. A control for antibiotic selection efficiency is recommended, e.g., by having an additional well containing non-transduced BM cells treated with selection antibiotics. These cells should die along the differentiation process. 13. Salmonella enterica serovar Typhimurium may cause illness if mishandled. Always use caution and wear gloves when handling. Be sure to dispose of everything as hazardous waste. 14. STm is a flagellated bacterium that expresses a type III secretion system (T3SS). Flagellin and T3SS components (rod and needle proteins) activate the NAIP/NLRC4 inflammasome [18]. However, STm only expresses flagellin under conditions that mimic the intestinal environment. These conditions are not met if bacteria are grown to a stationary phase to prevent flagellin expression and NAIP/NLRC4 inflammasome activation. In these conditions, STm will activate the NLRP3 inflammasome [38]. S. enterica with deletions in the genes fljB and fliC can be used to ensure flagellin is not expressed [39]. 15. Growth of S. enterica in hyperosmolar conditions without agitation induces the expression of the SPI-1 type-3 secretion system and flagellin [37, 38]. 16. OD 1 = 5 × 108 cells is a general rule of thumb, but this is not exact. However, for the purpose of this protocol, this approximation should be sufficient. 17. It is recommended to wash the pellet twice with PBS. 18. The pellet of flagellated bacteria should appear “fluffy” due to the flagella and will not be compact. Be especially careful when pipetting off the supernatant. 19. Harvesting and plating cells on 96-well plates may be done on the morning of the experiment. However, if collecting cells from p35 or p60 dishes for immunoblotting or performing immunofluorescence microscopy or live cell imaging, we recommend plating the cells the night before to give cells time to adhere to plates. If only the cell supernatants are assessed (e.g., for ELISA) and higher cell numbers are not required for immunoblotting, we recommend seeding the cells on the day of the experiment. 20. This step takes considerable time due to having to pipette the PBS onto every surface of the well. You should be able to see the adhered cells detach from the surface when pipetting. Be careful not to introduce bubbles during this step, as this may inadvertently lead to BMDC activation.

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21. The plate can be viewed under a light microscope to determine if the cells have been adequately detached. A small number of cells will most likely remain adhered to the plate (mainly macrophages). 22. For the detection of caspase-11 cleavage, a higher number of cells is required (a minimum of 3 × 106 cells/condition is recommended to perform TCA precipitation from cell supernatants). 23. If stimulating the inflammasome with different stimuli during the same experiment, you may time the stimulations so that the cells will all be collected simultaneously. For example, if stimulating the NLRP3 inflammasome with alum and the NLRC4 inflammasome with STm, add the STm during the last hour of alum incubation. 24. 50 μL of bacteria at a concentration of 2 × 107 bacteria/mL will give 1 × 106 bacteria, which aligns with an MOI of 5:1, assuming 2 × 105 cells were seeded in each well. The MOI is based on the ratio bacteria:cells and may be adjusted to achieve different degrees of inflammasome activation. 25. From the 200 uL of supernatant, you may separate 50 μL, and store at 4 °C to detect cell death by measuring lactate dehydrogenase release (according to manufacturer’s instructions). Store the remaining supernatants at -20 °C, and limit freeze– thaw cycles as much as possible. 26. Supernatants may be screened for proinflammatory cytokine secretion and IL-1β production (the hallmark of inflammasome activity) by ELISA or Luminex assays. For caspase-11 detection, follow Subheading 3.8. 27. Both MSU and alum will be clearly visible under a light microscope. Alum will make the image somewhat cloudy compared to MSU and may make imaging more challenging. MSU is recommended for NLRP3 stimulation if you plan to image the cells. 28. Additional antibodies may be added to detect inflammasome components or other desired proteins (such as anti-ASC or anti-NLRs). Incubation with anti-cytoskeleton probes such as phalloidin is recommended to visualize cell boundaries. 29. All BMDCs differentiated from ASC-citrine mice will express fluorescent ASC. If fluorescently tagged ASC or NLR constructs are transduced, transduction efficiency must be considered when calculating the percentage of activated cells, given that not all BMDCs may express the construct. Alternatively, the sorting of fluorescent cells may be performed in advance. The quantification of inflammasome-activated cells may also be performed by flow cytometry, gating on transduced cells, as described and shown in [39, 50, 51].

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30. The counting process can be automated by running these commands as a macro in ImageJ. The italicized text beneath each step is the macro command to run that step. The steps can be copied and pasted into a new macro in ImageJ. To use these macros, create a new macro [Plugins -> New -> Macro]. 31. Depending on the number of cells, the intensity of the fluorescence, and the exposure of the image, the “Window/Level” values may need to be changed. If there is significant noise in the image after adjusting the “Window/Level” values, try a higher “Window Center” value. If specks or nuclei are being excluded, try a lower “Window Center” value. 32. The previous step of setting the “Window/Level” effectively makes the image black-and-white. When thresholding, the “Auto-threshold” option is recommended. However, due to the “Window/Level” step, alternative threshold values do not result in significant changes. 33. The goal of this step is to remove particles and fragmented nuclei from the count, while still counting all nuclei. Depending on the size of the nuclei in the image, the radius for the outliers may need to be adjusted. To do so, measure the diameter of an average nucleus in your image (draw a straight line from one end of the nucleus to the other, and then go to Analyze -> Measure), and adjust the value accordingly. 34. This step may help reduce the counting of fragmented nuclei by combining them into single particles, but may not be necessary depending on the image. If you choose to use this step, try different “Iterations” and “Count” values to determine which work best for your image. Setting “Iterations” above 6 may change the size and circularity of the nuclei and may negatively impact counting accuracy.

Acknowledgments This work was supported by NIH grant R01 AI137173. The summary figure was created using Biorender.com through Thomas Jefferson University library portal. References 1. Steinman RM (2007) Dendritic cells: understanding immunogenicity. Eur J Immunol 37 (Suppl 1):S53–S60. https://doi.org/10. 1002/eji.200737400 2. Barton GM, Kagan JC (2009) A cell biological view of Toll-like receptor function: regulation through compartmentalization. Nat Rev

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Chapter 21 Analysis of LC3-Associated Phagocytosis and Antigen Presentation in Macrophages and B Cells Svenja Luisa Nopper, Katarina Wendy Schmidt, Laure-Anne Ligeon, and Christian Mu¨nz Abstract Canonical autophagy and the non-canonical autophagy pathway LC3-associated phagocytosis (LAP) play crucial roles in the immune system by processing antigens for major histocompatibility complex (MHC) class II restricted presentation to CD4+ T cells. Recent studies offer insight into the relationship between LAP, autophagy, and antigen processing in macrophages and dendritic cells; however their involvement during antigen processing in B cells is less well understood. In this chapter, we describe how to monitor, manipulate, and understand the role of LAP and classical autophagy during MHC-restricted antigen presentation by human monocyte-derived macrophages as well as in B cell lymphoblastoid cell lines (LCLs). It includes an explanation on how to generate LCLs and monocyte-derived macrophages from primary human cells. Then we describe two different approaches to manipulate the autophagy pathways: silencing of the atg4b gene using CRISPR/Cas9 technology and a lentivirus delivery system for specific ATG4B overexpression. We also propose a method for triggering LAP and measuring different ATG proteins using Western blot and immunofluorescence. Finally, we show an approach to investigate MHC class II antigen presentation by an in vitro co-culture assay that uses the measurement of secreted cytokines, released by activated CD4+ T cells, as readout. Key words Autophagy, Confocal microscopy, LC3-associated phagosome, CRISPR/Cas9, B cells

1

Introduction Both B cells and macrophages are professional antigen presenting cells (APCs) and are therefore a crucial component of the immune system [1, 2]. Antigen presentation on major histocompatibility complex (MHC) molecules for T lymphocyte activation is the key initiation switch for triggering an antigen-specific adaptive immune response. There are two types of MHC molecules which present

Svenja Luisa Nopper, Katarina Wendy Schmidt, Laure-Anne Ligeon, and Christian Mu¨nz contributed equally to this chapter. Roberto J. Botelho (ed.), Phagocytosis and Phagosomes: Methods and Protocols, Methods in Molecular Biology, vol. 2692, https://doi.org/10.1007/978-1-0716-3338-0_21, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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different antigens. Whereas MHC class I mainly presents peptides generated in the cytoplasm by the proteasome, MHC class II is loaded with exogenous peptides that have been endocytosed or phagocytosed and are degraded via the lysosomal degradation pathway [3]. Therefore, intracellular peptides are mainly presented by MHC class I to CD8+ cytotoxic T cells, and exogenous peptides are mainly presented by MHC class II to CD4+ helper T cells. However, there are other pathways which are involved in the lysosomal degradation and the presentation of antigens, such as autophagic pathways. The non-canonical autophagy pathway LC3-associated phagocytosis (LAP) as well as classical macroautophagy can transport extracellular or intracellular antigens to the MHC class II loading compartments, respectively, enabling MHC class II to present extracellular proteins more efficiently and cytoplasmic material to any extent [3]. Macroautophagy includes a stepwise formation of a doublemembrane vesicle, called the autophagosome, that sequesters diverse types of intracellular cargoes such as cytoplasmic protein aggregates, damaged organelles, or pathogens. Various AuTophaGy-related (ATG) proteins coordinate this process which concludes when the outer autophagosomal membrane fuses with a lysosome for cargo degradation [4]. The microtubule-associated protein light chain 3 (LC3/ATG8) is used as the main marker for autophagosomes. It is produced as pro-LC3, which is cleaved at the C-terminus by the cysteine protease ATG4 to form the cytosolic LC3-I form. Then, LC3-I is conjugated to phosphatidylethanolamine by the ATG12-ATG5-ATG16L conjugation complex to form LC3-II. The lipidated LC3-II form is embedded into the forming inner and outer autophagosomal membranes, where it facilitates cargo uptake, the recruitment of other ATG-related proteins, and membrane fusion [4]. Apart from mediating the transition from pro-LC3 to LC3-I, ATG4 is the sole ATG-related protein responsible for the deconjugation of LC3-II from the outer autophagosomal membrane during autophagosome maturation and lysosomal fusion, thereby allowing the recycling of LC3-II to the cytosolic LC3-I form [5]. Coupling of viral and tumor antigens to LC3 for autophagosome degradation has been shown to significantly increase their presentation on MHC class II molecules, which results in improved CD4+ T cell priming [3]. Indeed, several pathogen-derived proteins are processed by an autophagy-mediated mechanism for MHC class II presentation, including the Epstein-Barr virus (EBV) nuclear antigen 1, enhancing the adaptive immune response [6]. Additionally, LAP contributes to extracellular peptide presentation of MHC class II molecules by stabilizing pathogencontaining phagosomes [7]. During LAP, phagocytosed pathogens, which engage Toll-like receptor signaling, can trigger the recruitment and the direct lipidation of LC3 to the phagosomal

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membrane, in order to form the LC3-decorated single membrane vesicle called the LAPosome [8]. Although several ATG-related proteins are utilized by LAP as well as macroautophagy, there are distinct differences in their molecular machinery. For instance, LAP specifically requires NADPH oxidase 2 (NOX2) for the generation of reactive oxygen species (ROS). Oxidative modification can regulate the activity of ATG4B in both canonical macroautophagy and LAP [9]. Recently, it has been shown that in human macrophages, ROS regulates the activation of ATG4B during LAP by oxidative inactivation, which prevents ATG4B from cleaving LC3-II from the LAPosome membrane. This in turn stabilizes the LAPosome allowing a sustained antigen presentation on MHC class II and thereby enhancing CD4+ T cell activation [5]. These findings support the notion of macrophages as antigen presenting cells and strengthen their relevance in further investigations on the role of LAP in adaptive immunity. In B cells, however, the involvement of autophagy processes during antigen processing and presentation is less well understood, and many questions remain to be answered [7, 10]. An excellent tool to study B cell biology is the generation of lymphoblastoid cell lines (LCLs), which provide an easy-to-use mechanism for B cell immortalization. These immortalized and relatively stable cell lines can be generated from any donor by infecting human primary B cells with EBV. With the help of EBV oncoproteins, these cell lines circumvent apoptosis and maintain a state of constant activation, which enables unrestricted cell proliferation [11]. In comparison to other human cancer cells lines, which frequently undergo genetic rearrangements, LCLs have been reported to be extremely stable with regard to genomics, transcriptomics, and proteomics [12]. In the following chapter, we describe how to use LCLs as well as human monocyte-derived macrophages in different assays that manipulate the autophagy machinery involved in both macroautophagy and LAP to study MHC-restricted antigen presentation and subsequent T cell activation. First, we explain how to generate LCLs and monocyte-derived macrophages from primary human cells, isolated from blood. Secondly, we propose two different approaches to manipulate the autophagy machinery with one strategy focusing on deleting the atg4b gene using CRISPR/Cas9 technology and another using a lentivirus delivery system for specific ATG4B overexpression. Then, we discuss how to trigger LAP specifically and how to measure and visualize the level and localization of different ATG-related proteins using Western blot and immunofluorescence staining in B cells and macrophages. Finally, we describe a functional approach to investigate MHC class II antigen presentation by an in vitro co-culture assay that uses the measurement of secreted cytokines, released by activated CD4+ T cells, as readout. These techniques will provide valuable tools for

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further investigations regarding the influences of LAP or macroautophagy on the adaptive immune response.

2

Materials Cell Culture

All experiments are performed with human primary cells isolated from leukocyte concentrates of healthy donors (Zurich Blood Center).

2.1.1 Isolation of Human Peripheral Blood Mononuclear Cells

1. Leukocyte concentrate (buffy coat) 50 mL (Zurich Blood Center) (see Note 1).

2.1

2.1.2 Isolation and EBV Transformation of Human B Cells

2. Peripheral blood mononuclear cells (PBMCs) are isolated by Ficoll-Paque PREMIUM density-gradient centrifugation (GE Healthcare). 1. CD19+ B cells are isolated from PBMCs using positive selection by magnetic-activated cell sorting (MACS) (Miltenyi Biotec). 2. CD19+ B cells are cultured in lymphoblastoid cell lines (LCLs) RPMI-1640 culture medium supplemented with 10% fetal bovine serum (FBS) also referred to as “R10” (see Note 2). 3. CD19+ B cells are immortalized using EBV to produce LCLs (see Note 3).

EBV Production and Titration

1. EBV producer cell line: HEK293 stably expressing the p2089 EBV bacmid [13]. 2. The virus suspension is filtered through a 0.45 μm filter. 3. Virus concentration is performed in Nalgene® centrifugation tubes (Sigma Aldrich) by ultracentrifugation. 4. Virus titration is performed by transformation of CD19+ B cells into LCLs.

2.1.3 Isolation of Human Macrophages

1. Human PBMCs are isolated as described in Subheading 2.1.1. 2. The CD14+ positive cells are isolated from the PBMCs by positive selection using magnetic beads on an autoMACS cell separator (Miltenyi Biotec). 3. CD14+ monocytes are cultured in macrophage culture medium (see Subheading 2.7.1). 4. From the CD14+ monocytes, macrophages are derived through differentiation with granulocyte-macrophage colonystimulating factor (GM-CSF) as described in Subheading 3.1.4. Differentiated macrophages are used between day 6 and 10 of culture.

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Table 1 Sequence of crRNAs, targeting both DNA strands of the ATG4B gene

2.1.4 Isolation of Human Whole Blood CD4+ T Cells

Target gene

Target DNA strand

Sequence (5′-3′)

ATG4B

+

TCCTCAACGCATTCATCGAC

ATG4B

-

AGCAAACCGGAGAGTGTCGT

1. Proceed for the positive magnetic selection of the CD14+ cells from the PBMC as described in Subheading 2.1.3, but do not discard the CD14- fraction. 2. CD14- fraction is frozen using BAMBANKER according to manufacturer’s recommendations and will be used to isolate the autologous blood CD4+ T cells on the day of the antigen presentation assay. 3. Autologous human CD4+ T cells are positively selected using the CD4+ T cell isolation kit (Miltenyi Biotec) and MACS sorting (see Note 4). 4. The freshly isolated CD4+ T cells are co-cultured with stimulated macrophages (see Subheading 3.1.4).

2.2 CRISPR/Cas9 Knock-out of ATG4B in LCLs

1. The ribonucleic acid (RNA) duplex is composed of transactivating RNA (tracrRNA) (Alt-R® CRISPR-Cas9 tracrRNA, IDT) and target-specific CRISPR RNA (crRNA) (Alt-R® CRISPR-Cas9 crRNA, designed with and ordered from IDT). The tracrRNA and crRNAs are reconstituted in IDTE buffer (pH = 7.5, IDT) to a stock concentration of 200 μM. Two guide crRNAs were designed to target the plus and minus strand of the ATG4B gene, respectively (see Note 5). Exemplary sequences for an ATG4B-KO can be found in Table 1. 2. Cas9 nuclease (Alt-R® Cas9 Nuclease V3, IDT) is reconstituted in IDTE buffer (pH = 7.5, IDT) to a stock concentration of 61 μM. 3. The nucleofection medium and reaction 16-well strips are used from the Amaxa™ P3 Primary Cell 4D-Nuceleofector™ X Kit (Lonza). 4. The 4D Nucleofector technology (Lonza) is used for electroporation.

2.3 Lentiviral Constructs of ATG4Bwt or ATG4BC78S Overexpression

1. The genes carrying Flag-Flash-ATG4wt and Flag-FlashATG4C78S can be synthetized by GeneArt, Thermo Scientific. 2. The different ATG4B genes can be subcloned into a pENTR1A noccDB gene vector (Addgene) and introduced into the pLENTI-CMV-Puro DEST vector (Addgene) by site-specific recombination (Gateway Cloning Technology, Thermo Fisher

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Scientific). For production of lentiviral particles, lentiviral vectors are co-transfected with the helper plasmids pCMVΔR8.91 and pMDG into HEK293T cells by calcium phosphate transfection. Cell culture supernatants containing recombinant viral particles are then harvested on day 1, 2, and 3 after transfection, filtered through a 0.45 μm filter, and stored at -80 °C [14] (see Note 6). 3. PEG-it virus precipitation solution (System Biosciences) is used for concentrating the lentiviral particles according to the manufacturer’s instructions. 4. To increase transduction efficiency of the lentiviruses, primary human macrophages are infected together with 6 μg/mL polybrene (Merek), a cationic polymer which acts to neutralize the charge repulsion between virions and the cell surface. 2.4 2.4.1

Western Blot Reagents

1. 1× Laemmli sample buffer: 0.002% bromophenol blue, 10% glycerol, 0.07 M sodium dodecylsulfate (SDS), 0.0625 M Tris– HCl pH 6.8, ddH2O, 1% beta-mercaptoethanol freshly added before use). 2. ECL detection reagents. 3. 30% acrylamide/bis-acrylamide, 29:1 solution. 4. TEMED (N,N,N,N-Tetramethylethylthylenediamine). 5. Pre-stained protein marker. 6. 0.45 μm polyvinylidene difluoride (PVDF) transfer membrane. 7. Whatman filter paper. 8. Skim milk powder. 9. 0.1% TWEEN 20 detergent. 10. Isopropanol.

2.4.2

Buffers

1. Resolving gel buffer: 1.5 M Tris–HCl pH 8.8, for a 10% acrylamide gel (Bio-Rad). 2. Stacking gel buffer: 0.5 M Tris–HCl pH 6.8, for a 4% acrylamide gel (Bio-Rad). 3. 10% ammonium persulfate (APS); store aliquots at 4 °C. 4. SDS-PAGE Running buffer: 1× dilution of 10× Tris/glycine/ SDS (TGS) with ddH2O to obtain 1× TGS final—25 mM Tris, 192 mM glycine, 0.1% SDS, pH 8.3. 5. Transfer Buffer 1×: dilution of 20× NuPAGE transfer buffer (Thermo Fisher) with ddH2O; add 20% methanol. 6. 0.1% PBS-TWEEN-20 (PBS-T). 7. Blocking buffer: 5% skim milk powder in 0.1% PBS-T.

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2.5 Immunofluorescence and Confocal Microscopy

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1. Sterile, round glass coverslips (1.5 mm) are placed inside the well of a 24-well plate and used for seeding 1 × 106 LCLs or 2.5 × 105 human monocyte per well (see Note 7). 2. Fixation: 4% paraformaldehyde (PFA) solution in PBS. 3. Permeabilization: 0.1% Triton X-100 in PBS. 4. Blocking buffer: 1% FBS in sterile PBS. Freshly prepared each time. 5. Primary antibodies are diluted 1:100 (ATG4B) and 1:500 (LC3) in blocking buffer (see Note 8 and Subheading 3.6). 6. Secondary antibodies are diluted 1:500 in blocking buffer (see Note 9). 7. DAPI (4,6-diamidino-2-phenylindole) nucleic acid dye is diluted 1:5000 in PBS (stock concentration, 5 mg/mL). 8. Mounting medium: Dako Fluorescence Mounting Medium (Dako North America, Inc.). 9. Microscopy slides: micro slides, double frosted. 10. For image acquisition, a confocal laser scanning microscope (Leica Stellaris 5) is used with a 63× oil immersion objective (1.4 NA).

2.6 Role of LC3Associated Phagocytosis During MHC Class II Antigen Presentation

2.7 2.7.1

Reagents Cell Culture

1. Whole CD4+ T cells are co-cultured with their autologous human macrophages at a ratio (2:1) after previously being fed with soluble Candida albicans extract as described in Subheading 3.4. 2. Detection of IL-17A and IFN-ɣ in the supernatants after 5 days of co-culture using ELISA kits; see Subheading 2.7.3 (see Note 10). 1. MACS buffer: 500 mL sterile PBS, 1% (v/v) human serum, 2 mM EDTA filtered through 0.22 μm filter. 2. B cell and LCL culture medium: Roswell Park Memorial Institute medium (RPMI)-1640 (Gibco), supplemented with 10% heat-inactivated fetal bovine serum (FBS) and penicillin–streptomycin (ThermoFisher). From now on, this cell culture medium will be referred to as “R10.” 3. Macrophage culture medium: Dulbecco’s modified Eagle’s medium (DMEM, Gibco), 10% (v/v) heat-inactivated fetal bovine serum (FBS), 2 mM glutamine (Gibco), sodium pyruvate (Gibco), 100 U/mL penicillin–streptomycin. Add granulocyte-macrophage colony-stimulating factor (GM-CSF, Biolegend) immediately before use. 4. MicroBeads Kits (Miltenyi Biotec): human CD14+ kit; CD4+ T cell isolation kit and human CD19+ kit.

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1. Primary Antibodies (a) Immunofluorescence: mouse anti-ATG4B antibody (Medical and Biological Laboratories [MBL], N134/3) and rabbit antiLC3B antibody (MBL, PM036). (b) Western blot: rabbit anti-LC3B antibody (Novus NB1002220) antibody, rabbit anti-ATG4B (Abcam) antibody, mouse anti-GAPDH (Sigma) antibody, mouse anti-vinculin (Sigma) antibody. 2. Secondary Antibodies: (see Note 11). (a) Immunofluorescence: Alexa-Fluor® 488 or 555 conjugated goat anti-mouse immunoglobulin (Invitrogen) and AlexaFluor® 488 or 555 conjugated goat anti-rabbit immunoglobulin (Invitrogen). (b) Western blot: goat anti-rabbit immunoglobulin or goat antimouse immunoglobulin coupled to horseradish peroxidase (HRP) (Jackson).

2.7.3

Other Probes

1. Latex beads, amine-modified polystyrene, fluorescent blue (L0280, Sigma Aldrich) are stored in the dark at RT and used as a negative control for LAP induction (see Note 12). The beads can also be coated with Candida albicans extract (see Subheading 3.4.1). 2. Candida albicans extract, stock solution 0.4 mg/mL in sterile water, and aliquots of 125 μL are kept at -20 °C. 20–80 μg/ mL of C. albicans extract can be directly used to trigger LAP in human macrophages (see Subheading 3.4.2) or directly coupled to beads for LAP stimulation and antigen presentation assay (see Subheading 3.7). 3. ELISA kits to investigate CD4+ T cell activation: (a) Human IFN-ɣ ELISA kit (Mabtech). (b) Human IL-17A Quantikine High Sensitive ELISA kit (R & D). (c) Meso Scale Discovery (MSD) Th17 Panel 1 Human Kit (MSD).

3 3.1

Methods Cell Culture

3.1.1 EBV Production and Titration

1. EBV (strain B95-8) is constitutively produced by HEK293 cells with the p2089 bacmid and can be collected by harvesting the culture supernatant. The virus suspension is filtered through a 0.45 μm filter to eliminate cell remnants (see Note 13).

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2. The virus is concentrated in Nalgene® centrifugation tubes (Sigma Aldrich) by ultracentrifugation (15,800 rpm for 2 h at 4 °C). 3. After removing the supernatant, cold, sterile PBS is added on the pellet (see Note 14) and either left on ice for 20–30 min or at 4 °C overnight. Then, everything is resuspended well and combined in one tube. 4. The virus concentrate can be titrated by adding different volumes of the suspension (e.g., 2 μL, 5 μL, 10 μL) to CD19+ B cells to determine the necessary volume to produce outgrowing LCLs. However, the necessary volume of concentrated EBV may vary between different EBV productions. 3.1.2 Isolation of Human Peripheral Blood Mononuclear Cells (PBMCs)

The following steps are performed for the isolation of human B cells, T cells, and monocyte-derived macrophages from PBMCs. 1. For gradient centrifugation, 15 mL of Ficoll-Paque PREMIUM is gently and slowly overlaid with 25 mL blood and centrifuged for 25 min at 1000 g at room temperature (RT) (Acceleration 3, Deceleration 1). 2. The yellow leukocyte layer is carefully transferred to a new tube, and the cells are washed once with sterile cold PBS before to proceed for MACS sorting (see Note 15).

3.1.3 Isolation and EBV Transformation of Human B Cells

1. CD19+ B cells are isolated from PBMCs by positive magnetic cell separation using a CD19 MicroBeads kit (Miltenyi Biotec) (see Note 16). 2. The CD19+ B cells are counted and plated in a flat bottom 96-well plate at a density of 250,000 cells per well (see Note 17) in a final volume of 100 μL R10. 3. EBV virus concentrate is added to the cells in the necessary volume to result in LCL outgrowth (see Subheading 3.1.1). Then, the wells are filled up to a final volume of 200 μL with R10. 4. Optional: To increase infection efficacy, spin infection can be performed by centrifugation for 30 min at 800 g at 4 °C. 5. The cells are incubated at 37 °C, 5% CO2 for several days. When the medium gets yellow and cell clumps are observed, 100 μL of old medium is gently replaced by 100 μL of fresh R10. 6. Three weeks after EBV infection, the LCLs should be ready for expansion. LCLs grow in cell clumps and need to be gently resuspended prior to splitting. General tip: see Note 18.

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7. LCLs are considered an established cell line after approximately 3 weeks. • Immunofluorescence: 1 × 106 cells in a well of a 24-well plate in a final volume of 500 μL R10. • Western blot: Cell lysis at a concentration of 3000 cells per μL Laemmli buffer (e.g., 300,000 cells in 100 μL Laemmli buffer). 3.1.4 Isolation of Monocyte-Derived Macrophages

1. The PBMCs are collected according to Subheading 3.1.2, and then one can proceed for CD14+ magnetic MACS sorting with CD14 Microbeads (Miltenyi Biotec) according to the manufacturer’s instructions (see Note 19). 2. Resuspend 1 × 108 cells in 90 μL of MACS buffer (see Subheading 2.7.1). 3. Add 10 μL of CD14+ beads (per 1 × 108 cells), but not fewer than 100 μL of bead and cell mixture. 4. Mix well and incubate at 4–8 °C for 15 min (do not put on ice). 5. Wash with 10× MACS buffer volume more the initial staining volume, and centrifuge at 300 g for 5 min at 4 °C. In the meantime, equilibrate the MACS LS column (see Note 19) with 5 mL of MACS buffer. 6. Pipette off the supernatant, and resuspend cells at a concentration of 1.5 × 108 cells per mL. 7. Apply cell suspension onto the LS column. 8. Collect flowthrough and wash 3 times with MACS buffer, and collect the effluent which is the autologous CD14 negative fraction. This fraction can be frozen according to the BAMBANKER protocol to be later used for CD4+ T cell isolation in the antigen presentation assay (see Subheading 3.7 and see Note 20). 9. Remove column from the magnet, place column in a fresh tube, add 5 mL MACS buffer, insert plug, and flush out positive fraction using the plug. Wash with PBS. Centrifuge at 700 g for 3 min at 4 °C. Keep pellet in 3 mL (up to 15 mL) of D10 medium. Count cells and freeze or plate. 10. The CD14+ monocytes are plated using the appropriate plate for the experiment: • Immunofluorescence: 2.5–5 × 105 cells in a well of a 24-well plate in a final volume of 1 mL of macrophage medium • Western blot: 1 × 106 cells in a well of a 12-well plate with 1 mL of macrophage medium • Antigen presentation assay: 1.5 × 105 cells in a well of a 48-well plate with 500 μL of macrophage medium

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11. GM-CSF (1000 U/ml) is freshly added on day 0 as well as day 3 and 5 when macrophage medium is replaced by fresh medium. 12. After 6 days of culture, the monocytes have differentiated into macrophages and are ready to be used for experimentation until day 10 post isolation. 3.1.5 Isolation of Autologous CD4+ T Cells for Antigen Presentation

1. The autologous CD14- fraction frozen in the Subheading 3.1.4 should be thawed according to BAMBANKER recommendations on the day of experimentation (see Note 21). 2. Resuspend CD14- cell pellet in 5 mL of sterile PBS before determining the cell concentration. 3. Centrifuge at 300 g for 10 min at RT. 4. Gently discard the supernatant, and resuspend the CD14- cell pellet with 80 μL of MACS buffer per 107 total cells, and add 20 μL of CD4 Microbeads for every 107 total cells. 5. Incubate 15 min at 4 °C but do not put on ice. 6. Wash by adding 1 mL of MACS buffer per 107 total cells, and centrifuge at 300 g for 10 min. 7. Discard the supernatant, and resuspend up to 108 cells with 500 μL of MACS buffer. 8. Proceed for MACS sorting according to the CD4+ isolation kit recommendation. CD4 negative fraction is discarded. 9. Resuspend the autologous CD4+ T cells with macrophage medium, and determine cell concentration. 10. Whole blood CD4+ T cells will be co-cultured with pre-stimulated macrophages for antigen presentation assays, as described in Subheading 3.7.

3.2 CRISPR/Cas9 Knock-out of ATG4B in LCLs

The number of cell lines and controls that are used determines the scaling factor of the reaction. For the generation of one LCL ATG4B knock-out cell line with one mock control (without RNA duplex addition), the scaling factor would be 2× (see Note 22). It is strongly suggested to calculate all volumes for half a reaction more than necessary (see Note 23). 1. The RNA complex is generated by mixing tracrRNA with the two gene-specific crRNAs, targeting plus and minus strands of the DNA, respectively (see Note 24). The three RNAs are mixed in equimolar concentrations, to a final concentration of 66 μM for each component. The necessary volume of RNA complex for one reaction is 3.787 μL. 2. The RNA mix is incubated at 96 °C for 5 min and then cooled down to RT (see Note 25).

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3. The Cas9 nuclease is diluted in sterile PBS to a concentration of 50 μM (stock concentration, 61 μM). Per reaction, 1 μL of diluted Cas9 is needed, corresponding to 50 pmol. 4. The ribonucleoprotein (RNP) complex is generated by gently mixing the Cas9 enzyme (1 μL, step 3) with the RNA complex (3.787 μL, step 1) in a molar ratio of 1:5 (for the mock control, sterile PBS is used instead of RNA duplex). This corresponds to 50 pmol Cas9 enzyme and 250 pmol tracrRNA-crRNA complex. The final volume per reaction is 4.787 μL. Then, the RNP mixture is incubated at RT for 15 min. 5. To prepare 16.5 μL of P3 nucleofection medium per reaction, 2.97 μL nucleofection supplement is added to 13.53 μL nucleofection solution (P3 Primary Cell 4D-Nucleofector Kit). 6. Then, the RNP solution is added to the wells of a black 16-well strip (4.78 μL per well, which is provided by the Lonza Kit). It is possible to re-use these wells; however, sterilization with 70% ethanol for 10–20 min is necessary to prevent crosscontamination between cell lines. 7. After collecting and counting the LCLs, they are washed once in sterile PBS (centrifugation at 400 g, 5 min, 4 °C). For each reaction, 5 × 105 cells are transferred to a new Eppendorf tube and pelleted (centrifugation at 400 g, 5 min, 4 °C). 8. The supernatant is removed completely by careful aspiration. Then, the cells are resuspended in 15 μL P3 nucleofection medium (step 5) and transferred to the prepared 16-well strip. 9. The samples are electroporated immediately using the 4D Nucleofector technology (Lonza) (choose the correct wells, Solution P3, Code DN-100). 10. Directly after electroporation, 50 μL of warm R10 medium is added to each well, and the cells are left to rest for 10 min at RT. Then, each 16-well strip is completely transferred to a well in a flat-bottom 96-well plate. The 16-well strips are washed with 80 μL of R10, which is also transferred to the respective 96-well plate. The cells are incubated at 37 °C, 5% CO2. 11. The plate should not be disturbed for the first 3 to 4 days after electroporation. Then, the cells are expanded when they reach high density. 3.3 Lentivirus Transduction of Human Macrophages for ATG4 Overexpression

The autophagic protein ATG4 is the only ATG protease that plays a dual role during the autophagy pathway. Indeed, ATG4 is responsible for the cleavage of pro-LC3 into the cytosolic form LC3-I which will be later lipidated on the autophagosomal membranes, as well as it is the only ATG capable of cleaving of the lipidated LC3-II from the autolysosome. Furthermore, ATG4 has also been described to play a key role during LAP, and especially its RedOx

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regulation is involved in delaying LAPosome maturation allowing a sustained antigen presentation by APCs. Thereby, the ATG4B mutant (ATG4BC78S) insensitive for RedOx regulation is a great tool to characterize the LAP pathway during antigen presentation. 1. The human monocyte-derived macrophages are transduced with lentiviral construct for overexpression of ATG4Bwt or ATG4BC78S. 2. Primary human macrophages are infected with lentiviruses together with 6 μg/mL of polybrene, and then the plates are centrifuged at 1900 g for 45 min at 37 °C as described in Ligeon et al. [5]. 3. Twenty-four hours post-transduction, the medium is replaced, and the cells are incubated for an additional 24 h or directly used for experiments like in the case of antigen presentation assays. 3.4 Trigger LC3Associated Phagocytosis in Human Macrophages Using Candida albicans Extract

The non-canonical autophagy LAP pathway has been described to be involved in the presentation of fungal antigens on MHC class II molecules, thus becoming a key player during adaptive immune responses [5]. Therefore, to investigate the role of LAP in human antigen presenting cells (APC), we propose to take advantage of the well-described LAP trigger C. albicans extract, which is also known to be presented on MHC class II molecules to IL-17 producing helper CD4+ T cells [5].

3.4.1 Candida albicans Extract Coated Beads

C. albicans extract coated beads can be used to enhance LAP. Coated beads are prepared freshly at a final concentration of 50 μg of C. albicans extract per 108 beads. 1. Wash 108 beads in 1 mL of PBS. 2. Centrifuge the beads at 600 g for 5 min. 3. Gently discard the supernatant and resuspend the beads in 1 mL of PBS. 4. Repeat steps 4–5 three times. 5. After the last centrifugation, resuspend the beads in 375 μL of PBS. 6. Slowly add 125 μL of C. albicans extract at the initial concentration of 0.4 mg/mL to the beads. Keep this order to maximize the coupling efficiency. 7. Incubate under rotation for 2 h at RT and then rotate overnight at 4 °C.

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8. The next day, the coupled beads are centrifuged at 600 g for 5 min, the supernatant is discarded, and the beads gently resuspended in 1 mL of sterile PBS. 9. Repeat the washing step 8 five times to thoroughly remove the excess of ligands. 10. Finally the beads are counted and ready to be used to trigger LAP. 3.4.2 LAP Triggered with Candida albicans Extract Coated Beads

1. Human monocytes differentiated into macrophages as described in Subheading 3.1.4 and transduced with lentiviruses carrying either Flag-ATG4Bwt, Flag-ATG4BC78S, or the control Flag are ready to be stimulated: • With C. albicans extract coated beads to trigger LAP or with inert beads as negative control using a ratio 1:1 (bead/ macrophage) for immunofluorescence samples. • With soluble C. albicans extract 40 μg/mL for antigen presentation. • It is strongly recommended to always keep condition(s) for each transduced macrophages without C. albicans extract stimulation, which will be the negative control and used to assess the role of ATG4B during LAP. 2. Stimulated macrophages are incubated at 37 °C and 5% CO2 for up to 18 h to investigate sustained MHC class II presentation. 3. Cells can then be directly used for antigen presentation assays, fixed and processed for immunofluorescence, or lysed for a molecular analysis such as Western blot (see Subheadings 3.5, 3.6, and 3.7).

3.5 Western Blot to Validate ATG4 Knockout or Overexpression in Human Antigen Presenting Cells

1. All procedures are to be carried out at RT unless otherwise indicated. 2. Prepare a 12% resolving gel for analyzing LC3B and ATG4B: mix 2.4 mL resolving gel buffer, 4.1 mL 30% acrylamide/bisacrylamide, 29:1 solution (Bio-Rad), 3 mL H2O, 96 μL of 20% SDS, 72 μL APS, and lastly 9.6 μL of TEMED. Assemble the gel casting system, and pour 7.5 mL of the gel mixture in a 7.25 cm × 10 cm × 1.5 mm gel cassette (BioRad). Gently overlay with 100 μL isopropanol to remove bubbles and even out the gel surface. Wait for the gel to polymerize (see Note 26). 3. Next, prepare the stacking gel: mix 1.25 mL stacking gel buffer, 650 μL acrylamide, 3 mL H2O, 25 μL of 20% SDS, 18.75 μL APS, and lastly 5 μL of TEMED (see Note 27). 4. After the resolving gel has polymerized, remove the isopropanol from the gel cassette by flipping the system, and remove all

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residual isopropanol with a paper towel. Add the stacking gel mixture on top of the resolving gel, and insert a 10- or 15- well gel comb immediately into the gel, carefully checking to ensure there is no introduction of air bubbles. After polymerization, the gels can either be stored in running buffer at 4 °C for up to 2 weeks or used immediately. When the running cassette is assembled, the protein markers and samples are individually loaded into the wells of the stacking gel. 5. Perform electrophoresis at a constant voltage in SDS-PAGE running buffer. Start with 85 Volts (V) until the sample has entered the gel, and then continue at 100 V until the dye front has reached the bottom of the gel. 6. Immediately following electrophoresis, remove the gels from the glass plates with a spatula, paying attention not to rip the gel in the process. Remove the stacking gel and discard. 7. Cut a PVDF membrane to the size of the resolving gel, and activate it by immersion in 100% methanol for 5 min. Rinse once with transfer buffer. 8. Assemble the transfer stack with three filter papers, the gel, the PVDF membrane, and three more filter papers. Perform a wet transfer in transfer buffer at 110 V for 1 h and 20 min. 9. Block the membrane with blocking buffer for 1 h at RT. See Subheading 2.4.2. 10. Cut the membrane according to the molecular weight (MW) of the proteins of interest, and incubate the membranes in the corresponding primary antibody diluted 1:1000 (antiLC3B and anti-ATG4B) or 1:10000 (anti-GAPDH and antiVinculin; see Subheading 2.7.2) in PBS-T overnight at 4 °C under rotation. Primary antibodies: rabbit anti-ATG4B (MW 48 kDa), rabbit anti-LC3B (MW 15–18 kDa), and mouse anti-GAPDH (MW 35 kDa). See Subheading 2.7.2. 11. Wash with PBS-T twice for 10 min each. 12. Immerse the membrane in a solution containing the secondary antibody. Goat anti-rabbit immunoglobulin HRP-conjugate (see Subheading 2.7.2). Incubate for 1 h at RT under agitation. 13. Wash with PBS-T twice for 10 min each. Wash once with PBS for 5 min. 14. Reveal using the ECL kit following the manufacturer’s instructions. 15. Each Western blot band can then be quantified using the freely available software ImageJ. 16. The absence of the ATG4B band at 48 kDa confirms the CRISPR/Cas9 knockout of ATG4B in human B cells by Western blot (Fig. 1a). The lower ATG4B band is endogenous

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Fig. 1 Analysis of autophagy proteins by Western blot in human LCLs and macrophages. (a) LCLs were transfected with CRISPR/Cas9 and gRNA for ATG4B knock-out. Cells were then lysed in Laemmli buffer for Western blot analysis of ATG4B and LC3B protein expression. Vinculin served as a loading control. (b) Primary human macrophages were transduced with lentiviral vectors for the overexpression of Flag, FLAG-ATG4Bwt, and FLAG-ATG4BC78S and then stimulated with zymosan for 1 h up to 24 h. Cells were then lysed in Laemmli buffer for Western blot analysis of ATG4B and LC3B protein expression. Vinculin served as a loading control. The graph bar represents the level of LC3B-II/Vinculin normalized to the condition flag unstimulated 1 h

ATG4B, and the upper ATG4B band is the transduced FlagATG4B protein express (Fig. 1b). Zymosan simulation induces LAP in human macrophages, as shown by the increase in LC3B-II expression (Fig. 1b). 3.6 An Immunofluorescence Protocol to Investigate the Role of ATG4 During LAP in Human Macrophages and Its Role During Autophagy in Immortalized B Cells 3.6.1 Immunofluorescence Protocol for Adherent Cells

The same basic steps of the protocol apply to both monocytederived macrophages and primary B cells. However, several crucial steps are different due to the adherent or rather non-adherent nature of macrophages and B cells, respectively. General tips: see Notes 7–9 and 28–29.

1. CD14+ monocytes are plated directly on sterile, round 1.5 mm microscopy coverslips, placed in the well of a 24-well plate at a concentration of 2.5 × 105 cells per well (see Subheading 2.5). The cells are then treated according to the experimental outline, handling two wells for each condition and using at least

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two wells as untreated controls. Before fixation, cells are gently washed with sterile PBS twice. 2. The cells are fixed in 4% PFA (300 μL per well) for 15 min at RT (see Note 30). 3. The fixed cells are washed three times with PBS. This step and all further washing steps are performed with 500 μL PBS per well for 5 min each time. 4. Permeabilization is performed by adding 300 μL of 0.1% of Triton X-100 in PBS for 5 min at RT. Then the cells are washed three times with PBS as indicated in step 2. 5. Afterward, incubation with 300 μL of blocking buffer (see Subheading 2.5) per well is performed for 1 h at RT (see Note 31). 6. The primary antibody is diluted 1:100 (anti-ATG4B) or 1:500 (anti-LC3B) in blocking buffer (see Subheading 2.5/2.7.2 and Note 8) and added directly to the well for 1 h at RT. A sufficient volume is added to cover the coverslip completely (200 μL); to use a reduced volume, please see Note 32. For co-staining of different autophagy-related proteins or other protein target(s), see Note 33. Then, the cells are washed three times with PBS for 5 min each time. 7. The secondary antibody is diluted 1:500 in blocking buffer as well (see Subheading 2.5/2.7.2) and added to the cells as described in Step 5. Incubation is performed for 1 h at RT in the dark. Then, the cells are washed again three times with PBS for 5 min each time. 8. To stain cell nuclei, DAPI staining solution is added (1:5000 in PBS; see Subheading 2.5) and incubated for 5 min at RT in the dark. Finally, the cells are washed three times with PBS for 5 min each. 9. For mounting, one drop of Dako mounting medium (or 5 μL) is placed on a microscopy specific slide. The coverslip is then removed from the well of the 24-well plate and placed upside down on the liquid. The slides are left to dry overnight at RT in the dark (see Note 34). 3.6.2 Immunofluorescence Protocol for Cells in Suspensions

1. LCLs are treated according to the experimental setup. Then, cells are counted; resuspended in cold, sterile PBS; and plated onto sterile round coverslips placed into a well of 24-well plate at a density of 1 × 106 cells. 2. The cells are incubated at RT for 45 min, allowing them to settle to the bottom of the wells. The PBS is removed very carefully. As the cells are not adherent, thus not actually physically attached to the glass surface, they can be removed very

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easily. Only by pipetting very slowly and carefully, it is possible to leave the cells settled (see Note 29). 3. The cells are fixed in 4% PFA (300 μL per well) for 15 min at RT (see Note 30). 4. The fixed cells are washed once with PBS. This step and all further washing steps are performed with 500 μL PBS per well for 5 min each time. 5. Permeabilization is performed by adding 300 μL of 0.1% of Triton X-100 for 5 min at RT. Then, the cells are washed once with PBS as indicated in step 2. 6. Afterward, incubation with 300 μL of blocking buffer (see Subheading 2.5) per well is performed for 1 h at RT (see Note 31). 7. The primary antibody is diluted in blocking buffer (see Subheading 2.5 and Note 8) and added directly to the well for 1 h at RT. A sufficient volume is added to cover the coverslip completely (200 μL). For co-staining of different autophagyrelated proteins or other protein target(s), see Note 33. Then, the cells are washed twice with PBS for 5 min each. 8. The secondary antibody is diluted in blocking buffer as well (dilution 1:500; see Subheading 2.7.2) and added to the cells as described in step 5. Incubation is performed for 1 h at RT in the dark. Then, the cells are washed again twice with PBS for 5 min each. 9. To stain cell nuclei, DAPI staining solution is added (1:5000 in PBS; see Subheading 2.5) and incubated for 5 min at RT in the dark. Finally, the cells are washed three times with PBS for 5 min each. 10. For mounting, one drop of Dako mounting medium (or 5 μL) is placed on a microscopy specific slide. The coverslip is then removed from the well of the 24-well plate and placed upside down on the liquid. The slides are left to dry overnight at RT in the dark (see Note 34). 3.6.3 Image Acquisition and Analysis

1. Immunofluorescent images are acquired with a confocal laser scanning microscope (Leica Stellaris 5), using a 63× oil immersion objective (1.4 NA). Excitation is performed at 405 nm for DAPI fluorescence (blue emission), 488 nm for GFP or Alexa Fluor 488 (green emission), and 543 nm for Alexa Fluor 555 (red emission). After acquiring images with the Leica software (Leica), ImageJ is used for channel merging and image analysis. 2. A prominent representation of autophagy flux is the quantification of LC3 dots per cell, referring to the number of autophagosomes or LAPosomes per cell according to the experiment

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Fig. 2 The lack of ATG4B in human LCL impedes autophagosome formation. (a–b) LCLs CRISPR control (CTRL) (a) and knocked-out for ATG4B using the CRISPR/Cas9 technology (b) were fixed on glass cover slides and immunostained for LC3-AlexaFluor488, ATG4B-AlexaFluor555, and DAPI (nuclei). Scale bar, 5 μm. (c) Violin plot represents the average number of LC3 dots per cell quantified for CRISPR CTRL LCLs and ATG4B-KO LCLs. For both conditions, seven images with 20 to 30 cells were quantified. Each data point represents one representative image

set-up and stimulation. Using the ImageJ threshold analysis tool, the number of LC3 dots, as well as their average size, can be quantified for each image. Using the DAPI channel to determine the number of nuclei in each image, the average number LC3 dots per cell can be quantified. 3. In Fig. 2, images from a representative immunofluorescence experiment are shown, staining for ATG4B as well as LC3B in CRISPR CTRL LCLs and ATG4B-KO LCLs. In the CTRL LCLs, ATG4B and LC3B show a dotty staining corresponding to autophagosomes (Fig. 2a). In ATG4B-KO LCLs, the ATG4B staining is almost completely absent, and the LC3B staining appears to be dispersed in the cytoplasm (Fig. 2b, c). This indicates that the ATG4B-KO was successful as a defect in LC3B processing and autophagy is observed. 3.7 Assess the Involvement of ATG4 in LAPosome Stabilization to Allow Macrophages to Sustain MHC Class II Antigen Presentation

To assess how fungal antigen presentation by MHC class II molecules can be modulated by the LAP pathway and especially the importance of delaying the maturation of the LAPosomes, it is possible to use a Candida albicans specific CD4+ T cell clone as described in Ligeon et al. (2017). However, here we proposed an alternative assay by using whole blood CD4 + T cell, which will overcome the limitation of using a specific CD4+ T cell clone (see Note 35). Moreover, circulating C albicans specific T cells can be detected in almost all human healthy donors [15] and, thus, will provide more physiological antigen presentation conditions to study the involvement of LAP in antigen presentation by macrophages to their autologous CD4+ Th17 cells.

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1. Macrophages plated in 48-well plate at a cell concentration of 1.5 × 105 which were transduced with lentiviruses carrying Flag, Flag-ATG4wt, or Flag-ATG4BC78S mutant for 24 h as described in Subheading 3.3 (see Note 36). 2. Pulse the transduced macrophages with 80 μg/mL of soluble C. albicans extract for 4 h at 37 °C and 5% CO2. Keep four wells per condition without stimulus for control: • Two wells do not receive stimulus or CD4+ T cells as controls to show that the level of secreted cytokines is not only due to the added macrophages. • Two wells are co-cultured with CD4+ T cells and used as negative controls to show that the activation of CD4+ T cells is due to the presentation of fungal antigen peptides by the macrophages and not due to them being transduced. 3. Wash antigen-pulsed macrophages with warm macrophage medium 3 times to remove excess of stimuli. 4. Establish two conditions for each transduced macrophage: • 0 h = Condition without resting time: Directly add the autologous whole CD4+ T cells. • 18 h = Condition with resting time: Incubate macrophage at 37 °C and 5% CO2 for 18 h before adding the autologous CD4+ T cells. 5. Add whole blood CD4+ T cells prepared as described in Subheading 3.1.5 to the stimulated macrophages at a ratio 2:1 (see Note 21). 6. As a control, add the same number of CD4+ T cells to four empty wells, two wells will be treated with Phorbol 12-myristate 13-acetate (PMA)/Ionomycin as positive controls, and two wells will remain without treatment and used as negative controls. 7. After 5 days of co-culture at 37 °C and 5% CO2, centrifuge the plate at 500 g at 4 °C for 5 min prior to gently harvesting the supernatant (see Note 37). 8. Supernatants are diluted and plated directly on previously coated ELISA plates (see Note 38): Tested for IL-17A, IFN-ɣ, and V-plex TH17 panel 1 (human) kits by ELISA: • ELISA IFN-ɣ: ELISA plates are coated according to the manufacturer’s instructions. • ELISA IL-17A and V-plex TH17 panel human kit: the microplate strips are ready to be used according to the manufacturer’s instructions.

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9. Perform the three ELISA assays using the appropriate kits and following the manufacturer’s instructions. Recombinant human IL-17A and IFN-ɣ provided by the ELISA kits are used as standards. 10. Detection of IL-17A and IFN-ɣ in the supernatant of co-culture indicates that memory CD4+ T cells were activated by recognition of the MHC class II restricted fungal peptides presented by the macrophages. Indeed IL-17A and IFN-ɣ are two cytokines secreted by activated CD4+ T cells and play a role in adaptive immunity (see Note 10).

4

Notes 1. It is possible to use 2-day-old buffy coat. When handling human blood, appropriate biosafety rules should be respected as well as personal protective equipment should be worn. 2. It is important for thawed FBS to be heat inactivated at 56 °C for exactly 30 min. This ensures that any complement in the serum is denatured. Be sure to swirl and mix the FBS bottles thoroughly to ensure even temperature distribution and to reduce precipitation of FBS. 3. Working with EBV requires a biosafety level (BSL) 2 laboratory and appropriate training. 4. The selection of CD4+ T cells is a positive selection, and CD4+ T cells are retained in the MACS column during selection. 5. When designing crRNAs with IDT, the on- and off-target scores of the respective crRNAs are used as main indicator for target specificity. A combination of a very low off-target score with a very high on-target score is desirable. 6. Although there are risks associated with working with lentiviruses, lentiviral vector may be safely handled using Biosafety level 2 (BSL-2) conditions. In the system used here, the vector system is split into different plasmids, reducing the possibility of producing a replication-competent virus. 7. Round microscopy glass coverslips (1.5 mm) are sterilized with 70% ethanol. Ideally, the coverslips are placed in 24-well plate and incubated in a sufficient volume (approx. 500 μL) of 70% ethanol overnight or at least for 20 min at RT. Before plating cells, the coverslips are washed with sterile PBS twice. While washing, any traces of ethanol should be completely removed by aspiration. 8. For the initial establishment of a primary antibody, try the dilution recommended by the company. As an alternative we also recommend trying an initial dilution of 1:100. Afterward,

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the antibody concentration can be refined and decreased by titration if a very strong staining or a lot of background signal is observed with the initial dilution. 9. To choose appropriate fluorophore-coupled secondary antibodies, consider the limitations of your confocal microscope regarding laser and filter sets. Further, make sure to choose compatible fluorophores with a minimum or non-overlapping emission spectra for co-staining (e.g., a very well-established combination is DAPI (emits in the blue range), Alexa-488 (emits in the green range), and Alexa-555 (emits in the red range)). 10. IFN-ɣ and IL-17A secretions are used as direct readouts to assess the ability of human macrophages to present MHC class II restricted fungal peptides. Indeed, these two cytokines are secreted by activated CD4+ T cells, which play an important role in adaptive immunity. 11. Secondary antibodies must be carefully chosen according to the species of the primary antibody as well as the experimental set-up in case of performing several stainings at once. 12. Latex beads are known to be inert and not to trigger LAP, thus can be used as negative control. Latex beads are reconstituted by resuspending 1 × 109 inert beads in 1 mL PBS supplemented with 0.5% BSA into a protein low-binding 1.5 mL Eppendorf, and then incubate 1 h rotating at RT. Centrifuge 600 g at 5 min and then discard the supernatant. Wash 3 times with 1 mL PBS and then finally count the beads. 13. For some experiments it might be useful to produce GFP-positive EBV. For further information on this topic, see [16]. 14. The virus pellet may be very small or not visible to the naked eye. Thus, marking the theoretical location of the pellet on the outside of the tube helps to keep track of the tube orientation. 15. Four phases can be observed, the bottom layer is the red phase corresponding to the red blood cell layer, above is the FicollPaque which is transparent, the third phase is very thin and yellow corresponding to the PBMCs, and finally the top phase is the plasma. 16. CD19+ cells are magnetically retained inside the column and can be eluted as the positively selected cell fraction. To improve CD19+ cell purity, the magnetic separation can be repeated by passing the positively selected fraction through a second, new column. 17. Plate several wells to have replicates if one well does not grow out. We recommend a minimum of four wells. If the number of

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CD19+ B cells is not sufficient, the cell number can be decreased to 100,000 cells per well. However, this could reduce the outgrowth speed. 18. Feeder cells can be used to support vulnerable cells and yield a better outgrowth efficacy. For this purpose, CD40L transgenic fibroblasts are irradiated for 660 s at 17 mA, 250 V, and plated in a 96-well plate at a density of 125,000 cells per well. The CD19+ B cells are then added as described in step 2 of the Subheading 3.1.3 to the same wells. Ideally, both feeder cells and CD19+ B cells are added in 50 μL R10 to yield 100 μL per well before virus addition. When passaging the cells, new feeder cells are added to the new wells until the LCLs are sufficiently established to support their own growth. 19. If the concentration of PBMCs is higher than 2 × 108 total cells, it is recommended to use two columns to obtain a better separation (usually 10% of cells are CD14+). 20. The CD14 negative fraction is kept and frozen at the concentration of 10 × 106 cells/mL following BAMBANKER protocol and used later on to select the autologous CD4+ T cells. 21. It is essential to use the CD14- fraction that comes from the exact same buffy coat (autologous) as the monocyte-derived macrophages to avoid alloresponses. 22. Additional to a non-crRNA control, it is also recommended to include a non-Cas9 control or a condition with an unspecific crRNA that does not target any region in the human genome. 23. As the volumes in this protocol are very small, make sure to spin all tubes down before pipetting, and try to work as precisely as possible. It is not recommended to use less volume than indicated in this protocol. 24. By using two different crRNAs, targeting two different regions in the ATG4B gene on the plus and minus strand of the DNA, knock-out efficacy can be increased substantially compared to only using one gene-specific crRNA. 25. Let the sample cool down slowly by simply leaving it on the bench. Do not try to speed up cool-down. 26. Before casting the resolving gel, assemble the gel casting system, and fill the glass plates with H2O, and leave it for 5 min to ensure there are no leaks. If there is no leak, remove the water by inverting the system; otherwise reassemble the system anew. 27. By adding a few flakes of bromophenol blue to the stacking gel mixture, the blue color adds visual contrast to the wells, facilitating the accurate loading of the gel. 28. The following control conditions should be included to be able to interpret the results correctly:

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(a) Experimental controls should consist of wild-type (parental for CRISPR/Cas9 or non-transduced macrophages) and non-treated cells. (b) Include one coverslip that has only been incubated with blocking solution, without primary or secondary antibody. This will allow to determine the autofluorescence of the cells. (c) Include one coverslip that has only been incubated with the secondary antibody and not the primary antibody. (d) If co-staining is performed, include single staining for every marker, and check for spillover through other channels. For example, analyze the emission of Alexa-488 in the red channel when it is excited with the red laser, and observe Alexa-555 in the green channel. The staining is successful if no significant signal is detected in these channels for these fluorophores. 29. Pipetting carefully and gently throughout the entire protocol is crucial when handling non-adherent cells like B cells. Even after fixation, B cells can be detached from the glass coverslips very easily; thus always handle the coverslip with a lot of care and as gently as possible. 30. After fixation, a sterile cell culture hood is not necessary anymore, and the cell can be further processed on a laboratory bench. 31. Blocking incubation time can be reduced, but make sure to block at least for 30 min. 32. When working with adherent cells, the necessary volume of antibody can be reduced by placing a drop of the diluted antibody solution (30 μL) on parafilm, removing the coverslips from their wells, and inverting them on the antibody drop. Like this, the cells are in direct contact with the antibodies. For washing, the coverslips are placed back into the 24-well plate (inverting again so the cells face upward). Please do not perform this step with non-adherent cells as the cells will detach and will be lost. 33. Co-staining of different proteins is possible by using different antibody species for different proteins. For example, a rabbitLC3B primary antibody can be combined with a mouseATG4B primary antibody. To separate the protein signals into different fluorescent channels, two secondary antibodies are selected that are coupled to distinct fluorophores and that have defined specificities, however from the same species to avoid cross-reactivity, for example, a goat anti-mouse immunoglobulin Alexa-488 antibody and a goat anti-rabbit immunoglobulin Alexa-555 antibody.

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34. For long-term storage, the coverslips can be sealed with nail polish after the mounting medium is dry and stored at 4 °C in dark for several weeks. 35. Whole CD4+ T cells presumably contain polyclonal and polyepitope specific T cells for C. albicans from healthy donors. Thereby, their activation shown by the secretion of cytokines is due to the presentation of fungal antigen peptides via MHC class II molecules by the macrophages. The use of polyclonal CD4+ T cells allowed to overcome the limitation of using Candida albicans specific CD4+ T cell clones that have a distinct MHC class II restriction and need to be HLA matched to the macrophage preparations, as well as of the fitness of the respective clone after re-expansions. 36. For correct interpretation of the results, the following controls should be included: (a) Transduce at least 8 wells of macrophages for each lentivirus construct. (b) Keep 8 wells of macrophages without lentiviruses transduction. 37. The supernatant of the co-culture will be used to detect cytokine production. Thus, we recommend using a small volume of culture medium, which will facilitate the detection of the cytokines. If necessary, the supernatant can be frozen for further analysis; however we do not recommend having several freeze– thaw cycles. 38. To allow a more accurate determination of the concentration of the cytokine secreted, the supernatant can be diluted from 1: 2 to 1:10 in PBS.

Acknowledgments Research in our laboratory is supported by Cancer Research Switzerland (KFS-4962-02-2020), CRPP-ImmunoCure and HMZ ImmunoTargET of the University of Zurich, the Sobek Foundation, the Swiss Vaccine Research Institute, Roche, Novartis, the Swiss MS Society (2021-09), the Vontobel Foundation (1253/ 2022), and the Swiss National Science Foundation (310030_204470/1, 310030L_197952/1, and CRSII5_180323).

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References 1. Ghosh D, Jiang W, Mukhopadhyay D, Mellins ED (2021) New insights into B cells as antigen presenting cells. Curr Opin Immunol 70:129– 137. https://doi.org/10.1016/j.coi.2021. 06.003 2. Jakubzick CV, Randolph GJ, Henson PM (2017) Monocyte differentiation and antigenpresenting functions. Nat Rev Immunol 17: 349–362. https://doi.org/10.1038/nri. 2017.28 3. Mu¨nz C (2016) Autophagy beyond intracellular MHC class II antigen presentation. Trends Immunol 37:755–763. https://doi.org/10. 1016/j.it.2016.08.017 4. Mizushima N, Yoshimori T, Ohsumi Y (2011) The role of Atg proteins in autophagosome formation. Annu Rev Cell Dev Biol 27:107– 132. https://doi.org/10.1146/annurevcellbio-092910-154005 5. Ligeon L-A, Pena-Francesch M, Vanoaica LD et al (2021) Oxidation inhibits autophagy protein deconjugation from phagosomes to sustain MHC class II restricted antigen presentation. Nat Commun 12:1508. https:// doi.org/10.1038/s41467-021-21829-6 6. Paludan C, Schmid D, Landthaler M et al (2005) Endogenous MHC class II processing of a viral nuclear antigen after autophagy. Science 307:593–596. https://doi.org/10. 1126/science.1104904 7. Romao S, Gasser N, Becker AC et al (2013) Autophagy proteins stabilize pathogencontaining phagosomes for prolonged MHC II antigen processing. J Cell Biol 203:757– 7 6 6 . h t t p s : // d o i . o r g / 1 0 . 1 0 8 3 / j c b . 201308173 8. Sanjuan MA, Dillon CP, Tait SWG et al (2007) Toll-like receptor signalling in macrophages links the autophagy pathway to phagocytosis. Nature 450:1253–1257. https://doi.org/10. 1038/nature06421 9. Zheng X, Yang Z, Gu Q et al (2020) The protease activity of human ATG4B is regulated

by reversible oxidative modification. Autophagy 16:1838–1850. https://doi.org/10. 1080/15548627.2019.1709763 10. Ma J, Becker C, Lowell CA, Underhill DM (2012) Dectin-1-triggered recruitment of light chain 3 protein to phagosomes facilitates major histocompatibility complex class II presentation of fungal-derived antigens. J Biol Chem 287:34149–34156. https://doi.org/ 10.1074/jbc.M112.382812 11. Nilsson K, Klein G, Henle W, Henle G (1971) The establishment of lymphoblastoid lines from adult and fetal human lymphoid tissue and its dependence on EBV. Int J Cancer 8: 443–450. https://doi.org/10.1002/ijc. 2910080312 12. Gurwitz D (2016) Human lymphoblastoid cell lines: a valuable tool for personalized medicine research. Biochemist 38:6–9. https://doi.org/ 10.1042/BIO03801006 13. Delecluse H-J, Hilsendegen T, Pich D et al (1998) Propagation and recovery of intact, infectious Epstein–Barr virus from prokaryotic to human cells. Proc Natl Acad Sci 95:8245– 8250. https://doi.org/10.1073/pnas.95.14. 8245 14. Schmid D, Pypaert M, Mu¨nz C (2007) Antigen-loading compartments for major histocompatibility complex class II molecules continuously receive input from autophagosomes. Immunity 26:79–92. https://doi.org/ 10.1016/j.immuni.2006.10.018 15. Bacher P, Hohnstein T, Beerbaum E et al (2019) Human anti-fungal Th17 immunity and pathology rely on cross-reactivity against Candida albicans. Cell 176:1340–1355.e15. https://doi.org/10.1016/j.cell.2019.01.041 16. Antsiferova O, Mu¨ller A, R€amer PC et al (2014) Adoptive transfer of EBV specific CD8+ T cell clones can transiently control EBV infection in humanized mice. PLoS Pathog 10:e1004333. https://doi.org/10. 1371/journal.ppat.1004333

Chapter 22 Visualizing Phagocytic Cargo In Vivo from Engulfment to Resolution in Caenorhabditis elegans Gholamreza Fazeli, Julia Frondoni, Shruti Kolli, and Ann M. Wehman Abstract The nematode Caenorhabditis elegans offers many experimental advantages to study conserved mechanisms of phagocytosis and phagocytic clearance. These include the stereotyped timing of phagocytic events in vivo for time-lapse imaging, the availability of transgenic reporters labeling molecules involved in different steps of phagocytosis, and the transparency of the animal for fluorescence imaging. Further, the ease of forward and reverse genetics in C. elegans has enabled many of the initial discoveries of proteins involved in phagocytic clearance. In this chapter, we focus on phagocytosis by the large undifferentiated blastomeres of C. elegans embryos, which engulf and eliminate diverse phagocytic cargo from the corpse of the second polar body to cytokinetic midbody remnants. We describe the use of fluorescent time-lapse imaging to observe the distinct steps of phagocytic clearance and methods to normalize this process to distinguish defects in mutant strains. These approaches have enabled us to reveal new insights from the initial signaling to induce phagocytosis up until the final resolution of phagocytic cargo in phagolysosomes. Key words Phagocytosis, Phagosome maturation, Phagosome-lysosome fusion, Phagolysosome resolution, Time-lapse imaging, Fluorescent reporters, Caenorhabditis elegans embryos

1

Introduction Cells constantly clear their environment of harmful substances, whether self-produced or from external sources. Phagocytosis is the main cellular process that disposes of extracellular material, sequestering cell debris and pathogens in membrane-bound compartments for degradation. Phagocytosis is not only essential to clear exogenous pathogens during infection, but is also important for tissue homeostasis through the clearance of dying cells and cell fragments [1, 2]. Phagocytosis can be performed by professional phagocytes, such as macrophages, but also by diverse cell types, even undifferentiated cells, which use phagocytosis to engulf and degrade a variety of phagocytic cargos [3, 4]. The mechanisms of phagocytosis are conserved in animals from nematodes to mammals. Indeed, programed cell death and the

Roberto J. Botelho (ed.), Phagocytosis and Phagosomes: Methods and Protocols, Methods in Molecular Biology, vol. 2692, https://doi.org/10.1007/978-1-0716-3338-0_22, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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subsequent removal of cell corpses were discovered in the nematode model organism Caenorhabditis elegans, and C. elegans studies revealed many of the proteins involved in phagocytosis [5]. During the invariant development of C. elegans, many cells are programmed to die, including 113 of the 628 somatic cells born during embryogenesis, the linker cell that directs migration of the male gonad during larval development, almost half of adult germ cells, and two meiotic polar bodies that form during oocyte development [5–9]. In addition, C. elegans cells also engulf cell debris, including cell fragments such as residual bodies released by spermatocytes [10]. Even organelles are degraded by phagocytosis after serving their purpose, such as midbody remnants that contain the cytokinetic machinery released from the intercellular bridge of dividing cells [11]. In this chapter, we describe how time-lapse microscopy can be used to study the individual steps of phagocytosis and phagocytic clearance in early C. elegans embryos, focusing on the removal of the corpse of the second polar body and midbody remnants as examples of phagocytic cargos [6, 11–16]. These endogenous cargos are well suited for studying phagosome maturation because the phagocytic events occur with stereotyped timing and positioning. Furthermore, they are ~1–3 μm in diameter, while the engulfing cell can be >10 μm in diameter. We demonstrate how observing the dynamics of these cargos in early embryos can be used to discover defects in phagocytic processes in vivo, taking advantage of available mutant and transgenic worm strains, as well as the ease of knocking down specific genes by feeding worms doublestranded RNA [17]. These advantages have allowed this system to provide novel insights into cargo recognition, engulfment, phagosome maturation, fusion with lysosomes, phagolysosome tubulation, and the final resolution of phagolysosome cargo.

2

Materials

2.1 Worm Strains and Maintenance

1. C. elegans nematode worms. The wild-type Bristol N2 C. elegans strain in addition to mutant worm strains (see Note 1), transgenic worm strains (see Notes 2 and 3), or bacterial strains can be acquired from the Caenorhabditis Genetic Center (CGC) at the University of Minnesota (https://cgc.umn. edu). Deletion mutants are also available from the Japanese National Bioresource Project (https://shigen.nig.ac.jp/c. elegans). Worm strains are also shared among individual labs by request. 2. Temperature-regulated chamber or room to grow worms (see Note 4).

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Table 1 C. elegans promoters Expression

Promoter(s)

Germ line/early embryos

pie-1 mex-5

Coelomocytes

cc1

Phagocytes

ced-1

Heat shock-inducible

hsp-16.2 hsp-16.41

Strong ubiquitous

eft-3

Additional promoters can be found on WormBase.org

Table 2 Plasma membrane reporters Transgene

Localization

References

CED-1::GFP

C. elegans engulfment receptor on plasma membrane

[46]

GFP::PH(PLC1∂1) mCh::PH(PLC1∂1)

Mammalian PI4,5P2-binding domain on plasma membrane

[47, 48]

pie-1::GFP::ZF1::PH (PLC1∂1) pie-1::mCh::PH (PLC1∂1)::ZF1

Mammalian PI4,5P2-binding domain on plasma membrane in germ line and early embryos, but fluorescence becomes restricted to polar bodies in anterior cells after the 4-cell stage

[6, 49]

pie-1::mCh::PH (PLC1∂1)::CTPD

Mammalian PI4,5P2-binding domain on plasma membrane in germ line and early embryos, but fluorescence becomes restricted to polar bodies after the 2-cell stage

[36]

3. Nematode Growth Medium (NGM) plates (see Subheading 3.1.2) [18]. 4. E. coli strain OP50 (see Note 5) cultured in LB Broth (see Subheading 3.1.1) [19]. Common promoters and markers for phagocytic cargo are listed in Tables 1, 2, 3, and 4. 2.2 RNA Interference (RNAi) Reagents

1. NGM Lite plates for knockdown experiments using RNAi (see Subheading 3.1.3). 2. Bacterial strains for reverse genetics cultured in LB Broth (see Subheading 3.1.1). Strains can be obtained from genomic (JA or sjj, first generated in the lab of Julie Ahringer [20]) or cDNA (mv, generated in the lab of Marc Vidal [21]) RNAi libraries. Alternatively, gene-specific RNAi constructs can be subcloned into L4440 (Addgene plasmid #1654) and transformed into HT115 bacteria [17].

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Table 3 Midbody remnant reporters Transgene

Localization

References

NMY-2:: GFP NMY-2:: mKate2 NMY-2:: mCherry

C. elegans non-muscle myosin on the cell cortex, enriched in cytokinetic rings and midbody remnants

[50–52]

NMY-2:: GFP::ZF1

C. elegans non-muscle myosin on the cell cortex, enriched in cytokinetic rings and midbody remnants, but fluorescence becomes restricted to midbody remnants in anterior cells after the 4-cell stage

[51]

mCherry:: ZEN-4

C. elegans centralspindlin subunit localized to the central spindle and midbody remnants

[53]

GFP::MVB12

C. elegans ESCRT-I subunit trapped in midbody remnants

[54]

Table 4 Chromosomal reporters Transgene

Localization

References

GFP::H2B mCh::H2B

C. elegans histone

[55, 56]

pie-1::mKate2:: ZF1::npp-5

C. elegans nuclear pore protein in nuclear envelope of germ line and early embryos during interphase. During cell division, localizes around chromosomes. After the 4-cell stage, becomes restricted to polar bodies in anterior cells

[6]

pie-1::ZF1:: mCh::his-15

C. elegans histone in germ line and early embryos, but fluorescence becomes restricted to polar bodies in anterior cells after the 4-cell stage

[36]

2.3

Microscopy

1. M9 or egg salts buffer (see Subheadings 3.1.4 and 3.1.5). 2. Standard microscope slides and coverslips for imaging on agarose pads or μ-Slide 4-well glass bottom slide (ibidi) for imaging in wells. 3. Agarose pads to mount worms and embryos under microscope (see Subheading 3.1.6) [22]. 4. Dissection microscope to handle worms and prepare embryos, ideally with oblique transillumination using a mirror. 5. Worm pick: A 32 gauge platinum wire flattened on one end and fixed inside a Pasteur pipette using a flame.

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6. No. 15 scalpel blade in No. 3 holder or two gauge 16 needles to dissect gravid worms. 7. Watch glass (see Note 6). 8. Glass mouth pipette or albumin-coated plastic pipette to transfer embryos. 9. Vaseline/petroleum jelly to seal slides for imaging longer than 30 min. 10. A compound microscope to visualize fluorescence and DIC/Nomarski. 11. Software to view, acquire, analyze, and quantify imaging data.

3

Methods

3.1 Preparing Buffers and Media

1. To prepare 1 liter of LB medium, add 10 g of Bacto-tryptone, 5 g yeast extract, and 5 g NaCl to 800 mL of water.

3.1.1

2. Adjust pH to 7.0 and volume to 1 L.

Luria Broth (LB)

3. Sterilize by autoclaving. 3.1.2 Nematode Growth Media (NGM) Plates

NGM plates are used for maintenance of worm cultures on E. coli bacteria. 1. For 1 L of medium, add 3 g NaCl, 15 g agar, and 2.5 g tryptone to ~970 mL double distilled H2O, and autoclave. 2. Let cool to ~55 °C, and then add the following sterile additives: 1 mL of filter-sterilized 5 mg/mL cholesterol in ethanol (EtOH), 1 mL of autoclaved 1 M MgSO4, 25 mL of autoclaved K-Salts (0.28 M K2HPO4 + 0.72 M KH2PO4, pH 6.0), 1 mL of filter-sterilized 1 mg/mL uracil, and 1 mL of autoclaved 1 M CaCl2. 3. Pour in 60 × 15 mm plastic dishes and let dry overnight. 1 L will produce ~80 plates. 4. Seed plates with 200 μl of overnight culture of E. coli OP50 strain grown in LB medium. 5. Let bacteria grow on plates overnight at room temperature. Seeded plates can be stored at 4 °C for months.

3.1.3

NGM Lite Plates

NGM Lite plates are used for RNA interference (RNAi) experiments where specific mRNA levels are knocked down by feeding worms bacteria expressing double-stranded RNA (dsRNA). This is typically performed using the tetracycline-resistant nuclease-deficient HT115 E. coli strain transformed with a variant of the L4440 plasmid carrying ampicillin resistance and the gene of interest flanked by T7 promoters, which allows dsRNA expression to be induced by ß-lactose or IPTG.

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1. For 1 L, add 1.5 g NaCl, 20 g agar, and 4 g tryptone to ~950 mL double distilled H2O, and autoclave. 2. Let cool to ~55 °C, and then add the following sterile additives: 1 mL cholesterol from filter-sterilized 5 mg/mL stock in EtOH, 1 mL from 1 M MgSO4, 25 mL K-Salts (see above), 1 mL of 1 M CaCl2, 1 mL ampicillin from 100 mg/mL stock, 1 mL tetracycline from 12.5 mg/mL stock in 70% EtOH, and 10 mL ß-lactose from 20% stock solution. 3. Pour in 60 × 15 mm plastic dishes. 1 L will produce ~80 plates. 4. Let dry overnight and store at 4 °C. 5. Plates are seeded before experiments with 200 μL of genespecific RNAi culture or of empty vector control culture. 3.1.4

M9 Buffer

M9 buffer can be used to transfer or culture all stages of worms [19]. 1. Dissolve 3 g of KH2PO4, 6 g of Na2HPO4, and 5 g of NaCl in 1 L of water, and autoclave. 2. After cooling, add 1 mL of sterile 1 M MgSO4.

3.1.5

Egg Salts

Egg salts is a buffer for handling young or mutant embryos sensitive to osmolarity changes [23]. 1. Mix 18.8 mL 5 M NaCl, 32 mL 1 M KCl, 2.8 mL 1 M MgCl2, 2.8 mL 1 M CaCl2, 4 mL 0.5 M HEPES pH 7.5, and 939.6 mL ddH2O. 2. Filter-sterilize.

3.1.6

Agarose Pads

1. Prepare a 4% agarose solution in water and boil until clear (see Note 7). Incubate at 65 °C until needed (see Note 8). 2. Place three slides in parallel on a flat surface. 3. Add a drop of agarose solution to the middle slide, and immediately cover the drop with a fourth slide in a perpendicular orientation. 4. After the agarose has solidified, remove the fourth slide by sliding it to the side and exposing the agarose pad.

3.2 Mounting Embryos for TimeLapse Imaging

3.2.1 Mounting Embryos for Time-Lapse Imaging (Agarose Pad)

We suggest two methods for mounting embryos, depending on the illumination used. Mounting worms on agarose pads (Subheading 3.2.1) allows imaging on upright or inverted widefield or confocal microscopes but compresses the embryos (see Note 9). Mounting worms in well slides (Subheading 3.2.2) is useful for light-sheet imaging on inverted microscopes. 1. Pick three to four gravid hermaphrodite worms into a 2–3 μL drop of M9 or egg salts on a coverslip.

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Fig. 1 Dissecting adult worms. Diagram of a gravid C. elegans adult showing the best location for dissection (green lines through red spermatheca) to release young embryos. Older embryos are found near the vulva (vertical black line in center of worm)

2. Dissect the worms using a #15 scalpel or two needles to release embryos. If very young embryos are required, aim to dissect worms around the spermatheca, which is one third of the animal’s length away from the head or tail (Fig. 1). 3. Flip the coverslip so that the embryos are facing downward, and gently place the coverslip on top of the agarose pad on the slide. 4. Fill the space between the coverslip and the slide with additional M9 or egg salts using a pipette. 5. Seal the boundaries of the coverslip with petroleum jelly to avoid evaporation during long-term imaging. 3.2.2 Mounting Embryos for Time-Lapse Imaging (Well)

1. Pipette 1.8–2 mL of egg salts into a watch glass and 0.8 mL of egg salts into one chamber of a glass bottom slide. 2. Pick two to five gravid worms into the watch glass, and dissect using a #15 scalpel blade or two needles to release embryos (Fig. 1). 3. Mouth pipette embryo of desired stage from the watch glass into the glass bottom slide.

3.3 Time-Lapse Imaging

For time-lapse imaging, photobleaching of the fluorophore and phototoxicity are important to avoid, especially with blue and ultraviolet illumination. Microscopes that only illuminate the part of the sample being imaged (i.e., light sheet or confocal microscopes) are preferred, but long-term time-lapse imaging can also be performed on widefield microscopes if light exposures are minimized (see Notes 10 and 11), and samples are given time to recover between acquisitions (see Note 12). 1. Place the slide on the microscope stage with coverslip facing the objective lens. 2. Find the embryo of interest using a low magnification objective (5× or 10×) and bright-field or DIC optics. 3. Change to desired high magnification objective (40×). 4. Define the region of interest (XY) on the acquisition software controlling the microscope. 5. Define exposure times for each channel (see Note 10).

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6. Define Z boundaries and Z steps (see Note 11). 7. Define duration of the time-lapse series and the time intervals according to the dynamics of the process to be analyzed (see Note 12). 8. Start acquiring multicolor 3D time-lapse images (see Notes 13 and 14). If necessary, include a bright-field or DIC image for orientation. 9. Save the time-lapse series in a format readable by your analysis software. 3.4 Normalization of Time-Lapse Series

1. Open the time-lapse series with analysis software such as ImageJ/FIJI, Imaris, or acquisition software. 2. Annotate a zero timepoint for each analysis. This can be based on a developmental hallmark (i.e., a cell stage; see Note 15) or a process of interest (i.e., internalization; see Note 16 and Subheading 3.6).

3.5 Cargo Signaling for Engulfment

Phagocytic cargos typically present an “eat-me” signal that is recognized by receptors on the engulfing cell. One of the best characterized “eat-me” signals is the disruption of membrane lipid asymmetry and exposure of phosphatidylserine (PtdSer) on the surface of the cell targeted for disposal [24]. In C. elegans, PtdSer is usually confined to the cytofacial membrane leaflet by TAT-1, an aminophospholipid translocase or flippase [25]. Indeed, loss of PtdSer asymmetry in tat-1 mutants led to misrecognition of neurons as dying cells and their subsequent clearance. Similarly, multiple phagocytic events by different cells engulf a single midbody remnant in tat-1 mutants [26]. Multiple proteins have been shown to bind PtdSer, creating an array of secreted reporters for detecting PtdSer exposure before, during, and after engulfment (Table 5).

Table 5 Secreted reporters for monitoring PtdSer exposure Transgene

PtdSer detection method

ssGFP::C2(mfge8) C2 domain of mammalian milk fat globule epidermal growth factor ssGFP::Lact-C2 VIII (MFGE8), also called lactadherin

References [6, 57]

Annexin V::GFP

Ca2+-dependent binding by mammalian Annexin A5 (ANXA5)

[57]

TTR-52::mCh

C. elegans transthyretin-related protein expressed from the intestine that bridges PtdSer and the engulfment receptor CED-1

[58]

ssGFP::mNRF-5

C. elegans lipid-binding serum glycoprotein expressed from body wall [57] muscle and thought to transfer PtdSer to engulfing cells

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1. Prepare embryos expressing a secreted PtdSer reporter (Table 5) and a cargo reporter (see Note 18) for microscopy according to Subheading 3.2. 2. Image according to Subheading 3.3 with desired time intervals to observe PtdSer dynamics (see Note 19). 3. Annotate timing of PtdSer externalization, defined as the first frame the PtdSer reporter appears as a ring around cargo brighter than other cell–cell contacts (see Note 20). 4. Annotate timing of PtdSer disappearance. PtdSer disappearance is defined as the first frame the PtdSer reporter is no longer visible as a ring in the internalized phagosome. 5. Normalize timing to relevant time zero, such as cell stage (see Notes 15 and 21). 3.6

Engulfment

Multiple receptor pathways recognize PtdSer or other signals to regulate the actin cytoskeleton to induce phagocytic engulfment [27]. For example, stimulation of the Rac1 homolog CED-10 initiates actin filament formation under the plasma membrane, driving the formation of a phagocytic cup and engulfment of the cargo [28]. The dynamin protein DYN-1 accumulates with actin along the phagocytic cup and eventually forms a helix around the neck of the forming phagosome [6, 29]. DYN-1 promotes membrane fission and the release of the phagosome into the engulfing cell cytosol. Therefore, actin and dynamin enrichment are good markers of phagocytic cup formation and engulfment when expressed in the engulfing cell (Table 6), but internalization can also be visualized with cargo markers (Fig. 2). 1. Prepare embryos expressing a cargo marker (see Note 18) and an engulfment reporter (Table 6) for microscopy according to Subheading 3.2. 2. Image with desired time intervals to analyze engulfment dynamics (see Note 22). 3. Annotate timing of internalization (see Note 16). 4. Measure actin or dynamin fluorescence intensity over time in a circle around the cargo using analysis software (i.e., FIJI/ImageJ) (see Note 23). Also measure the fluorescence intensity Table 6 Reporters for monitoring corpse engulfment Transgene

Function

References

Lifeact::GFP Lifeact::RFP

Yeast F-actin-binding peptide

[59]

dyn-1b::gfp

C. elegans dynamin

[60]

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mCh::PH::ZF1

346

2 min

3 min Minutes past 4-cell stage

Fig. 2 Polar body internalization. The second polar body is a cell corpse found between the large anterior cells and engulfed shortly after the 4-cell stage in most C. elegans embryos. Scale bars are 10 μm in the whole embryo images and 2 μm in the polar body insets

over time in nearby cell cortex to normalize the cargo values to any cortical background (see Note 24). 5. Normalize the cargo measurements to the cortical measurements, and plot the normalized fluorescence intensity in relation to internalization as timepoint 0 (see Note 16) to determine the first and the last time engulfment reporters enrich around the cargo over background (>1). This will show timely recruitment of the reporters and whether the process is completed with typical timing. 3.7 Phagosome Maturation

Phagosome maturation is a rapid series of events that ultimately facilitates phagosome fusion with lysosomes. Phagosome maturation is characterized by changes in the phosphatidylinositol composition of the phagosome membrane and the exchange of cytosolic proteins that temporarily associate with the phagosome membrane, including Rab GTPases and LC3 family proteins. The phagosome membrane originates from the plasma membrane, which is enriched in the lipid phosphatidylinositol 4,5 bisphosphate (PI4,5P2). During phagosome maturation, PI4,5P2 is depleted from the phagosome by the lipid phosphatase OCRL-1 [30, 31], and phosphatidylinositol 3-monophosphate (PI3P) is formed by the class III PI3Kinase VPS-34, thereby converting PI4,5P2 to PI3P. To monitor this conversion, lipid-binding protein domains are used. For example, PI4,5P2 reporters include the PH domain of PLC1∂1 (Table 2) [32], while PI3P reporters include the FYVE domain of EEA-1 (Table 7) [33]. The small GTPase RAB-5 accumulates rapidly on early phagosomes. RAB-5 and VPS-34 are required for SNX-1/6-dependent

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Table 7 Reporters for monitoring phagosome maturation Transgene

Function

References

GFP::2xFYVE PI3P-binding FYVE domain of EEA-1 found on maturing phagosomes mCh::2xFYVE YFP::2xFYVE

[28]

gfp::rab-5 mCherry:: RAB-5

Small GTPase found on early phagosomes

[61, 62]

GFP::snx-1

Sorting nexin involved in receptor recycling from early phagosomes

[63]

gfp::rab-7

Small GTPase found on late phagosomes

[64]

GFP::LGG-1 mCh::LGG-2

Atg8/LC3 family proteins conjugated to lipids on phagosomes and phagolysosomes

[65]

recycling of engulfment receptors back to the plasma membrane to maintain the phagocytic capacity of the engulfing cell [6, 34]. RAB-5 is then rapidly exchanged for RAB-7 to define the late phagosome ([28], Fig. 3). RAB-7 also eventually disassociates (Fig. 3). Autophagy-related proteins also decorate maturing phagosomes and early phagolysosomes, including the GABARAP and LC3 homologs LGG-1 and LGG-2 [6, 35]. These proteins are amenable to fluorescent tagging (Table 7), allowing their recruitment and disassociation to be monitored directly. 1. Prepare embryos expressing a cargo marker (see Note 18) and reporters for different stages of maturation (Table 7; see Note 25) for microscopy according to Subheading 3.2. 2. Image with desired time intervals to analyze maturation dynamics (see Note 22). 3. Annotate the first and the last time a marker colocalizes with the cargo to plot the duration of maturation factors. 4. Graphing the appearance and disappearance of maturation markers on the phagosome in relation to internalization (see Note 16) will show whether recruitment and/or removal of the markers is normal, delayed, or premature. 3.8 Phagosome– Lysosome Fusion

Phagosome fusion with lysosomes leads to the formation of the phagolysosome. Lysosome fusion contributes lysosomal transmembrane proteins to the phagosome membrane and lysosomal hydrolases to the phagosome lumen. Unlike maturation factors, lysosomal proteins remain associated with the phagolysosome through cargo degradation. However, these proteins differ in their amenability to fluorescent tagging (Table 8). For example, the lysosome-associated membrane protein LMP-1 is normally

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mCh::H2B

GFP::RAB-7

Merge

348

1 min

5 min Time past engulfment

23 min

Fig. 3 Phagosome maturation. After engulfment, phagosomes recruit maturation factors, like GFP::RAB-7 localizing to the second polar body phagosome (5 min). GFP::RAB-7 is later turned over and lost from the phagolysosome surface (23 min). Scale bar is 10 μm Table 8 Reporters for monitoring phagosome fusion with lysosomes Transgene

Function

References

arl-8::mCit

Small GTPase associated with the surface of lysosomes and other organelles

[16]

lmp-1::GFP

Transmembrane protein trafficked to late endosomal and lysosomal membranes (also weakly labels the plasma membrane and thereby phagosome membrane)

[6]

ctns-1:: mScarlet-I ctns-1:: mCitrine

Cystinosin family cystine transporter located in lysosomal membrane

[16]

laat-1::GFP

Lysosome-associated amino acid transporter (arginine/lysine) located in [37] lysosomal membrane

nuc-1::mCh

Nuclease involved in DNA fragmentation located in lysosome lumen

cpl-1::mChOint Cathepsin protease located in lysosome lumen

[66] [38]

trafficked to lysosomes through the plasma membrane, but tagging a cytosolic domain with a fluorescent protein often leads to it being retained on the plasma membrane in addition to lysosomes. This can make ascertaining the timing of lysosome fusion more

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challenging given that the phagosome membrane is derived from the plasma membrane. Furthermore, luminal domains of transmembrane proteins or luminal hydrolases must be tagged with acid-tolerant fluorophores, such as mCherry, to be able to fluoresce in the acidic environment of the lysosome lumen (see also Subheading 3.9). 1. Prepare embryos expressing a cargo marker (see Note 18) and lysosome reporters (Table 8) for imaging according to Subheading 3.2. 2. Image with desired time intervals to analyze lysosome fusion (see Notes 22 and 26). 3. Annotate the first time the lysosome marker appears as a ring around the cargo (see Note 27). 4. Graphing the appearance of lysosome markers on the phagosome in relation to internalization (see Note 16) will show whether lysosome fusion is normal. 3.9 Phagosome and Phagolysosome Acidification

The lumen of the phagosome and phagolysosome acidify progressively. Fluorescent proteins have a range of pKa values (see Note 3) and can lose their fluorescence intensity significantly in the lumen as the surrounding pH approaches or is lower than their pKa. A combination of reporters with higher (i.e., EGFP, pKa 6.0) and lower (i.e., mCherry, pKa 4.5) pH sensitivities can be used to assess acidification of the phagosomal and phagolysosomal lumen (Table 9). 1. Prepare embryos expressing cargo or luminal reporters with different pH sensitivities (Table 9) for microscopy according to Subheading 3.2. 2. Image with desired time intervals to visualize the persistence of fluorescence (see Note 28).

Table 9 Strains for monitoring phagolysosome acidification Transgene

Function

References

GFP::H2B; mCh:: H2B

Histone reporters using fluorescent proteins with different pKa values [6]

GFP::H2B; ZF1:: mCh::his-15

Histone reporters using fluorescent proteins with different pKa values. [36] The ZF1 degron will restrict mCherry::H2B to the polar bodies after the 4-cell stage in anterior cells

nmy-2::mCh; nmy-2::GFP::ZF1

Non-muscle myosin reporters using fluorescent proteins with different [36] pKa values. The ZF1 degron will restrict NMY-2::GFP to midbody remnants after the 4-cell stage in anterior cells

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3. Plot the time of disappearance of the pH-sensitive marker over time, and compare to internalization as timepoint 0 (see Note 16). Longer persistence of pH-sensitive markers would indicate disturbed lumen acidification (see Note 29). Phagocytic cargo is often confined in its own membrane before phagocytosis, such as a cell corpse’s plasma membrane. The cargo membrane must be disrupted inside the phagolysosome in order for lysosomal hydrolases to degrade the internal protein or DNA cargo (Fig. 4a). If a membrane reporter is expressed in both the cargo and engulfing cells (Table 2), it can be difficult to distinguish the cargo membrane from the neighboring phagosome membrane by standard resolution microscopy. However, degron protection assays can degrade fluorescent reporters on the cytosolic surface of the phagosome membrane, leaving internal membrane markers inside the limiting membrane of the phagosome to fluoresce [36].

A

L

X

XX

X X

XX

X X

XX

XX

X

mCherry:: PH::CTPD

7 min

8 min

B

C

D

E 8 min

mCh::H2B

3.10 Cargo Membrane Breakdown

F

9 min G

Minutes past engulfment

Fig. 4 Cargo membrane breakdown. (a) Diagram of phagosome-lysosome fusion (left) to release lysosomal hydrolases into the phagolysosome lumen (center). However, the phagocytic cargo still has an intact membrane (center), which is broken down to give hydrolases access to cargo contents (right), such as chromosomes in dying cells (X). (b) Intact second polar body membrane inside a phagolysosome. (c) The membrane reporter disperses throughout the phagolysosome after membrane breakdown. (d) Line scan of intact cargo membrane with highest fluorescence at the rim of the second polar body. (e) Line scan of dispersed membrane with highest fluorescence in the core of the phagolysosome. (f) A histone reporter labels the compact C-shaped chromosomes before membrane breakdown. (g) The histone reporter disperses throughout the phagolysosome after membrane breakdown. Scale bars are 2 μm

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Breakdown of the cargo membrane inside the phagolysosome is followed rapidly by degradation of DNA by the lysosomal nuclease NUC-1 [6, 37, 38]. In a phagocytosed cell corpse, this also leads to a change in the compact morphology of chromosomal reporters (Table 4, Fig. 4a). Therefore, both cargo membrane breakdown and nuclease activity can be assayed indirectly by observing chromosomal morphology after phagocytosis [6]. 3.10.1 Direct Membrane Breakdown Assay

1. Prepare embryos expressing a membrane marker with a degron tag (ZF1 or CTPD in Table 2) for microscopy according to Subheading 3.2 (see Note 30). 2. Image with desired time intervals to visualize the morphology of the corpse membrane (see Note 31). 3. Annotate timing of change in membrane shape from a hollow to a filled sphere (Fig. 4b–c), corresponding to the membrane marker dispersing throughout the phagolysosome lumen. Compare timing to internalization as timepoint 0 (see Note 16). 4. Using analysis software (i.e., FIJI), draw a line across the center of the phagolysosome, and measure the profile of fluorescence intensity along the line. A line scan of an intact cargo membrane shows two peaks at the edge of the phagolysosome and a valley in the center (Fig. 4d). After cargo membrane breakdown, a line scan shows one peak in the center of the phagolysosome (Fig. 4e).

3.10.2 Indirect Chromosome Assay for Membrane Breakdown

1. Prepare embryos expressing a chromosomal reporter (Table 4) for microscopy according to Subheading 3.2. 2. Image with desired time intervals to visualize the morphology of the corpse membrane (see Note 30). 3. Annotate timing of change in morphology from a compact C-shape to a filled sphere (Fig. 4f–g), corresponding to the chromosome marker dispersing throughout the phagolysosome lumen. Compare timing to internalization as timepoint 0 (see Note 16).

3.11 Phagolysosome Shrinkage

Once lysosomal hydrolases have access to phagocytic cargo, breakdown products will be formed inside the phagolysosome. Lysosomal transporters then export macromolecules and solutes out to the cytosol. Export leads to a visible decrease in the size of the phagolysosome, which can be used to assay shrinkage [39]. 1. Prepare embryos expressing a pH-insensitive cargo reporter (Tables 2, 3, and 4; see Note 32) for microscopy according to Subheading 3.2. 2. Image with desired time intervals to visualize the phagocytic cargo (see Note 33).

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3. Using analysis software, measure the perimeter and/or area of the phagolysosome in the z-plane showing its maximum width (see Note 34), and compare over time using internalization or cargo membrane breakdown as timepoint 0 (see Note 16 or Subheading 3.10). 3.12 Phagolysosome Tubulation, Vesiculation, and Resolution

While DNA degradation occurs within minutes after cargo membrane breakdown (Subheading 3.10), the degradation of cargo protein content takes hours. During this time, large phagolysosomes go through multiple rounds of tubulation to form smaller phagolysosomal vesicles containing cargo proteins (Fig. 5). After vesiculation, protein cargos are degraded, thereby resolving the phagolysosome [6]. 1. Prepare embryos expressing pH-insensitive cargo markers (Tables 2, 3, and 4; see Notes 32 and 35) according to Subheading 3.2. 2. Image with desired time intervals to visualize the phagolysosome over hours (see Note 36). 3. Using visualization software (i.e., Imaris), observe each time when the phagolysosome elongates or forms a tubule. Also note each time a vesicle separates from the original phagolysosome (fission event). Comparing the timing of the first fission event in comparison to internalization (see Note 16) or the number of fission events can demonstrate the ability of a given strain to vesiculate. 4. Continue to monitor each individual vesicle until they disappear, including the original phagolysosome (see Note 37).

mCh::PH::ZF1

5. Data can be plotted as branched trees for a simultaneous visualization of fission events and cargo protein persistence, as the timing of individual vesicle disappearance, or the timing of phagolysosome resolution in comparison to internalization (see Note 16) [6].

Phagolysosome

42

Tubulation

Vesiculation

45 50 Minutes past engulfment

Resolution

87

Fig. 5 Cargo tubulation, vesiculation, and resolution. Long after engulfment of the second polar body, the phagolysosome tubulates (45 min) to release vesicles (50 min). This continues until eventually the polar body proteins are degraded (87 min), resolving the phagolysosome cargo. Scale bar is 2 μm

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Notes 1. Forward genetics can be used to generate new mutations using the mutagen ethyl methanesulfonate (EMS) [40] or by CRISPR/Cas9-mediated genome editing [41, 42]. 2. Reporter constructs can be expressed under the endogenous or an alternative promoter, but overexpression of tagged proteins can also disrupt cell biology. Table 1 summarizes common promoters used for heterologous expression in C. elegans. Transgenes can be generated by microinjection or by bombardment to generate extrachromosomal arrays or stable, integrated transgenes. Details for transgenesis can be found here [43]. To modify endogenous gene loci, CRISPR/Cas9 approaches can be applied by microinjection [41, 42]. 3. Reporter strains are typically made by tagging a protein of interest with a fluorescent protein. Fluorescent proteins are available with a variety of characteristics, including wavelength, fluorescence intensity, photostability, pKa, maturation time, etc. The choice of fluorescent tag can therefore affect analyses, especially in acidic organelles. Properties of fluorescent proteins can be checked here (https://www.fpbase.org). Fluorescent proteins are large tags that can disrupt protein dynamics, localization, activity, or function based on their size, placement, or oligomerization. Fluorescent tags should be placed on a position where no vital domain of the target protein is hindered. Generally, adding a floppy linker of around 20 amino acids between the target and the fluorescent tag is recommended. Dynamic properties of the tagged protein, such as cleavage for activity, should also be considered. 4. Worms grow best in a temperature range of 15–25 °C. Temperature-sensitive mutants may need more specific temperatures, and multi-copy transgenes often require higher temperatures (23–25 °C) to maintain expression. 5. The growth of the uracil auxotroph OP50 bacteria can be controlled by the concentration of uracil in the NGM plates. A thin bacterial lawn allows better observation of embryos on the plate. For growing worms in large scale, the NA22 bacteria strain is used for a thick bacterial lawn. 6. Pre-coat the watch glass with Sigmacote or another siliconizing reagent for glass and other surfaces. This thin film will prevent embryos from sticking to the glass. 7. Avoid repeated boiling of the agarose solution as boiling leads to evaporation, concentrating the agarose solution. 8. Freshly made agarose is less likely to swell or shrink during time-lapse imaging. See also Note 13.

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9. Compressing the embryo improves image quality in deeper tissue but may alter physical properties of the membrane. For better control of compression, embryos can also be embedded in a polymer bead solution [44], where 20 μm beads offer similar compression to an agarose pad, while 30 μm beads avoid compression. 10. We generally set exposure times and light intensities to allow 2000–3000 gray values on the camera, well under detection limits. For widefield imaging, we turn the light source down to 25%. For light sheet imaging, we use laser power set to 150 mW and 50% for the 488 nm laser or 100 mW and 70% for the 568 nm laser. We also align the light sheet on the embryo using the 568 nm laser to avoid phototoxicity from the 488 nm laser. Data sets are excluded if there is a delay or arrest in embryonic development during imaging, which is a sign of phototoxicity. 11. A wild-type embryo is ~20 um thick when compressed by an agar pad or >25 μm uncompressed. Since 2 μm corpses and 1 μm midbody remnants can move extensively, a balance between covering the whole embryo while not losing information between Z steps should be found. For polar body imaging, we recommend 16 × 1.2 μm Z steps for widefield imaging to avoid photobleaching and phototoxicity. For less phototoxic methods, such as light sheet or spinning disk confocal, more Z steps with smaller Z intervals can be used. 12. For rapid processes, intervals are often limited by the microscope but can routinely reach 10–15 s for whole embryo capture. For imaging over hours, time intervals are more commonly 1–2 min to avoid photobleaching and phototoxicity. 13. If the agarose solution is old or the cover slip is not sealed well, the agarose pad can lose water content, causing the embryo to sink during image acquisition. This is most noticeable during the first 10 min of the time lapse. Check the Z range during this time, and correct the Z range to cover the whole region of interest. 14. For a given analysis, use the same settings (exposure time, fluorescence channels, number of Z steps, and time intervals) to minimize the effects of variables such as photobleaching or phototoxicity. 15. In C. elegans embryos, the first timepoint of a new cell stage occurs with the onset of cytokinetic furrow ingression. This is visible with DIC imaging, using a plasma membrane reporter (see Table 2) or a non-muscle myosin reporter (see Table 3). If using a chromosomal marker (see Table 4), anaphase onset can be considered the first timepoint of a new cell stage.

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16. Internalization timing may be defined as the first timepoint when the cargo moves away from the plasma membrane (Fig. 2) or when the phagosome first becomes spherical instead of elongated (see also Note 17). 17. It is important to find the z-plane with the largest diameter of the cargo to ensure that the cargo is not above or below the curved cells of the embryo. 18. We use plasma membrane reporters to label both polar bodies and midbody remnants (see Table 2). Chromosome reporters only label polar bodies (see Table 4), while the reporters in Table 3 most brightly label midbody remnants. 19. PtdSer exposure on the second polar body occurs during the 2to 4-cell stage, making 1-min timepoints starting at the 2-cell stage practical. 20. PtdSer reporters need to be stably expressed and visible. Additionally, they cannot be too highly expressed, or they will label other structures, such as the eggshell. 21. The second polar body externalizes PtdSer after the 2-cell stage, making the 2-, 3-, or 4-cell stage recommended timepoints for normalization. Midbody remnants externalize PtdSer after their release by abscission, so each remnant can be normalized to the cell stage that formed it. 22. Due to the rapid dynamics of engulfment, phagosome maturation, and lysosome fusion, time intervals of 10–20 s are recommended for these analyses. The second polar body is internalized at the 2- or 4-cell stage, while midbody remnants are internalized starting at the 4- or 6-cell stage. 23. For midbody remnants or the second polar body, we used a 2 μm2 circle starting at the 2-cell stage and proceeding until after internalization [6, 11]. 24. Under widefield illumination, the embryo is typically brighter in the thicker center than at the edges, making it important to normalize to a similarly sized circle of cortex at a similar position within the embryo that does not include the phagocytic cargo. 25. Combination of two sequential maturation reporters (e.g., green RAB-5 and red RAB-7) can also be used to better understand the dynamics. 26. Phagosomes and phagolysosomes can fuse with multiple lysosomes, making it important to distinguish between phagolysosome formation and later lysosome fusion events. 27. Lysosomes can be recruited to the phagosome and appear as puncta surrounding the phagosome before fusion of the phagosome and lysosome membranes to disperse the lysosomal proteins in the phagolysosome.

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28. As acidification of the lumen proceeds gradually and it is important to avoid photobleaching, 1-min time intervals starting before cargo engulfment are sufficient for the analysis of acidification. 29. Acidification data should be interpreted together with other assays, such as Subheadings 3.7, 3.8, or 3.10. Persistence of a pH-sensitive marker may indicate a disturbed proton pump, defective lysosome fusion, or delayed cargo membrane breakdown. 30. Fluorescent reporters with low cytoplasmic background, such as mCherry, are preferable for this assay. 31. Cargo membrane breakdown typically occurs within 10 min of engulfment in C. elegans embryos, making 30-s intervals useful for this assay. Membrane breakdown is easier to observe when the phagolysosome is found in the half of the embryo closest to the objective, as yolk in embryos distorts fluorescence. 32. Long-term assays are facilitated by specifically labeling the phagolysosome, such as using degron-tagged markers (ZF1 or CTPD) where any cytosolic reporter is degraded and only the phagolysosome and derived vesicles are fluorescently labeled. 33. Shrinkage is easier to visualize with larger cargos like cell corpses when using standard resolution imaging. For the second polar body, phagolysosome shrinkage is observed ~20 min after phagocytosis and continues, making 1- or 2-min timepoints starting at the 3- to 6-cell stage practical for this longterm analysis. 34. The inclusion of more Z steps during imaging will enable measurement of phagolysosome volume and decrease the variability in measurements caused by phagolysosome movement between z-planes. Otherwise, individual measurements can be averaged over time to smooth the curve, i.e., the average of 3–5 timepoints. 35. Combination of this assay with lysosome fusion assays (Subheading 3.8) can give information about new rounds of fusion with lysosomes. Phagolysosome vesiculation has also been observed to form new lysosomes from tubules devoid of cargo [45]. 36. Phagolysosome vesiculation starts 30+ min after engulfment of the second polar body, and cargo resolution occurs over 1–2 h after engulfment [6], making 1-min intervals preferable to avoid photobleaching. However, it can be challenging to track individual phagolysosomal vesicles with large Z steps or longer time intervals.

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37. This assay measures disappearance of a fluorescent protein, which may not be the same as protein degradation. Factors that can affect the visibility of the marker include the short exposure times and low light levels used to allow long-term imaging (see Subheading 3.3), photobleaching of fluorescent protein over time, and the pH-sensitivity of the fluorescent protein in the acidifying phagolysosome (see Subheading 3.9).

Acknowledgments The authors would like to thank David Michaelson for writing assistance, Alyanna Ferrer for assistance with shrinkage data analysis, Maurice Stetter for RAB-7::GFP data, and Riley Harrison for membrane breakdown data. GF was funded by Deutsche Forschungsgemeinschaft (DFG) grant FA1046/3-1 to GF, and all authors were funded by NIH grant R15 GM143727-01 to AMW and GF. References 1. Flannagan RS, Jaumouille V, Grinstein S (2012) The cell biology of phagocytosis. Annu Rev Pathol 7:61–98. https://doi.org/ 10.1146/annurev-pathol-011811-132445 2. Shklover J, Levy-Adam F, Kurant E (2015) Apoptotic cell clearance in development. Curr Top Dev Biol 114:297–334. https://doi.org/ 10.1016/bs.ctdb.2015.07.024 3. Fond AM, Ravichandran KS (2016) Clearance of dying cells by phagocytes: mechanisms and implications for disease pathogenesis. Adv Exp Med Biol 930:25–49. https://doi.org/10. 1007/978-3-319-39406-0_2 4. Ghose P, Wehman AM (2021) The developmental and physiological roles of phagocytosis in Caenorhabditis elegans. Curr Top Dev Biol 144:409–432. https://doi.org/10.1016/bs. ctdb.2020.09.001 5. Sulston JE, Schierenberg E, White JG, Thomson JN (1983) The embryonic cell lineage of the nematode Caenorhabditis elegans. Dev Biol 100(1):64–119 6. Fazeli G, Stetter M, Lisack JN, Wehman AM (2018) C. elegans blastomeres clear the corpse of the second polar body by LC3-associated phagocytosis. Cell Rep 23(7):2070–2082. https://doi.org/10.1016/j.celrep.2018. 04.043 7. Gumienny TL, Lambie E, Hartwieg E, Horvitz HR, Hengartner MO (1999) Genetic control of programmed cell death in the Caenorhabditis elegans hermaphrodite germline. Development 126(5):1011–1022

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Chapter 23 Assessing the Phagosome Proteome by Quantitative Mass Spectrometry Maria Emilia Duen˜as, Jose´ Luis Marı´n-Rubio, Julien Peltier-Heap, Anetta Hartlova, and Matthias Trost Abstract The process of phagocytosis involves a series of defined steps, including the formation of a new intracellular organelle, i.e., the phagosome, and the maturation of the phagosome by fusion with endosomes and lysosomes to produce an acidic and proteolytic environment in which the pathogens are degraded. Phagosome maturation is associated with significant changes in the proteome of phagosomes due to the acquisition of new proteins or enzymes, post-translational modifications of existing proteins, as well as other biochemical changes that ultimately lead to the degradation or processing of the phagocytosed particle. Phagosomes are highly dynamic organelles formed by the uptake of particles through phagocytic innate immune cells; thus characterization of the phagosomal proteome is essential to understand the mechanisms controlling innate immunity, as well as vesicle trafficking. In this chapter, we describe how novel quantitative proteomics methods, such as using tandem mass tag (TMT) labelling or acquiring label-free data using data-independent acquisition (DIA), can be applied for the characterization of protein composition of phagosomes in macrophages. Key words Macrophages, Quantitative proteomics, Phagosomes, Tandem mass tag (TMT), Mass spectrometry (MS), Data-independent acquisition (DIA), Data-dependent acquisition (DDA)

1

Introduction Phagocytosis, the mechanism by which particles are internalized, leads to the formation of a specialized organelle called the phagosome. After uptake, phagosomes mature by the fusion of the nascent organelle with early endosomes, late endosomes, and ultimately lysosomes forming a phagolysosome in which the engulfed material is degraded [1]. This process is important for tissue homeostasis, innate immunity, and adaptive immunity through antigen presentation from the phagosome. Numerous studies

˜ as and Jose´ Luis Marı´n-Rubio contributed equally to this work. Maria Emilia Duen Roberto J. Botelho (ed.), Phagocytosis and Phagosomes: Methods and Protocols, Methods in Molecular Biology, vol. 2692, https://doi.org/10.1007/978-1-0716-3338-0_23, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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have contributed to the understanding of the importance of many factors in phagosome maturation including small GTPases, phosphoinositol kinases, signalling, and actin dynamics [2, 3]. However, the mechanisms of uptake and maturation, as well as signalling from the phagosome, are still relatively poorly understood. In recent years, proteomics approaches of phagosomes has led to a better understanding of these molecular mechanisms occurring during phagosome maturation [4]. For example, proteomic analysis of phagosomes showed for the first time translocation of the endoplasmic reticulum (ER) to the phagosome membrane [5, 6], providing new potential mechanisms of antigen cross-presentation [7, 8]. It further provided clues to how macrophage activation changed phagosome dynamics for the gain of antigen presentation [9]. Moreover, proteomics of phagosomes from Dictyostelium, Drosophila, and mouse showed strong conservation of a core phagosome proteome over evolution [10]. Two-dimensional gel electrophoresis (2DE) coupled to matrixassisted laser desorption/ionization time-of-flight (MALDI-TOF) or electrospray ionization mass spectrometry (ESI-MS) was the first method to study complex protein mixtures like phagosomes [11, 12]. However, 2DE-MS had inherent limitations such as limited sample capacity and sensitivity, restricting the identification of low abundance proteins, as well as an incompatibility for transmembrane proteins. To overcome these critical limitations, the last decade has seen the development of reproducible quantitative proteomic approaches using sensitive data-dependent and dataindependent tandem mass spectrometry analysis (LC-MS/MS) that now allow the systematic identification and quantification of thousands of proteins in complex proteomes. Quantitative data from proteomics come in two forms, the absolute quantity of the protein or the relative change of the protein in different samples. Relative quantification is mostly used in the discovery phase, comparing levels of a protein in different states with results being expressed as a relative fold change of protein abundance. Absolute quantification refers to the exact determination of the amount of a protein in question (e.g., ng/mL) and is widely used in targeted proteomics approaches. Differential analysis is generated from LC-MS/MS experiments and can be carried out using either label-free or isotope labelling approaches such as stable isotope labelling by amino acids in cell culture (SILAC) [13], dimethyl labelling [14], or tandem mass tag (TMT) reagents [15]. Application of such analytical methods could resolve fundamental questions about molecular mechanisms in phagosome biogenesis [9, 16–19]. In this chapter, we describe reliable quantitative proteomic strategies for studying the phagosome proteome [20, 21]. Important considerations in every step of the sample preparation, TMT labelling, and mass spectrometry analysis are described and discussed in further details.

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2 2.1

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Materials Equipment

1. Low protein binding microcentrifuge tubes (1.5 mL or 2.0 mL). 2. Suspension trapping (S-Trap™) spin columns (see Note 1). 3. Vacuum centrifuge with cold trap. 4. Thermal mixer. 5. Benchtop centrifuge. 6. Probe sonicator. 7. Peptide desalting spin columns. 8. C18 solid phase extraction columns (C18 SPE). 9. Off-line liquid chromatography (LC): Dionex Ultimate 3000 off-line LC system. 10. Mass spectrometer with electrospray ion source (see Note 2). 11. On-line LC: Dionex Ultimate 3000 RSLC-nano system. 12. C18 LC trap column: Acclaim PepMap® (100 μm ID × 20 mm, 3 μm, 100 Å). 13. C18 analytical column: EASY-Spray nanoLC C18 column (75 μm ID × 750 mm, 2 μm, 100 Å). 14. Basic reverse phase column: Phenomenex Gemini C18 (3 μm particle size, 110 Å pore, 3 mm internal diameter, 250 mm length).

2.2 Cell Lysis, Reduction, Alkylation, and Digestion

1. Lysis buffer: 5% sodium dodecyl sulfate (SDS) (see Note 3), 50 mM triethylammonium bicarbonate buffer (TEAB), pH 7.55 in high-performance liquid chromatography (HPLC) grade water with protease inhibitor cocktail, phosphatase inhibitor cocktail (see Note 4), 100 mM N-Ethylmaleimide (NEM) (see Note 5), and 1 unit/μL of Pierce™ Universal Nuclease. 2. Bicinchoninic acid (BCA) protein assay kit. 3. 500 mM Tris(2-carboxyethyl)phosphine hydrochloride solution) (TCEP). 4. 1× phosphate-buffered saline (PBS), pH 7.4: 137 mM NaCl, 2.7 mM KCl, 8 mM Na2HPO4, and 2 mM KH2PO4. 5. 12% phosphoric acid solution: 1.4 mL H3PO4 (85% w/w) in 8.6 mL of HPLC grade water. 6. S-Trap biding buffer: 90% methanol, 100 mM TEAB in HPLC grade water, pH 7.1. 7. 50 mM TEAB solution, pH 8.0: 0.5 mL of 1 M TEAB, pH 8.5 in 9.5 mL of in HPLC grade water.

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8. Trypsin solution, pH 8.0: 1 μg/μL MS grade trypsin in 50 mM acetic acid. 9. 500 mM iodoacetamide (IAA) (see Note 6): 92.5 mg IAA in 1 mL of HPLC grade water. 2.3 Sample Cleanup, TMT Labelling, and MS

1. 100 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) solution, pH 8.5: 1.19 g of HEPES in 50 mL of HPLC grade water. 2. 5% hydroxylamine solution: 50 μL of 50% hydroxylamine (NH2OH) in 450 μL of 100 mM HEPES. 3. 20 mM ammonium formate solution, pH 8.0: 1.26 g ammonium formate (NH4HCO2) in 1 L of HPLC grade water. 4. Isobaric tandem mass tag (TMT) reagents: TMTpro 16plex label reagents (see Note 7). 5. Washing buffer: 90% (v/v) HPLC grade acetonitrile (MeCN), 0.1% (v/v) trifluoroacetic acid (TFA) in HPLC grade water. 6. Loading buffer: 0.1% (v/v) TFA in HPLC grade water. 7. Elution buffer: 50% (v/v) MeCN, 0.1% (v/v) TFA in HPLC grade water. 8. Buffer A: 0.1% formic acid (FA) in HPLC grade water. 9. Buffer B: 80% HPLC grade MeCN, 0.1% FA in HPLC grade water. 10. Solubilization buffer for mass spectrometry analysis: 2% HPLC grade MeCN, 0.1% TFA in HPLC grade water.

2.4

Data Analysis

1. Software packages for data-dependent analysis (DDA): Proteome Discoverer (Thermo Scientific), Mascot (Matrix Science), Scaffold Q+ (Proteome software), or MaxQuant. 2. Software packages for data-independent analysis (DIA): DIA-NN or Spectronaut (Biognosys). 3. Data analysis software: R and RStudio.

3

Methods An overview of the common quantitative mass spectrometry workflows used for the study of the phagosomal proteome is outlined in Fig. 1.

3.1 Cell Lysis, Reduction, Alkylation, and Digestion for Quantitative Proteomic Analysis

1. Isolate latex/polystyrene bead phagosomes as described in Chapter 16. 2. To remove the remaining sucrose, wash pellet phagosomes with 7 mL of 1× PBS, and centrifugate at 15,000 × g for 15 min.

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Fig. 1 Schematic of phagosome preparation and proteomics. (a) Sample preparation scheme for phagosome proteomics. After solubilization of phagosome pellets, the lysate is reduced, alkylated, and digested with trypsin using the suspension trapping (S-Trap) sample preparation. (b) Quantitative mass spectrometry workflows used for studying phagosome proteomes. Horizontal lines indicate when samples are combined. For label-free mass spectrometry analysis, all experimental procedures are processed in parallel until data analysis. In TMT mass spectrometry experiments, experimental procedures are performed in parallel until proteins have been digested. Each experimental condition is individually labelled by a different chemical stable isotope. From this point, samples are combined for mass spectrometry and data analyses. Label-free samples are acquired in DIA, while TMT-labelled samples are acquired using SPS-MS3

3. Remove the supernatant and beads sticking to the side of the tube. Wipe dry the wall of the tube before adding the lysis buffer. 4. Lyse phagosome pellets by adding 50–100 μL lysis buffer (see Note 8). 5. Use probe sonication. 6. Perform a BCA protein assay or similar according to manufacturer’s instructions to determine protein concentration.

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7. Dilute 10–50 μg of protein (see Note 9) in 25 μL of lysis buffer (see Note 10). 8. Reduce disulfide bonds by adding a final concentration of 20 mM TCEP (1 μL of 500 mM TCEP), and incubate for 30 min at 37 °C with agitation (600 rpm). 9. Cool the protein solution to room temperature, and alkylate cysteines by adding 1 μL of freshly prepared 500 mM IAA (final concentration of 20 mM IAA), and incubate for 30 min at room temperature in the dark (see Note 11). 10. Acidify samples adding 2.5 μL of 12% phosphoric acid solution to the sample. 11. Perform protein digestion using the suspension trapping sample preparation method according to the manufacturer’s guidelines (see Note 12). Add 165 μL S-Trap binding buffer to the sample, and add mix into the S-Trap Micro spin column (see Note 13). 12. Centrifuge the samples at 4000 × g for 2 min until all the solution passed through the filter. 13. Wash each S-Trap micro-spin column with 150 μL S-trap binding buffer by centrifugation at 4000 × g for 1 min. Repeat this process for four washes. 14. To digest, add 25 μL of 50 mM TEAB, pH 8.0 containing trypsin (1:10–1:20 trypsin-to-protein, wt:wt) to each sample (see Note 14). 15. Cap the spin column loosely, and incubate in a clean tube for 2 h at 47 °C using a thermomixer without shaking (see Note 15). 16. Add 40 μL of 50 mM TEAB pH 8.0 to the spin column, and elute the peptides by centrifuging at 4000 × g for 2 min. Elute peptides with 40 μL of 0.2% FA and centrifuge 2 min at 4000 × g. 17. Elute hydrophobic peptides with 35 μL 50% MeCN, 0.2% FA by centrifuging at 4000 × g for 2 min. 18. Combine the three eluates, and dry down peptides using a vacuum centrifuge with cold trap. 19. Store sample at -80 °C. 20. For label free-based quantitative proteomic strategy, resuspend samples, and perform mass spectrometry analysis as described in Subheading 3.5, step 1.

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1. Prepare TMT reagents as per manufacturer’s instruction. Equilibrate the TMTpro label reagent to room temperature, add anhydrous acetonitrile to each vial to have 25 μg/μL final concentration, and allow the reagent to dissolve for 5 min with occasional vortexing (see Note 16). 2. Reconstitute 25–100 μg protein digest (from Subheading 3.1, step 19) in 25 μL of 100 mM HEPES solution, pH 8.5. 3. Assign each sample to a TMT tag, and add 5 μL of the corresponding TMT tag to each 25 μL sample (see Note 17). 4. Incubate the reaction for 1 h at room temperature. 5. Pool a small aliquot of each sample (corresponding to 1 μg protein digest) for labelling efficiency and for ratio check. For the test pool, quench with 1 μL of 5% hydroxylamine solution, incubated for 15 min at room temperature, dry using vacuum centrifuge, clean samples using a C18 spin column as per the manufacturer’s guidelines, and subsequently dry sample using vacuum centrifuge (see Note 18). 6. Store the remaining unquenched samples at -80 °C until check is completed. 7. Thaw unquenched samples and add 5 μL of 5% hydroxylamine to the sample, and then incubate for 15 min to quench the reaction. 8. Based on ratio check, pool each sample in a new microcentrifuge tube, and then dry samples using vacuum centrifuge. 9. Clean up the samples using the peptide desalting spin columns, as per the manufacturer’s guidelines (see Note 19). 10. Dry samples using vacuum centrifuge and store samples at -80 °C.

3.3 Offline HPLC Fractionation for TMTpro 16plex Labelled Samples

1. Resuspend TMTpro 16plex labelled peptides in 20 mM ammonium formate, pH 8.0 at final concentration of 2.5 μg/μL. 2. Fractionate peptides on a Basic Reverse Phase column on an off-line LC system. 3. Set-up method as follows: Load 40 μL of peptides on column for 1 min at 250 μL/min using 99% Buffer X (20 mM ammonium formate, pH 8.0), and elute for 40 min on a linear gradient from 1 to 90% Buffer Y (100% MeCN). Monitor peptide elution by UV detection at 214 nm. Collect fractions every 60 s from 2 to 38 min for a total of 36 fractions. Pool fractions using non-consecutive concatenation to obtain 18 pooled fractions (e.g., pooled fraction 1: fraction 1 + 19). 4. Acidify each fraction to a final concentration of 1% TFA, and dry using vacuum centrifuge, and store at -80 °C.

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3.4 Mass Spectrometry Analysis for TMT Samples

Each sample is independently analyzed on a mass spectrometer connected to an LC system. 1. Resuspend each sample at 0.5 μg/μL using solubilization buffer for mass spectrometry analysis; load 1–2 μg of peptide digest per injection. 2. Set up LC method as follows: Load peptide digest on a C18 trap column followed by separation on a C18 analytical column at a flow rate of 250 nL/min. The gradient used for analysis of proteome samples is as follows: buffer B is maintained at 3% for 5 min, followed by an increase of buffer B from 3% to 35% in 120 min, 35% to 90% B in 30 s, maintained at 90% B for 4 min, followed by a decrease to 3% in 30 s and equilibration at 3% for 20 min. 3. Perform mass spectrometric identification and quantification operating in data-dependent, positive ion mode (see Note 20). 4. Set up Orbitrap Fusion Lumos as follows: acquire full scan spectra in a range from m/z 375–1500, at a resolution of 120,000, with a standard automated gain control (AGC) (Tune 3.3) (see Note 21) and a maximum injection time of 50 ms. 5. Isolate precursor ions with a quadrupole mass filter width of 0.7 m/z, and perform CID fragmentation (see Note 22) in one-step collision energy of 30% and 0.25 activation Q. 6. Acquire detection of MS/MS fragments in the linear ion trap in a rapid mode using a Top 3 s method, with a standard AGC target and a maximum injection time of 50 ms. Enable dynamic exclusion of previously acquired precursor for 60 s with a tolerance of ±10 ppm. 7. Perform quantitative analysis of TMT-tagged peptides using FTMS3 acquisition in the Orbitrap mass analyzer operated at 60,000 resolution, with a standard AGC target and maximum injection time of 118 ms. 8. Perform HCD fragmentation on MS/MS fragments in one-step collision energy of 55% to ensure maximal TMT reporter ion yield and enable synchronous-precursor-selection (SPS) to include 10 MS/MS fragment ions in the FTMS3 scan.

3.5 Mass Spectrometry Analysis for Label-Free Samples

For label-free samples: Perform mass spectrometric identification and quantification operating in data-independent acquisition (DIA), positive ion mode [22]. Each sample is independently analyzed on a mass spectrometer connected to an LC system 1. Repeat steps 1 and 2 from Subheading 3.4. Set up Orbitrap Fusion as follows: Acquire full scan spectra in a range from m/z 390–1010, at a resolution of 60,000, with an AGC target of 100% at a maximum injection time of 55 ms.

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2. Collect DIA scans using 8 m/z staggered windows, with loop count set to 75, Orbitrap resolution of 30,000, AGC target of 100%, and maximum injection time of 23 ms (see Note 23). 3.6 Data Processing and Analysis

The primary requirement of data processing is to convert raw files from mass spectrometry analysis into a list of identified peptides, coupled with quantitative information given by the precursor intensity of the identified peptides (current method used for label-free) or given by the reporter masses for TMT labelling. Many software packages have been developed to handle both identification and quantification of proteins such as Proteome Discoverer, Mascot, Scaffold Q+, and MaxQuant [23] for DDA and DIA-NN [24] and Spectronaut for DIA. Mass spectrometry data analysis was performed similarly as described previously [17, 21], but TMT modification on the peptide N-termini or lysine residues were enabled for the 16-plex TMT reagents was included as variable modification in MaxQuant and carbamidomethyl (C) as fixed modification. TMT data analysis was performed using the R package Limma [25]. The final outcome of these proteomic approaches is a list of proteins and their quantitative information. A brief overview of expected phagosome proteins are displayed in summary tables and in [21].

3.7

It is important to check phagosome purity and check for marker proteins. From experience, latex bead phagosome proteomes usually contain at least 2500–3000 proteins, although the core proteome is made up of ~800–1000 [10]. We have listed the 20 most abundant proteins according to the intensity-based absolute quantitation (iBAQ) (Table 1) from a recent paper (see Note 24) [21, 26]. Latex bead phagosomes are generally rich in endosomal, lysosomal, plasma membrane proteins and contain subsets of ER and Golgi proteins and usually few proteins with described mitochondrial localization [6]. Common potential contaminants found in phagosome proteome analyses are fetal bovine serum proteins, keratins, histones, and ribosomes.

4

Quality Control

Notes 1. Different S-Trap cartridges provided by Profit can be used: micro columns (≤100 μg protein), mini columns (100–300 μg protein), midi columns (≥300 μg protein), and 96-well plate (100–300 μg protein per well). 2. While the pipeline described here utilizes the Orbitrap Fusion Tribrid Mass Spectrometer (Orbitrap Fusion Tribrid Mass Spectrometer with an EASY-Spray ion source), experiments can also be performed on other high-resolution mass

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Table 1 Identification of the 20 most abundant phagosome proteins according to the intensity-based absolute quantification (iBAQ), i.e., estimated absolute abundance

Protein name

Mol. weight Peptides (kDa)

Proteomics iBAQ

P63254

Cysteine-rich protein 1

9

8.5

8.02E + 09

Gapdh

P16858

Glyceraldehyde-3-phosphate dehydrogenase

38

35.8

5.62 E + 09

Rpl38

Q9JJI8

60S ribosomal protein L38

8

8.2

4.41E + 09

Lyz1

P17897

Lysozyme C-1

15

16.8

3.33E + 09

Lyz2

P08905

Lysozyme C-2

5

16.7

3.11 E + 09

Lamtor5

Q9D1L9

Regulator complex protein LAMTOR5

6

9.6

2.57 E + 09

Ifitm3

Q9CQW9

Interferon-induced transmembrane protein 3

6

15

2.35 E + 09

Vim

P20152

Vimentin

82

53.7

2.21 E + 09

Stom

P54116

Erythrocyte band 7 integral membrane protein

29

31.4

2.00 E + 09

Atp6v0c

E9Q9C5

V-type proton ATPase 16 kDa proteolipid 3 subunit

15.3

1.83 E + 09

Actb

P60710

Actin, cytoplasmic 1

32

41.7

1.62 E + 09

Ubc

P62984

Ubiquitin-60S ribosomal protein L40

12

14.7

1.55 E + 09

Rab7a

P51150

Ras-related protein Rab-7a

27

23.5

1.51 E + 09

V-type proton ATPase subunit B

48

56.6

1.40 E + 09

Uniprot ID

Uniprot accession

Crip1

Atp6v1b2 P62814 Ctsz

Q9WUU7 Cathepsin Z

19

34

1.30 E + 09

Cd68

P31996

Macrosialin

7

34.8

1.20 E + 09

Gnai2

P08752

Guanine nucleotide-binding protein G (i) subunit alpha-2

31

40.5

1.06 E + 09

Gpnmb

Q99P91

Transmembrane glycoprotein NMB

23

63.7

9.41 E + 08

Atp6v1a

P50516

V-type proton ATPase catalytic subunit A

53

68.3

9.40 E + 08

Gnb2

P62880

Guanine nucleotide-binding protein G(I)/ 21 G(S)/G(T) subunit beta-2

37.3

8.99 E + 08

spectrometers such as quadrupole-time of flight or quadrupole-Orbitrap instruments. The major advantages of the hybrid Quadrupole-Orbitrap such as the Fusion Tribrid mass spectrometer are the ability to select ion for fragmentation with high specificity and perform MS3 experiments to improve quantitative performance (see Note 20).

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3. If PBS remains at the bottom of the tube, 10% SDS can be used. However, the final concentration of SDS has to be kept below 5%. 4. Protease inhibitor cocktail cOmplete™ tablets are used. Phosphatase inhibitor cocktail consists of 1.2 mM sodium molybdate, 1 mM sodium orthovanadate, 4 mM sodium tartrate dihydrate, 5 mM glycerophosphate, and 20 mM NEM. 5. NEM is used to inhibit deubiquitylases. 6. The 500 mM IAA solution should be made fresh just prior to use. Alternately, 500 mM NEM can be used for ubiquitin peptides analysis for absolute quantification by parallel reaction monitoring (Ub-AQUA-PRM) assay [27]. 7. TMTpro 18-plex Label Reagent is also available in Thermo Fisher Scientific. 8. If possible, pellet should be frozen prior to protein extraction. Freezing pellets inactivate most enzymatic activity and lead to higher protein recovery and identification of more modification sites. 9. Proteins in solution are stable for years at -80 °C. Protein modifications such as phosphorylation and acetylation can be lost during storage. Ubiquitin-like modifications are stable. 10. The sample must be in at least 5% SDS present. If necessary, add concentrated SDS solution to ensure 5% SDS final concentration. 11. After this step, proteins can be stored at -80 °C. 12. S-Trap micro columns (≤100 μg protein) were used for proteome analysis as we described in this chapter. Manufacturer’s instruction and S-Trap Micro high recovery protocol is also available in https://protifi.com/pages/protocols. 13. It will not flow through. Do not fill the column higher than the narrow “stem.” 14. The protein trapping matrix is highly hydrophilic and will absorb the solution. However, ensure there is no bubble atop the protein trap. 15. Incubate from 1 to 3 h at 47 °C for trypsin or trypsin/Lys-C or overnight at 37 °C. Ensure the entire column is exposed to heat. Do not shake. 16. Reagents dissolved in acetonitrile are stable for 1 week when stored at -20 °C and warmed to room temperature before opening. Anhydrous ethanol can be used as an alternative solvent to dissolve reagents. 17. For complete labelling of lysine and N-terminal, use a minimum ratio of 1:5–1:10, sample to tag (wt:wt).

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18. For the ratio check, each protein is binary logarithm (log2) transformed, and each sample (corresponding to a single TMT channel) is normalized to the median intensity of all sample within its pool. A scaling factor is calculated. 19. Wash samples with the two additional column washes to remove excess TMT reagent. 20. An established limitation of isobaric tagging strategies occurs during mass spectrometry analyses of samples. Precursor ions selected for MS/MS fragmentation in complex peptide mixtures are typically co-selected with a population of interfering ions. Co-selection of such ion background contributes to the isobaric tag reporter ion ratios [28]. Setting the ion precursor selection window to 0.5 m/z notably improves the accuracy and precision of quantification but lowers spectral quality and sensitivity. Reciprocally, applying a higher precursor ion selection window of 1.6 m/z will allow higher rate of identification, but the precision of quantification is typically poorer. Similar approaches are also applicable to other mass spectrometers used for isobaric tag experiments, such as hybrid quadrupole time of flight and quadrupole-Orbitrap mass spectrometers. An alternative approach using MS3 on Orbitrap mass spectrometers has been proposed and indicates that the precursor co-isolation effect in complex samples can be addressed to a large extent by relying on the reporter ion intensities extracted from MS3 spectra [29, 30]. A novel Fourier Transform MS3 Synchronous Precursor Selection method (SPS-MS3 fragmentation) on the Orbitrap Fusion showed around 20% reduction in the number of identified proteins but improved quantitation accuracy and quantitation of 95% of the identified proteins [31]. 21. Automatic gain control (AGC) is a software feature in mass spectrometers that predicts the time (in ms) required to fill an ion trap with a certain number of ions using the intensities of the ions from the precursor scan. This is important as underfilling of the ion trap could result in low sensitivity while overfilling could lead to space-charge effects. 22. The most commonly used fragmentation techniques are collision-induced dissociation (CID) and higher-energy collisional dissociation (HCD). HCD is a CID technique specific to Orbitrap and Thermo Scientific ion trap instruments that allow making use of the high resolution of the Orbitrap mass spectrometer or—when used in ion traps—allows the detection of lower m/z fragments that are otherwise usually lost in CID MS/MS experiments in ion traps. TMT-based quantification requires the detection of low masses (126–135). Therefore, HCD is essential for these types of experiments in ion trap/ Orbitrap hybrid instruments.

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23. Label-free samples can also be acquired in data-dependent acquisition mode. Set up the Orbitrap Fusion as follows: set up the instrument in “Top Speed” data-dependent mode, operated in positive ion mode. FullScan spectra are acquired in a range from 400 to 1600 m/z, at a resolution of 120,000 (at 200 m/z), with an automated gain control (AGC) (see Note 21) of 3.0 × 105 and a maximum injection time of 50 ms. Charge state screening is enabled to exclude precursors with a charge state of 1. The intensity threshold for a MS/MS fragmentation is set to 104 counts. The most intense precursor ions are isolated with a quadrupole mass filter width of 1.6 m/z, and collision-induced dissociation (CID fragmentation) (see Note 22) is performed in one-step collision energy of 32% and activation Q of 0.25. MS/MS fragments ions are analyzed in the segmented linear ion trap with a normal scan range, in a rapid mode. The detection of MS/MS fragments is set up as the “Universal Method,” using a maximum injection time of 300 ms and a maximum AGC of 103 ions. 24. The iBAQ (intensity-based absolute quantification) method is an algorithm implemented in MaxQuant search engine that estimates the absolute abundance of a protein by taking the sum of intensities of identified peptides of proteins, normalized to the length and theoretical number of possible peptides.

Acknowledgments This research was partly funded by a Wellcome Trust Investigator Award (215542/Z/19/Z) to M.T and Knut and Alice Wallenberg Foundation, University of Gothenburg (Sweden), to A.H. M.E.D. is a Marie Sklodowska Curie Fellow within the European Union’s Horizon 2020 research and innovation program under the Marie Skłodowska-Curie grant agreement No. 890296. References 1. Kinchen JM, Ravichandran KS (2008) Phagosome maturation: going through the acid test. Nat Rev Mol Cell Biol 9(10):781–795 2. Gutierrez MG (2013) Functional role(s) of phagosomal Rab GTPases. Small GTPases 4(3):148–158 3. Levin R, Grinstein S, Schlam D (2015) Phosphoinositides in phagocytosis and macropinocytosis. Biochim Biophys Acta Mol Cell Biol Lipids 1851(6):805–823 4. Pauwels A-M, H€artlova A, Peltier J et al (2019) Spatiotemporal changes of the Phagosomal proteome in dendritic cells in response to LPS

stimulation. Mol Cell Proteomics 18(5): 909–922 5. Gagnon E, Duclos S, Rondeau C et al (2002) Endoplasmic reticulum-mediated phagocytosis is a mechanism of entry into macrophages. Cell 110(1):119–131 6. Campbell-Valois F-X, Trost M, Chemali M et al (2012) Quantitative proteomics reveals that only a subset of the endoplasmic reticulum contributes to the phagosome. Mol Cell Proteomics 11(7):M111.016378 7. Houde M, Bertholet S, Gagnon E et al (2003) Phagosomes are competent organelles for

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antigen cross-presentation. Nature 425(6956): 402–406 8. Guermonprez P, Saveanu L, Kleijmeer M et al (2003) ER–phagosome fusion defines an MHC class I cross-presentation compartment in dendritic cells. Nature 425(6956):397–402 9. Trost M, English L, Lemieux S et al (2009) The Phagosomal proteome in interferon-γ-activated macrophages. Immunity 30(1): 143–154 10. Boulais J, Trost M, Landry CR et al (2010) Molecular characterization of the evolution of phagosomes. Mol Syst Biol 6(1):423 11. Desjardins M, Huber LA, Parton RG et al (1994) Biogenesis of phagolysosomes proceeds through a sequential series of interactions with the endocytic apparatus. J Cell Biol 124(5): 677–688 12. Garin J, Diez R, Kieffer S et al (2001) The phagosome proteome: insight into phagosome functions. J Cell Biol 152(1):165–180 13. Ong SE, Blagoev B, Kratchmarova I et al (2002) Stable isotope labeling by amino acids in cell culture, SILAC, as a simple and accurate approach to expression proteomics. Mol Cell Proteomics 1(5):376–386 14. Boersema PJ, Raijmakers R, Lemeer S et al (2009) Multiplex peptide stable isotope dimethyl labeling for quantitative proteomics. Nat Protoc 4(4):484–494 15. Li J, Van Vranken JG, Pontano Vaites L et al (2020) TMTpro reagents: a set of isobaric labeling mass tags enables simultaneous proteome-wide measurements across 16 samples. Nat Methods 17(4):399–404 16. Dill BD, Gierlinski M, H€artlova A et al (2015) Quantitative proteome analysis of temporally resolved phagosomes following uptake via key phagocytic receptors. Mol Cell Proteomics 14(5):1334–1349 17. Guo M, H€artlova A, Dill BD et al (2015) High-resolution quantitative proteome analysis reveals substantial differences between phagosomes of RAW 264.7 and bone marrow derived macrophages. Proteomics 15(18): 3169–3174 18. Dean P, Heunis T, H€artlova A et al (2019) Regulation of phagosome functions by posttranslational modifications: a new paradigm. Curr Opin Chem Biol 48:73–80 19. Breyer F, H€artlova A, Thurston T et al (2021) TPL-2 kinase induces phagosome acidification to promote macrophage killing of bacteria. EMBO J 40(10):e106188 20. Guo M, H€artlova A, Gierlin´ski M et al (2019) Triggering MSR1 promotes JNK-mediated

inflammation in IL-4-activated macrophages. EMBO J 38(11):e100299 21. Bilkei-Gorzo O, Heunis T, Marı´n-Rubio JL et al (2022) The E3 ubiquitin ligase RNF115 regulates phagosome maturation and host response to bacterial infection. bioRxiv 2022: 20210713452284 22. Pino LK, Just SC, MacCoss MJ et al (2020) Acquiring and analyzing data independent acquisition proteomics experiments without Spectrum libraries. Mol Cell Proteomics 19(7):1088–1103 23. Cox J, Mann M (2008) MaxQuant enables high peptide identification rates, individualized p.p.b.-range mass accuracies and proteomewide protein quantification. Nat Biotechnol 26(12):1367–1372 24. Demichev V, Messner CB, Vernardis SI et al (2020) DIA-NN: neural networks and interference correction enable deep proteome coverage in high throughput. Nat Methods 17(1): 41–44 25. Ritchie ME, Phipson B, Wu D et al (2015) Limma powers differential expression analyses for RNA-sequencing and microarray studies. Nucleic Acids Res 43(7):e47–e ˜ as ME et al 26. Hatton CF, Botting RA, Duen (2021) Delayed induction of type I and III interferons mediates nasal epithelial cell permissiveness to SARS-CoV-2. Nat Commun 12(1):7092 27. Heunis T, Lamoliatte F, Marı´n-Rubio JL et al (2020) Technical report: targeted proteomic analysis reveals enrichment of atypical ubiquitin chains in contractile murine tissues. J Proteome Res 229:103963 28. Savitski MM, Sweetman G, Askenazi M et al (2011) Delayed fragmentation and optimized isolation width settings for improvement of protein identification and accuracy of isobaric mass tag quantification on Orbitrap-type mass spectrometers. Anal Chem 83(23):8959–8967 29. McAlister GC, Nusinow DP, Jedrychowski MP et al (2014) MultiNotch MS3 enables accurate, sensitive, and multiplexed detection of differential expression across cancer cell line proteomes. Anal Chem 86(14):7150–7158 30. Ting L, Rad R, Gygi SP et al (2011) MS3 eliminates ratio distortion in isobaric multiplexed quantitative proteomics. Nat Methods 8(11):937–940 31. Erickson BK, Jedrychowski MP, McAlister GC et al (2015) Evaluating multiplexed quantitative Phosphopeptide analysis on a hybrid quadrupole mass filter/linear ion trap/Orbitrap mass spectrometer. Anal Chem 87(2): 1241–1249

Chapter 24 Using Ion Substitution and Fluid Indicators to Monitor Macropinosome Dynamics in Live Cells Guillermo A. de Paz Linares, Spencer A. Freeman, and Ruiqi Cai Abstract All forms of endocytosis involve the incidental uptake of fluid (pinocytosis). Macropinocytosis is a specialized type of endocytosis that results in the bulk ingestion of extracellular fluid via large (>0.2 μm) vacuoles called macropinosomes. The process is a means of immune surveillance, a point of entry for intracellular pathogens, and a source of nutrients for proliferating cancer cells. Macropinocytosis has also recently emerged as a tractable system that can be experimentally exploited to understand fluid handling in the endocytic pathway. In this chapter, we describe how stimulating macropinocytosis in the presence of extracellular fluids of a defined ionic composition can be combined with high-resolution microscopy to understand the role of ion transport in controlling membrane traffic. Key words Endocytosis, Pinocytosis, Macrophage, Ion channels, Ion transporters

1

Introduction Macropinocytosis was first described nearly 100 years ago [1]. The advent of model organisms [2] and advanced microscopy [3, 4] has further enabled its characterization beyond the seminal studies in myeloid cells first initiated by Steinman and Cohn [5, 6] and followed by those in transformed cells, macrophages, and dendritic cells by Bar-Sagi [7], Swanson [8, 9], and Watts [10]. Generally, macropinocytosis follows a systematic template largely conserved through evolution [4, 11]. Macropinosomes form when F-actindriven ruffles that extend out from the body of the cell fold back to the membrane and fuse, resulting in a membrane-bound vacuole [12]. The resultant macropinosome is >0.2 μm in size [12] and can reach more than 5 μm in diameter, thus containing large amounts of extracellular fluid. Macropinocytosis can be made constitutively active in myeloid cells when supported by G-protein-coupled receptors (GPCRs) that sense readily available ligands [13]. Alternatively, macropinocytosis can be induced when cells are stimulated

Roberto J. Botelho (ed.), Phagocytosis and Phagosomes: Methods and Protocols, Methods in Molecular Biology, vol. 2692, https://doi.org/10.1007/978-1-0716-3338-0_24, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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Fig. 1 Membrane remodelling and ion fluxes during macropinocytosis. Macropinocytosis is important for immune surveillance, is a route of entry for pathogens, and is employed by malignant cells to support their growth (left). In all cases, the signaling pathways that stimulate macropinocytosis converge on the robust, localized assembly of submembranous branched F-actin networks that propel membrane ruffling (right). Ras-GTP and class I PI3 kinases are recruited in order to generate phosphoinositides that favor docking of Rac and the SCAR/WAVE complex to the membrane. These interactions are stabilized by electrostatically attracting moieties, i.e., cationic stretches in the proteins and negatively charged head groups of the phosphoinositides. Obstructing the electrostatic binding of Rho GTPases and negatively charged inner leaflet lipids, as occurs upon the inhibition of Na+-H+ exchangers and the build-up of H+, arrests macropinocytosis. Macropinosomes entrap a variety of solutes, though the predominant osmolytes are Na+ and Cl-. Upon scission from the membrane, Na+ and Cl- are effluxed through TPCs and ASOR, respectively. Water follows by osmosis, though the permeating channels are not known

by growth factors, including epithelial growth factor (EGF), macrophage colony-stimulating factor (M-CSF), and platelet-derived growth factor (PDGF), pathogens, or soluble stimuli like microbial-associated molecular patterns (e.g., lipopolysaccharide) [14] (see Fig. 1). Activation of GPCRs, receptor tyrosine kinases, or Toll-like receptors all leads to the recruitment and activation of phosphatidylinositol-3-kinases (PI3K) as well as Ras and Rho GTPases [3, 15, 16]. In turn, these signaling hubs fuel the activity of actin nucleation factors that branch actin filaments and buttress the membrane dynamics and fusion required to complete macropinocytosis. Not surprisingly then, a wide segment of cancer cells expressing oncogenic forms of RAS or active PI3K undergo macropinocytosis constitutively [7, 17], which has been proposed to provide proliferating tumors with nutrients from their fluid environment [18–20]. Similarly, viruses that enter cells by triggering macropinocytosis do so by activating PI3K, Rac, and Ras [21] (see Fig. 1).

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The uptake of fluid, together with cell surface membrane, is an efficient means to survey the microenvironment and internalize receptors and their ligands without the need for restricting endocytic programs to small regions. Macropinocytosis is thus envisaged to be an important ongoing program that supports immune surveillance [10, 13]. In addition, and most relevant to this chapter, since all forms of endocytosis involve pinocytosis, macropinocytosis provides a tractable system to study fluid handling in the endocytic pathway as it results in non-diffraction limited endosomes. Additionally, we [22] and others [23] have reported that the robust macropinocytic programs of macrophages can proceed despite acute changes in the ionic composition of the extracellular fluid. This has enabled ion substitution experiments where new vacuoles formed (indicated with fluorescent dyes) contain fluid that is initially identical to that of the extracellular fluid engulfed. Macropinosomes can in fact be delineated from other endosomes derived from types of endocytosis by the same means as they accommodate larger macromolecules, i.e., fluorescently labelled 70 kDa dextran, while other forms do not. Below, we describe a method in which the major osmolytes of macropinosomes, Na+ and Cl-, are substituted. Therefore, a final and important consideration for these experiments is that mammalian membranes are generally made permeable to water, mediated in part by aquaporins, and cannot maintain an osmotic gradient. While the nature of organellar aquaporins is poorly understood [24], endomembranes are also considered to be permeable to water. Like virtually all interstitial fluids, the osmolarity of organelles and the cytosol is ~325 mOsm (see Fig. 1). As macropinosomes mature, they lose fluid volume by expelling their major osmolytes through channels down their concentration gradient. For Na+, the efflux occurs via two-pore channels (TPCs) [22] which are gated by the phosphoinositide PtdIns(3,5)P2. The parallel efflux of Cl-, which maintains electroneutrality to the process, occurs via the acid-activated anion channel (ASOR), composed of TMEM206 [23]. As its name implies, ASOR is activated by H+. The early acidification of the macropinosome is critical for the resolution of its internalized volume and seems to be mediated by another Cl- transport pathway involving the H+:Cl- exchanger, ClC-5 [23]. ClC-5, in principle, would support both Cl- efflux and early acidification [25]. These initial studies using macropinosomes have interrogated mechanisms of fluid handling in the early endocytic pathway and served to uncover the transport pathways for major osmolytes. Similar approaches can be taken to analyze a variety of solutes and solute transporters. Therefore, while we describe how ion substitutions for Na+ and Cl- can be done below, the same experimental approach can grossly be applied to better elucidate the role of solute transport in organelles.

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Materials Cell Culture

1. Bone marrow-derived macrophages (BMDM). 2. Culture medium: Dulbecco’s Modified Eagle Medium supplemented with 4 g/L glucose, 4 mM L-glutamine, and 1 mM pyruvate, stored at 4 °C. 3. Culture medium: Low glucose Dulbecco’s Modified Eagle Medium supplemented with 1.0 g/L glucose, 4 mM L-glutamine, and 1 mM pyruvate, stored at 4 °C. 4. Heat inactivated FBS. 5. 10% L929 cell fibroblast-conditioned medium for M-CSF. 6. 0.45 μm membrane filter. 7. Microcentrifuge tubes (autoclaved) and microcentrifuge. 8. Non-tissue culture treated Petri dishes (100 × 15 mm) and tissue culture plates (12 wells). 9. Pipette tips and serological pipettes. 10. Round glass coverslips (18 mm; autoclaved). 11. Cell culture incubator with water tray maintained at 37 °C and 5% CO2.

2.2 Primary Macrophage Derivation

1. Sterilized phosphate-buffered saline (PBS) without calcium and magnesium. 2. 5 mM EDTA in PBS (filtered with a 0.22 μm membrane), stored at 4 °C. 3. 15 mL centrifuge tubes. 4. 400 μL PCR tubes. 5. Sterilized transfer pipette. 6. 100× antibiotic–antimycotic. 7. 10 cm3 syringes. 8. M-CSF ELISA kit (R & D systems, #MMC00B) (see Note 1). 9. Surgical instruments.

2.3

Buffers

1. Control buffer: 150 mM NaCl, 1 mM MgCl2, 1 mM CaCl2, 5 mM KCl, 20 mM HEPES, pH 7.2. 2. Na+-substitution buffer: 150 mM N-methyl-glucamine-chloride (NMG-Cl), 1 mM MgCl2, 1 mM CaCl2, 5 mM KCl, 20 mM HEPES, pH 7.2. 3. Cl--substitution buffer: 150 Na-gluconate, 1 mM MgCl2, 1 mM CaCl2, 5 mM KCl, 20 mM HEPES, pH 7.2.

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2.4

Chemicals

379

1. NaCl, NMG-Cl, Na-gluconate, MgCl2, CaCl2, KCl, HEPES. 2. Recombinant murine M-CSF (PeproTech, #315-02). 3. Dextran: 70,000 MW TRITC-dextran (TdB labs) (see Notes 2, 3, and 4). 4. Wheat germ agglutinin (WGA) Fluor™ 647 conjugate (ThermoFisher, W32466). 5. Pattern Recognition Receptors (PRR) ligands: muramyl dipeptide (MDP).

2.5 Confocal Microscope

1. A spinning disk microscope equipped with a 63× magnifying objective with numeric aperture 1.4. An EM-CCD camera is used to capture the images (see Note 5). 2. Chamlide magnetic chambers (Quorum Technologies, CM-B18-1) which fit 18 mm rounded coverslips were used for live cell imaging. 3. Computer installed with imaging software, i.e., Volocity 6.3, FIJI (ImageJ).

3 3.1 3.1.1 CSF

Methods Deriving BMDM Preparation of M-

1. Plate L292 cells in round 100 mm × 15 mm cell culture Petri dishes in DMEM supplemented with 10% FBS (see Note 6). 2. Incubate cells for 2–3 days at 5% CO2 and 37 °C, until the cells have reached 90–100% confluency. 3. Aspirate medium from the plates, and add 10 mL of fresh low glucose DMEM without FBS. 4. Incubate cells for another 7–10 days at 5% CO2 and 37 °C without changing the medium. 5. Harvest the supernatant when the cells are just starting to become dislodged from the plate. 6. Filter the supernatant with a 0.45 μm filter into 15 mL centrifuge tubes, and centrifuge at 300 g for 5 min to remove any leftover cellular debris. 7. Perform an assay to determine the concentration of M-CSF using an ELISA kit (use according to manufacturer’s instructions). The final concentration of M-CSF should be 100 ng/ mL for deriving macrophages: bring the M-CSF containing solution to a 10× working concentration by diluting M-CSF in complete DMEM. 8. Store M-CSF-containing medium at -20 °C for up to 1 month (see Note 7).

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3.1.2 Isolation of BMDMs from Murine Long Bones

1. After sacrificing animals (see Note 8), isolate the femur, tibia, and humerus, and keep bones at 4 °C until ready to use. Ensure that bones have not been broken, which may result in contamination. 2. Cut the bottom of a 400 μL PCR tube, and place inside a 1.5 mL microcentrifuge tube, ensuring the bottom hole is not big enough to allow for the bones to pass through. 3. Pipette 300 μL of cold PBS supplemented with antibiotic– antimycotic into the 1.5 mL tube. 4. Cut one end of each bone, and place one to two bones inside each of the 400 μL PCR tubes. Each mouse will yield six bones in total, which equals to three to six 400 μL PCR tubes each inside a 1.5 mL microfuge tube. The humerus will yield the least amount of bone marrow, so harvesting from these long bones is optional. 5. Place in a centrifuge and spin for 10 s (pulse spin, reaching ~9000 Gs). Following centrifugation, a red pellet should form at the bottom of the 1.5 mL tube, inside the PBS. Ensure that no solid debris from the bones escaped into the 1.5 mL tube. 6. Discard bones along with 400 μL tube, and bring the 1.5 mL microcentrifuge tube with the pellet to a BSC for sterile work environment (see Note 6). 7. Using a sterile transfer pipette, resuspend the pellet in the PBS, and transfer to a 15 mL conical tube. 8. Add enough complete DMEM to the 15 mL conical tube to bring the volume to ~10 mL. 9. Centrifuge at 1500 rpm for 5 min at room temperature. 10. Aspirate supernatant, and resuspend pellet in 6 mL of complete DMEM to yield six plates of BMDM. 11. In 100 mm × 15 mm non-tissue culture-treated Petri dishes, add 8 mL of complete DMEM, 1 mL of M-CSF (L929 medium), and 1 mL of resuspended cells (the final volume in each plate should be 10 mL). 12. Store in a 37 °C incubator at 5.0% CO2 for 3–4 days before replating and using for experiments.

3.2 Seeding BMDM onto Coverslips

1. After 4–5 days of culturing in complete medium supplemented with 10% M-CSF on Petri dishes, BMDM should be lifted using 5 mM EDTA in PBS, pH 7.4. This takes ~5 min and can require gentle scraping. 2. Lifted BMDMs can then be centrifuged at 1000 rpm for 3 min. 3. Discard the supernatant, and resuspend the pellet in pre-warmed complete medium with 10% M-CSF. 4. Seed BMDM onto coverslips in a 12-well plate for 24–48 h. 5. Incubate cells in incubator with 5% CO2 at 37 °C.

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3.3 Inducing Macropinocytosis

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1. To induce macropinocytosis with M-CSF, add 1:40 of the M-CSF L929 medium in culture medium or control, Na+substituted or Cl--substituted buffer, and along with 70 kDa TRITC-dextran. The L929 medium will add 3–4 mM concentration of Na+ and Cl- to the final concentration. This can be avoided by using recombinant M-CSF resuspended in the defined medium (see Note 9). 2. Put the plate in incubator for 5 min. 3. Wash the cells with ice-cold HBSS 3 times. 4. Transfer the coverslip to cold HBSS with 0.1% (v/v) WGA on ice for 1 min. 5. Wash the cells with ice-cold HBSS 3 times. 6. Mount the coverslip onto magnetic imaging chambers. Switch the medium to pre-warmed complete DMEM if using a CO2controlled imaging system or HBSS with 5 mM glucose if this is not available.

3.4 Imaging and Recording

1. Switch on the spinning disk microscope, lamp, lasers, and camera in advance, especially if using an EM-CCD camera. 2. Launch the relevant imaging software (i.e., Volocity). 3. Set up file name and acquisition parameters including exposure time, laser intensity, and camera intensity for each channel. 4. Set the shutter to Balanced Sample Protection. 5. Set timelapse to minimize photobleaching and phototoxicity, but to acquire at a sufficient frame rate to capture the relevant kinetics. One can bin the pixels to increase the image acquisition speed and decrease photobleaching/phototoxicity. We opt to bin 4× and acquire a stacked image every 10 s. 6. Add a small amount of oil to the 63× objective, and mount imaging chamber onto the objective. 7. Select fields of interest and record (Fig. 2).

3.5

Data Analysis

1. Open image files with Volocity. 2. Select Measurement and then Find Objectives. 3. Assign the corresponding channel, i.e., Cy3 Confocal, Cy5 confocal. 4. Set the minimum objective size with diameter >0.2 μm. 5. Automatically count the number of macropinosomes (Nm) and, if desired, sum of dextran intensity (Id). 6. Manually count the cell number depending on WGA staining (Nc). 7. Calculate the number of macropinosomes per cell as NNmc and the dextran mean intensity per cell as NI dc (see Note 10).

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Fig. 2 Confocal images showing macropinosomes in BMDM containing indicated buffers. WGA, wheat germ agglutin Alexa-647. Scale bar, 10 μm. Images are taken 10 min after changing the medium

4

Notes 1. There are other sources of this product. We use this specific product in our lab. The corresponding company names and catalogue numbers are listed hereafter where applicable. 2. Multiple molecular weights of dextran are commercially available, e.g., 10,000, 70,000, and 150,000 Da. Use of 70,000 Da dextran is the gold standard to distinguish macropinosomes from other endosomes. The use of small, bright dyes may be useful to observe vacuole substructures (tubules and vesicles). 3. Charged dextrans can be clumpy and have their own caveats vis-a`-vis charge density in the macropinosome and potential unwanted effects on membrane potential. We have also noticed that even uncharged, conjugated dextrans from ThermoFisher tend to clump. Clumped dextrans will skew data considerably: either sonicate the aggregated dextrans, or, preferably, find a source of conjugated dextrans that are not aggregated. 4. Red (TMR/TRITC) dextrans are preferred since they do not readily photobleach and are less pH sensitive. On the other hand, FITC-dextrans have been used extensively for ratiometric measurements, including in the endocytic pathway. Far red dextrans are easily bleached so we tend to avoid using these. 5. We used an Axiovert 200 M spinning disk microscope with an EM-CCD camera (Hamamatsu). This system has a XY-stage with a Piezo1 Z-focus drive and a set of diode pumped solidstate lasers emitting in the wavelengths of 405, 440, 561, 638, and 655 nm. For this method, however, only the 561 and 655 nm lasers were used.

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6. All procedures for cell culturing must be conducted in a BSC to maintain a sterile work environment and avoid contamination of BMDMs. 7. Ready-to-use MCSF may also be purchased directly from scientific vendors. If so, begin the protocol at Subheading 3.1.2. 8. All animals utilized in the described methods are subjected to approval by Research and Ethics Board from individual institutions. 9. Drugs such as PIKfvye inhibitors (e.g., YM201636), the two pore channel (TPC) inhibitor tetrandrine, the PAC/ASOR channel blockers 4,4’-Diisothiocyanatostilbene-2,2’-disμLfonate (DIDS), niflumic acid (NFA), or 5-Nitro-2(3-phenylpropylamino)benzoic acid (NPBB) have all been used to prevent macropinosome shrinkage. These could be used as controls in experiments. 10. This method is also applicable for other myeloid cells or cancer cells undergoing macropinocytosis. References 1. Lewis WH (1937) Pinocytosis by malignant cells. Am J Cancer 29(4):666–679 2. Kay RR, Lutton J, Coker H, Paschke P, King JS, Bretschneider T (2022) The Amoebal Model for macropinocytosis. Subcell Biochem 98:41–59. https://doi.org/10.1007/978-3030-94004-1_3 3. Quinn SE, Huang L, Kerkvliet JG, Swanson JA, Smith S, Hoppe AD et al (2021) The structural dynamics of macropinosome formation and PI3-kinase-mediated sealing revealed by lattice light sheet microscopy. Nat Commun 12(1):4838. https://doi.org/10.1038/ s41467-021-25187-1 4. Veltman DM, Williams TD, Bloomfield G, Chen BC, Betzig E, Insall RH et al (2016) A plasma membrane template for macropinocytic cups. elife:5. https://doi.org/10.7554/eLife. 20085 5. Steinman RM, Swanson J (1995) The endocytic activity of dendritic cells. J Exp Med 182(2): 283–288. https://doi.org/10.1084/jem.182. 2.283 6. Steinman RM, Brodie SE, Cohn ZA (1976) Membrane flow during pinocytosis. A stereologic analysis. J Cell Biol 68(3):665–687. https://doi.org/10.1083/jcb.68.3.665 7. Bar-Sagi D, Feramisco JR (1986) Induction of membrane ruffling and fluid-phase pinocytosis in quiescent fibroblasts by ras proteins. Science 233(4768):1061–1068. https://doi.org/10. 1126/science.3090687

8. Racoosin EL, Swanson JA (1993) Macropinosome maturation and fusion with tubular lysosomes in macrophages. J Cell Biol 121(5): 1011–1020. https://doi.org/10.1083/jcb. 121.5.1011 9. Swanson JA (1989) Phorbol esters stimulate macropinocytosis and solute flow through macrophages. J Cell Sci 94(Pt 1):135–142 10. Norbury CC, Hewlett LJ, Prescott AR, Shastri N, Watts C (1995) Class I MHC presentation of exogenous soluble antigen via macropinocytosis in bone marrow macrophages. Immunity 3(6):783–791. https://doi. org/10.1016/1074-7613(95)90067-5 11. King JS, Kay RR (2019) The origins and evolution of macropinocytosis. Philos Trans R Soc Lond Ser B Biol Sci 374(1765):20180158. https://doi.org/10.1098/rstb.2018.0158 12. Swanson JA, Watts C (1995) Macropinocytosis. Trends Cell Biol 5(11):424–428. https:// doi.org/10.1016/s0962-8924(00)89101-1 13. Canton J, Schlam D, Breuer C, Gutschow M, Glogauer M, Grinstein S (2016) Calciumsensing receptors signal constitutive macropinocytosis and facilitate the uptake of NOD2 ligands in macrophages. Nat Commun 7: 1 1 2 8 4 . h t t p s : // d o i . o r g / 1 0 . 1 0 3 8 / ncomms11284 14. Marques PE, Grinstein S, Freeman SA (2017) SnapShot: macropinocytosis. Cell 169(4): 766–7e1. https://doi.org/10.1016/j.cell. 2017.04.031

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15. Swanson JA (2008) Shaping cups into phagosomes and macropinosomes. Nat Rev Mol Cell Biol 9(8):639–649. https://doi.org/10. 1038/nrm2447 16. Araki N, Johnson MT, Swanson JA (1996) A role for phosphoinositide 3-kinase in the completion of macropinocytosis and phagocytosis by macrophages. J Cell Biol 135(5): 1249–1260. https://doi.org/10.1083/jcb. 135.5.1249 17. Bar-Sagi D, McCormick F, Milley RJ, Feramisco JR (1987) Inhibition of cell surface ruffling and fluid-phase pinocytosis by microinjection of anti-ras antibodies into living cells. J Cell Physiol Suppl Suppl 5:69–73. https://doi.org/10.1002/jcp.1041330414 18. Commisso C, Davidson SM, SoydanerAzeloglu RG, Parker SJ, Kamphorst JJ, Hackett S et al (2013) Macropinocytosis of protein is an amino acid supply route in Ras-transformed cells. Nature 497(7451): 6 3 3 – 6 3 7 . h t t p s : // d o i . o r g / 1 0 . 1 0 3 8 / nature12138 19. Palm W, Park Y, Wright K, Pavlova NN, Tuveson DA, Thompson CB (2015) The utilization of extracellular proteins as nutrients is suppressed by mTORC1. Cell 162(2):259–270. https://doi.org/10.1016/j.cell.2015.06.017 20. Kamphorst JJ, Nofal M, Commisso C, Hackett SR, Lu W, Grabocka E et al (2015) Human

pancreatic cancer tumors are nutrient poor and tumor cells actively scavenge extracellular protein. Cancer Res 75(3):544–553. https:// doi.org/10.1158/0008-5472.CAN-14-2211 21. Mercer J, Helenius A (2009) Virus entry by macropinocytosis. Nat Cell Biol 11(5): 5 1 0 – 5 2 0 . h t t p s : // d o i . o r g / 1 0 . 1 0 3 8 / ncb0509-510 22. Freeman SA, Uderhardt S, Saric A, Collins RF, Buckley CM, Mylvaganam S et al (2020) Lipidgated monovalent ion fluxes regulate endocytic traffic and support immune surveillance. Science 367(6475):301–305. https://doi.org/ 10.1126/science.aaw9544 23. Zeziulia M, Blin S, Schmitt FW, Lehmann M, Jentsch TJ (2022) Proton-gated anion transport governs macropinosome shrinkage. Nat Cell Biol 24(6):885–895. https://doi.org/10. 1038/s41556-022-00912-0 24. Nozaki K, Ishii D, Ishibashi K (2008) Intracellular aquaporins: clues for intracellular water transport? Pflugers Arch 456(4):701–707. https://doi.org/10.1007/s00424-0070373-5 25. Gunther W, Piwon N, Jentsch TJ (2003) The ClC-5 chloride channel knock-out mouse - an animal model for Dent’s disease. Pflugers Arch 445(4):456–462. https://doi.org/10.1007/ s00424-002-0950-6

INDEX A Acidification....................................... 103, 140, 154, 159, 163, 199, 261, 348–350, 356, 377 Actin...........................................2, 25, 26, 62, 63, 80, 87, 89, 102, 122, 245, 345, 362, 370, 376 Antibodies ........................................8, 16, 18–21, 26, 28, 29, 31–34, 36–38, 45, 50, 52, 57, 66, 68, 72, 73, 76, 80–83, 87, 88, 92, 98, 101, 102, 104–106, 122–128, 130–133, 135, 166, 223, 226, 233, 234, 240, 244, 263–266, 269–272, 276, 278, 281, 283, 294, 305, 317, 318, 325, 327, 328, 330, 332, 334 Antigen presentation.................................. 277, 280–283, 311, 313, 315, 317, 318, 320, 321, 323, 324, 329, 361, 362 Apoptosis .................................................... 41, 42, 49, 50, 56, 62, 153, 291, 313 ASC speck ............................................290, 295, 300, 301 Autophagy ........................................................... 171, 210, 312, 313, 322, 323, 325–329

B Bacterial killing.................................................4, 5, 10, 13 B cells .......................................................... 311, 313, 314, 317, 319, 325–329, 333, 334

C Caenorhabditis elegans embryos ................................... 338 Chloroquine resistance assay ............................... 213–216 Colony forming units (CFUs) .................................... 7–9, 11–13, 212–216, 218 Conditional immortalization............................... 109, 174 Confocal microscopy ................................ 35, 36, 80, 133 Co-occurrence ..................................................... 222, 223, 227–232, 234 CRISPR/Cas9..................................................... 313, 315, 321, 325, 326, 329, 334, 353 CypHer5e ............................................................. 158–160

D Data-dependent acquisition (DDA).............................................. 364, 369, 373

Data-independent acquisition (DIA) .......................................364, 365, 368, 369 Dendritic cells (DCs) ............................................ 25, 132, 141, 174, 182, 247, 275–285, 289, 292–300, 375 Differentiation........................ 46, 65, 67, 111, 116, 117, 120, 189, 192–195, 278, 295, 297, 304, 314 Diffusion.......................................................63, 70–72, 76 Disulfide reduction ....................139, 140, 142, 146, 147 Dual fluorophore ratio imaging .......................... 159, 162 Dual wavelength ratio imaging ........................... 161, 162

E Eα .........................................................276, 278, 280–285 Efferocytosis ................................. 41–58, 62–64, 76, 153 Endocytosis ............................................................ 76, 377 Epithelial cells....................................................... 209–218 Eructophagy ............................... 171–174, 178, 181–184

F Filamentous bacteria .................................. 91, 92, 94, 97, 100–102, 105, 106, 223, 225, 227 Flow cytometry ................................................... 1, 15–17, 19–22, 118, 122, 130, 131, 135, 156, 188, 276, 278, 281, 283, 284, 294, 297, 305 Fluorescein ........................ 155, 156, 158–162, 166, 167 Fluorescence microscopy .................................... 1, 36, 62, 156, 158, 160, 250, 256, 264, 270–271, 291, 300 Fluorescent reporters ........................................... 350, 356 Frustrated efferocytosis.............................................62–76 FYVE domain .............................................. 263, 346, 347

G Gentamicin protection assay....................................5, 7–9, 11, 12, 209–217

H HoxB8 .................................................................. 109, 118

I Imaging................................................. 15, 45, 50, 53–56, 62, 63, 76, 80, 83–86, 89, 101–103, 127, 150, 156–158, 160, 163, 164, 167, 172, 173, 175,

Roberto J. Botelho (ed.), Phagocytosis and Phagosomes: Methods and Protocols, Methods in Molecular Biology, vol. 2692, https://doi.org/10.1007/978-1-0716-3338-0, © The Editor(s) (if applicable) and The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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178, 179, 181, 182, 184, 197, 200, 202, 204, 205, 223, 225–227, 233, 240, 244, 292, 293, 300, 305, 340–342, 349, 354, 356, 357, 379, 381 Immunity ...........................................................2, 90, 153, 275, 313, 331, 332, 361 Immunofluorescence ........................................ 25–38, 72, 73, 100, 122, 127, 128, 131, 188, 238, 263, 265, 266, 270, 273, 282–285, 297, 304, 313, 318, 320, 324–329 Immunostaining......................................... 45, 52, 53, 57, 66, 68, 134, 222, 226, 232, 234, 270 Infection ............................................ 5, 6, 8, 11, 56, 187, 196–198, 204, 212, 213, 217, 248, 283, 319 Inflammasome ....................290–293, 298–300, 303–305 Ion channels .................................................................. 375 Ion transporters............................................................. 377

L LAMP1 ...................................................45, 54, 122, 123, 126–128, 130–133, 222, 232–234, 238, 245 LC3-associated phagosome .......................................... 312 Lipid phosphatases ........................................................ 346 Live cell imaging ........................................ 45, 50, 52–57, 66, 69, 80, 81, 83, 93, 102, 103, 106, 162, 166, 182, 188, 192, 195, 197, 200, 202, 204, 223, 225, 228, 233, 300, 304, 379 Lysosomes ................................................ 25, 26, 47, 100, 102, 103, 121, 122, 139, 140, 154, 155, 157, 173, 174, 209, 221, 232, 238, 247–258, 261, 262, 312, 338, 346–349, 355, 356, 361

M Macrophages ......................................1–3, 5, 6, 8, 10, 11, 13, 22, 25, 26, 30–35, 37, 42, 43, 45, 46, 49, 50, 52, 54–57, 62, 65, 67–69, 72, 76, 79–82, 84–88, 90–94, 98–101, 104, 109–111, 117–120, 122–124, 127–129, 131, 132, 140, 141, 145, 151, 154, 158, 159, 162, 166, 171, 173–175, 182, 187, 189, 191–195, 197–199, 202, 204, 210, 223–226, 228, 232–234, 237, 238, 240, 242, 247, 249, 250, 252, 257, 267–269, 272, 305, 311, 313–335, 337, 362, 375–379 Major Histocompatibility Complex Class II (MHC-II) ................................................. 275–285 Mass spectrometry .............................................. 263, 362, 364–366, 368, 369, 372 Membrane fusion ........................................ 102, 248, 312 Method ...........................................1, 2, 4–11, 15, 17–20, 26, 41–58, 67–75, 81, 84–87, 94–104, 111–117, 124–131, 142–150, 163–165, 175–182, 188, 191–203, 212–216, 221–235, 238, 240, 249, 263, 266–271, 294–303, 318–330, 341–352, 364–369, 377, 379–381

Microscopy .......................................................... 2, 11, 15, 25–38, 42, 45, 50, 57, 64, 66, 70–72, 80, 83, 85, 89, 98, 103, 104, 122, 126–130, 133, 163, 238, 241, 249, 263, 265–266, 273, 299, 304, 317, 325, 327, 328, 330, 338, 340, 345, 347, 348, 350, 351 Mycobacterium ..................................................... 122, 187, 189, 195, 196, 202, 247

N Neutrophils............................................. 1, 15–19, 21, 22, 25, 44, 49–51, 55, 56, 92, 132, 158, 159, 247 NLR family CARD containing 4 (NLRC4) ............... 290–293, 298–300, 304, 305 NLR family pyrin domain containing 3 (NLRP3) .................................................. 290, 292, 293, 299, 300, 304, 305

O Oregon Green ............................155, 158, 159, 162, 167 Organelles................................................... 122, 132, 140, 154, 155, 157, 187, 188, 190, 197–200, 238, 289, 312, 338, 348, 353, 361, 377 Oxidation........................................................58, 147, 152

P Particle binding ...........................................................8, 37 Pattern recognition receptors (PRRs) ........................237, 275–278, 283, 289, 290, 292, 379 Peritonitis ........................................................................ 16 pH ...................................... 251, 262–264, 272, 339, 346 Phagocytes .........................................................1, 2, 8, 25, 26, 35, 41, 42, 45, 79, 91, 92, 126–129, 132, 139, 140, 148–150, 154, 155, 165, 182, 187, 221, 257, 261, 271, 275, 289, 337, 339 Phagocytic cup formation................................54, 92, 345 Phagocytic index .................................................... 35, 100 Phagocytosis ........................................... 1, 2, 5, 8, 15–21, 25–38, 58, 62, 79, 80, 83–85, 87, 90–106, 121, 125, 128, 132, 133, 147, 153, 154, 162, 164–166, 172, 187, 191, 195, 221, 224–225, 228, 232, 233, 237–241, 245, 247, 258, 261, 262, 272, 276–278, 280–282, 285, 289, 312, 317, 323–324, 337, 338, 350, 351, 356, 361 Phagolysosome.................................................... 121, 122, 171–174, 178, 181, 184, 188, 221, 222, 224, 225, 234, 238, 245, 247, 251, 255, 258, 261–263, 265, 268–269, 271–273, 338, 347–352, 355–357, 361 Phagolysosome resolution ............................................ 352 Phagosome-derived vesicles (PDVs)...........................221, 222, 224–235 Phagosome lysosome fusion......................................... 350

PHAGOCYTOSIS Phagosome maturation......................................... 92, 102, 121, 122, 126–128, 132, 133, 160, 187–205, 238, 244, 247–250, 261, 262, 275, 277, 280, 338, 346–348, 355, 362 Phagosome resolution ........................122, 221–235, 262 Phagosomes .......................................................25, 26, 32, 80, 91, 92, 98, 102–104, 121–123, 126–135, 139, 140, 147, 149, 150, 154, 156–159, 162, 164, 165, 167, 172–174, 180, 182–184, 187, 188, 191, 197, 199, 200, 210, 221–224, 227–230, 233, 234, 237, 238, 242–245, 247–258, 261–264, 268–273, 275–285, 289, 290, 312, 345–350, 355, 361, 362, 364, 365, 367, 369, 370 Phosphatidylinositol phosphates (PIPs) ................................................248, 261–273 Phosphoinositide detection ................................. 261–273 Phosphoinositide kinases ............................ 262, 264, 269 Phosphoinositide lipids........................................ 262, 263 Phosphoinositide metabolism ............................. 261–273 pHrodo .............................................................. 17–20, 22, 158–160, 174, 175, 177, 183 Pinocytosis ..................................................................... 377 Podosomes ................................................................80–82 Proteolysis.................................................... 140, 172, 175 Pyroptosis ............................................................. 210, 290

Q Quantification ....................................................... 1, 3, 29, 34–35, 38, 42, 48, 50, 53, 87, 126, 127, 131, 134, 175, 181, 198, 221–235, 264, 270–271, 276, 305, 328, 362, 368–373 Quantitative proteomics .....................362, 364, 366, 367

R Rab GTPases .......................................248, 264, 265, 346 Redox................ 139, 140, 142–144, 146–150, 322, 323

AND

PHAGOSOMES: METHODS

AND

PROTOCOLS Index 387

S Salmonella-containing vacuole (SCV) .......................................210, 215, 217, 218 Salmonella enterica ............................................. 209–218, 248, 292, 298, 304 Secretion ............................ 172, 289, 304, 305, 332, 335 Single particle tracking (SPT) ....................................... 63, 64, 66, 70, 72, 76 SNARF1 .....................................155, 158, 159, 162, 167 SNARE proteins............................................................ 248 Super-resolution.............................................................. 80

T Tandem Mass Tag (TMT) ...........................................362, 364, 365, 367–369, 372 Target morphology ................................................ 92, 105 Tethering factors ........................................................... 248 Time-lapse imaging......................................342–343, 353 Type III secretion system (T3SS) ...............................210, 291, 303, 304

V Vacuole lysis.......................................................... 209–218 Vinculin ............................................................80, 88, 326 Volume...................................................... 6–9, 11, 12, 17, 18, 20, 21, 26, 31, 37, 46, 47, 49, 50, 67, 84, 89, 94–96, 98, 115, 132, 135, 145, 146, 182, 193, 194, 197, 200, 204, 212, 217, 226–230, 232, 234, 240, 241, 247–249, 256, 257, 265, 267, 269, 279, 280, 319–322, 327, 328, 330, 333–335, 341, 356, 377, 380

W Western blotting................................................... 237, 238