The Immune Synapse: Methods and Protocols [2 ed.] 1071631349, 9781071631348

Table of contents :
Preface
Contents
Contributors
Chapter 1: Measuring the Co-Localization and Dynamics of Mobile Proteins in Live Cells Undergoing Signaling Responses
1 Introduction
2 Materials
2.1 Cell Culture and Transfection
2.2 Dye Conjugation of Labeling Proteins
2.3 Cell Plating and Sample Preparation
2.4 Imaging Reagents
2.5 Microscope and Imaging Software
2.6 Post-Processing and Data Analysis
3 Methods
3.1 Preparation of Fluorescent Fab Fragments
3.2 Cell Preparation
3.3 Imaging
3.4 Post-Processing: Single Molecule Localization
3.5 Post-Processing: Cross-Correlation Analysis
3.6 Post-Processing: Diffusion Analysis
4 Notes
References
Chapter 2: Fluorescence-Based Measurements of Two-Dimensional Affinity in Membrane Interfaces
1 Introduction
2 Materials
2.1 Lipids, Reagents, and Supplies
2.2 Cell Culturing
2.3 Proteins and Labels
2.4 Measurement Cell
2.5 Microscope Setup (TIRF)
3 Methods
3.1 Cell Culturing
3.2 Vesicle Making
3.3 Preparation of Measurement Samples
3.4 SLB Formation
3.5 Functionalizing the SLB with Fluorescently Labeled His-Tagged Proteins
3.6 Adding Cells
3.7 Imaging of Cell Contacts
3.8 Imidazole Addition
3.9 Converting Between Intensity and Protein Density
3.10 Image Analysis and Detection
3.11 Zhu-Golan Analysis
4 Notes
References
Chapter 3: Surfaces for Study of Receptor Dynamics on T Cells
1 Introduction
2 Materials
2.1 Materials for Adhesive Surfaces
2.2 Additional Materials for Activating Surfaces
2.3 Additional Materials for Cell Mimetic Lipid Bilayers
2.4 Imaging T Cell Receptors
3 Methods
3.1 Preparations of Adhesive Surface
3.2 Preparations of Activating and Nonactivating Surface
3.3 Preparations of Cell Mimetic Lipid Bilayers
3.4 Imaging T Cell Surface Receptors
4 Notes
References
Chapter 4: Molecular Dynamics Simulations of Immune Receptors and Ligands
1 Introduction
2 Software
3 Methods
3.1 Protein Structure
3.2 Membrane Embedding
3.3 Equilibration and Simulations
4 Notes
References
5: Investigating Diffusion Dynamics and Interactions with Scanning Fluorescence Correlation Spectroscopy (sFCS)
1 Introduction
2 Materials
2.1 Hardware and Software
2.2 Calibration and Alignment
2.3 Model Membrane Sample Preparation
2.4 Live Cell Samples
3 Methods
3.1 Microscope Calibration Using Point FCS in Solution
3.2 Fluorophore Optimization with FCS Excitation Scans
3.3 Cross-Correlation Controls and Calibrations
3.4 Model Membrane Systems: GUVs
3.5 Model Membrane Systems: SLBs
3.6 Live Cells (Transient Transfection)
3.7 Data Acquisition (sFCS)
3.8 Data Analysis
3.9 Downstream Analysis
3.10 Example Data: Lck Diffusion in T Cells Is Linked to Activatory Signaling
4 Notes
References
Chapter 6: Combined FRET-FLIM and NAD(P)H FLIM to Analyze B Cell Receptor Signaling Induced Metabolic Activity of Germinal Cen...
1 Introduction
2 Materials
2.1 Mice
2.2 Liquids (Sterile)
2.3 Single B Cell Suspensions
2.4 Cell Transfer and Immunization
2.5 Drugs for Anesthesia
2.6 Surgery
2.7 Microscopic Equipment
2.8 Software and Code
3 Methods
3.1 Surgical Preparation of Mice
3.2 System Setup
3.3 Surgical Preparations
3.4 Data Acquisition
3.5 Data Analysis
3.6 Cell Segmentation
3.6.1 Imaris
3.6.2 CellProfiler
4 Notes
References
Chapter 7: Measurement of Molecular Height Using Cell Surface Optical Profilometry (CSOP)
1 Introduction
2 Materials
2.1 Microscopy
2.2 Spherical CSOP Targets
2.3 A Membrane Labeling Reagent
2.4 A Protein Labeling Reagent
2.5 Imaging Buffer
3 Methods
3.1 Preparation of CSOP Samples
3.2 Calibration of the CSOP Imaging System
3.3 Height Measurement
3.4 Image Analysis
4 Notes
References
Chapter 8: Observing Membrane and Cell Adhesion via Reflection Interference Contrast Microscopy
1 Introduction
2 Materials
3 Methods
3.1 Microscope Alignment
3.2 Observation and Recording
3.3 Image Processing and Data Analysis
Box 1
3.4 Dy-RICM Analysis
4 Notes
References
Chapter 9: En-Face Imaging of T Cell-Dendritic Cell Immunological Synapses
1 Introduction
2 Materials and Equipment
2.1 PDMS Device
2.2 Cell Culture
3 Methods
3.1 Photolithography of Wafers
3.2 Production of Soft PDMS Pistons
3.3 Preparation of Bottom Dish with a Glass Slide
3.4 Production of PDMS Micro Pillars
3.5 Preparation of Cells
3.5.1 Isolation and Staining of T Cells
3.5.2 Preparation of Dendritic Cells
3.6 Assembly of Confiner Setup
4 Notes
References
Chapter 10: High- and Super-Resolution Imaging of Cell-Cell Interfaces
1 Introduction
2 Materials
3 Method
3.1 Preparations
3.2 Small Glasses and Chamber Wash and Coating
3.3 Cell Staining (See Note 3)
3.4 Cell-on-Cell Engagement
3.5 Imaging
4 Notes
References
Chapter 11: Separation of Single Core and Multicore Lytic Granules by Subcellular Fractionation and Immunoisolation
1 Introduction
2 Materials
2.1 Primary Murine T Cells
2.2 Cell Homogenization
2.3 Subcellular Fractionation
2.4 Immunoisolation of MCG and SCG
3 Methods
3.1 Cell Homogenization
3.2 Subcellular Fractionation
3.3 Immunoisolation of MCG and SCG
4 Notes
References
Chapter 12: Microvillar Cartography: A Super-Resolution Single-Molecule Imaging Method to Map the Positions of Membrane Protei...
1 Introduction
2 Materials
2.1 Labeling
2.2 Imaging
2.3 Software
2.4 Buffer Preparation
2.5 Microscope
3 Methods
3.1 Labeling of the Cell Membrane and Membrane Proteins (Timing: 1 Day)
3.2 Labeling of Two Different Membrane Proteins for Co-localization Probability (CP) Analysis (Timing: 1 Day)
3.3 Sample Preparation for Microscopy
3.4 Imaging (Timing ~1 h for Imaging of One Cell)
3.4.1 Variable Angle-Total Internal Reflection Microscopy
3.4.2 Stochastic Localization Nanoscopy (SLN)
3.4.3 Analysis (Timing 15-30 min)
4 Notes
References
Chapter 13: T Cell Immunological Synaptosomes: Definition and Isolation
1 Introduction
2 Materials
2.1 Materials for Preparation of T Cell Blasts
2.2 Materials for Bone Marrow-Derived DC (BMDC) Preparation
2.3 Materials for Preparation of Ab-Immobilized CNBr-Activated Sepharose Beads
2.4 Materials for TIS Purification
2.5 Materials for TIS Measurement by Flow Cytometry
2.6 Materials for Evaluation of TIS Activity on DCs
2.7 Regeneration of Ab-Immobilized CNBr-Activated Sepharose Beads
3 Methods
3.1 Preparation of T Cell Blasts
3.2 Preparation of Bone Marrow-Derived DCs (BMDCs)
3.3 Preparation of Ab-Immobilized CNBr-Activated Sepharose Beads
3.4 TIS Purification (Fig. 1)
3.5 TIS Measurement by Flow Cytometry
3.6 Evaluation of TIS Activity on DCs
3.7 Regeneration of Ab-Immobilized CNBr-Activated Sepharose Beads
4 Notes
References
14: Dynamics of Immune Cell Microvilli
1 Introduction
2 Materials
2.1 Microscope and Supplies
2.2 Cell Culture
2.3 Staining and Fixation (see Note 3)
3 Methods
3.1 Cell Culture
3.2 Live Cell Imaging
3.3 Fixed Cell Imaging (Fig. 1e)
3.4 Data Acquisition (Fig. 1f)
3.5 Data Processing (Fig. 1g)
4 Notes
References
Chapter 15: Visualization of Myddosome Assembly in Live Cells
1 Introduction
2 Materials
2.1 Purification and Labeling of His10-IL-1
2.2 Supported Lipid Bilayers Functionalized with IL-1
3 Methods
3.1 Purification and Labeling of IL-1
3.1.1 Labeling HaloTag in His10-Halotag-IL-1β with JF646 Halo Ligand
3.2 Preparing Small Unilamellar Vesicles (SUVs) Suspension for SLB Formation
3.2.1 Formation of SUVs by Freeze-Thaw and Bath Sonication
3.3 Cleaning 96-Well Glass Bottom Plates
3.4 Formation of Supported Lipid Membranes, Labeling SLB with His10-IL1β, Total Internal Reflection Microscopy of SLBs and Ima...
3.4.1 Formation of SLBs in Glass-Bottom 96-Well Plates
3.4.2 Preparing GFP and mScarlet Calibration Wells
3.4.3 Quantification of IL-1 Density and SLB Mobility and Live-Cell Imaging of Myddosome Formation
4 Notes
References
Chapter 16: Detection of Telomere Transfer at Immunological Synapse
1 Introduction
2 Materials
2.1 APCs Purification from Human Primary Peripheral Blood Mononucleate Cells (PBMCs)
2.2 TelC-PNA Probe Live Labeling
2.3 Fluorescence-Activated Vesicle Sorting (FAVS)
2.4 Antigen-Specific Conjugates
2.5 Tzap Overexpression for Alternative Telomere Detection Method
3 Methods
3.1 APCs Purification from Human Primary Peripheral Blood Mononucleate Cells (PBMCs)
3.2 TelC-PNA Probe Live Labeling
3.2.1 Preparation of Glass Beads
3.2.2 APC Telomere Live Labeling
3.2.3 PKH67 Staining and Ionomycin Stimulation
3.3 Fluorescence-Activated Vesicle Sorting (FAVS)
3.4 Antigen-Specific Conjugates
3.4.1 Flowcytometry Analysis to Detect T Cells Acquiring Telomere from APCs
3.4.2 Telomere Immunofluorescence In Situ Hybridization FISH (IF-FISH)
3.5 Tzap Overexpression for Alternative Telomere Detection Method
4 Notes
References
17: Bottom-Up Assembly of Bioinspired, Fully Synthetic Extracellular Vesicles
1 Introduction
2 Materials
2.1 SUV Production
2.2 LUV Production by Emulsification
2.3 Evaluation of Lipid Concentration in LUV Suspension
2.4 Evaluation of Vesicle Concentration and Size by NTA
2.5 Evaluation of Vesicle Size by DLS
2.6 Functionalization of Vesicles with Proteins
3 Methods
3.1 Assembly of synEVs in the Form of SUVs
3.2 Assembly of synEVs in the Form of LUVs by Emulsification
3.3 Evaluation of Lipid Concentration in LUV Suspension
3.4 Evaluation of the Vesicle Concentration and Size by NTA
3.5 Evaluation of the Vesicle Concentration and Size by DLS
3.6 Functionalization of Vesicle Surface with Proteins
4 Notes
References
Chapter 18: A DNA Origami-Based Biointerface to Interrogate the Spatial Requirements for Sensitized T-Cell Antigen Recognition
1 Introduction
2 Materials
2.1 Protein Purification
2.2 Microscopy Setup
2.3 Other Components
3 Methods
3.1 Expression of Streptavidin and I-Ek Subunits as Insoluble Inclusion Bodies in E. coli
3.2 Refolding and Purification of Trans dSav
3.3 Refolding I-Ek in Complex with a Placeholder Peptide
3.4 Purification of I-Ek/ANP
3.5 Site-Specific Biotinylation of I-Ek/ANP Using the BirA Biotin Ligase
3.6 Site-Specific Labeling of a Peptide with Maleimide-Conjugated Dyes
3.7 Exchange of the I-Ek-Associated ANP Placeholder Peptide with Site-Specifically Labeled Peptides
3.8 DNA Origami Preparation
3.9 DNA Origami Purification
3.10 DNA Origami Functionalization Strategy Using Divalent Streptavidin
3.11 DNA Origami Quality Control: Gel Electrophoresis
3.12 SLB Preparation
3.13 SLB Functionalization
3.14 Diffusion Analysis of DNA Origami Structures
3.15 Determining the Fraction of DNA Origami Structures Devoid of pMHC
3.16 Determining the Number of pMHCs on a DNA Origami Structure
3.17 Determination of pMHC Surface Density
4 Notes
References
Chapter 19: Leveraging DNA Origami to Study Phagocytosis
1 Introduction
2 Materials
3 Methods
3.1 Prepare Origami Pegboards
3.2 Couple Origamis to Glass and Quantify Relative Origami Concentration
3.3 Generate Supported Lipid Bilayer Coated Beads
3.4 Quantify Origami Conjugation to Supported Lipid Bilayer-Coated Beads
3.5 Phagocytosis Assays
3.6 TIRF Assays
4 Notes
References
Chapter 20: Fabrication of Nanoscale Arrays to Study the Effect of Ligand Arrangement on Inhibitory Signaling in NK Cells
1 Introduction
2 Materials
2.1 Mold Fabrication
2.2 Array Fabrication by Nanoimprint
2.3 Array Fabrication by E-Beam Lithography
2.4 Chemical Functionalization
2.5 Biofunctionalization
2.6 Verification of the Functionalization Site-Specificity by Fluorescent Staining and Microscopy
2.7 NK Cell Stimulation on the Arrays
3 Methods
3.1 Mold Fabrication
3.2 Array Fabrication by Nanoimprint
3.3 Array Fabrication by E-Beam Lithography
3.4 Chemical Functionalization
3.5 Biofunctionalization
3.6 Verification of the Functionalization Site-Specificity by Fluorescent Staining and Microscopy
3.7 NK Cell Degranulation Assay
3.8 NF-κB Activation
3.9 Interferon-γ Secretion by NK Cells
4 Notes
References
Chapter 21: Measurement of Forces for Trans-Endocytosis at Dorsal and Ventral Sides of the Cell
1 Introduction
2 Materials
2.1 TGT
2.2 Micropillar Arrays
2.3 Microfluidics
3 Methods
3.1 Force Measurement Using TGT Substrate
3.1.1 Protein G-ssDNA Conjugation
3.1.2 Hybridization of Protein G-ssDNA and Complementary ssDNA-Biotin
3.1.3 Conjugation of Ligand (CD80-Fc) to the Protein G-TGT
3.1.4 Preparation of TGT-Coated Substrate for Force Measurement
3.1.5 TGT Image Acquisition
3.1.6 TGT Image Analysis
3.2 Micropillar-Based Traction Force Microscopy
3.2.1 Fabrication of Template of Micropillar Arrays
3.2.2 Fabrication of Micropillar Arrays
3.2.3 Coating Micropillar Arrays for Trans-endocytosis Measurement
3.2.4 Coating Micropillar Arrays for Reference Measurement
3.2.5 Micropillar Array Image Acquisition
3.2.6 Micropillar Array Image Analysis
3.3 Microfluidic-Based Force Sensor
3.3.1 Microfluidic Apparatus
3.3.2 Preparation for Ligand-Conjugated Particles
3.3.3 Microfluidic Force Sensor Image Acquisition
3.3.4 Microfluidic Force Sensor Image Analysis
4 Notes
References
Chapter 22: Functionalized Lipid Droplets and Microfluidics Approach to Study Immune Cell Polarity In Vitro
1 Introduction
2 Materials
2.1 Microfabrication
2.2 Microfluidic Chip Assembly
2.3 Droplet Formulation and Functionalization
2.4 Microfluidic Pairing to Reconstitute Immune Synapses
3 Methods
3.1 Microfabrication
3.2 Microfluidic Chip Assembly
3.3 Droplet Formulation
3.4 Droplet Functionalization
3.5 Immune Synapse Reconstitution in the Microfluidic Chip
3.6 Final Remarks
4 Notes
References
Chapter 23: Quantifying Immune Cell Force Generation Using Traction Force Microscopy
1 Introduction
2 Materials
2.1 Cover Glass Preparation
2.2 Gel Preparation and Polymerization
2.3 Gel Functionalization
2.4 TFM Acquisition
3 Methods
3.1 Glass Preparation
3.2 Gel Preparation & Polymerization
3.3 Gel Functionalization
3.4 TFM Acquisition
3.5 Data Analysis
4 Notes
References
Chapter 24: Characterizing Biophysical Parameters of Single TCR-pMHC Interactions Using Optical Tweezers
1 Introduction
2 Materials
2.1 PEG Slides
2.2 3500 bp DNA Linker with 20 Base Overhang
2.3 Functionalizing Beads with DNA Linker
2.4 Cleaving Antibody and Crosslinking to 20 Base ssDNA Compliment to Overhang
2.5 Ligating ssDNA-Half 2H11 to Beads with Overhang DNA
2.6 Single-Molecule Slide Preparation
2.7 Optical Tweezers Instrumentation
3 Methods
3.1 PEG Slides
3.2 3500 bp DNA Linker with 20 Base Overhang
3.3 Functionalizing Beads with DNA Linker
3.4 Cleaving Antibody and Crosslinking to 20 Base ssDNA Compliment to Overhang
3.5 Ligating ssDNA-Half 2H11 to Beads with Overhang DNA
3.6 Single-Molecule Slide Preparation
3.7 Single-Molecule Force Spectroscopy Measurement
4 Notes
References
Chapter 25: Isolation of the B Cell Immune Synapse for Proteomic Analysis
1 Introduction
2 Materials
2.1 Preparation of the Beads Coated with Activatory and Non-activatory Ligands
2.2 Primary Mouse B Cell Isolation
2.3 Conjugate Formation
2.4 Synapse Isolation
2.5 In-Gel Digestion
3 Methods
3.1 Coating of Magnetic Beads
3.2 Primary Mouse B Cell Isolation (Sterile Work)
3.3 Conjugate Formation
3.4 Synapse Isolation
3.5 In-Gel Digestion
4 Notes
References
Chapter 26: Analyzing Single Cell Secretions by ``Shadow Imaging´´
1 Introduction
2 Materials
2.1 Disposables
2.2 Equipment
2.3 Reagents
2.4 Biological Materials
3 Methods
3.1 Primary NK Cell Isolation
3.2 Sample Preparation
3.3 Imaging Setup
3.4 Data Analysis
4 Notes
References
Chapter 27: Exploiting the RUSH System to Study Lytic Granule Biogenesis in Cytotoxic T Lymphocytes
1 Introduction
2 Materials
2.1 Cloning of GZMB in a RUSH Construct
2.2 CD8+ Cell Purification from Peripheral Blood and CTL Differentiation
2.3 CTLs Nucleofection with the RUSH-GZMB Construct
2.4 Evaluation of RUSH on Fixed Cells by Confocal Microscopy
2.5 Evaluation of RUSH in Live Cells
3 Methods
3.1 Generation of the RUSH-GZMB Construct
3.1.1 GZMB Amplification by PCR
3.1.2 Insert and Vector Restriction Enzyme Digestion
3.1.3 DNA Ligation and Transformation
3.1.4 Plasmid DNA Preparation and Colony Screening
3.2 CD8+ Cell Purification from Peripheral Blood and CTL Differentiation
3.3 CTL Nucleofection with the RUSH-GzmB Construct
3.4 Evaluation of RUSH on Fixed Cells by Confocal Microscopy
3.5 Evaluation of RUSH on Live Cells
4 Notes
References
28: Interactions of Tissue-Resident T Cells
1 Introduction
2 Materials
2.1 Mice
2.2 Preparation of T Cells for Adoptive Transfer
2.3 Models for Generation of Cutaneous CD8+ TRM
2.4 Skin Preparation and Staining Methods for Analysis by Microscopy
3 Methods
3.1 Preparation of CD8+ T Cells for Adoptive Transfer-Naïve CD8+ T Cell Purification
3.2 Preparation of CD8+ T Cells for Adoptive Transfer-Generation of Early Effector CD8+ T Cells
3.3 Models for Generation of Cutaneous CD8+ TRM- Zosteriform Model of HSV Infection
3.4 Models for Generation of Cutaneous CD8+ TRM- OVA Plasmid Tattoo
3.5 Models for Generation of Cutaneous CD8+ TRM- DNFB-Induced Skin Inflammation
3.6 Skin Preparation and Staining Methods for Analysis by Microscopy- Cryosectioning
3.7 Skin Preparation and Staining Methods for Analysis by Microscopy- Skin Whole Mount
3.8 Skin Preparation and Staining Methods for Analysis by Microscopy- Epidermal Sheet
4 Notes
References
Chapter 29: Live Imaging of CAR T Cell Ca2+ Signals in Tumor Slices Using Confocal Microscopy
1 Introduction
2 Materials
2.1 Machines and Equipment
2.2 Buffers and Powders
2.3 Instruments and Tools
2.4 Antibodies
3 Methods
3.1 CAR-T Production and Culture
3.2 Labeling CAR-T and Non-transduced T Cells with Fluo-4
3.3 Obtaining Human Tumor Samples
3.4 Tissue Processing
3.5 Preparation of Agarose Gel
3.6 Embedding of Tumor Samples in Agarose Gel
3.7 Vibratome Slicing of Human Tumor Samples
3.8 Immunostaining of Tumor Slices
3.9 Microscope Preparation
3.10 Imaging of Ca2+ Levels of CAR-T
3.11 Analyzing CAR-T Activation
4 Notes
References
Chapter 30: Measuring CTL Lytic Granule Secretion and Target Cell Membrane Repair by Fluorescent Lipophilic Dye Uptake at the ...
1 Introduction
2 Materials
3 Methods
3.1 Cell Culture
3.2 Procedure for Time-Lapse Microscopy
3.3 Procedure for Flow Cytometry
4 Notes
References
Chapter 31: In Vitro Generation of Human Tolerogenic Monocyte-Derived Dendritic Cells
1 Introduction
2 Materials
2.1 Peripheral Blood Mononuclear Cell (PBMC) Isolation
2.2 Monocyte Isolation
2.3 Generation and Culture of Monocyte-Derived Dendritic Cells (moDC)
3 Methods
3.1 Peripheral Blood Mononuclear Cell (PBMC) Isolation
3.2 Monocyte Isolation
3.3 Generation and Culture of tolDC
4 Notes
References
Chapter 32: Methods of Machine Learning-Based Chimeric Antigen Receptor Immunological Synapse Quality Quantification
1 Introduction
2 Materials
2.1 Recommended Hardware Configuration
2.2 Software Installation
3 Methods
3.1 Immunofluorescence Imaging of CAR IS
3.2 Automated CAR IS Quantification Using Machine Learning
3.3 Alternative Method: Manual Evaluation of CAR IS Using ImageJ
4 Notes
References
Chapter 33: Imaging CAR-T Synapse as a Quality Control for CAR Engineering
1 Introduction
2 Materials
3 Methods
3.1 Lentivirus Production in HEK293T Cells
3.2 Estimation of Virus Titer
3.3 Generation of Primary CAR T Cells
3.4 Culturing Primary CAR T Cell and Raji B-mCherry-CAAX Cells
3.5 Imaging CAR T Cell-Tumor Cell Conjugation and CD45 Exclusion in the Synapse
4 Notes
References
Index

Citation preview

Methods in Molecular Biology 2654

Cosima T. Baldari Michael L. Dustin  Editors

The Immune Synapse Methods and Protocols Second Edition

METHODS

IN

MOLECULAR BIOLOGY

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

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

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

The Immune Synapse Methods and Protocols Second Edition

Edited by

Cosima T. Baldari Department of Life Sciences, University of Siena, Siena, Italy

Michael L. Dustin Kennedy Institute of Rheumatology, University of Oxford, Oxford, UK

Editors Cosima T. Baldari Department of Life Sciences University of Siena Siena, Italy

Michael L. Dustin Kennedy Institute of Rheumatology University of Oxford Oxford, UK

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-3134-8 ISBN 978-1-0716-3135-5 (eBook) https://doi.org/10.1007/978-1-0716-3135-5 © The Editor(s) (if applicable) and The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023 Chapters 5, 11, 18, 27, 30 and 32 are licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/). For further details see license information in the chapters. 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. Cover Illustration Caption: The cover artwork depicts a dendritic cell and a T cell that have formed an immunological synapse. Cells were stained for F-actin (yellow), nuclei (cyan) and LFA-1 (magenta). Image credit: Dr. Alexander Leithner. 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 We are excited to introduce the second edition of The Immune Synapse: Methods and Protocols. The first edition, published in 2017, covers methods and protocols from nearly 20 years of primary publications since the first peer-reviewed publications on the immunological synapse in the late 1990s. This 2nd edition needed only to cover the last 5 years, but it has nearly the same number of totally new chapters, demonstrating the vigorous state of the field! Thus, the two editions together cover over 25 years of know-how in the field. The first eight chapters focus on topics in biophysical measurements, Chapters 9, 10, 11, 12, 13, 14, 15, and 16 cover cell biology of synapses, Chapters 17, 18, 19, and 20 introduce methods for advanced substrate engineering, Chapters 21, 22, 23, and 24 grapple with mechanobiology topics, Chapters 25, 26, and 27 present new technologies to describe and manipulate synaptic components, and Chapters 28, 29, 30, 31, 32, and 33 present methods related to sites of action and immunotherapy. Image correlation methods provide powerful tools to look at overall associations between proteins on the cell surface as illustrated for the B cell receptor and its partners in Chapter 1. The measurement of 2D affinity has been improved by the development of methods that enable affinity measurements from single cells, which removes a major source of variability in the classical approach that required measurements from hundreds of cells, as described in Chapter 2. Chapter 3 provides an excellent introduction for generating classical surface used for the study of T cell receptor dynamics, including a microscale Piranha cleaning protocol. Chapter 4 explores molecular dynamics simulations of the T cell receptor complex in a membrane environment, which may help to interpret measurements such as those made in Chapter 2. Chapter 5 provides an introduction to scanning fluorescence correlation spectroscopy, a method well adapted to investigate the diffusion and interactions of membrane proteins. Chapter 6 provides protocols for in vivo analysis of metabolism and activation using two photon fluorescence lifetime imaging. Chapter 7 provides a general description of approaches to study the height of extracellular domains and the depth of cytoplasmic domains on spherical cells or giant vesicles using cell surface optimal profilometry, and Chapter 8 presents a classical method for measuring distances at membrane interfaces with a microscope cover-glass. Chapters 9 and 10 provide methods for confining T cell-antigen presenting cell synapses to a 2D imaging plane to time lapse and superresolution imaging. Chapter 11 provides methods for isolation of different types of granules from cytotoxic T cells, one that releases soluble perforin and granzymes and the other that releases supramolecular attack particles, among other species. Chapters 12, 13, and 14 provide a suite of methods to study microvilli, the small projections on the surface of T cells that are key sites of sensing and signal initiation. Chapter 15 looks at the assembly of innate signaling complexes using ligands in supported lipid bilayers. Chapter 16 provides methods to investigate telomere transfer through trans-synaptic extracellular vesicles originating from antigen presenting cells. Chapter 17 provides methods to generate biomimetic synthetic extracellular vesicles. Chapters 18, 19, and 20 provide methods for nanoscale positioning of ligands on substrates using DNA origami and bifunctional lithographic methods. The mechanobiology chapters provide a diverse suite of approaches from liquid droplet

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surfaces to the detection of single molecule mechanics with laser tweezers. Chapters 25 and 26 provide methods to identify proteins in the synapse, and Chapter 27 delivers a method to control the release of proteins from the endoplasmic reticulum. Chapter 28 introduces methods to isolate elusive tissue resident cells. Chapter 29 describes a powerful approach to image T cells in ex vivo tumor slices. Chapter 30 provides methods to investigate active resistance of tumor cells to killing by T cells. Chapter 31 looks at tolerogenic dendritic cells, a holy grail in treatment of autoimmunity and a bane of cancer immunotherapy. Chapters 32 and 33 anchor the book with methods to study synapse formation by chimeric antigen receptor expressing T cells. We very much hope that the book will provide insights well beyond the typical published methods sections, particularly the notes, where the tricks of the trade are shared openly. Siena, Italy Oxford, UK

Cosima T. Baldari Michael L. Dustin

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

1 Measuring the Co-Localization and Dynamics of Mobile Proteins in Live Cells Undergoing Signaling Responses. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sarah A. Shelby, Thomas R. Shaw, and Sarah L. Veatch 2 Fluorescence-Based Measurements of Two-Dimensional Affinity in Membrane Interfaces. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Tommy Dam, Manto Chouliara, and Peter Jo¨nsson 3 Surfaces for Study of Receptor Dynamics on T Cells . . . . . . . . . . . . . . . . . . . . . . . . James McColl and David Klenerman 4 Molecular Dynamics Simulations of Immune Receptors and Ligands . . . . . . . . . . Prithvi R. Pandey, Bartosz R
ozycki, and Thomas R. Weikl 5 Investigating Diffusion Dynamics and Interactions with Scanning Fluorescence Correlation Spectroscopy (sFCS) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Alexander M. Mørch and Falk Schneider 6 Combined FRET-FLIM and NAD(P)H FLIM to Analyze B Cell Receptor Signaling Induced Metabolic Activity of Germinal Center B Cells In Vivo . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Carolin Ulbricht, Ruth Leben, Yu Cao, Raluca A. Niesner, and Anja E. Hauser 7 Measurement of Molecular Height Using Cell Surface Optical Profilometry (CSOP) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sungmin Son and Daniel A. Fletcher 8 Observing Membrane and Cell Adhesion via Reflection Interference Contrast Microscopy. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ahmed Abdelrahman, Ana-Suncˇana Smith, and Kheya Sengupta 9 En-Face Imaging of T Cell-Dendritic Cell Immunological Synapses . . . . . . . . . . . Alexander Leithner, Jack Merrin, and Michael Sixt 10 High- and Super-Resolution Imaging of Cell-Cell Interfaces . . . . . . . . . . . . . . . . . Julia Sajman and Eilon Sherman 11 Separation of Single Core and Multicore Lytic Granules by Subcellular Fractionation and Immunoisolation . . . . . . . . . . . . . . . . . . . . . . . . . . Claudia Schirra, Nadia Alawar, Ute Becherer, and Hsin-Fang Chang 12 Microvillar Cartography: A Super-Resolution Single-Molecule Imaging Method to Map the Positions of Membrane Proteins with Respect to Cellular Surface Topography. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Shirsendu Ghosh, Andres Alcover, and Gilad Haran

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T Cell Immunological Synaptosomes: Definition and Isolation . . . . . . . . . . . . . . . Hye-Ran Kim, Jeong-Su Park, Na-Young Kim, and Chang-Duk Jun Dynamics of Immune Cell Microvilli . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . En Cai Visualization of Myddosome Assembly in Live Cells . . . . . . . . . . . . . . . . . . . . . . . . Fakun Cao and Marcus J. Taylor Detection of Telomere Transfer at Immunological Synapse . . . . . . . . . . . . . . . . . . Alessio Lanna and Clara D’Ambra Bottom-Up Assembly of Bioinspired, Fully Synthetic Extracellular Vesicles . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Meline Macher, Ilia Platzman, and Joachim P. Spatz A DNA Origami-Based Biointerface to Interrogate the Spatial Requirements for Sensitized T-Cell Antigen Recognition . . . . . . . . . . . . . . . . . . . . Joschka Hellmeier, Rene´ Platzer, Johannes B. Huppa, and Eva Sevcsik Leveraging DNA Origami to Study Phagocytosis . . . . . . . . . . . . . . . . . . . . . . . . . . . Wyatt D. Miller, Nadja Kern, Shawn M. Douglas, and Meghan A. Morrissey Fabrication of Nanoscale Arrays to Study the Effect of Ligand Arrangement on Inhibitory Signaling in NK Cells . . . . . . . . . . . . . . . . . . . . . . . . . . Guillaume Le Saux, Esti Toledo-Ashkenazi, and Mark Schvartzman Measurement of Forces for Trans-Endocytosis at Dorsal and Ventral Sides of the Cell . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Seungman Park and Yun Chen Functionalized Lipid Droplets and Microfluidics Approach to Study Immune Cell Polarity In Vitro . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Judith Pineau, Le´a Pinon, Jacques Fattaccioli, and Paolo Pierobon Quantifying Immune Cell Force Generation Using Traction Force Microscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Marcel Issler, Huw Colin-York, and Marco Fritzsche Characterizing Biophysical Parameters of Single TCR-pMHC Interactions Using Optical Tweezers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Hannah M. Stephens, Evan Kirkpatrick, Robert J. Mallis, Ellis L. Reinherz, and Matthew J. Lang Isolation of the B Cell Immune Synapse for Proteomic Analysis . . . . . . . . . . . . . . Diogo M. Cunha, Sara Herna´ndez-Pe´rez, and Pieta K. Mattila Analyzing Single Cell Secretions by “Shadow Imaging” . . . . . . . . . . . . . . . . . . . . . Ashley R. Ambrose, Khodor S. Hazime, and Daniel M. Davis Exploiting the RUSH System to Study Lytic Granule Biogenesis in Cytotoxic T Lymphocytes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Nagaja Capitani, Chiara Cassioli, Keerthana Ravichandran, and Cosima T. Baldari

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Interactions of Tissue-Resident T Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Rut Mora-Buch, Hasan Akbaba, and Shannon K. Bromley Live Imaging of CAR T Cell Ca2+ Signals in Tumor Slices Using Confocal Microscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . David Espie, Sarah Barrin, Irena Rajnpreht, Lene Vimeux, and Emmanuel Donnadieu Measuring CTL Lytic Granule Secretion and Target Cell Membrane Repair by Fluorescent Lipophilic Dye Uptake at the Lytic Synapse . . . . . . . . . . . . ¨ ller, Liza Filali, Marie-Pierre Puissegur, Sabina Mu and Salvatore Valitutti In Vitro Generation of Human Tolerogenic Monocyte-Derived Dendritic Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Catharien M. U. Hilkens, Julie Diboll, Fiona Cooke, and Amy E. Anderson Methods of Machine Learning-Based Chimeric Antigen Receptor Immunological Synapse Quality Quantification . . . . . . . . . . . . . . . . . . . . Julian Gan, Jong Hyun Cho, Ryan Lee, Alireza Naghizadeh, Ling Yue Poon, Ethan Wang, Zachary Hui, and Dongfang Liu Imaging CAR-T Synapse as a Quality Control for CAR Engineering . . . . . . . . . . Qian Xiao and Xiaolei Su

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

ix

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Contributors AHMED ABDELRAHMAN • Aix Marseille University, CNRS, CINAM, Turing Centre for Living Systems, Marseille, France HASAN AKBABA • Center for Immunology and Inflammatory Diseases, Division of Rheumatology, Allergy and Immunology, Massachusetts General Hospital, Harvard Medical School, Boston, MA, USA; Department of Pharmaceutical Biotechnology Faculty of Pharmacy, Ege University, Bornova, Izmir, Turkey NADIA ALAWAR • Cellular Neurophysiology, Center for Integrative Physiology and Molecular Medicine (CIPMM), Saarland University, Homburg, Germany ANDRES ALCOVER • Institut Pasteur, Universite´ Paris Cite´, INSERM U1224, Unite´ Biologie Cellulaire des Lymphocytes, Paris, France ASHLEY R. AMBROSE • The Lydia Becker Institute of Immunology and Inflammation, University of Manchester, Manchester, UK AMY E. ANDERSON • Translational & Clinical Research Institute, Newcastle University, Newcastle-upon-Tyne, UK COSIMA T. BALDARI • Department of Life Sciences, University of Siena, Siena, Italy SARAH BARRIN • Centre de recherche en cance´rologie de Lyon, CNRS 5286, Centre Le´on-Be´ rard, Lyon, France UTE BECHERER • Cellular Neurophysiology, Center for Integrative Physiology and Molecular Medicine (CIPMM), Saarland University, Homburg, Germany SHANNON K. BROMLEY • Center for Immunology and Inflammatory Diseases, Division of Rheumatology, Allergy and Immunology, Massachusetts General Hospital, Harvard Medical School, Boston, MA, USA EN CAI • Department of Biological Sciences, Carnegie Mellon University, Pittsburgh, PA, USA FAKUN CAO • Max Planck Institute for Infection Biology, Berlin, Germany a tsmedizin Berlin, corporate member of Freie Universit€ at YU CAO • Charite´—Universit€ Berlin and Humboldt-Universit€ at zu Berlin, Department of Rheumatology and Clinical Immunology, Berlin, Germany; Immune Dynamics, Deutsches RheumaForschungszentrum (DRFZ), a Leibniz Institute, Berlin, Germany NAGAJA CAPITANI • Department of Life Sciences, University of Siena, Siena, Italy CHIARA CASSIOLI • Department of Life Sciences, University of Siena, Siena, Italy HSIN-FANG CHANG • Cellular Neurophysiology, Center for Integrative Physiology and Molecular Medicine (CIPMM), Saarland University, Homburg, Germany YUN CHEN • Department of Mechanical Engineering, Johns Hopkins University, Baltimore, MD, USA; Center for Cell Dynamics, Johns Hopkins University, Baltimore, MD, USA; Institute for NanoBio Technology, Johns Hopkins University, Baltimore, MD, USA JONG HYUN CHO • Department of Pathology, Immunology and Laboratory Medicine, Rutgers University-New Jersey Medical School, Newark, NJ, USA; Center for Immunity and Inflammation, New Jersey Medical School, Newark, NJ, USA MANTO CHOULIARA • Department of Chemistry, Lund University, Lund, Sweden HUW COLIN-YORK • Kennedy Institute for Rheumatology, University of Oxford, Oxford, UK FIONA COOKE • Translational & Clinical Research Institute, Newcastle University, Newcastle-upon-Tyne, UK

xi

xii

Contributors

DIOGO M. CUNHA • Institute of Biomedicine, MediCity Research Laboratories, and InFLAMES Research Flagship, University of Turku, Turku, Finland; Turku Bioscience, University of Turku and Åbo Akademi University, Turku, Finland CLARA D’AMBRA • Sentcell UK laboratories, Rome, Italy TOMMY DAM • Department of Chemistry, Lund University, Lund, Sweden DANIEL M. DAVIS • The Lydia Becker Institute of Immunology and Inflammation, University of Manchester, Manchester, UK; Department of Life Sciences, Imperial College London, London, UK JULIE DIBOLL • Translational & Clinical Research Institute, Newcastle University, Newcastle-upon-Tyne, UK EMMANUEL DONNADIEU • Universite´ Paris Cite´, CNRS, INSERM, Equipe Labellise´e Ligue Contre le Cancer, Institut Cochin, Paris, France SHAWN M. DOUGLAS • Department of Cellular and Molecular Pharmacology, University of California San Francisco, San Francisco, CA, USA DAVID ESPIE • Universite´ Paris Cite´, CNRS, INSERM, Equipe Labellise´e Ligue Contre le Cancer, Institut Cochin, Paris, France; Invectys, Paris, France JACQUES FATTACCIOLI • Laboratoire P.A.S.T.E.U.R., De´partement de Chimie, E´cole Normale Supe´rieure, PSL Research University, Sorbonne Universite´, CNRS, Paris, France; Institut Pierre-Gilles de Gennes pour la Microfluidique, Paris, France LIZA FILALI • INSERM UMR1037, CNRS UMR5071, Centre de Recherche en Cance´rologie de Toulouse (CRCT), Universite´ de Toulouse III-Paul Sabatier, Toulouse, France; Luxembourg Institute of Health, Department of Cancer Research, Strassen, Luxembourg DANIEL A. FLETCHER • Department of Bioengineering and Biophysics Program, University of California, Berkeley, CA, USA; Division of Biological Systems and Engineering, Lawrence Berkeley National Laboratory, Berkeley, CA, USA; Chan Zuckerberg Biohub, San Francisco, CA, USA MARCO FRITZSCHE • Kennedy Institute for Rheumatology, University of Oxford, Oxford, UK; Rosalind Franklin Institute, Harwell Campus, Didcot, UK JULIAN GAN • Department of Pathology, Immunology and Laboratory Medicine, Rutgers University-New Jersey Medical School, Newark, NJ, USA SHIRSENDU GHOSH • Department of Chemical and Biological Physics, Weizmann Institute of Science, Rehovot, Israel GILAD HARAN • Department of Chemical and Biological Physics, Weizmann Institute of Science, Rehovot, Israel ANJA E. HAUSER • Charite´—Universit€ a tsmedizin Berlin, corporate member of Freie Universit€ at Berlin and Humboldt-Universit€ a t zu Berlin, Department of Rheumatology and Clinical Immunology, Berlin, Germany; Immune Dynamics, Deutsches RheumaForschungszentrum (DRFZ), a Leibniz Institute, Berlin, Germany KHODOR S. HAZIME • The Lydia Becker Institute of Immunology and Inflammation, University of Manchester, Manchester, UK; Department of Life Sciences, Imperial College London, London, UK JOSCHKA HELLMEIER • Institute of Applied Physics, TU Wien, Vienna, Austria; Max Planck Institute of Biochemistry, Planegg, Germany SARA HERNA´NDEZ-PE´REZ • Institute of Biomedicine, MediCity Research Laboratories, and InFLAMES Research Flagship, University of Turku, Turku, Finland; Turku Bioscience, University of Turku and Åbo Akademi University, Turku, Finland

Contributors

xiii

CATHARIEN M. U. HILKENS • Translational & Clinical Research Institute, Newcastle University, Newcastle-upon-Tyne, UK ZACHARY HUI • Department of Pathology, Immunology and Laboratory Medicine, Rutgers University-New Jersey Medical School, Newark, NJ, USA JOHANNES B. HUPPA • Medical University of Vienna, Center for Pathophysiology, Infectiology and Immunology, Institute for Hygiene and Applied Immunology, Vienna, Austria MARCEL ISSLER • Kennedy Institute for Rheumatology, University of Oxford, Oxford, UK; Institute of Biology, Humboldt Universit€ at zu Berlin, Berlin, Germany PETER JO¨NSSON • Department of Chemistry, Lund University, Lund, Sweden CHANG-DUK JUN • School of Life Sciences, Immune Synapse and Cell Therapy Research Center, Gwangju Institute of Science and Technology (GIST), Gwangju, South Korea NADJA KERN • Department of Cellular and Molecular Pharmacology, University of California San Francisco, San Francisco, CA, USA HYE-RAN KIM • School of Life Sciences, Immune Synapse and Cell Therapy Research Center, Gwangju Institute of Science and Technology (GIST), Gwangju, South Korea; Division of Rare and Refractory Cancer, Immuno-oncology, Research Institute, National Cancer Center, Goyang, South Korea NA-YOUNG KIM • School of Life Sciences, Immune Synapse and Cell Therapy Research Center, Gwangju Institute of Science and Technology (GIST), Gwangju, South Korea EVAN KIRKPATRICK • Department of Chemical and Biomolecular Engineering, Vanderbilt University, Nashville, TN, USA DAVID KLENERMAN • Department of Chemistry, University of Cambridge, Cambridge, UK MATTHEW J. LANG • Department of Chemical and Biomolecular Engineering, Vanderbilt University, Nashville, TN, USA; Department of Molecular Physiology and Biophysics, Vanderbilt University, Nashville, TN, USA ALESSIO LANNA • Sentcell UK laboratories, Rome, Italy GUILLAUME LE SAUX • Department of Materials Engineering, Isle Katz Institute for Nanoscale Science and Technology, Ben-Gurion University of the Negev, Beer-Sheva, Israel RUTH LEBEN • Biophysical Analysis, Deutsches Rheuma-Forschungszentrum (DRFZ), a Leibniz Institute, Berlin, Germany; Dynamic and functional in vivo imaging, Freie Universit€ at Berlin, Veterinary Medicine, Berlin, Germany RYAN LEE • Department of Pathology, Immunology and Laboratory Medicine, Rutgers University-New Jersey Medical School, Newark, NJ, USA ALEXANDER LEITHNER • Kennedy Institute of Rheumatology, University of Oxford, Oxford, UK DONGFANG LIU • Department of Pathology, Immunology and Laboratory Medicine, Rutgers University-New Jersey Medical School, Newark, NJ, USA; Center for Immunity and Inflammation, New Jersey Medical School, Newark, NJ, USA MELINE MACHER • Max Planck Institute for Medical Research, Heidelberg, Germany; Institute of Molecular Systems Engineering, Heidelberg, Germany; Max Planck School Matter to Life, Heidelberg, Germany; Max Planck-Bristol Center for Minimal Biology, University of Bristol, Bristol, UK ROBERT J. MALLIS • Laboratory of Immunobiology and Department of Medical Oncology, Dana-Farber Cancer Institute, and Department of Dermatology, Harvard Medical School, Boston, MA, USA PIETA K. MATTILA • Institute of Biomedicine, MediCity Research Laboratories, and InFLAMES Research Flagship, University of Turku, Turku, Finland; Turku Bioscience, University of Turku and Åbo Akademi University, Turku, Finland

xiv

Contributors

JAMES MCCOLL • Department of Chemistry, University of Cambridge, Cambridge, UK JACK MERRIN • Institute of Science and Technology Austria (ISTA), Klosterneuburg, Austria WYATT D. MILLER • Department of Molecular, Cellular and Developmental Biology, University of California Santa Barbara, Santa Barbara, CA, USA; Department of Biomolecular Science and Engineering, University of California Santa Barbara, Santa Barbara, CA, USA RUT MORA-BUCH • Center for Immunology and Inflammatory Diseases, Division of Rheumatology, Allergy and Immunology, Massachusetts General Hospital, Harvard Medical School, Boston, MA, USA; Cell Therapy Services, Blood and Tissue Bank, Barcelona, Spain ALEXANDER M. MØRCH • Kennedy Institute of Rheumatology, University of Oxford, Oxford, UK MEGHAN A. MORRISSEY • Department of Molecular, Cellular and Developmental Biology, University of California Santa Barbara, Santa Barbara, CA, USA SABINA MU¨LLER • INSERM UMR1037, CNRS UMR5071, Centre de Recherche en Cance´ rologie de Toulouse (CRCT), Universite´ de Toulouse III-Paul Sabatier, Toulouse, France ALIREZA NAGHIZADEH • Department of Pathology, Immunology and Laboratory Medicine, Rutgers University-New Jersey Medical School, Newark, NJ, USA RALUCA A. NIESNER • Biophysical Analysis, Deutsches Rheuma-Forschungszentrum (DRFZ), a Leibniz Institute, Berlin, Germany; Dynamic and functional in vivo imaging, Freie Universit€ at Berlin, Veterinary Medicine, Berlin, Germany PRITHVI R. PANDEY • Max Planck Institute of Colloids and Interfaces, Potsdam, Germany JEONG-SU PARK • School of Life Sciences, Immune Synapse and Cell Therapy Research Center, Gwangju Institute of Science and Technology (GIST), Gwangju, South Korea SEUNGMAN PARK • Department of Mechanical Engineering, Johns Hopkins University, Baltimore, MD, USA; Center for Cell Dynamics, Johns Hopkins University, Baltimore, MD, USA; Institute for NanoBio Technology, Johns Hopkins University, Baltimore, MD, USA PAOLO PIEROBON • Institut Curie, PSL Research University, INSERM U932, Paris, France JUDITH PINEAU • Institut Curie, PSL Research University, INSERM U932, Paris, France; Universite´ Paris Cite´, Paris, France LE´A PINON • Institut Curie, PSL Research University, INSERM U932, Paris, France; Laboratoire P.A.S.T.E.U.R., De´partement de Chimie, E´cole Normale Supe´rieure, PSL Research University, Sorbonne Universite´, CNRS, Paris, France; Institut Pierre-Gilles de Gennes pour la Microfluidique, Paris, France RENE´ PLATZER • Medical University of Vienna, Center for Pathophysiology, Infectiology and Immunology, Institute for Hygiene and Applied Immunology, Vienna, Austria ILIA PLATZMAN • Max Planck Institute for Medical Research, Heidelberg, Germany; Institute of Molecular Systems Engineering, Heidelberg, Germany; Max Planck-Bristol Center for Minimal Biology, University of Bristol, Bristol, UK LING YUE POON • Department of Pathology, Immunology and Laboratory Medicine, Rutgers University-New Jersey Medical School, Newark, NJ, USA MARIE-PIERRE PUISSEGUR • INSERM UMR1037, CNRS UMR5071, Centre de Recherche en Cance´rologie de Toulouse (CRCT), Universite´ de Toulouse III-Paul Sabatier, Toulouse, France IRENA RAJNPREHT • Universite´ Paris Cite´, CNRS, INSERM, Equipe Labellise´e Ligue Contre le Cancer, Institut Cochin, Paris, France

Contributors

xv

KEERTHANA RAVICHANDRAN • Cellular Neurophysiology, Center for Integrative Physiology and Molecular Medicine (CIPMM), Saarland University, Homburg, Germany ELLIS L. REINHERZ • Laboratory of Immunobiology and Department of Medical Oncology, Dana-Farber Cancer Institute and Department of Medicine, Harvard Medical School, Boston, MA, USA BARTOSZ RO´ZYCKI • Institute of Physics of the Polish Academy of Sciences, Warszawa, Poland JULIA SAJMAN • Racah Institute of Physics, The Hebrew University, Jerusalem, Israel; Jerusalem College of technology, Jerusalem, Israel CLAUDIA SCHIRRA • Cellular Neurophysiology, Center for Integrative Physiology and Molecular Medicine (CIPMM), Saarland University, Homburg, Germany FALK SCHNEIDER • Translational Imaging Center, University of Southern California, Los Angeles, California, USA MARK SCHVARTZMAN • Department of Materials Engineering, Isle Katz Institute for Nanoscale Science and Technology, Ben-Gurion University of the Negev, Beer-Sheva, Israel KHEYA SENGUPTA • Aix Marseille University, CNRS, CINAM, Turing Centre for Living Systems, Marseille, France EVA SEVCSIK • Institute of Applied Physics, TU Wien, Vienna, Austria THOMAS R. SHAW • Program in Applied Physics, University of Michigan, Ann Arbor, MI, USA SARAH A. SHELBY • Program in Biophysics, University of Michigan, Ann Arbor, MI, USA EILON SHERMAN • Racah Institute of Physics, The Hebrew University, Jerusalem, Israel MICHAEL SIXT • Institute of Science and Technology Austria (ISTA), Klosterneuburg, Austria ANA-SUNCˇANA SMITH • PULS Group, Department of Physics, Centre for Computational Materials and Processes, Friedrich Alexander University Erlangen-Nu¨rnberg, IZNF, Erlangen, Germany; Group for Computational Life Sciences, Division of Physical Chemistry, Ruđer Bosˇkovic´ Institute, Zagreb, Croatia SUNGMIN SON • Department of Bioengineering and Biophysics Program, University of California, Berkeley, CA, USA; Department of Bio and Brain Engineering, KAIST, Daejeon, Republic of Korea JOACHIM P. SPATZ • Max Planck Institute for Medical Research, Heidelberg, Germany; Institute of Molecular Systems Engineering, Heidelberg, Germany; Max Planck School Matter to Life, Heidelberg, Germany; Max Planck-Bristol Center for Minimal Biology, University of Bristol, Bristol, UK HANNAH M. STEPHENS • Department of Chemical and Biomolecular Engineering, Vanderbilt University, Nashville, TN, USA XIAOLEI SU • Department of Cell Biology, Yale School of Medicine, New Haven, CT, USA; Yale Cancer Center, Yale University, New Haven, CT, USA MARCUS J. TAYLOR • Max Planck Institute for Infection Biology, Berlin, Germany ESTI TOLEDO-ASHKENAZI • Department of Materials Engineering, Isle Katz Institute for Nanoscale Science and Technology, Ben-Gurion University of the Negev, Beer-Sheva, Israel CAROLIN ULBRICHT • Charite´—Universit€ a tsmedizin Berlin, corporate member of Freie Universit€ at Berlin and Humboldt-Universit€ a t zu Berlin, Department of Rheumatology and Clinical Immunology, Berlin, Germany; Immune Dynamics, Deutsches RheumaForschungszentrum (DRFZ), a Leibniz Institute, Berlin, Germany

xvi

Contributors

SALVATORE VALITUTTI • INSERM UMR1037, CNRS UMR5071, Centre de Recherche en Cance´rologie de Toulouse (CRCT), Universite´ de Toulouse III-Paul Sabatier, Toulouse, France; Department of Pathology, Institut Universitaire du Cancer-Oncopole de Toulouse (IUCT), Toulouse Ce´dex, France SARAH L. VEATCH • Program in Biophysics, University of Michigan, Ann Arbor, MI, USA LENE VIMEUX • Universite´ Paris Cite´, CNRS, INSERM, Equipe Labellise´e Ligue Contre le Cancer, Institut Cochin, Paris, France ETHAN WANG • Department of Pathology, Immunology and Laboratory Medicine, Rutgers University-New Jersey Medical School, Newark, NJ, USA THOMAS R. WEIKL • Max Planck Institute of Colloids and Interfaces, Potsdam, Germany QIAN XIAO • Department of Cell Biology, Yale School of Medicine, New Haven, CT, USA; Duncan and Nancy MacMillan Cancer Immunology and Metabolism Center of Excellence, Rutgers Cancer Institute of New Jersey, New Brunswick, NJ, USA; Department of Medicine, Robert Wood Johnson Medical School, Rutgers University, New Brunswick, NJ, USA

Chapter 1 Measuring the Co-Localization and Dynamics of Mobile Proteins in Live Cells Undergoing Signaling Responses Sarah A. Shelby, Thomas R. Shaw, and Sarah L. Veatch Abstract Single molecule imaging in live cells enables the study of protein interactions and dynamics as they participate in signaling processes. When combined with fluorophores that stochastically transition between fluorescent and reversible dark states, as in super-resolution localization imaging, labeled molecules can be visualized in single cells over time. This improvement in sampling enables the study of extended cellular responses at the resolution of single molecule localization. This chapter provides optimized experimental and analytical methods used to quantify protein interactions and dynamics within the membranes of adhered live cells. Importantly, the use of pair-correlation functions resolved in both space and time allows researchers to probe interactions between proteins on biologically relevant distance and timescales, even though fluorescence localization methods typically require long times to assemble well-sampled reconstructed images. We describe an application of this approach to measure protein interactions in B cell receptor signaling and include sample analysis code for post-processing of imaging data. These methods are quantitative, sensitive, and broadly applicable to a range of signaling systems. Key words Fluorescence localization microscopy, Super-resolution microscopy, Pair distribution, Co-localization, Single molecule tracking, Diffusion analysis

1

Introduction The invention of super-resolution microscopy has revolutionized biological imaging, allowing researchers to directly visualize the organization and interactions of proteins on distance scales relevant to cellular processes. In the field of immune receptor signaling, super-resolution imaging methods have, for example, helped to define cellular contacts between immune cells and antigenpresenting cells as well as the composition of signaling platforms formed after immune receptors engage with antigens or crosslinkers [1–7]. Single molecule fluorescence localization microscopy (SMLM) is one method within the broader class of super-resolution imaging that achieves high spatial resolution by imaging sparsely distributed fluorophores over time [8–11]. Fluorophores

Cosima T. Baldari and Michael L. Dustin (eds.), The Immune Synapse: Methods and Protocols, Methods in Molecular Biology, vol. 2654, https://doi.org/10.1007/978-1-0716-3135-5_1, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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labeling proteins of interest stochastically transition between emitting and transiently dark states, allowing for the ensemble of molecules to be fully sampled over time. Single and multi-color SMLM has been used to image components of the immune response in a wide variety of contexts [12–21]. Our own work has largely focused on probing early events in B cell receptor signaling [22–24]. An important limitation of SMLM, especially when applied to living systems, is that it typically takes seconds to minutes to acquire an image because many (typically 100 s to 1000s) individual images of single molecules are needed to reconstruct a well-sampled superresolved image. Over these extended time periods, molecules of interest tend to diffuse over distances much larger than the biological structures under investigation, washing out the superresolved information researchers seek to extract. One approach to overcoming this limitation is to focus on systems with slow dynamics [25, 26], or to push the limits of camera frame rates and fluorophore photophysics [27]. In our own past work, we have sought to overcome this limitation by extracting information from SMLM datasets without reconstructing images, by interrogating the relative distributions of molecules with pair correlation functions. Pair correlation functions are distributions of pair-wise distances between localized molecules imaged over time. Correlation functions can take the form of the autocorrelation, which tabulates pairwise distances between molecules of the same type, or the crosscorrelations, which tabulate pairwise distances between molecules of the different types. Autocorrelation functions provide information on how labeled proteins move and how they self-associate, while cross-correlation functions probe the co-localization of proteins, as well as the dynamics and effective energy of these associations [15, 28–33]. Pair-correlation methods can also be used as part of a general image processing pipeline to enhance or characterize SMLM datasets [34, 35]. Previously, we described how a steady-state cross-correlation can extract information relevant to interactions between proteins in live cells undergoing signaling responses after the B cell receptor (BCR) is engaged with a multivalent cross-linker [22, 31]. We have continued to further optimize and extend these approaches, enabling the detection of weak signals over time [24]. This protocol chapter describes optimized experimental approaches including sample preparation, fluorescent probe selection, and microscopy imaging methods. While the protocol focuses on the specific application of BCR activation in CH27 B cells, the imaging methods described can be applied to study interactions and dynamics of plasma membrane signaling proteins in a variety of cell types and signaling processes. We also include protocols for single fluorophore localization in raw images, the evaluation of properly normalized pair correlation functions, and use of correlation functions to characterize co-localization and molecular motions in live cells. Analysis code is provided to facilitate the application of these methods by other groups.

Quantifying Dynamic Signaling Interactions in Cell Membranes

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Materials

2.1 Cell Culture and Transfection

The methods presented can be applied to a variety of cell types and signaling systems, the primary requirement being that a relatively flat membrane surface can be imaged. 1. CH27 mouse B cells [36]. 2. Low-glucose DMEM (Life Technologies, Carlsbad, CA or equivalent) with 15% fetal bovine serum (Mediatech, Manassas, VA or equivalent), 10 mM HEPES, 110 mg l-1 sodium pyruvate, 50 μM BME, and 1% Pen/Strep [31]. 3. Lonza 4D Nucleofector X unit or equivalent. 4. High-quality plasmid DNA encoding proteins of interest fused to a protein tag (such as GFP, SNAP or HALO) or a photoswitchable fluorescent protein (such as mEos3.2 [37]) with a strong promoter.

2.2 Dye Conjugation of Labeling Proteins

In-house dye conjugation of labeling proteins (e.g., antibody Fab fragments) is often required because commercial reagents are not always available. For situations where protein clustering is to be avoided or controlled, labeling proteins should bind monovalently to their targets. 1. Goat anti-Mouse IgM(μ) Fab fragments (Jackson ImmunoResearch 115-007-020 or equivalent). 2. Silicon rhodamine (SiR) NHS esters (Invitrogen, B1582; Spirochrome, SC003 or similar). Solubilize dye NHS esters in anhydrous DMSO at 10 mM. Store stock solution aliquots at -80 °C with desiccation, and avoid freeze thaw. We use biotin and SiR NHS esters to label and functionalize anti-BCR Fab fragments. 3. Conjugation reaction buffer: 0.01 M NaH2PO4 with 0.01 M NaH2CO3 at pH 8.5. 4. Gel filtration column, e.g., GE Healthcare illustra NAP-5 Columns (Fisher 45-000-151 or equivalent). 5. Column equilibration and elution buffer: PBS (1.1 mM KH2PO4, 2.9 mM Na2HPO4, 155 mM NaCl, pH 7.4) with 1 mM EDTA. 6. Vivaspin-500 Polyethersulfone concentration spin column (Vivaproducts: VS0191, VS0121, VS0131, VS0141 or equivalent) with an appropriate molecular weight cutoff (3–100 k) of approximately half the protein molecular weight or lower.

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2.3 Cell Plating and Sample Preparation

Cells are prepared on glass substrates conducive to optical microscopy. Non-adherent cell lines can be attached to coverslips functionalized with adhesion molecules such as integrin ligands or poly-Llysine. We use VCAM-1-coated dishes to promote attachment of CH27 B cells. 1. Cell culture dishes with #1.5 glass coverslips (Mattek P35G-1.5-14-C or equivalent). 2. VCAM-1/Fc chimeric proteins and anti-human Fc antibodies (recombinant human VCAM-1/Fc chimera protein, Fisher 862-VC-100). 3. Fcγ-specific goat anti-human IgG, Jackson ImmunoResearch 109-005-008 or equivalent). 4. Oxygen plasma equivalent).

2.4 Imaging Reagents

cleaner

(e.g.,

Harrick

PDC-32G

or

1. Fiducial markers: 0.1 μm TetraSpeck beads, ThermoFisher T7279 or equivalent. 2. L-glutathione reduced (Sigma G6013 or equivalent). 3. 4 mg/mL Catalase stock solution in PBS. 4. 10 mg/mL Glucose oxidase stock solution in PBS. 5. Imaging buffer: 30 mM Tris, 9 mg/mL glucose, 100 mM NaCl, 5 mM KCl, 1 mM KCl, 1 mM MgCl2, 1.8 mM CaCl2, 10 mM glutathione, 8 μg/mL catalase, 100 μg/mL glucose oxidase, pH 8 (see Note 1). 6. Streptavidin or other reagents to cluster cell surface proteins or receptors, antigens to stimulate cell signaling responses, etc.

2.5 Microscope and Imaging Software

Many available commercial systems that are designed for single molecule imaging are compatible with super-resolution localization microscopy. In general, relatively high-power laser illumination is needed for imaging of organic dyes, and multiple laser lines are needed for multicolor imaging. To image the plasma membrane selectively, illumination is set up in a total internal reflection (TIR) configuration. A high numerical aperture (NA) objective lens is used to maximize photons collected from single molecules. Multiple color channels are split using emission splitting optics and recorded on a high sensitivity EMCCD or sCMOS camera. The camera control computer and software must be capable of acquisition and storage of a high volume of image data. Acquisition software should record image metadata including acquisition timestamp information. For our setup, we use an Olympus IX81 inverted microscope outfitted with the following:

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1. Coherent OBIS fiber-pigtailed solid-state lasers: 405 nm (50 mW), 561 nm (120 mW), and 647 nm (100 mW) or equivalent. 2. Olympus cellTIRF module to adjust laser incident angle or equivalent. 3. 100× UAPO TIRF objective (NA = 1.49) or equivalent. 4. LF405/488/561/647 quadband filter cube (TRF89902, Chroma or equivalent). 5. Olympus active Z-drift correction (ZDC) module or equivalent. 6. DV2 (Photometrics) or Gemini (Hamamatsu) emission splitter or equivalent. 7. Andor iXon-897 EMCCD camera or equivalent. 2.6 Post-Processing and Data Analysis

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Software and computing resources are needed for post-processing of raw image data. Handling large amounts of image data and determining single molecule localizations from raw data are computationally intensive. Numerous commercial and freely available software packages are available for single molecule localization. We find that the imageJ plugin ThunderSTORM [38] is flexible and accessible. We use MATLAB for all subsequent analysis of single molecule localization to quantify protein co-localization and dynamics.

Methods All steps are carried out at room temperature unless otherwise noted.

3.1 Preparation of Fluorescent Fab Fragments

Labeling proteins such as antibodies, Fab fragments, toxins, or other binding proteins can be conjugated with organic dyes suitable for super-resolution imaging (see Note 2) via NHS ester chemistry. In these reactions, a reactive NHS ester group commonly available on organic dyes is used to covalently attach the dye to amines on lysine residues or N-termini of purified proteins. After conjugation, unreacted dye is separated from labeled proteins by size exclusion chromatography. For applications where the labeling protein is cross-linked during the imaging experiment, for example, to study the effects of protein clustering, labeling proteins can be conjugated with both dye and biotin groups using the same NHS ester chemistry. In our specific application, we label goat anti-Mouse IgM(μ) Fab fragments with biotin and silicon rhodamine (SiR) NHS esters.

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1. Conjugation reactions should be carried out near pH 8.5 under buffer conditions where proteins are stable. Exchange purified proteins into the reaction buffer through dialysis. If the initial protein storage buffer contains free amines such as Tris or glycine, several rounds of dialysis may be necessary to remove them. Final protein concentrations between 0.5 and 2 mg/mL are optimal for labeling. 2. Add amine-reactive dye NHS ester DMSO stock solutions to labeling protein solutions in 2–8× molar excess (see Note 3). Immediately mix the protein solution by inversion or by flicking the tube to avoid exposing proteins to high local concentrations of DMSO. 3. Incubate the reaction for 1 h at room temperature with rotation. Protect from light, e.g., by enclosing the reaction tube in a larger foil-covered tube. 4. After incubation, pass the reaction solution through a NAP-5 gel filtration column using PBS with 1 mM EDTA as the equilibration and elution buffer. 5. Modified proteins can be further purified and concentrated by centrifugation in a Vivaspin-500 polyethersulfone concentration spin column. 6. Estimate the degree of labeling using absorbance measurements of the dye-conjugated protein at the dye’s absorption maximum and 280 nm. We aim for a dye-to-protein ratio around 2. If the protein is under-modified, the reaction can be repeated to increase the dye-to-protein ratio. 7. For proteins modified with both fluorescent dyes and biotin, we use a two-step procedure. First, react unlabeled proteins with a mixture of fluorophore and biotin NHS esters in 5× and 12× molar excess, respectively. Purify as described above. Second, repeat the labeling procedure with only fluorophore NHS ester in 5× molar excess. 3.2

Cell Preparation

Cells can express tagged proteins, e.g., through transient transfection or through the preparation of stable cell lines through antibiotic selection or viral transduction. Expressible fluorescent tags can take the form of photo-activatable/photo-switchable fluorescent proteins or self-labeling protein tags that covalently bind to organic dyes such as SNAP tags and HaloTags. Proteins on the cell surface can be labeled with antibodies or toxins conjugated to organic fluorophores (see Subheading 3.1). Labeling strategies that use transient binding and exchange of fluorescent conjugates to achieve stochastic blinking, as in PAINT and DNA-PAINT, are also compatible. Our most common two-color labeling approach is to pair BCR labeling using SiR-biotin-conjugated Fab fragments with transient expression of an mEos3.2 fusion protein. Prior to

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imaging, cells should be plated on dishes or wells with #1.5 glass coverslips under conditions that facilitate uniform adhesion. 1. Transiently transfect cells with fluorescent fusion constructs, and allow them to recover in growth media in the 37 °C CO2 incubator for 18–24 h before preparation for imaging. 2. Prepare VCAM-1-coated dishes which can be prepared as follows. (i) Plasma clean imaging chambers using an oxygen plasma cleaner at the highest power setting for 3 min. (ii) Immediately incubate cleaned dishes with a solution of Fcγ-specific goat anti-human IgG antibodies in PBS at 100 μg/mL for 30 min at room temperature. (iii) Rinse dishes with PBS and block for 30 min with a 5% BSA solution in PBS. (iv) Rinse dishes with PBS, and incubate dishes for 1 h at room temperature or overnight at 4 °C with recombinant VCAM-1/human Fc chimera protein at 10 μg/mL. (v) Rinse dishes thoroughly in PBS before use. (vi) Coated dishes can be stored in PBS at 4 °C for up to 1 week. 3. Cells can be grown overnight on untreated glass, but should be plated soon before imaging if (1) cells secrete an expressed fluorescent protein which can become deposited on the glass (see Note 4) or (2) a surface treatment such as VCAM-1 coating is used. CH27 cells adhere to untreated glass within 2–4 h at 37 °C and VCAM-1-coated dishes within 10 min at room temperature. 4. Cells should be plated at a density such that they remain isolated from one another for imaging. 5. A day of imaging usually involves multiple replicates of a live cell imaging measurement. Prepare as many dishes of cells as replicates are desired. 6. Label cells with dye-conjugated proteins at room temperature or on ice if needed to minimize probe internalization. We label BCR by incubation with 5 μg/mL SiR-biotin-Fab in culture medium for 10 min at room temperature. Cells should be labeled shortly (within 30 min) before imaging to prevent internalization. 7. At this stage and prior to imaging, cells can be pre-treated with drugs or other perturbations as needed for the experiment.

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Imaging

Sparse, single molecules are simultaneously imaged in two color channels over time. 1. Configure microscope such that two fluorophores can be imaged simultaneously. This likely will involve illumination from several excitation lasers (e.g., 405 nm, 561 nm, 647 nm) paired with a multi-band filter cube that reflects laser lines and passes fluorophore emission. An emission splitter unit is used to separate emission from distinct fluorophores and direct them onto separate cameras or separate sides of the same EMCCD or sCMOS camera (see Note 5). Lasers and emission optics should be aligned prior to sample preparation. We recommend using an automated correction for drift in the axial direction (e.g., ZDC from Olympus). Total internal reflection (TIR) illumination is commonly used for selective imaging of cell surface proteins and structures on the plasma membrane. 2. Prepare a fiducial marker sample by first cleaning a #1.5 glass coverslip cell culture dish in an oxygen plasma cleaner at full power for 3 min. A 1:100 dilution of beads in PBS is immediately added to the dish and incubated for 30 min–2 h at room temperature. Unbound beads are removed with gentle rinsing. This sample can be stored in PBS at 4 °C and reused over several months. 3. Acquire images of fiducial markers to register color channels: Take many (>100) images of the fiducial marker bead sample using the same illumination laser lines and filters to be used in the cellular imaging measurement. Between acquisition of individual bead images, translate the stage so that the field of view is uniformly sampled by bead positions. It is important that the splitting optics are stable throughout the course of the imaging experiment. We recommend acquiring bead images before and after a cell measurement, to ensure that the relative positions of the two color channels do not shift during a measurement, impairing channel registration. 4. Prepare imaging buffer for SiR/mEos3.2 two-color imaging. Add oxygen-scavenging enzymes glucose oxidase and catalase to the imaging buffer (see Note 1), and replace buffer in cells with fresh imaging buffer (see Note 6). 5. Select cells for imaging. Cells should be firmly adhered to the dish to minimize membrane topography. It can be helpful to image cells using an interference reflection microscopy (IRM) [39] filter cube to screen for uniform membrane flatness across the cell footprint. Alternately, dSTORM probes can be viewed with low laser power with TIR illumination to assess the uniformity of the signal (Fig. 1a), or the blue excitation/green emission of un-converted mEos3.2 can be used for the same purpose. Cells should also be chosen based on expression levels of fluorescent constructs or labeling density.

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Fig. 1 Sample raw and reconstructed images. (a). Total internal reflection (TIR) and interference reflection microscopy (IRM) images of an adherent CH27 B cell after incubation with a Fab fragment αIgMμ conjugated to both SiR and biotin. When imaged with low-intensity 647 nm excitation, SiR should exhibit a uniform and bright distribution over the cell surface when imaged in TIR, and the IRM image should also be uniform indicating minimal membrane topography. (b) (top) Raw TIR image frames of SiR imaged at high 647 nm laser power and mEos3.2 imaged with both 561 nm excitation and 405 nm activation lasers. Imaging SiR with high laser power converts some probes into a reversible dark state allowing the visualization of isolated single

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6. Adjust laser intensities and frame rate. The goal for imaging is to view sparse single molecules in each color channel (Fig. 1b) over an extended period of time (10 min). High laser intensity is needed to induce dSTORM probe photoswitching, and the probe density is inversely dependent on illumination intensity. 405 nm laser intensity can be used to control mEos3.2 probe density. For SiR/mEos3.2 imaging, we use the following range of laser powers during single molecule data collection: 5–20 kW/cm2 for 561 nm and 647 nm lasers and 100–200 W/cm2 for the 405 nm laser. Frame rates should be chosen such that exposure times are as long as possible to maximize signal-to-noise but short enough to minimize motion blur and accurately localize mobile single molecules. Exposure times in the tens of milliseconds corresponding to frame rates between 20 and 100 Hz are typical, depending on the capabilities of the camera and mobility of proteins of interest. 7. Acquire single molecule images over time. Acquire several minutes of images of the untreated cell prior to carefully adding a treatment or activator (see Note 7). We add streptavidin during the experiment to cluster SiR-biotin-Fab-labeled BCR and induce cellular activation. Image acquisition timestamps can be useful for keeping track of changing behavior relative to the time of treatment addition. When adding treatments during imaging, add relatively large volumes to facilitate fast mixing. Add treatments diluted in imaging buffer. We recommend saving images in successive multistack or movie files with a set number of images per file (e.g., 500 frames corresponding to 10 s). Smaller file size speeds saving during acquisition and loading during post-processing. 8. Independent measurements should be repeated over multiple days using multiple biological replicates. 3.4 Post-Processing: Single Molecule Localization

Single molecules can be localized using a wide variety of software packages. The end goal is to generate a list containing the positions and other attributes of single molecules over time in each color channel.

ä Fig. 1 (continued) molecules. (bottom) Super-resolution image of SiR and mEos3.2 from localizations acquired from 4500 image frames over 2.9 min. Note the presence of a gradient in the SiR image. This is due to mobile probes diffusing from the cell edges that are not initially in a reversible dark state since the TIR field does not extend above the ventral cell surface. (c) The same conditions as B but images are acquired after the addition of streptavidin to induce clustering and activation of BCR. Super-resolved images are generated from 8500 raw images frames acquired over 5.8 min

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1. Localize single molecule positions. This can be accomplished in a wide range of software packages, including imageJ using the Thunderstorm plug-in [38]. This particular software package has implemented multi-emitter fitting, which allows for the accurate processing of images with a high density of single molecules per frame. Localized positions and other fit parameters can be exported for further processing (see Notes 8 and 9). 2. Generate a spatial mapping between image channels using fiducial markers, for example, using the procedures described in [40]. To accomplish this, fiducial markers are matched across image channels and used as control points to generate a transform. Once the positions of the control points are established, this map can be generated from control points using the function fitgeotrans() in MATLAB. The transform can then be applied to points using the function transformPointsForward () in MATLAB. 3. Cull localizations to remove poor fits. This is typically done by accumulating distributions of fit parameter values and removing the top and bottom 565% for parameters such as intensity or width. 4. Localized positions can be corrected for stage drift or collective cellular motions using a drift correction algorithm, for example, using the procedures described in [41]. Although drift correction is not strictly required for the cross-correlation or diffusion analysis because it applies at slower timescales than are typically of interest, it can be important when establishing a region of interest (ROI) that is appropriate for localizations acquired over the entire dataset. 5. Images representing the time-averaged positions of localized molecules can be rendered by generating two-dimensional (2D) histograms from the localized positions. This can be accomplished using the histcounts2() function in MATLAB. Histograms can be presented as is or blurred using a Gaussian filter for display purposes. 3.5 Post-Processing: Cross-Correlation Analysis

Cross-correlation analysis is conducted using the locations of molecules in both space and time. Cross-correlation functions are generated by tabulating pairwise distances between localizations in different color channels, assembling these distance into histograms, and then normalizing these histograms to account for the expected number of pairs if probes are randomly co-distributed over the specified region of interest. We have assembled a collection of MATLAB functions to perform these calculations from lists of localizations, as well as sample analysis scripts, publically available at https://github.com/VeatchLab/smlm-analysis. Scripts and localizations used to generate the remaining figures are also included in the SMLM-analysis distribution.

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These methods derive their power from the assumption that the effective interactions between labeled proteins are similar at different locations across the region of interest, so that localizations from different locations on the cell can be thought of as samples from one statistical distribution. By combining all of these observations, the poor sampling at any one location adds up to excellent sampling of the pair distribution overall. Note that this assumption may not hold for some structures of interest – for example, a polarized cell may give rise to different effective interactions on the two poles. Functions within this publically available package are referenced below. 1. Specify the region of interest (ROI) for further analysis. ROIs should be placed within the cellular footprint and not contain cell edges. ROIs should also exclude other clear topographical features. Since localizations are analyzed as projections on two dimensions, features that contain more membrane per area (e.g., cell edges, filopodia) will lead to trivial correlations. A tool to draw ROIs on reconstructed images called spacewin_gui() is included in the SMLM-analysis distribution. This tool allows a user to draw polygons on images reconstructed from lists of localized positions. Polygons can be drawn that either include or exclude points, allowing for ROIs with complicated topologies. Examples of different ROIs and their impact on the resulting cross-correlation functions are shown in Fig. 2. 2. Tabulate cross-correlations that properly normalize for ROI boundaries. This can be accomplished using the MATLAB function spacetime_xcor() posted in the SMLM-analysis distribution. This base function includes an efficient code to tabulate the pairwise distances between localizations over time, adapted from [42] and described previously [41], as well as boundary conditions for arbitrary spatial ROIs and temporal windows, as described [34]. In some cases, long-range gradients in images can obscure results when an analysis of short-range correlations is desired. To reduce the impact of long-range gradients, a local density correction can be applied, tabulated using spatial_gradient_correction() in the SMLM-analysis distribution, following an approach described previously [43]. We find that this correction is often necessary when imaging mobile (d)STORM probes such as SiR that diffuse into the illuminated area from the cell edges in an activated state. These probes tend to take some time to be converted into a reversible dark state, leading to a larger density of probes at the cell periphery (evident in Fig. 1b for SiR). A properly normalized correlation function should decay to 1 when probed at large separation distance or large separation times. Examples of correlation functions tabulated from the same set of localizations but with different normalizations are shown in Fig. 3.

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Fig. 2 Defining an appropriate region of interest (ROI). (a) Reconstructed image (left) of SiR-tagged BCR and transiently expressed Src15-mEos3.2 in an adhered CH27 B cell imaged after BCR clustering with streptavidin. The yellow polygon indicates the region of interest (ROI) used for the cross-correlation analysis (right). This ROI contains areas inside and outside the cell footprint, including membrane topographical features of the cell boundary itself and thin protrusions. Because both probes reside on the cell membrane, the crosscorrelation takes on values >1, indicating co-localization of the two probes. (b) (left) The same cell shown in (a) with a ROI defined within the cell boundary. (right) The cross-correlation function tabulated using this ROI indicates exclusion of the two probes (values 90% purity. 4. In our lab, we recently changed the immunization protocol for popliteal lymph node imaging from footpad immunization [16] to hock immunization [17] to reduce the burden on the mice. For this purpose, the injection is placed on the inner side of the (right) leg, close to the ankle joint. We confirmed that animals treated this way show the same immune reaction as classically immunized ones, but show reduced pain stress and do not suffer from reduced mobility. 5. We inject into the right leg, because our imaging set up restricts us to imaging the right lymph node [16]. Adjustments to the customizable stage can be made anytime, but always keep in mind that the spatial arrangements under the objective or accessibility of connective adapters (e.g., anesthesia, heating) will also change. 6. If working with a restrainer, animals should be trained to the procedure at least four times within the week prior to the experiment, in accordance with refinement measures. This will reduce stress and improve the cooperation of mice. 7. A data set of several FLIM movies in two wavelengths can easily amount to 100 GB or more.

ä Fig. 3 (continued) (3) 3-hydroxyacyl-CoA dehydrogenase (HADH); (4) lactate dehydrogenase; (5) glucose-6phosphate dehydrogenase (G6PDH); (6) succinate dehydrogenase, binding NADH (SDH); (7) glyceraldehyde-3phosphate dehydrogenase (GAPDH); (8) itaconate dehydrogenase; (9) SDH binding NADPH; (10) pyruvate dehydrogenase (PDH)/C-terminal binding protein 1 (CTBP1); (11) inducible nitric oxide synthase (INOS); (12) alcohol dehydrogenase (ADH); (13) NADPH oxidase (NOX). Lower right: Activity map color-coded according to LUT “royal,” 0–100%. (b) Median Ca2+ concentration quantification of B cells shown in (a), segmented with CellProfiler. Red line is indicating overall mean of total cells. (c) Histogram of mean enzyme distribution of the B cells in (a), segmented with CellProfiler. (d) In-depth analysis of enzyme distribution and activity of five single TNXXL+ GC B cells. Quantification as pixel histograms and frequency in the metabolic sub-groups “anaerobic glycolysis and ß-oxidation,” “aerobic glycolysis and oxidative phosphorylation (OXPHOS)/tricarboxylic acid cycle (TCA),” and “oxidative burst” (see also [21])

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8. If you possess more than one objective for your system, make sure to choose the right one from the drop-down menu in panel “Objectives,” since this will affect your field of view and resolution options. 9. We recommend acquiring all free fluorescent channels, also those not necessarily needed for detection of the fluorophores, because it will help to discern macrophages from the rest of the “true” signal. Macrophages, unlike most other cells, will be visible in all channels due to autofluorescence. 10. Increase TCSPC voltage until the decay curve shows no more afterpulses (Fig. 1) and before the detector saturates. If you pass this point, blue pixels will appear in the live image and the decay curve will collapse; decrease TCSPC gain by 1% again. This will be the optimal gain. With the TCSPC warming up, this optimum might increase further. To achieve good SNR for FLIM, it is recommended to reach >20 on the maximum of the intensity scale (depicted as heat map legend below the FLIM image). Carefully turn up laser power if this is not the case. 11. The kinetics and effects of injection anesthesia are very different in individual mice. A young and slim female Balb/c mouse will need less initial dosage, but more frequent reinjections (re-dose ketamine only, 50% of initial) than an older C57BL6J male with significant body fat. Together with your facility’s animal welfare officer, you should determine which dosage applies for the mouse strains in your lab. Additionally, i.p. injection subjects the mice to stress and hormone release, which can reduce the effect of the drugs. Ideally, work with trained animals that are used to the experimenter’s handling and have received sham injections. Calculate 1–2 h extra time before start of measurements. Do not try to speed up the process if you do not observe a reaction to the drugs right away, or even an opposite effect (excitation stage). 12. Depends on cell type, motility, desired total size of the final data stack, and acquisition (wait) time. A typical distance is 4 μm. 13. There are considerations to be made that will affect (a) acquisition and (b) data analysis. Concerning (a) the time between object excitation (sampling rate) should be long enough for you to be able to check on mice (ca. all 15 min). Also, image acquisition and especially the tuning between 760 nm and 850 nm takes time. In other words, the wait time has to be longer than the acquisition takes. If the acquisition time exceeds the wait time, decrease the number of z layers, and consider separate acquisition of the wavelengths or decreasing resolution. Concerning (b) you should adapt the

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sampling rate to known properties of the imaged cells. Movement of the cells during the unobserved times can only be approximated from mathematical models (most likely an option in your analysis software). It is therefore worthwhile considering with which average velocity and directionality your cells of interest are moving. Furthermore, sampling also causes loss of fluorescent lifetime data, which cannot be prevented. For FLIM data acquisition of B cells in and around germinal centers, a frame rate of 1/60 s has proven a good compromise. See also [20]. 14. In ImageJ: To determine the overall SNR values of your TCSPC sum image, randomly choose five regions of interest in the background, and measure (STRG+M) their mean gray value and standard deviation. Average each set of values. Subtract the mean background gray value from all pixels in the image: “Process” → “Math” → “Subtract. . .”. Divide the entire image by the mean background standard deviation: “Process” → “Math” → “Divide. . .”. Now cut off all values under 5. 15. An alternative would be Volocity (Quorum Technologies). 16. You don’t want Imaris to segment B cells from a signal that actually originates from macrophages, but is visible in the same channel. To prevent this, you will have to mask out the voxel belonging to macrophages from the rest of the signal. After having created the “Macrophage” Surface, select the surface. In the creation wizard, click on the pencil tab → “Mask all.” In the pop-up window, you will be asked which values you want to set to zero. Uncheck the box “Set values outside surface to 0.0,” and check “Set values inside surface to 0.0.” This will erase all macrophage-owned voxel from your B cell signal. Imaris is generating a new (masked) channel popping up in the “Display adjustment” Panel. Proceed with this channel for B cell segmentation. 17. In the creation wizard of the selected surface, navigate to the color tab (represented by a rainbow-colored hexagon). Here you can set the coloring to “statistics-based,” and choose a parameter from the drop-down menu. Select “Intensity in Channel 4” (assuming your tau values are in channel 4), and adjust the margins. For Ca2+, the range should include the values from 700 to 2300 ps, for NAD(P)H, from 450 to 3700 ps. This might help finding regions of interest with significantly lower (or higher) lifetimes than average by eye. 18. The Imaris Learning Center is updated regularly: https:// imaris.oxinst.com/learning/. The Imaris Reference Manual is provided upon request and purchase of the software license. The recent edition is Imaris 9.9 (as of October 2022).

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19. “Play” with the values and test specific results within the “Test Mode” to inspect the quality of your adjustments visually. 20. Tau values can always be calculated from real and imaginary values but do not have to be, because they are also exported by the algorithm. This means the information from tau, real, and imaginary channels are redundant to some degree. However, if you want to reconstruct the phasor plots, real and imaginary channels are needed to extract the x and y pixel coordinates, respectively. 21. Visualize the pixel intensity range by right-clicking on original PMT image (pop-up window after testing “RescaleIntensity” in “Test Mode”) → “Show image histology.” 22. “Mean” might generate not-a-number entries (NaNs). In this case, “Median” is a better choice.

Acknowledgments We thank Robert Gu¨nther, Ralf Uecker, and Peggy Mex for excellent technical assistance. A.E.H. was supported by Deutsche Forschungsgemeinschaft (DFG) collaborative research grant TRR130, project P17, and by grant HA5354/12-1. A.E.H. and R.A.N. were supported DFG TRR130, project C01 and DFG CRC1444, project 14, and a grant from the Einstein Foundation Berlin (A-2019-559). References 1. Choi SC, Morel L (2020) Immune metabolism regulation of the germinal center response. Exp Mol Med 52:348. https://doi.org/10.1038/ s12276-020-0392-2 2. Shlomchik MJ, Luo W, Weisel F (2019) Linking signaling and selection in the germinal center. Immunol Rev 288:49–63. https://doi. org/10.1111/imr.12744 3. Zeng Q, Zhou Z, Qin S, Yao Y, Qin J, Zhang H, Zhang R, Xu C, Zhang S, Huang S, Chen L (2020) Rapamycin inhibits B-cell activating factor (BAFF)-stimulated cell proliferation and survival by suppressing Ca2+CaMKII-dependent PTEN/Akt-Erk1/2signaling pathway in normal and neoplastic B-lymphoid cells. Cell Calcium 87:102171. https://doi.org/10.1016/j.ceca.2020. 102171 4. Kwak K, Akkaya M, Pierce SK (2019) B cell signaling in context. Nat Immunol 20:963. https://doi.org/10.1038/s41590-0190427-9

5. Ulbricht C, Leben R, Rakhymzhan A, Kirchhoff F, Nitschke L, Radbruch H, Niesner RA, Hauser AE (2021) Intravital quantification reveals dynamic calcium concentration changes across B cell differentiation stages. elife 10. https://doi.org/10.7554/eLife.56020 6. Leben R, Ko¨hler M, Radbruch H, Hauser AE, Niesner RA, Leben R, Ko¨hler M, Radbruch H, Anja E, RAN H, Leben R, Ko¨hler M, Radbruch H, Hauser AE, Niesner RA (2019) Systematic enzyme mapping of cellular metabolism by phasor-analyzed label-free NAD(P)H fluorescence lifetime imaging. Int J Mol Sci 20: 1 – 1 9 . h t t p s : // d o i . o r g / 1 0 . 3 3 9 0 / ijms20225565 7. Leben R, Ostendorf L, Van Koppen S, Rakhymzhan A, Hauser AE, Radbruch H, Niesner RA (2018) Phasor-based endogenous NAD(P)H fluorescence lifetime imaging unravels specific enzymatic activity of neutrophil granulocytes preceding NETosis. Int J Mol S c i 1 9 . h t t p s : // d o i . o r g / 1 0 . 3 3 9 0 / ijms19041018

Combined Calcium and Metabolic FLIM in B cells 8. Wheeler ML, Defranco AL (2012) Prolonged production of reactive oxygen species in response to B cell receptor stimulation promotes B cell activation and proliferation. J Immunol 189:4405–4416. https://doi.org/ 10.4049/jimmunol.1201433 9. Feng Y-Y, Tang M, Suzuki M, Gunasekara C, Anbe Y, Hiraoka Y, Liu J, Grasberger H, Ohkita M, Matsumura Y, Wang J-Y, Tsubata T (2019) Essential role of NADPH oxidase– dependent production of reactive oxygen species in maintenance of sustained B cell receptor signaling and B cell proliferation. J Immunol 202:2546–2557. https://doi.org/10.4049/ jimmunol.1800443 10. Holmstro¨m KM, Finkel T (2014) Cellular mechanisms and physiological consequences of redox-dependent signalling. Nat Rev Mol Cell Biol 15:411–421. https://doi.org/10. 1038/nrm3801 11. Tsubata T (2020) Involvement of reactive oxygen species (ROS) in BCR signaling as a second messenger BT. In: Wang J-Y (ed) B cells in immunity and tolerance. Springer Singapore, Singapore, pp 37–46 12. Akkaya M, Traba J, Roesler AS, Miozzo P, Akkaya B, Theall BP, Sohn H, Pena M, Smelkinson M, Kabat J, Dahlstrom E, Dorward DW, Skinner J, Sack MN, Pierce SK (2018) Second signals rescue B cells from activation-induced mitochondrial dysfunction and death. Nat Immunol 19:871–884. https://doi.org/10.1038/s41590-0180156-5 13. Digman MA, Caiolfa VR, Zamai M, Gratton E (2008) The phasor approach to fluorescence lifetime imaging analysis. Biophys J 94:L14– L16. https://doi.org/10.1529/biophysj.107. 120154 14. Mank M, Direnberger S, Mrsic-flogel TD, Hofer SB, Ferra A, Stein V, Hendel T, Reiff DF, Levelt C, Borst A, Bonhoeffer T, Griesbeck O (2008) A genetically encoded calcium indicator for chronic in vivo two-photon imaging. Nat Methods 5:805–811. https://doi. org/10.1038/NMETH.1243

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15. Rickert RC, Roes J, Rajewsky K (1997) B lymphocyte-specific, Cre-mediated mutagenesis in mice. Nucleic Acids Res 25:1317–1318. https://doi.org/10.1093/nar/25.6.1317 16. Ulbricht C, Lindquist RL, Tech LM, Hauser AE (2017) Tracking plasma cell differentiation in living mice with two-photon-microscopy. Methods Mol Biol 1623:37–50. https://doi. org/10.1007/978-1-4939-7095-7_3 17. Kamala T (2007) Hock immunization: a humane alternative to mouse footpad injections. J Immunol Methods 328:204–214. https://doi.org/10.1016/j.jim.2007.08.004 18. Rinnenthal JL, Bo¨rnchen C, Radbruch H, Andresen V, Mossakowski A, Siffrin V, Seelemann T, Spiecker H, Moll I, Herz J, Hauser AE, Zipp F, Behne MJ, Niesner R (2013) Parallelized TCSPC for dynamic Intravital fluorescence lifetime imaging: quantifying neuronal dysfunction in neuroinflammation. PLoS One 8:e60100. https://doi.org/10.1371/ journal.pone.0060100 19. Geiger A, Russo L, Gensch T, Thestrup T, Becker S, Hopfner K-PP, Griesinger C, Witte G, Griesbeck O (2012) Correlating calcium binding, Fo¨rster resonance energy transfer, and conformational change in the biosensor TN-XXL. Biophys J 102:2401– 2410. https://doi.org/10.1016/j.bpj.2012. 03.065 20. Frattolin J, Watson DJ, Bonneuil WV, Russell MJ, Fasanella Masci F, Bandara M, Brook BS, Nibbs RJB, Moore JEJ (2021) The critical importance of spatial and temporal scales in designing and interpreting immune cell migration assays. Cell 10. https://doi.org/10. 3390/cells10123439 21. Liublin W, Rausch S, Leben R, Lindquist RL, Fiedler A, Liebeskind J, Beckers IE, Hauser AE, Hartmann S, Niesner RA (2022) NAD (P)H fluorescence lifetime imaging of live intestinal nematodes reveals metabolic crosstalk between parasite and host. Sci Rep 12: 7264. https://doi.org/10.1038/s41598022-10705-y

Chapter 7 Measurement of Molecular Height Using Cell Surface Optical Profilometry (CSOP) Sungmin Son and Daniel A. Fletcher Abstract The plasma membrane of cells is covered by proteins, glycoproteins, and glycolipids with molecular heights ranging from just a few nanometers to hundreds of nanometers. Formation of cell-cell contacts and signal transduction by individual receptors can be dependent on both the average height of a cell’s glycocalyx and the specific height of individual receptors, sometimes with nanometer-scale sensitivity. While superresolution imaging techniques allow molecular distances to be measured with the sub-diffraction limited resolution, typically 10 nm in the lateral direction and 100 nm in the axial direction, measurements of molecular heights at the single nanometer scale on native cell membranes have been difficult to obtain. Cell surface optical profilometry (CSOP) is a simple and rapid method that achieves nanometer height resolution by localizing fluorophores at the tip and base of cell surface molecules and determining their separation with high precision by radially averaging across many molecules. Here we describe how to make CSOP measurements of multi-domain proteins on model membrane surfaces as well as native cell surfaces. Key words Plasma membrane, Glycocalyx, Glycoproteins, Glycolipids, Protein height, Super-resolution microscopy, Cell surface proteins, Antibody epitopes, Cell-cell contacts

1

Introduction Cell surface optical profilometry (CSOP) is based on precise localization of fluorophores [1]. However, rather than localizing in three dimensions like in conventional super-resolution imaging, CSOP achieves nanometer-scale resolution of cell surface molecules in one dimension, height, by foregoing positional information in the other two dimensions. Height is determined using two fluorophores, one at the tip of the cell surface molecule to be measured and the other on the membrane in which the molecule is anchored. In the simplest implementation of CSOP, the fluorescently labeled molecules are arranged in a spherical geometry, such as on a glass bead, giant vesicle, or swollen cell, so that all molecules are oriented axisymmetrically and can be averaged to obtain a measurement of fluorophore radius with high precision (see Note 1). The radius of

Cosima T. Baldari and Michael L. Dustin (eds.), The Immune Synapse: Methods and Protocols, Methods in Molecular Biology, vol. 2654, https://doi.org/10.1007/978-1-0716-3135-5_7, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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Fig. 1 Principle of CSOP. (a) CSOP measures protein height by radially localizing two fluorophores, one labeling the protein tip and the other labeling the membrane surface. (b) Radial averaging of multiple fluorophores (right) improves the localization accuracy compared to the simple line scan (left). (Adapted from Ref. [1] with permission)

each fluorophore is measured separately, and height is then quantified as the difference in radii of the two fluorophores. This approach overcomes the typical ~10 nm resolution limit of single molecule localization [2] by increasing the number of fluorophores used in the radius localization by orders of magnitude (Fig. 1). As in the super-resolution imaging, CSOP resolution is dependent on size of the point-spread function, number of total photons collected, and camera pixel size [3]. For a measurement of fluorescently labeled proteins bound to the surface of a glass bead, CSOP achieves height measurements with 1 nm resolution. For a protein in a native cell membrane, CSOP achieves 2–3 nm resolution. This resolution remains the same for the broad range of molecular heights and is not limited to small distances as in the case of FRET-based distance measurements [4]. The height measured by CSOP is not the fully extended length of a molecule but its average height while freely rotating, tilting, and diffusing on a surface – a measurement that is relevant to a cell surface molecule’s function and can be used to back out properties such as persistence length of multi-domain proteins [1]. CSOP can measure both the intracellular and extracellular heights of a molecule by fluorescently labeling either or both intracellular and extracellular ends of the proteins (Fig. 2). The method can also measure the average height of other parts of a cell surface molecule, such as the location of the domain targeted by an antibody. Furthermore, it can determine the average heights of the entire cell surface proteome and glycome on native cell membranes, as well as the osmotic effects of soluble molecules on molecular heights.

2 2.1

Materials Microscopy

1. A selective z-plane imaging system (e.g., a spinning disk confocal microscopy (Yokogawa CSU-X), a laser scanning confocal microscopy (Zeiss 880 LSM), iSPIM, or diSPIM (ALI)) (see Note 2).

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Fig. 2 CSOP measurement with a GPMV. The height of either the C-terminus of FcүRIIA or the N-terminus of CEACAM5 is measured with respect to the plasma membrane surface. The C-terminus of FcүRIIA is labeled with GFP, the N-terminus of CAECAM 5 is labeled with SPOT-label, the plasma membrane surface is labeled with fluorescent cholera toxin subunit b. The picture (top) shows a typical GPMV image used for CSOP measurement

2. A high magnification immersion objective with low spherical aberrations (e.g., 60× WI NA 1.20 (Nikon) or Apo TIRF 100×, NA 1.49, oil (Nikon)). 3. A multi-spectral laser illumination module (e.g., ILE-400 with 488/561/641 nm solid-state lasers (Andor Technologies)). 4. A filter wheel with emission filters (e.g., ET525/36 M, ET605/52 M, ET705/72 M (Chroma Technology) or an optic-splitter (e.g., W-View Gemini, Hamamatsu). 5. A sensitive camera with a small pixel size (e.g., Zyla 4.2, 6.5 μm/pixel (Andor Technologies)). 6. A z-piezo stage or a motorized z-stage. 2.2 Spherical CSOP Targets

1. 1,2-Dioleoyl-sn-glycero-3-phosphocholine (DOPC) in chloroform (Avanti Polar Lipids or equivalent). 2. 1,2-Dioleoyl-sn-glycero-3-[(N-(5-amino-1-carboxypentyl) iminodiacetic acid)succinyl] (DOGS-NTA) in chloroform (Avanti Polar Lipids or equivalent). 3. 1,2-Dioleoyl-sn-glycero-3-phosphoethanolamine-N-(lissamine rhodamine B sulfonyl) (RhodPE) in chloroform (Avanti Polar Lipids or equivalent). 4. MOPS buffer 10×: 500 mM MOPS pH 7.4, 1 M NaCl. 5. HEPES buffer 1×: 50 mM HEPES pH 7.4, 100 mM NaCl. 6. 293 T cell line ATCC CRL-3216 or equivalent. 7. Complete cell culture media and transfection reagents. 8. Plasmids encoding tagged proteins for transfection of 293 T cells.

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9. Phosphate-buffered saline (PBS): 137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 1.8 mM KH2PO4. 10. 2 mM Latrunculin A in DMSO. 11. 10 mM Y27632 in PBS. 12. Swelling buffer A: PBS with 1 μM latrunculin A and 10 μM Y27632. 13. Swelling buffer B: 10 mM HEPES, pH 7.4, 1 μM latrunculin A and 10 μM Y27632. 14. Lab-Tek chamber slide, Thermo Fisher or equivalent. 15. Poly-L-lysine. 2.3 A Membrane Labeling Reagent

1. Membrane dyes (e.g., HCS CellMask stain, Thermo Fisher). 2. Cell surface label (e.g., labeled cholera toxin subunit B, Thermo Fisher, or equivalent). 3. Fluorescent cholesterol (e.g., cholesterol-PEG-FITC, Nanocs, or equivalent). 4. A fluorescent histidine-repeat peptide that binds to a Ni-NTAcontaining supported lipid bilayer (SLB) or GUV [1].

2.4 A Protein Labeling Reagent

1. SNAP- or CLIP-tag-based labeling reagents (New England Biolabs). 2. Enzymatic labeling reagents [5, 6] (see Note 3). 3. Labeled nanobodies targeting a peptide tag (e.g., SPOT-tag, ChromoTek). 4. Labeled antibodies targeting a protein epitope. 5. A labeled lectin targeting a glycan. 6. Fluorescent fusion proteins.

2.5

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Imaging Buffer

1. A range of neutral-pH, isosmotic media works well for CSOP measurement. For example, HEPES, PBS, MOPS, TRIS buffers are compatible with CSOP.

Methods

3.1 Preparation of CSOP Samples

CSOP uses either a reconstituted lipid bilayer presenting a purified protein such as lipid-coated bead or GUV or a circularized native cell membrane containing a protein of interest such as GPMV or swollen cell. The protocols for preparing GUV and GPMV are extensively described elsewhere [7, 8]. Here we describe protocols for preparing a lipid-coated bead and a swollen cell.

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Preparation of lipid-coated bead coated with a purified protein: 1. Mix lipids: Mix DOPC, 2.5 mol% DOGS-NTA-Ni2+, and 0.15 mol% RhodPE in chloroform, and dehydrate to a glass tube under strong vacuum. 2. Form small unilamellar vesicles (SUVs): Rehydrate with MQ water to 0.22 mg/mL (e.g., 200 μg of dried lipids in 0.9 mL of MQ). Transfer the lipid solution to a 1.5 mL microcentrifuge tube, fix in fine ice, and sonicate with a microtip sonicator (Branson) at low power (20% of max, 1 s ON and 2 s OFF for 3 min). Add 1 part 10× MOPS buffer to 9 part SUV solution to make the final 0.2 mg/mL SUV in 1X MOPS (sonicating lipids in pure MQ help form uniform small vesicles). 3. Piranha clean beads (CAUTION: this solution is dangerous if mishandled. Register use with your local safety office and perform this step under a fume hood, and follow all recommended precautions): Add 10 μL of 6.8 μm glass bead slurry (10% solid, Bands laboratory) to a 10 mL glass tube. Put the tube in a tube holder, and place it in a glass tray for safety. Using a glass Pasteur pipette and a bulb, carefully add 10 drops of H2O2 (equivalent to 200 μL) and 15 drops of H2SO4 (equivalent to 300 μL) to the bead. Mount the bottom portion of tube in a floating tube rack, balance carefully, and sonicate for 30 min in a bath sonicator. To rinse, put the glass tube in a 15 mL Falcon tube, fill the glass tube with ~9 mL MQ, and mix thoroughly using a glass Pasteur pipette. Pellet the bead by centrifuging for 1 min, and aspirate the diluted piranha solution. Repeat the rinsing step for two more times, and, after the final aspiration, add 100 μL MQ to prepare the bead suspension at 1/tenth of original concentration. 4. Form a lipid-coated bead: Add 5 μL of clean bead slurry to 50 μL of SUV solution, and mix gently by pipetting. Incubate the bead/SUV mixture for 15 min at room temperature, laying still on its side. Wash the mixture with HEPES buffer for five times or more. To wash, hold the tube horizontally, and add 500 μL 1× MOPS to the bead settled on sidewall, and, without agitating the bead, gently syphon the solution while leaving the original 50 μL solution in the tube. Be careful not to dry the bead pellet. One can also wash by spin-pelleting the bead. A very slow spin is recommended (e.g., 50 rcf) to prevent bead aggregation and lipid scratch. 5. Coat a protein to the lipid-coated bead: Incubate the lipidcoated bead with the purified, labeled target protein for 30 min. Wash away excess protein as in step 4, and proceed to imaging.

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Preparation of swollen cell presenting a protein of interest: 6. Lift cells by shear or mild trypsinization (e.g., 0.05% at room temperature), and transfer to an imaging chamber coated with Cell-Tak cell Adhesive (Corning) at a density where single cells are separated. Allow cells to gently adhere to the glass for 10 min at room temperature (see Note 4). 7. Wash cells with PBS, and add Swelling Buffer A for 15 min at room temperature or 37 °C. 8. Wash cells three times with PBS, and add labeling reagents, such as a labeled antibody, lectin, or CellMask in Swelling Buffer A for 15 min at 4 °C. We typically add labeled proteins at 1 μg/mL and achieve clean background without washing. 9. Add Swelling Buffer B dropwise to swell cells while observing them under the microscope. Typically, maximal cell swelling is achieved at 1/4 X PBS to 1/10 X PBS (see Note 5). Proceed to imaging. 3.2 Calibration of the CSOP Imaging System

Calibration procedure accounts for the chromatic aberrations specific to the sample and to the microscope configuration (see Note 6). Same samples are used for both the calibration run and the height measurement, except that for the calibration run, two fluorophores are labeling the same lipid membrane (Fig. 3). The imaging method is identical for both calibration and real measurement.

Fig. 3 CSOP image analysis workflow. (a) CSOP calibration measurement using a lipid-coated bead and two fluorophores labeling its membrane surface. Radius of individual fluorophores are quantified at z-positions below and above the equator and a circle fit estimated the maximum radius of each fluorophore, Rred and Rgreen. The offset between two radii hoffset is caused by chromatic aberration. (b) Measurement of CEACAM 5 with either a ybbR-tag (bead) or SPOT-tag (GPMV) added to the tip of protein (red dot). The difference between hmeasured and hoffset reports the actual height of a protein . (c) A representative CSOP result obtained with a lipid-coated bead. Results from a calibration run (blue circle) and a measurement run (black circle) are combined. Each datapoint corresponds to an individual bead. (d) Same as C, except the measurement is obtained with a GPMV. (e) Height of CEACAM5 measured in a bead (C) or a GPMV (D). n = 30 and 40, respectively. P-value based on a two-sample Student’s t-test is 0.56

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1. Prepare a glass-bottom imaging chamber. The surface was passivated by forming an SLB or by adsorbing BSA to prevent nonspecific adsorption of labeled proteins, which can add a background signal. In addition to passivation, a binding molecule is added to the surface to immobilize a CSOP sample (e.g., a low amount of streptavidin that can bind across two biotinylated SLBs or Cell-Tak solution (Corning) that adheres to a GPMV or a swollen cell to the surface) (see Note 7). 2. Prepare a calibration sample with the two fluorophores to be used for the height measurement both labeling the same lipid bilayer, which should give a height measurement of zero (see Note 8). Single field of view can contain more than one CSOP target particle. 3. Prior to CSOP imaging, adjust the laser power and exposure time in both channels using a fixed z-position. If possible, keep the signal of two fluorescence channels comparable to avoid any potential bias to CSOP measurement (see Note 9). 4. Manually determine the equator of samples and acquire approximately 20 slices of images with a 50–100 nm step size using the z-piezo stage. 5. Calculate the radius of fluorophores in each z-slice, and determine the maximum radius in each color at the equator (see Image Processing). The offset between the measured fluorescent radii, hoffset, results solely from the chromatic aberration, providing a parameter that can be used to compensate subsequent measurements. 3.3 Height Measurement

1. Prepare a sample with the two fluorophores labeling the membrane surface and the tip of a protein. Make sure that the background fluorescence either inside or outside the target particle is negligible compared to the signal from the surfacebound protein (see Note 10). 2. Perform imaging by repeating the steps 3 and 4 in Subheading 3.2. 3. Calculate the radial offset between the two fluorophores at the equator, hmeasured. To determine the protein’s average height , compensate the chromatic aberration by subtracting hoffset from hmeasured (see Note 11). Report the error of by calculating the standard deviation of hmeasured (see Note 12).

3.4

Image Analysis

1. Use a custom MATLAB (Mathworks) script (https://github. com/smson-ucb/CSOP) to estimate the radius of samples from CSOP images. The algorithm radially averages fluorescent signals from the center of a circle to construct the 1D signal distribution in the radial direction. For signals that are

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not uniformly distributed around the circular cross-section (e.g., due to clustering), the radial averaging can be limited to portions of the circle that contain the molecules of interest. 2. To determine the center of a circle accurately, we performed the radial averaging in 900 different center locations and quantified the height of radial profile at its peak. We then created the 2D map of peak heights versus centers and fit the 2D Gaussian surface to estimate the true center, where the peak height becomes maximized. 3. To determine the peak location of the radial profile from the center, which corresponds to the radius of CSOP sample, we fit the Gaussian curve to the top 10% region of the peak. 4. To determine the maximum radius at the equator, the radii estimated in individual planes were plotted along the z-axis and fit to a circle. The radius of the circle corresponds to the maximum radius at the equator.

4

Notes 1. Although the symmetry of a spherical target geometry simplifies CSOP image analysis and provides the highest resolution, CSOP also works with a noncircular shape by performing, for example, topological averaging with a polar transformation [9]. 2. In most cases, CSOP measurement with a typical confocal microscope provides the resolution at a single-nanometer scale. In cases where the protein of interest is present at a very low concentration, a photon-efficient high-magnification light-sheet microscope can provide more accurate CSOP measurement by collecting more photons from the limited fluorophores. 3. The choice of fluorophore is important to prevent its membrane interaction and permeation [10]. 4. Extensive trypsinization should be avoided since it can digest the tagged protein of interest. If trypsin is necessary to lift cells, its dose should be optimized so that the protein-tag is preserved for CSOP measurement after successful lift-off. 5. The buffer osmolarity should be decreased slowly to prevent rupturing a cell. Depending on cell type, the swelling rate and the extent of swelling at a given buffer osmolarity widely varies. Empirically, 1/10 X PBS was enough to swell most of cells to a full sphere even though, for some cells, it took tens of minutes to swell them. Diluting the buffer to less than 1/10 X PBS is not recommended as certain antibodies and lectins have been observed to unbind from their targets at higher dilutions.

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6. Typically, a 6.8 μm bead imaged with a 100× Apochromat objective provides the CSOP measurement offset of approximately 10 nm between adjacent channels (e.g., 488–561 or 561–647) or more for distant channels. The offset remains almost constant for the same imaging system (~1 nm drift over a week). Since this offset increases with the diameter of sample sphere (i.e., the depth of sample mid-plane) and the refractive index mismatch between the sphere and the buffer, the calibration run should be performed every time a new sample type is used. 7. For a fast imaging system like spinning disk confocal microscope or light-sheet microscope, the sample vibration does not cause a significant CSOP measurement error. However, for a laser scanning confocal microscope, sample vibration within one imaging frame can distort the image shape, leading to an error. 8. Even though height measured for two fluorophores labeling the same surface should be 0 when no chromatic aberration exists, fluorescent polarization can introduce an additional height offset [11]. Fluorophores are often highly polarized on a membrane surface due to its interaction with lipid. If a fluorophore labeling the tip of a protein is not polarized (during the measurement run) but the same fluorophore is polarized when labeling a membrane surface (during the calibration run), the height measured between the two locations become inaccurate. For this reason, we carefully chose a membrane labeling strategy that is not polarized. For example, we used an Alexa Fluor dye-conjugated hexa-histidine peptide that binds to the Ni-containing lipid bilayers or cholera toxin subunit B that binds to gangliosides on the plasma membrane. 9. If fluorescence from one channel is substantially brighter than the other, a localization error is prone to occur in the dim channel because of the potential spectral bleed-through from the bright channel to the dim one. 10. If a significant background exists either in solution or within a CSOP target, for example, from a residual fluorophore in sample buffer or from a soluble fluorescent fusion protein in the cytosol, it can result in an asymmetric baseline to the radially averaged line scan. An asymmetric baseline can introduce a significant bias to the height measurement. 11. The chromatic aberration changes with sample size. When working with a sample with heterogenous size (e.g., GUVs, GPMVs, swollen cells), care must be taken to measure both hmeasured and hoffset from the similar size samples (Fig. 3d).

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12. The accuracy of CSOP measurement is dependent on both hmeasured and hoffset. However, the error in hmeasured and hoffset are independent, and the accuracy of hoffset can be improved by averaging from a large number of samples. On the other hand, hmeasured can be subjected to a biological variability since the same protein’s height can change across different cell types or cell states. Therefore, we treat that the accuracies of hmeasured and are identical.

Acknowledgments We would like to thank Sho C. Takatori, Brian Belardi, Marija Podolski, and Matthew H. Bakalar for feedback. The FcγR data in Fig. 2 was collected with Emily Sutter. References 1. Son S, Takatori SC, Belardi B, Podolski M, Bakalar MH, Fletcher DA (2020) Molecular height measurement by cell surface optical profilometry (CSOP). Proc Natl Acad Sci U S A 117(25):14209–14219. https://doi.org/10. 1073/pnas.1922626117 2. Huang B, Wang W, Bates M, Zhuang X (2008) Three-dimensional super-resolution imaging by stochastic optical reconstruction microscopy. Science 319(5864):810–813. https:// doi.org/10.1126/science.1153529 3. Thompson RE, Larson DR, Webb WW (2002) Precise nanometer localization analysis for individual fluorescent probes. Biophys J 82(5): 2775–2783. https://doi.org/10.1016/ S0006-3495(02)75618-X 4. Roy R, Hohng S, Ha T (2008) A practical guide to single-molecule FRET. Nat Methods 5(6):507–516. https://doi.org/10.1038/ nmeth.1208 5. Yin J, Straight PD, McLoughlin SM, Zhou Z, Lin AJ, Golan DE, Kelleher NL, Kolter R, Walsh CT (2005) Genetically encoded short peptide tag for versatile protein labeling by Sfp phosphopantetheinyl transferase. Proc Natl Acad Sci 102(44):15815–15820. https://doi.org/10.1073/pnas.0507705102 6. Theile CS, Witte MD, Blom AE, Kundrat L, Ploegh HL, Guimaraes CP (2013) Site-specific N-terminal labeling of proteins using sortasemediated reactions. Nat Protoc 8(9): 1800–1807. https://doi.org/10.1038/nprot. 2013.102

7. Schmid EM, Richmond DL, Fletcher DA (2015) Chapter 17 – Reconstitution of proteins on electroformed giant unilamellar vesicles. In: Ross J, Marshall WF (eds) Methods in cell biology, vol 128. Elsevier, Amsterdam, The Netherlands, pp 319–338. https://doi.org/ 10.1016/bs.mcb.2015.02.004 8. Sezgin E, Kaiser HJ, Baumgart T, Schwille P, Simons K, Levental I (2012) Elucidating membrane structure and protein behavior using giant plasma membrane vesicles. Nat Protoc 7(6):1042–1051. https://doi.org/10.1038/ nprot.2012.059 9. Kumra Ahnlide V, Kumra Ahnlide J, Wrighton S, Beech JP, Nordenfelt P (2022) Nanoscale binding site localization by molecular distance estimation on native cell surfaces using topological image averaging. elife 11: e64709. https://doi.org/10.7554/eLife. 64709 10. Hughes LD, Rawle RJ, Boxer SG (2014) Choose your label wisely: water-soluble fluorophores often interact with lipid bilayers. PLoS One 9(2):e87649. https://doi.org/10.1371/ journal.pone.0087649 11. Lew MD, Backlund MP, Moerner WE (2013) Rotational mobility of single molecules affects localization accuracy in super-resolution fluorescence microscopy. Nano Lett 13(9): 3967–3972. https://doi.org/10.1021/ nl304359p

Chapter 8 Observing Membrane and Cell Adhesion via Reflection Interference Contrast Microscopy Ahmed Abdelrahman, Ana-Suncˇana Smith, and Kheya Sengupta Abstract Reflection interference contrast microscopy (RICM) is an optical microscopy technique ideally suited for imaging adhesion. While RICM (and the closely related interference reflection microscopy (IRM)) has been extensively used qualitatively or semiquantitatively to image cells, including immune cells, it can also be used quantitatively to measure membrane to surface distance, especially for model membranes. Here, we present a protocol for RICM and IRM imaging and the details of semiquantitative and quantitative analysis. Key words Interferometry, Adhesion, Fluctuations, Membranes, Antiflex

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Introduction Reflection interference contrast microscopy (RICM) and its variant interference reflection microscopy (IRM) have become indispensable tools to observe and quantify adhesion of cells and membranes. As the names suggest, they are based on interpreting interference images and thus require no fluorescent labeling. Herein lies the great advantage of IRM/RICM. IRM of cells was first proposed and performed by Curtis [1, 2]. The essential idea was to image the interaction of a cell and a functionalized surface in interference mode. Light reflected from the ventral surface of the cell interferes with light reflected from the glass-buffer interface and the resulting interference pattern can, in principle, be analyzed to obtain a map of cell membrane to surface distance. However, in spite of intense immediate interest to use technique quantitatively, it fell into obscurity, probably because the optical complexity of a cell meant that true quantification was very difficult [2–6]. In the context of cell adhesion, IRM has however been used extensively in a qualitative [7–10] and sometimes semiquantitative manner to measure the extent to cell spreading.

Cosima T. Baldari and Michael L. Dustin (eds.), The Immune Synapse: Methods and Protocols, Methods in Molecular Biology, vol. 2654, https://doi.org/10.1007/978-1-0716-3135-5_8, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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The next breakthrough came when Radler and Sackmann used the antiflex technique to improve contrast and signal-to-noise ratio in IRM, called it RICM, and applied it to optically well-defined objects like colloidal beads and synthetic membranes in the form of supported lipid bilayers or giant unilamellar vesicles [11] The antiflex technique is essentially a trick to cut off all extraneous reflections. It consists of using polarized incident light coupled to an analyzer before the recording device and inserting a lambdaquarter plate in the light-path at the exit/entry of the objective [12]. Soon afterward, the same group refined the modeling to account for tilt, curvature, and divergence of light [13–15]. With the crucial improvement in contrast, RICM was able to measure membrane to surface distances with truly nanometric accuracy and was typically used to reconstruct the shape of the membrane of a giant unilamellar vesicle close to a surface [14, 16]. Later improvements, like use of two wavelengths to improve the range of absolute height measurements [17, 18] and measure the fluctuations of membrane close to a wall [19–22] and use of a new way to calculate height taking advantage of independently measure refractive index of the buffers [23], further improved the application to membranes. Notably, with introduction of dy-RICM (dynamic RICM), it was possible to infer bond organization by analysis of RICM movies [24–26]. Meanwhile, IRM as well as RICM was applied to different cell types, usually to detect adhesion, sometimes for quantitative measurement of adhesion area [27–29], and also for measuring relative height variations [30–32]. More recently, multi-wavelength and multi-aperture RICM could reconstruct both the topography of the ventral surface of an adherent cell and the shape of the entire lamellipodium [33]. Here, however, we shall restrict ourselves to single wavelength RICM, which is sufficient to measure the extent of cell adhesion. The theoretical basis of RICM and IRM has been extensively reviewed before [4, 34–36], as has been detailed on hardware and light path for RICM. Here, we shall provide a detailed protocol on how to use IRM and RICM on microscopes that are already appropriately equipped. Note that IRM can be performed on any standard inverted microscope equipped with a semi-coherent light source of the type used for epi-fluorescence imaging. Already in the single wavelength form described here, RICM and IRM are an indispensable tool to study adhesion of cells to mimic surfaces. The reason is that an excellent vertical resolution can be achieved without fluorescent staining. If the distances do not exceed half the wavelength of the incoming light, relative heights can be resolved with the resolution of down to 1 nm, while the absolute vertical resolution for a fluctuating membrane is 4–5 nm. For fully adhered membranes, 1 nm absolute vertical resolution is achievable [37]. Larger separations can be resolved with multi-

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wavelength techniques, which may be important in highly fluctuating systems. The fluctuations themselves can be harnessed to combat the optical lateral resolution. For small and dilute adhesive contacts, localization precisions comparable to super-resolution microscopy can be obtained. This is ideal for measuring binding and unbinding rates for protein-mediated formation of adhesive contacts in a fully unperturbed manner [24]. In the case of GUVs, which have recently seen a revival, including in the context of immunology [38], RICM can be used quantitatively to determine membrane surface distance to the resolution of down to 1 nm [21, 23, 24, 27, 39]. For cells, several open issues persist, providing only semiquantitative information. One of more difficult challenges in imaging of cells is the index of refraction that is variable in both time and spatial domain, which could be resolved by combining RICM with the emergent techniques capable of providing this information independently. Alternatively, multiple illumination conditions become necessary [33]. Consequently, quantitative information about cell adhesion and local flows of matter could be accessible. These are important perspectives for RICM, which will allow taking this method to a new level. Meanwhile, here, we have provided an easy to use, point-by-point protocol for the uninitiated.

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Materials Our microscope configuration for RICM is schematized in Fig. 1. 1. Inverted microscope (Axiovert, Zeiss, Goettingen, Germany or equivalent). 2. Epi-illumination source we use is a 100 W Hg Arc lamp but can be substituted with appropriate light-emitting diodes. 3. Epi-illumination light path with aperture and field-stop diaphragms and a light source with monochromatic filter. 4. Epi-illumination filter cube with crossed polarizer and analyzer filters and a 50–50 beam splitter. 5. Objective with low reflections and quarter wave plate (preferably with NA > 1.2, oil immersion, for example, Antiflex EC “Plan-Neofluar” 63×/1.25 Oil Ph3 M27 (Zeiss, Goettingen, Germany) (see Note 1). 6. Camera with high sensitivity and low noise. The background should be roughly set in the middle of the detection range. For measurements of substrate–sample separation distance, the number of gray levels is important (usually encoded on 8–16 bits). A good example is an electron multiplication charge coupled device (EMCCD) (iXon, Andor, Belfast, UK or equivalent).

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Fig. 1 Setup of RICM microscope

For sample preparation the cover slide quality is critical: 7. Microscope coverslips of accurate and precise thickness (typically 0.17 mm, e.g., we use μ-Slide I Luer, Cat.No:80176 Ibidi) (see Note 2). 8. Sample-adherent cells—live or fixed, or giant unilamellar vesicles (GUVs) (see Notes 3 and 4).

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Methods

3.1 Microscope Alignment

For both IRM and RICM, Kohler illumination in reflection needs to be ensured. For this: 1. Ensure that the microscope is roughly aligned. Place the sample on the x-y stage, put immersion oil on the objective as appropriate, switch on the fluorescent lamp, and focus as best as possible. 2. If available, use a Bertrand lens to project the image of the aperture diaphragm on the camera and center aperture diaphragm using appropriate keys/screwdrivers to move it. Alternatively, put the objective in place and focus on the sample. Next, remove one eyepiece, and ensuring that the lamp is on and with monochromatic filter in the light path, look into the objective and carefully center the image of the aperture diaphragm. 3. For final alignment, put the objective in place and project the image on the camera. Close the aperture diaphragm as much as possible while still getting enough light. Close the field stop

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such that its image is visible on the camera. Focus on the object of interest—the field stop and the image should be in focus at the same time. 4. The image should not move (only blur) when focusing and defocusing slightly by moving the objective incrementally up and down. Make final small adjustment to center the diaphragm to ensure this. 5. To ensure the best focus, it is helpful to draw the intensity profile across the edge of the field stop in real-time and ensure that this step function is as sharp as possible. 6. The field stop not being in focus at once along all its edges implies that the sample holder is not fully flat. 7. The final adjustment is to the antiflex system. If a pre-adjusted polarization filter cube is used (recommended), the crossing of the polarizer and analyzer is already correct. Else, this needs to be ensured. Next, the lambda quarter plate of the antiflex objective needs to be rotated till the background intensity is maximum. 3.2 Observation and Recording

1. Once the final adjustments including focusing are done, the observation and recording proceed in the same way as traditional microscopy (see Note 5). The advantage of RICM is its capacity to image fast dynamical processes, with 100 ms illumination being a common choice. Fast imaging requires minimizing the exposure time, which comes on the expense of a poorer signal-to-noise ratio. This can be compensated by increasing the illumination through the aperture diaphragm or in post-treatment, when the shot noise needs to be accounted for. 2. The RICM imaging mode can be coupled with other modes like phase contrast or fluorescence. It is usually useful to record a bright-field image by default (Fig. 2). 3. To improve signal-to-noise ratio, short movies can be recorded in RICM mode—the frames are averaged before analysis.

3.3 Image Processing and Data Analysis

Both RICM and IRM images can be used semiquantitatively to measure the adhesion area, and RICM can additionally be used to quantify the size of the gap between the membrane and the surface to which it adheres. Here, the processing was done using in-house Python scripts. However, all the algorithms mentioned are standard and available, for example, in the image processing software Fiji [40].

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Fig. 2 Examples of images obtained in RICM (a–c) or bright-field (d) channel. (a) A nonadherent GUV, (b) GUV is adhered but not fully spread, and (c) GUV is strongly adhered and spread. The state of adhesion of the GUVs cannot be interfered from the transmission bright field image (d)

Fig. 3 Workflow for area and height determination: (a) Original image. Note that sufficient blank background needs to be included for fitting in later step. (b) Denoised image (used exclusively for ease of segmentation). (c) Edge detected image. (d) Mask image (e) fitted background obtained by fitting all area outside the mask and extrapolating into the masked region. (f) Background corrected image obtained by subtracting e from a and adding a constant to restore the mean intensity to its original value. (g) Normalized image obtained by normalizing f with respect to the background. (h, i) Calculated intermediate and final height image, respectively. (j) Histogram of height distribution

The steps, summarized in Fig. 3, are as follows: 1. Adhesion zone: To identify the adhesion zone, first the raw image is de-noised, and then it is segmented, and an edge detection filter (“sobel” algorithm in “stikit-image” Python library) is used, and the edge is detected by using a threshold (using “otso” algorithm in same library). Finally, the detected hole is filled using a filling algorithm (binary fill holes in SciPy open-source Python library). The detected ROI serves as mask and helps in the next step for background subtraction.

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2. Background subtraction: Since the RICM image is very sensitive to small differences in focus, even a tiny tilt in the sample holder results in inhomogeneous background, which needs to be corrected. This is done by first fitting a 2D parabola on the image excluding the adhesion zone, creating a background image from the fitted parabola, subtracting this from the whole image including the adhesion zone, and then adding its average again. 3. Adhesion area: The adhesion area can be easily determined by simply thresholding the intensity. 4. Image normalization: The image is normalized with respect to the background by subtracting and dividing the whole image by the average background intensity. 5. Determination of height versus intensity relation: Separately from image analysis, the parameters in the cosine relation between height and intensity needs to be determined. For this, the theoretical relation between the intensity (I) and the height (h) is calculated using Fresnel formalism for reflection from successive transparent layers with known refractive index and thickness as described in [4] (see Box 1). 6. Height image: The height at each pixel of the image is calculated from the normalized intensity to determine the height image. For this, the normalized intensity in each pixel is used in the arccosine function described in Box 1. It should be mentioned that the argument of the arccosine function needs to be between -1 and 1. If this is not the case, for example, due to noise, an error is generated (see the white dots in the height image in Fig. 3). 7. Average height: To have a good estimate of the height, first a histogram of all the height values inside the adhesion zone is constructed. Next, a Gaussian is fitted to this distribution. The height of the GUV membrane can be defined to be the mean of this Gaussian. This procedure to demine the absolute height avoids strong impact of small defects inside or around the adhesion zone.

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Box 1

Height-intensity relation. The Python code to generate the expected normalized intensity values for heights between 0 and 200 nm can be found here: https://github.com/ahmed344/RICM.git

3.4 Dy-RICM Analysis

For dy-RICM analysis, the previously described height analysis is done on each frame of a RICM movie, consisting of at least 100 frames (Fig. 4). Next steps are described below: 1. Do height analysis on each time frame and store as a stack. 2. Do a time average of the stack to obtain the average height map (average intensity project). 3. At each pixel, calculate the standard deviation of the height to make the fluctuation map (standard deviation project) (see Note 6).

4

Notes 1. IRM can be performed well with nearly any oil immersion objective and most commercial confocal microscopes can be configured to obtain semiquantitative results. Currently, there is only one commercial choice for antiflex objective enabling

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Fig. 4 Fluctuation analysis by dy-RICM: Left to right: bond configurations inferred from the analysis, colorcoded fluctuation map showing bound zones corresponding to the different bond configuration types with RICM intensity image as inset, 3D average height and 3D fluctuation maps

RICM, which is indicated here. In principle, any oil immersion objective with a lambda quarter plate integrated can be used. 2. For quantitative RICM with the aim of determining membrane to substrate distance, a thickness corrected glass cover slide is recommended as support. 3. In the context of immunology, relevant samples consist of mimetic GUVs and cells (see Chapter 5 for a typical GUV preparation protocol). The lipid/cholesterol composition of GUVs can be complex in which case special care is needed to control the phase behavior of the membrane. The preparation technique is often adjusted to the functional units that are incorporated on the GUV surface (mimic glycocalyx, transmembrane proteins or their fragments, ion channels) or in its interior (actin filaments, microtubule, linker proteins, etc.). GUVs are often used to reconstitute adhesion [41, 42]. In this case, they are slightly deflated by using the outer buffer with a somewhat larger osmolality than the inner buffer to create surplus area. The inner buffer is often chosen to promote sedimentation of GUVs toward the functionalized substrate. The substrate is often decorated with ligands or a receptor for the functional unit on the GUV. The binding partner for the GUV on the substrate can be either immobilized due to a deposition on a pattern or simple physisorption. Alternatively, it can be incorporated in a supported lipid bilayer, when it maintains lateral mobility. In all these systems, to get reliable data from RICM, it is necessary to control the spreading of the GUV in a nonspecific adhesive potential with the substrate. Nonspecific adhesion is omnipresent and is controlled by the passivation layer on the substrate [43]. Ideally, it is adjusted such that without proteins of interest promoting adhesion, the vesicle spontaneously maintains a distance of 60–120 nm from the interface. For studies of cell adhesion, similar preparation of the substrate is expected.

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4. Model membranes of GUVs reflect around 0.1% of the light in aqueous solution due to the small variation in refractive index, which make it challenging to image them with acceptable contrast using an interference method. A way to increase the image contrast is to eliminate the stray light, and for that, the antiflex technique was introduced by Ploem in 1975 [12]. Antiflex technique uses a pair of crossed polarizers together with a lambda/4 plate to eliminate most of the stray light that doesn’t come from the sample. When the incident linearly polarized light passes through the lambda/4 plate, it gets circularly polarized. After reflection at various interfaces, when the circularly polarized light passes the lambda/4}-plate a second time, the polarization state of the reflected light is changed back to linear polarized but perpendicular to the initial polarization. The second polarization filter, the analyzer, blocks off light that did not pass twice through the lambda/4 plate, for example, that reflected within the objective. 5. The choice of aperture diaphragm opening depends on the experiment. A closed aperture allows greater imaging depth but less light and hence poorer contrast. However, this may be beneficial when the fluctuations of the membrane of a very floppy vesicle or a cell should be detected. However, if this type of information is recorded, the simple analysis described above may not hold, and dual wave length RICM may need to be used to deconvolve the heights. Larger aperture gives slightly better lateral resolution and improves contrast, and in thin samples such as in cells, it may prevent a second reflection from the upper surface. This is particularly suitable in imaging adhesive contacts and outer regions of the spreading cell. In any case, the alignment needs to be strictly correct, and the Ko¨hler illumination needs to be secured for the RICM image to be usable [4]. 6. Two important points must be taken into account while doing fluctuation analysis. First, it is not correct to work with intensities rather than height. The conversion of intensity to height is essential because of the sinusoidal shape of the intensity height relation. Otherwise, equivalent height variations may give rise to reduced intensity variations close to intensity extrema. Second, even with correct height determination, the intrinsic noise depends on the intensity. In a typical RICM experiment recorded with modern standard cameras, the main source of camera noise is the intrinsic shot noise, also called counting noise, which is proportional to the square root of the intensity. Thus, bright regions may appear to fluctuate more than dark regions simply because of shot noise. This effect needs to be correctly taken into account [24].

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Chapter 9 En-Face Imaging of T Cell-Dendritic Cell Immunological Synapses Alexander Leithner, Jack Merrin, and Michael Sixt Abstract Imaging of the immunological synapse (IS) between dendritic cells (DCs) and T cells in suspension is hampered by suboptimal alignment of cell-cell contacts along the vertical imaging plane. This requires optical sectioning that often results in unsatisfactory resolution in time and space. Here, we present a workflow where DCs and T cells are confined between a layer of glass and polydimethylsiloxane (PDMS) that orients the cells along one, horizontal imaging plane, allowing for fast en-face-imaging of the DC-T cell IS. Key words Dendritic cell, T cell, Immunological synapse, Imaging, PDMS

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Introduction Dendritic cells (DCs) are the most potent antigen-presenting cells [1]. Owing to a unique combination of co-stimulatory ligands, secreted cytokines, and cytoskeleton-regulated biophysical properties, they provide optimal stimulation for naive T cells at the immunological synapse (IS) [2–5]. However, relatively little is known about the structure and dynamics of the DC IS. This is mainly due to the fact that cell-cell interactions in suspension primarily form along the vertical axis, requiring optical sectioning that does not yield the spatiotemporal resolution required to capture events that are often at the timescale of seconds. The advent of lattice-lightsheet-microscopy is, in principle, well suited to address these issues [6, 7] but requires specialized equipment and sufficient IT infrastructure for data storage and analysis that is often not readily available to many laboratories. Additionally, the IS of DCs and T cells is topologically highly complex [8], further complicating the analysis of any given 3D dataset. To circumvent these problems, IS formation between DCs and T cells needs to be oriented along the horizontal plane [9, 10]. This en-face view allows imaging of the

Cosima T. Baldari and Michael L. Dustin (eds.), The Immune Synapse: Methods and Protocols, Methods in Molecular Biology, vol. 2654, https://doi.org/10.1007/978-1-0716-3135-5_9, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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interaction plane in one or only a few optical sections and utilizes the excellent x-y resolution of conventional confocal microscopes. Here, we describe a simple microfluidics-based workflow to confine DCs and T cells between a layer of glass and polydimethylsiloxane (PDMS) [11, 12]. In this assay, DCs are migratory and form dynamic cell-cell contacts with T cells. Synapses preferentially form in the horizontal imaging plane, allowing high-resolution imaging of the interaction plane.

2 2.1

Materials and Equipment PDMS Device

1. LINKCAD (http://www.linkad.com) and either Coreldraw (https://www.coreldraw.com) or Autocad (http://www. autodesk.com) software. 2. 5 inch photomask class 4, 1 μm resolution (JD Photo Data). 3. 4 inch (~10 cm) diameter silicon wafer (Si-Mat). 4. Wafer carrier. 5. Spin coater. 6. EVG Mask Aligner 610 (EVG group). 7. PL-360LP UV filter (Omega Optical). 8. 6005 TF SU8 photoresist (micro resist technologies). 9. SU-8 developer Propylene glycol monomethyl ether acetate (PGMEA) (Merck). 10. Trichloro(1H,1H,2H,2H-perfluorooctyl) (Merck).

silane,

97%

11. Hot plate that can reach 135
C. 12. Vacuum desiccator. 13. Vacuum pump. 14. 100 mm diameter plastic or aluminum casting disk, 1 cm thickness. 15. PDMS Sylgard 184 Kit (Dow). 16. Planetary centrifugal mixer (Thinky, ARE250) and corresponding adaptor and cups; we use 200 mL cups, but other volumes work as well. 17. Aluminum mold for PDMS pistons of 12 mm diameter and 13 mm height; alternatively, a 48-well plate with single-well dimensions of ~12 mm diameter and ~ 16.5 mm height can be used as a mold (e.g., Greiner). 18. An oven that can reach 80
C. 19. Bath sonicator.

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20. 60  15 mm non-pyrogenic polystyrene tissue culture dish (e.g., Falcon Easy-grip, the actual dimensions of these dishes are 54.74  13.13 mm). 21. 22  22 mm glass slides, #2 (e.g., Mentzel). 22. Transparent aquarium silicone sealant (e.g., Marina). 23. Drill with 17 mm drill bits that are suitable for plastic. 24. Round cover glasses, 12 mm diameter, #1 (e.g., Mentzel). 25. Plasma cleaner (Harrick). 26. Adhesive tape. 27. Cloth tape, 100 N/cm tensile strength (e.g., Tesa extra Power Perfect). 28. Ethanol, pure. 29. Isopropanol. 30. Milli-Q water. 31. Scalpel blade. 2.2

Cell Culture

1. 1 PBS, cell culture grade without MgCl2 and CaCl2. 2. Bone marrow-derived dendritic cells from Lifeact-eGFP transgenic mice on a C57BL/6 background [13], see Lutz et al., 1999 [14] for DC differentiation protocols. 3. OT-II transgenic mice on a C57BL/6 background (Charles River or the Jackson Laboratory). 4. CD4+ untouched immunomagnetic T cell isolation kit (e.g., Stemcell). 5. A magnet for immunomagnetic cell separation (e.g., Stemcell). 6. T cell isolation buffer for immunomagnetic cell separation: 1 PBS, cell culture grade without MgCl2 and CaCl2, supplemented with 2% fetal calf serum and 1 mM EDTA. 7. 70 μm cell strainer. 8. Cell culture incubator (37
C, 5% CO2). 9. Cell culture media: RPMI 1640 with 10% fetal calf serum, 2 mM L-glutamine, 100 U/mL penicillin, 100 μg/mL streptomycin, 50 μM 2-mercaptoethanol. 10. LPS. 11. OVA 323–339 peptide (e.g., Invivogen). 12. TAMRA, mixed isomers (e.g., Thermo Fisher), prepare a 10 mM stock solution in DMSO, store at 80
C.

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Methods

3.1 Photolithography of Wafers

Note: The production of wafers by photolithography requires specialized equipment that is relatively expensive. As wafers can in principle be used indefinitely and unless it is not planned to establish photolithography in the laboratory, we recommend obtaining wafers through more specialized collaboration partners. Alternatively, there are a number of companies and foundries that produce customized wafers (e.g., https://www.microfludicsfoundry.com) 1. Design the photomask in Coreldraw or Autocad. The individual micropillars should have a diameter of 500 μm, with a spacing of 500 μm between the micropillars (Fig. 1c). This pattern is supposed to be repeated over the whole photomask. Convert the.dxf files with the design to Gerber format using LINKCAD. 2. Order a 5 inch photomask of class 4, 1 μm resolution of your design in Gerber format from JD Photo Data (https://www.jdphotodata.co.uk). 3. Bake the wafer at 110
C for 5 min. 4. Spin coat 6005 TF SU8 to 4 μm on the wafer. The exact conditions to achieve the desired height differ slightly from machine to machine and relying on the datasheet of the photoresist manufacturer does not always produce satisfying results. We recommend to find the optimal conditions by performing some test-runs and to measure the height with a profilometer. As a guidance, accelerate for 10 s at 500 rpm/s to reach 5025 rpm and spin for 30 s. 5. Bake at 110
C for 4 min. 6. Place the photomask on the wafer and expose at 200 mJ/cm through the UV filter. 7. Bake at 110
C for 5 min, and then develop by submerging the wafer for 1 min in SU-8 developer. 8. Inspect and bake at 135
C for 5 min. 9. Put the wafer in a wafer carrier and place it inside a vacuum desiccator, together with a 15 mL tube, inside a fume hood. Pipette 20 μL of silane into the tube, eject tip in it, and close the desiccator. 10. Apply vacuum, seal the desiccator, and switch off the vacuum pump. 11. Let the silane vaporize for 1 h. 12. Vent the desiccator and close the 15 mL tube with the cap. Dispose of the tube as hazardous waste. Silanes are toxic! Always use a fume hood, check the safety datasheet, and follow local safety guidelines. The wafer is now coated with a thin layer of silane that will allow PDMS to be peeled off easily.

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A.

1. glue piston

2. dry glue

1. drill hole

2. glue glass slide

B.

3. dry glue

C. 500µm 500µm

D.

1. attach micropillars

2. add cells

3. push down confiner

4. attach tape

Fig. 1 Schematic overview of cell confiner production and application. (a) PDMS piston is glued into the lid of cell culture dish. (b) A hole is drilled into the bottom of a cell culture dish, and an imaging glass slide is glued onto the dish. (c) Schematic overview of the micropillar design. Individual micropillars (black squares) have a diameter of 500 μm and are spaced 500 μm apart. (d) A micropillar-bearing small round glass slide is placed on the PDMS piston. DC-T cell suspension is pipetted onto the micropillar. The micropillar is pressed down on the glass slide and kept in place by strong tape

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3.2 Production of Soft PDMS Pistons

Note: The soft PDMS pistons should be 0.5–1 mm longer than the distance between the inner side of the lid and the glass slide at the bottom, which has to be considered when other dishes than the ones recommended are used. This ensures that enough pressure is applied for confinement. 1. Mix silicone elastomer and curing agent in a 30:1 ratio inside a 200 mL centrifugal mixer cup. A single 12  13 mm PDMS piston will require a 1.5–2 mL PDMS mixture. Note that the ratio of elastomer and curing agent determines the stiffness of the PDMS. A high ratio, as is used here, will result in soft PDMS. Mix and degas in the planetary centrifugal mixer. 2. Fill the mold with the mixture. Aluminum molds can be designed in a way to produce soft PDMS pistons of a defined height. Alternatively, or when using a 48-well plate as a mold, the volume of PDMS mixture that is required to achieve a certain height should be determined empirically, for example, by simply marking the required filling level on the mold. Degas in a vacuum desiccator until no more air bubbles emerge from the mixture. 3. Bake PDMS for 6 h or overnight at 80
C. As the curing time primarily depends on the temperature, higher temperatures for a shorter time can be tested if desired. 4. Carefully remove PDMS pistons from the mold, using copious amounts of isopropanol. 5. Glue soft PDMS pistons into the middle of the lids of 60  15 mm non-pyrogenic polystyrene tissue culture dishes using aquarium silicone sealant. Let the sealant dry overnight. (Fig. 1a).

3.3 Preparation of Bottom Dish with a Glass Slide

1. Drill a hole with a diameter of 17 mm in the bottom of a 60  15 mm non-pyrogenic polystyrene tissue culture dish. 2. Clean 22  22 mm, #2 glass slides (see Note 1) by sonicating them first in isopropanol and then in pure ethanol for 20 min. Each with sweeping. Rinse the glass slide in milli-Q water. Cleaned slides can be stored in milli-Q water. 3. Blow dry glass slides with N2 or compressed air. 4. Apply aquarium silicone sealant around the rim of the hole of the cell culture dish. 5. Put the cleaned glass slide onto the cell culture dish. Carefully press down the glass slide with forceps. Excess silicone sealant should be removed with a tissue (Fig. 1b).

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1. Mix silicone elastomer and curing reagent in a 7:1 ratio for a total volume of 10 mL inside a 200 mL cup for the centrifugal mixer. 10 mL is enough to cover one wafer. Mix and degas in a planetary centrifugal mixer. 2. Clean the silicone waver with canned air. Pour the PDMS mixture onto the wafer so that it is completely covered. 3. Plasma clean 12 mm diameter round cover glasses at high intensity for 2 min. Place them on the PDMS mixture covered wafer with the plasma-cleaned side facing downward. As the pillar pattern is repeated over the whole wafer, the exact placement is irrelevant. Carefully press the cover glasses on the wafer using forceps. PDMS on the non-pillar facing side of the cover glasses is not a problem and will be removed after baking. 4. Bake PDMS by placing the wafer on a heating plate for 15 min at 95
C. 5. To remove PDMS micropillars from the wafer, carefully lift the PDMS layer on the edges using a razor blade. Then squirt copious amounts of isopropanol between the wafer and PDMS layer. Repeat this process until the PDMS layer detaches, usually as a single sheet, and PDMS micropillar carrying round glass slides can be peeled off. Glass slides that stick to the wafer can be carefully removed using a razor blade and copious amounts of isopropanol. 6. Allow isopropanol to evaporate and then store round glass slides, PDMS micropillars facing up, in a closed container to protect them from dust (e.g., in an empty cell culture dish with lid) (Fig. 1c).

3.5 Preparation of Cells

Note: Different cells and labeling strategies might be used. For example, OT-II cells can be easily replaced by any other TCR transgenic T cells with or without fluorescently labeled reporters. Unspecific labeling of T cells can be achieved with any other suitable dye or can be replaced with specific probes to report on the position of surface molecules on T cells during synapse formation. Likewise, Lifeact-eGFP expressing DCs can be replaced with other transgenic or otherwise labeled DCs from any other source.

3.5.1 Isolation and Staining of T Cells

1. Sacrifice 6- to 12-week-old OT-II mouse, extract spleen and lymph nodes. Make a single-cell suspension, for example, by squashing spleen and lymph nodes between two microscopy glass slides and filter through a 75 μm cell strainer. Spin down at 300 g for 10 min, resuspend cells in T cell isolation buffer, and isolate CD4+ T cells according to the manual provided by the manufacturer of the T cell isolation kit. The purity, which should be above 90%, can be checked by staining for CD4 followed by analysis on a flow cytometer.

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2. Count CD4+ T cells and bring them to a concentration of 20  106/mL in PBS. It is usually sufficient to stain 10  106 cells. Prepare a 5 μM TAMRA working solution and add a volume that is equivalent to the volume of the cell suspension to the cells to reach an end concentration of 2.5 μM. 3. Stain cells for 10 min at room temperature and protect them from light. 4. Add warm cell culture media to stop the staining and wash cells twice with media. 5. Allow cells to recover for at least 30 min in a cell culture incubator. 3.5.2 Preparation of Dendritic Cells

1. Bring LPS matured DCs to a concentration of 1  106/mL in cell culture media and add OVA peptide to an end concentration of 0.1 μg/mL. 2. Incubate DCs with the peptide for 1 h in a cell culture incubator.

3.6 Assembly of Confiner Setup

1. Clean a PDMS micropillar-bearing round glass slide with adhesive tape to remove any potential dust (see Note 2). 2. Place the cleaned glass slide onto a PDMS piston with the micropillars facing upward. 3. Pipette up to 1 mL of cell culture media into the outer edge of the bottom dish with the glass slide. This will ensure that the cells under confinement will not dry out during the experiment. 4. Bring T cells to a concentration of 200  106/mL and DCs to a concentration of 50  106/mL. 5. Mix the two cell suspensions in a ratio of 1:1 to have at least 5 μL. 6. Pipette 5 μL (~12.500 DCs and 50.000 T cells) of the mixture in the center of the PDMS micropillar (Fig. 1d). 7. Assemble the confiner by pressing the PDMS piston on the glass slide of the bottom dish. Keep pressing down while you use cloth tape to secure the lid in place (Fig. 1d). 8. Tilt the dish slightly until the cell culture media that was pipetted into the bottom dish with the glass slide in 3. makes contact with the outside of the micropillar bearing coverslip. 9. Place the dish in a cell culture incubator for ~30 min before you start imaging (see Notes 3–6).

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Notes 1. It is paramount to use #2 glass slides. Glass slides below this thickness bend and will not allow cell confinement. In our experience, this does not interfere with imaging using water or oil objectives. 2. The micropillars should be completely dry before cells are added. Due to the hydrophobic nature of the PDMS micropillar array, a clear drop will form that retains the cells in the center of the micropillar until confinement and thus ensures that the maximum number of cells is confined. 3. What to expect: Typical results of DC-T cell confinement can be seen in Fig. 2. DCs exhibit a polarized morphology, are migratory, and form contacts with T cells. Importantly, DC-T cell synapses preferentially form in the horizontal plane, making them ideal for fast imaging with confocal microscopy. 4. We usually use PDMS devices whose micropillars consist of multiple channels instead of single PDMS structures (Fig. 2a). Although we found that this makes cell confinement more efficient, it is not an essential design principle. 5. One indication of successful cell confinement is the absence of cells underneath the micropillars. If cells are visible below the micropillars, foster confinement by lightly tapping the glass bottom with metal forceps. 6. In the setup described here, T cells are only weakly migratory. However, the bottom glass slide can be pre-absorbed, for example, with ICAM1 and CCL21 to trigger robust migration of T cells if required.

Acknowledgments A.L. was funded by an Erwin Schro¨dinger postdoctoral fellowship of the Austrian Science Fund (FWF, project number: J4542-B) and is an EMBO non-stipendiary postdoctoral fellow. This work was supported by a European Research Council grant ERC-CoG72437 to M.S. We thank the Imaging & Optics facility, the Nanofabrication facility, and the Miba Machine Shop of ISTA for their excellent support.

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Fig. 2 Exemplary images of confined DC-T cell immunological synapses. (a) Overview image of a typical cell confiner experiment with antigen-loaded DCs and T cells. Yellow ¼ pillars of PDMS confiner showing channels (see Note 4), magenta ¼ T cells, cyan ¼ DCs. Yellow arrows highlight synapses that have formed in the horizontal imaging plane. Scale bar ¼ 50 μm. (b) Exemplary spinning disc confocal image of an antigenloaded Lifeact-eGFP-expressing DC (cyan) that has formed synapses with several TAMRA-stained T cells (magenta). Yellow arrows highlight a synapse that has formed in the horizontal imaging plane. Note the Lifeact-eGFP accumulation in the DC at the synapse. Scale bar ¼ 10 μm. (c) Orthogonal view of the DC-T cell synapse highlighted in (b). Scale bar ¼ 5 μm. (d) and (e) Exemplary time-lapse imaging of synapses formed between antigen-loaded Lifeact-eGFP-expressing DCs (cyan) and TAMRA stained T cells (magenta). Yellow arrows in (e) highlight the Lifeact-eGFP dynamics in the DC as the T cell moves underneath it. Imaging interval ¼ 20 s, scale bar ¼ 10 μm

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References 1. Banchereau J, Steinman RM (1998) Dendritic cells and the control of immunity. Nature 392: 245–252. https://doi.org/10.1038/32588 2. Comrie WA, Li S, Boyle S, Burkhardt JK (2015) The dendritic cell cytoskeleton promotes T cell adhesion and activation by constraining ICAM-1 mobility. J Cell Biol 208: 457–473. https://doi.org/10.1083/jcb. 201406120 3. Leithner A, Altenburger LM, Hauschild R et al (2021) Dendritic cell actin dynamics control contact duration and priming efficiency at the immunological synapse. J Cell Biol 220. https://doi.org/10.1083/JCB.202006081 4. Malinova D, Fritzsche M, Nowosad CR et al (2016) WASp-dependent actin cytoskeleton stability at the dendritic cell immunological synapse is required for extensive, functional T cell contacts. J Leukoc Biol 99:699–710. https://doi.org/10.1189/jlb.2a0215-050rr 5. Blumenthal D, Chandra V, Avery L, Burkhardt JK (2020) Mouse t cell priming is enhanced by maturation-dependent stiffening of the dendritic cell cortex. elife 9:1–44. https://doi. org/10.7554/eLife.55995 6. Chen BC, Legant WR, Wang K et al (2014) Lattice light-sheet microscopy: imaging molecules to embryos at high spatiotemporal resolution. Science 1979:346. https://doi.org/10. 1126/science.1257998 7. Ritter AT, Asano Y, Stinchcombe JC et al (2015) Actin depletion initiates events leading to granule secretion at the immunological synapse. Immunity 42:864–876. https://doi.org/ 10.1016/j.immuni.2015.04.013

8. Fisher PJ, Bulur PA, Vuk-Pavlovic S et al (2008) Dendritic cell microvilli: a novel membrane structure associated with the multifocal synapse and T-cell clustering. Blood 112: 5037–5045. https://doi.org/10.1182/ blood-2008-04-149526 9. Oddos S, Dunsby C, Purbhoo MA et al (2008) High-speed high-resolution imaging of intercellular immune synapses using optical tweezers. Biophys J 95:L66–L68. https://doi.org/ 10.1529/biophysj.108.143198 10. Biggs MJP, Milone MC, Santos LC et al (2011) High-resolution imaging of the immunological synapse and T-cell receptor microclustering through microfabricated substrates. J R Soc Interface 8:1462–1471. https://doi. org/10.1098/rsif.2011.0025 11. le Berre M, Zlotek-Zlotkiewicz E, Bonazzi D et al (2014) Methods for two-dimensional cell confinement. Methods Cell Biol 121:213–229. https://d oi.org/10.1016 /B97 8-0-12800281-0.00014-2 12. Schwarz J, Sixt M (2016) Quantitative analysis of dendritic cell haptotaxis, 1st edn. Elsevier Inc. 13. Riedl J, Flynn KC, Raducanu A et al (2010) Lifeact mice for studying F-actin dynamics. Nat Methods 7:168–169. https://doi.org/10. 1038/nmeth0310-168 14. Lutz MB, Kukutsch N, Ogilvie ALJ et al (1999) An advanced culture method for generating large quantities of highly pure dendritic cells from mouse bone marrow. J Immunol Methods 223:77–92. https://doi.org/10. 1016/S0022-1759(98)00204-X

Chapter 10 High- and Super-Resolution Imaging of Cell-Cell Interfaces Julia Sajman and Eilon Sherman Abstract Physical interfaces mediate interactions between multiple types of cells. Despite the importance of such interfaces to the cells’ function, their high-resolution optical imaging has been typically limited due to poor alignment of the interfaces relative to the optical plane of imaging. Here, we present a simple and robust method to align cell-cell interfaces in parallel to the coverslip by adhering the interacting cells to two opposing coverslips and bringing them into contact in a controlled and stable fashion. We demonstrate aberration-free high-resolution imaging of interfaces between live T cells and antigen-presenting cells, known as immune synapses, as an outstanding example. Imaging methods may include multiple diffractionlimited and super-resolution microscopy techniques (e.g., bright-field, confocal, STED, and dSTORM). Thus, our simple and widely compatible approach allows imaging with high- and super-resolution the intricate structure and molecular organization within a variety of cell-cell interfaces. Key words Imaging, Microcopy, Super-resolution, Diffraction limit, Immune synapse, Cell-to-cell interaction, T cells

1

Introduction Cell-cell interactions are an important physiological process in both unicellular and multicellular organisms. Such interfaces convey mechanical support, signaling cues, and exchange of nutrients between the interacting cells [1]. Some interactions are stable and long-lasting, while others demonstrate fast dynamics. Long-lasting interactions include, for instance, cell-cell junctions in epithelial and muscle tissues [2], neuronal synapses and interactions with glial ending [3], cell lacunas in connective tissues [4], etc. Transient cell-cell interactions may involve interactions of blood and lymphoid cells, bacteria, and others [5–8]. Important examples of highly dynamic cell-cell interfaces are immune synapses—the interfaces that form between CD4+ T cells and antigen-presenting cells, or between CD8+ T cells and infected or malignant cells [6]. In response to pathogen recognition in the body, these specialized immune cells get activated and help to

Cosima T. Baldari and Michael L. Dustin (eds.), The Immune Synapse: Methods and Protocols, Methods in Molecular Biology, vol. 2654, https://doi.org/10.1007/978-1-0716-3135-5_10, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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mount an immune response [9]. Subsequently, most of these immune cells continue and meet other cells and test if they are infected, transformed, or intact. Each of such encounters can form very quickly, within seconds to minutes, and dynamically evolve till separation [10]. For example, activated T or NK cells meet infected somatic cells, sensitively recognize these cells as infected, and promptly, selectively, and directionally attack them toward their killing [11, 12]. In such short and transient encounters, two cells meet each other, form an intricate interface, and exchange information through substantial molecular mobilization, reorganization, and signaling, along with major geometrical modifications of the interface before each cell goes on its own way. The cell-cell interaction then leaves a relatively more prolonged mark on the cells, as they get influenced in multiple ways. Such cellular modifications and responses should be studied in detail in order to better understand mechanism and function of the immune cells. For instance, molecular interactions occur between surface cells receptors and molecules. Molecules involved in cell-cell interactions move toward the synapse, as the immune cells become polarized [6]. Other molecules move outside the synapse in order not to interfere with the cell-cell interaction [13]. Some proteins undergo conformational changes, get activated, and activate other membrane and cytoplasmic proteins in their surroundings. Other proteins recruit vesicular proteins to the plasma membrane and initiate or propagate intracellular signaling pathways through activating kinases, phosphatases, or secondary messengers. Later on, gene expression changes (being either upregulated or downregulated), molecules get synthetized, and ions and molecules exit or enter the cell through carrier proteins or via exo/endocytosis [13]. The latter facilitate the targeted cell killing of the infected cells. High-resolution optical microscopy allows the detailed study of such cells and their interfaces in fixed and live cells (e.g., [10]). In fixed cells, these interfaces can be conveniently imaged with essentially no time constraints [14]. Still, for short-lasting cell-cell interactions, live cell imaging is much more challenging. Moreover, microscopy has been significantly improved in recent decades [10, 15, 16]. Specific super-resolution microscopy techniques currently allow visualization and localization of single fluorescent molecules in intact cells, well beyond the diffraction limit of light [17]. One can image static protein organization patterns in the single cell [18]. Dynamics of certain proteins can be followed over time through live cell imaging [19]. Molecular (re)organization can be visualized during the cell-cell interaction (e.g., [20]). In spite of recent advancements, high- and super-resolution imaging of cell-cell conjugates have been hindered by the inappropriate orientation of the interface in typical microscopy. Visualizing the interface between cell-cell conjugates by optical microscopy

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suffers from several limitations. Most microscopes are planned for imaging in horizontal (X-Y) planes with higher resolution (~200 nm) and less optical aberrations than in a perpendicular plane (i.e., along the Z-axis; ~750 nm) [21]. Also, most superresolution methods improve the resolution primarily in the X-Y planes [17]. Due to this difficulty in harnessing the highest resolution possible in imaging cell-cell interfaces, multiple artificial interfaces have been developed to mimic the target cells [especially antigenpresenting cells (APCs)]. Such model systems utilize glasssupported lipid bilayer with embedded molecules of interest [22] and coverslips coated with stimulatory antibodies [23]. Such interfaces have been highly successful in synchronizing the cell interaction with the surface and high-resolution imaging of the interface over time. Thus, intricate features of the IS have been resolved and studied (e.g., as reviewed in [10]). Still, there is a need for more physiological description of the dynamic 3D architecture of the IS and molecular organization within, yet with minimal loss of spatial and temporal resolution. Such properties cannot be fully captured using the cell mimics. Recently, we developed a simple way to form the interface between the cells in an orientation that is parallel to the coverslip and thus favorable for high- and super-resolution fluorescence microscopy [24]. We showed that this method is compatible with a wide range of microscopy modalities and imaging techniques. These include wide-field and confocal arrangements, diffractionlimited and super-resolution microscopy, as well as fixed- and livecell imaging. Imaging covers spatial ranges from large-scale microscopy for quickly identifying forming cell conjugates, to single cellcell conjugates, and even to single molecules at their interface. Demonstrated super-resolution imaging and reconstruction techniques include stimulated emission depletion (STED) [25], single molecule localization microscopy (SMLM) such as direct stochastic optical reconstruction microscopy (dSTORM) and photoactivated localization microscopy (PALM) (reviewed in [26]), super-resolution optical fluctuation imaging (SOFI) [27], and super-resolution radial fluctuations (SRRF) [28]. The spatial resolutions obtained were comparable to imaging of single cells using these techniques [24]. The high resolution of our imaging attests to the lack of optical aberrations and high mechanical stability of our approach. Our imaging typically includes two colors, for distinguishing various molecules and entities on the two interacting cells, yet can be easily expanded to more colors. Thus, our simple and widely compatible approach allows imaging with high- and super-resolution the intricate structure and molecular organization within immune synapses and in a variety of other cell-cell interfaces.

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Materials 1. Glass or polymer (170 ± 5 μm).

bottom

sterilized

chamber

#1.5

2. 1 mm Microscope slide glasses. 3. Glass cutter. 4. Phosphate-buffered saline (PBS). 5. Acidic ethanol: Fresh made 1 M HCL in 70% Et-OH solution for chamber clearing. 6. 0.01% Poly-L-Lysine (PLL) in double-distilled water (DDW) for antibody adhesion. 7. Non-stimulatory antibody for slides coating: anti-CD45 and anti-CD11 for lymphocytes. 8. Cell culture medium appropriate to your cell types. 9. Two cell types of interest. 10. Antibodies/dyes for fluorescent imaging specific for your experiment. 11. Fluorescence-assisted cell sorting (FACS) buffer: 10% fetal bovine serum, 0.02% NaN3 in PBS. 12. Tweezers. 13. 20 mg/mL silicon beads, 20 μm diameter. 14. 4% Formaldehyde solution in PBS: Dissolve 40 g of paraformaldehyde powder in 800 mL PBS upon 1 N NaOH titration, heating to 60 °C and stirring. Cool and filter, adjust the pH to 6.9 with HCl and to 1 L volume of PBS. Store at 4 °C for a few weeks or at-20 °C for longer periods. 15. Imaging buffer: RPMI without phenol red, 10% serum, 25 mM Hepes (filter sterilized). 16. dSTORM buffer: 50 mM Tris pH 8.0, 10 mM NaCl, 0.5 mg/ mL glucose oxidase, 40 μg/mL catalase, 10% (w/v) glucose, 10 mM 2-aminoethaneyhiol (MEA) [29].

3 3.1

Method Preparations

1. Choose your cells of interest for studying cell-cell interaction. Grow each cell type separately in an appropriate growth medium. 2. Prepare a chamber for microscopic imaging. We used #1.5 slide with 8 wells (glass or polymer bottom). 3. Prepare small glasses for carrying the opposite cells to the cells in the chambers. We cut 1 mm microscope slide glasses into ~0.8 × ~0.8 mm2 squares (a size that fits into single 1 × 1 mm2 chamber wells).

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1. Wash chambers and small glasses with acidic ethanol at room temperature (RT) for 10 min; wash with ethanol; aspirate the liquid and dry the coverslips at 37 °C for 1 h (see Note 1). 2. Incubate cleaned coverslips and small glasses at RT for 15 min with 0.01% poly-L-lysine diluted in water. Aspirate the liquid and let dry at 37 °C for 2 h (see Note 1). 3. If one of the cell types grows in suspension, perform this step. Incubate small glasses with non-stimulatory antibodies at a concentration of 10 μg/mL overnight at 4 °C or 2 h at 37 ° C. Wash three times with PBS (see Note 2). 4. Cell types that grow adherent to a plate can be seeded directly in a chamber or on a small glass a day prior to the experiment in high confluence.

3.3 Cell Staining (See Note 3)

1. On the day of the experiment, label proteins of interest, in each of the cell types separately using immuno-fluorescence (IF) protocol (see Note 4) or stain with some general fluorescent markers (e.g., specific to certain cellular organelles). See Fig. 1a. 2. If one of your cell types grows in suspension, drop the cells after staining onto the coverslip covered with non-stimulating antibody and let it adhere to the coverslip for 10 min in 37 °C, 5% CO2 incubator. See Fig. 1b.

3.4 Cell-on-Cell Engagement

1. Add 10 μL silicon beads to the chamber well for keeping a predetermined separation distance (in height) between the opposite slides (see Note 5). The size of the beads should be about the sum of the average diameters of the two cell types. In the case of imaging T cell interaction with APC, we typically used 20 μm beads. 2. Add (d)STORM buffer for SMLM imaging or imaging buffer for applying other methods of fluorescent microscopy. 3. Flip small glass with one cell type onto the chamber, which carries the other cell type (see Note 6). Use tweezers for handling the small glass. See Fig. 1c.

3.5

Imaging

1. Using bright-field (BF) imaging, find a pair of cells that reside one on top of the other (see Note 7). 2. Using proper laser excitation and fluorescence imaging, find the interface between the cells. Start focusing on the lower cell and then focus higher until finding the interface between the two cells (see Note 8). See Fig. 1d. 3. Take an image/movie of fluorescent markers in the interface (see Note 9). Additional planes of interest across the cell-cell conjugate may be captured by shifting or scanning the focal plane in the Z-axis.

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Staining for IF

A. Adherent cells

B. Suspension cells Adhering to surface

Staining for IF

C. Cell-cell conjugation

D. Imaging cell-cell interfaces

Silicon bead

Silicon bead

Fig. 1 Schematic representation of a method for high- and super-resolution imaging of cell-cell conjugates in a favorable orientation, explained step by step. (a) Adherent cell staining. (b) Suspension cells staining and attachment to glass slide. (c) Cell-cell engagement. (d) Imaging of cell-cell interactions

4. We used for imaging a total internal reflection (TIRF) N-STORM Nikon microscope. For staining, we used photoactivatable fluorophores and captured each cell-to-cell contact by taking fast 2000 frames of each fluorophore channel separately. Image analysis and reconstruction were performed using the ThunderSTORM plugin [30] of ImageJ on each channel in separate. Then, we overlaid green and red channels (either with or without bright-field image). See Figs. 2 and 3 for examples.

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A. DiD-stained (red) CD8+ T cells

DiO-stained (green) APC cells

C. Green (DiO)

B. Bright field

D. Red (DiD)

1.5 μm

E. Overlay (all)

F. Overlay (Green+Red)

G. Zoom in

0.5 μm

Fig. 2 dSTORM images of cell conjugates of CD8+ T cells with T2 antigen-presenting cells (APC). T2 cells were loaded with the activating peptide NY-ESO-1. The plasma membrane (PM) of the CD8+ T cells was stained with DiD (red) and the PM of the T2 APC cells was stained with DiO (green). (a) Schematic description of the sample. (b) Bright-field image. (c) DiO stained bottom cell. (d) DiD stained top cell. (e) Fluorescence and BF overlaid images. (f) Merged image of DiO and DiD fluorescence. (g) Zoom in image of panel f

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Notes 1. Chambers and glasses after this stage can be kept at RT for a few weeks. 2. Slides coated with antibodies can be kept for ~1 week in 4 °C in PBS.

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A. CD45-Atto488 CD8+ T cells (green)

CD45-Alexa647 APC cells (red) C. Green (CD45)

B. Bright field

D. Red (CD45)

1.5 μm

E. Overlay (all)

F. Overlay (Green+Red)

G. Zoom in

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Fig. 3 dSTORM images of cell conjugates of CD8+ T cells with T2 antigen-presenting (APC) cells. T2 cells were loaded with the activating peptide NY-ESO-1. The plasma membrane (PM) of the T2 APC cells was stained with CD45-Alexa647 (red) and the PM of the CD8+ T cells was stained with CD45-Atto488 (green). (a) Schematic description of the sample. (b) Bright-field image. (c) CD45-Atto488 stained top cell. (d) CD45Alexa647 stained bottom cell. (e) Fluorescence and BF overlaid images. (f) Merged image of CD45 fluorescence. (g) Zoom in image of panel f

3. If your proteins are genetically fused to fluorescent proteins (FPs) proceed to Subheading 3.4. 4. We used primarily surface staining (including membrane dyes or IF labeling of surface proteins) of the live cells, without fixation, and permeabilization prior engagement. This option often yields higher fluorescent signals relative to imaging proteins tagged with FPs.

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5. The recommended size of beads is about the sum of average heights of the two cell types (~10 μm each). 6. If your interaction is fast, perform the last step (flip and place the small glass on the surface of the chamber and its adherent cells) under the microscope. If the interaction is prolonged or if you fix the cells, this last step can be performed in a hood prior fixation. 7. If the two types of cells are significantly different in size or morphology, this difference can help in recognizing a heterotypic cell-cell conjugate (of interest). If there are a low number of cell-cell interactions, next time seed/attach higher confluence of one of the cell types for achieving higher probability of cell-cell engagements—and thus higher yield of productive cell-cell conjugates. It is useful to quickly scan multiple fields in this stage (using both bright-field and fluorescence imaging) for finding and selection of candidate cell-cell conjugates for further highand super-resolution imaging. 8. If the two types of cells are stained with different colors, it is easier to focus on the right plane when both colors simultaneously, in the same plane. 9. For super-resolution imaging, it is advantageous to stain the cells adhering to the bottom coverslip with the more stable fluorophore. This helps in reducing photo bleaching of the less stable fluorescent stains. References 1. Rusu AD, Georgiou M (2020) The multifarious regulation of the apical junctional complex. https://doi.org/10.1098/rsob.190278 2. Cavey M, Lecuit T (2009) Molecular bases of cell-cell junctions stability and dynamics. Cold Spring Harb Perspect Biol 1: a002998. 10.1101/cshperspect.a002998 3. Nicholls JG (2012) From neuron to brain. Sinauer Associates: Oxford University Press 4. Helfrich MH, Horton MA (2006) Integrins and other adhesion molecules. Dyn Bone Cartil Metab:129–151. https://doi.org/10. 1016/B978-012088562-6/50009-1 5. Minetti G, Campbell R, Barshtein G, Pretini V, Koenen MH, Kaestner L, A M Fens MH, Schiffelers RM, Bartels M, Van Wijk R (2019) Red blood cells: chasing interactions. Front Physiol 10:945. https://doi.org/10.3389/fphys. 2019.00945 6. Trautmann A, Valitutti S (2003) The diversity of immunological synapses. Curr Opin

Immunol 15:249–254. https://doi.org/10. 1016/S0952-7915(03)00040-2 7. Garnett JA, Matthews S (2012) Interactions in bacterial biofilm development: A structural perspective, Current Protein & Peptide Science 13. https://dx. doi.org/10.2174/138920312804871166 8. Abisado RG, Benomar S, Klaus JR, Dandekar AA, Chandler JR, Garsin DA (2018) Bacterial quorum sensing and microbial community interactions. mBio 9:e02331-17 https://doi. org/10.1128/mBio.02331-17 9. Choquet D, Triller A (2013) The dynamic synapse. Neuron 80:691–703. https://doi.org/ 10.1016/J.NEURON.2013.10.013 10. Balagopalan L, Sherman E, Barr VA, Samelson LE (2011) Imaging techniques for assaying lymphocyte activation in action. Nat Rev Immunol 11:21. https://doi.org/10.1038/ nri2903 11. Actor JK (2012) T-cell immunity. Elsevier’s Integr Rev Immunol Microbiol 25–32.

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https://doi.org/10.1016/B978-0-32307447-6.00004-1 12. Vivier E, Tomasello E, Baratin M, Walzer T, Ugolini S (2008) Functions of natural killer cells. Nat Immunol 9:503–510. https://doi. org/10.1038/ni1582 13. Smith-Garvin JE, Koretzky GA, Jordan MS (2009) T cell activation. Annu Rev Immunol 27:591. https://doi.org/10.1146/annurev. immunol.021908.132706 14. Green MV, Pengo T, Raybuck JD, Naqvi T, McMullan HM, Hawkinson JE, Marron Fernandez de Velasco E, Muntean BS, Martemyanov KA, Satterfield R, Young SM, Thayer SA (2019) Automated live-cell imaging of synapses in rat and human neuronal cultures. Front Cell Neurosci 13:1–14. https://doi. org/10.3389/fncel.2019.00467 15. Okabe S (2012) Fluorescence imaging of synapse formation and remodeling. Microscopy (Oxf) 62:51–62. https://doi.org/10.1093/ jmicro/dfs083 16. Ng MR, Besser A, Brugge JS, Danuser G (2014) Mapping the dynamics of force transduction at cell–cell junctions of epithelial clusters. elife 3:e03282. https://doi.org/10. 7554/eLife.03282 17. Hell SW (2007) Far-field optical nanoscopy. Science 316:5828 18. Stender AS, Marchuk K, Liu C, Sander S, Meyer MW, Smith EA, Neupane B, Wang G, Li J, Cheng J-X, Huang B, Fang N (2013) Single cell optical imaging and spectroscopy. Chem Rev. 113:2469–2527. https://doi.org/ 10.1021/cr300336e 19. Ettinger A, Wittmann T (2014) Fluorescence live cell imaging. In: Methods in cell biology, pp 77–94 20. Razvag Y, Neve-Oz Y, Sajman J, Reches M, Sherman E (2018) Nanoscale kinetic segregation of TCR and CD45 in engaged microvilli facilitates early T cell activation. Nat Commun 9:732. https://doi.org/10.1038/s41467018-03127-w 21. Nasse MJ, Woehl JC (2010) Realistic modeling of the illumination point spread function in confocal scanning optical microscopy. J Opt Soc Am A 27:295. https://doi.org/10.1364/ JOSAA.27.000295 22. Grakoui A, Bromley SK, Sumen C, Davis MM, Shaw AS, Allen PM, Dustin ML (1999) The

immunological synapse: a molecular machine controlling T cell activation. Science 285: 221–227. https://doi.org/10.1126/science. 285.5425.2 23. Bunnell SC, Barr VA, Fuller CL, Samelson LE (2003) High-resolution multicolor imaging of dynamic signaling complexes in T cells stimulated by planar substrates. Sci STKE 2003:pl8. https://doi.org/10.1126/stke.2003.177.pl8 24. Sajman J, Razvag Y, Schidorsky S, Kinrot S, Hermon K, Yakovian O, Sherman E (2021) Adhering interacting cells to two opposing coverslips allows super-resolution imaging of cell-cell interfaces. Commun Biol 4:439. https://doi.org/10.1038/S42003-02101960-2 25. Willig KI, Harke B, Medda R, Hell SW (2007) STED microscopy with continuous wave beams. Nat Methods 4:915-8. https://doi. org/10.1038/NMETH1108 26. Khater IM, Nabi IR, Hamarneh G (2020) A review of super-resolution single-molecule localization microscopy cluster analysis and quantification methods. Patterns 1:100038. https://doi.org/10.1016/J.PATTER.2020. 100038 27. Dertinger T, Colyer R, Iyer G, Weiss S, Enderlein J (2009) Fast, background-free, 3D superresolution optical fluctuation imaging (SOFI). Proc Natl Acad Sci U S A 106:22287–22292. https://doi.org/10.1073/pnas.0907866106 28. Culley S, Tosheva KL, Matos Pereira P, Henriques R (2018) SRRF: Universal live-cell superresolution microscopy. Int J Biochem Cell Biol 101:74–79. https://doi.org/10.1016/J.BIO CEL.2018.05.014 29. Dempsey GT, Vaughan JC, Chen KH, Bates M, Zhuang X (2011) Evaluation of fluorophores for optimal performance in localization-based super-resolution imaging. Nat Methods 8:1027. https://doi.org/10. 1038/nMeth.1768 30. Ovesny´ M, Krˇ´ızˇek P, Borkovec J, Sˇvindrych Z, Hagen GM (2014) ThunderSTORM: a comprehensive ImageJ plug-in for PALM and STORM data analysis and super-resolution imaging. Bioinformatics 30:2389–2390. https://doi.org/10.1093/bioinformatics/ btu202

Chapter 11 Separation of Single Core and Multicore Lytic Granules by Subcellular Fractionation and Immunoisolation Claudia Schirra, Nadia Alawar, Ute Becherer, and Hsin-Fang Chang Abstract Subcellular fractionation is an important tool used to separate intracellular organelles, structures or proteins. Here, we describe a stepwise protocol to isolate two types of lytic granules, multicore (MCG), and single core (SCG), from primary murine CTLs. We used cell disruption by nitrogen cavitation followed by separation of organelles via discontinuous sucrose density gradient centrifugation. Immunoisolation with a Synaptobrevin 2 antibody attached to magnetic beads was then used to harvest Synaptobrevin 2 positive granules for immunoblotting, mass spectrometry, electron, and light microscopy. Key words Lytic granule, Subcellular fractionation, Sucrose density gradient centrifugation, Immunoisolation, Magnetic beads, Nitrogen cavitation, Mass spectrometry

1

Introduction Cytotoxic granules (CGs) are specialized secretory lysosomes of cytotoxic T lymphocytes (CTL) and NK cells [1–5]. Over decades, cell biologists and immunologists have tried to isolate these secretory lysosomes and their luminally active components such as Granzymes and Perforin, to characterize the granules and understand their role in T cell function [6–11]. The presence of a dense core was demonstrated using electron microscopy [1, 12]. The protein composition of isolated secretory lysosomes was first reported in human NK cells [10] and later on in human CTLs [8, 9]. Methods for granule isolation differ, with various protocols that result in uneven purity and granule subpopulations, although the typical lysosomal markers such as LAMP-1, CD63, Cathepsin variants, and Granzymes [8–10] are present. Electron microscopic analysis revealed the heterogeneity of secretory lysosomes, suggesting subclasses of cytotoxic granules though they all contain cyto-

Cosima T. Baldari and Michael L. Dustin (eds.), The Immune Synapse: Methods and Protocols, Methods in Molecular Biology, vol. 2654, https://doi.org/10.1007/978-1-0716-3135-5_11, © The Author(s) 2023

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toxic molecules and are released at the synaptic cleft adjacent to target cells [8, 13]. Since Synaptobrevin 2 (Syb2) is a specific marker for CGs in mouse CTLs [14], we used Syb2-mRFP knock-in mice (Syb2 KI) for organelle purification and proteomics. CGs are endogenously labeled and can therefore be visualized by red fluorescence during the different isolation steps [15]. To avoid chemical and physical stress on enzymes or subcellular compartments and to preserve the integrity of the organelles, homogenization of the cultivated cells was done by nitrogen cavitation [16]. After several low-speed differential centrifugation steps to separate nuclei and intact cells, cellular fractionation was done by high-speed discontinuous sucrose density gradient centrifugation. We selectively chose the sucrose gradient composition (0.3 M, 0.8 M, 1.0 M, 1.2 M, 1.4 M, and 1.6 M), speed and time of ultracentrifugation were able to enrich Syb2-mRFP positive organelles in two fractions at the interface between 1.0 and 1.2 M (fraction 6) and 1.2 and 1.4 M sucrose (fraction 8) [15]. Sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) was performed directly on the collected gradient fractions. Western blots were then probed for markers specific for CG (Syb2, Perforin, Granzyme B), plasma membrane (Na+/K+-ATPase), mitochondria (SDHA), and cytoskeleton (β-Actin, α-Tubulin). Magnetic beads coupled to anti-Syb2 antibody were used for immunoisolation of organelles in fractions 6 and 8. After organelle binding, the magnetic beads can be immediately used for high-resolution microscopy, immunoblotting, electron microscopy, and mass spectrometry. Thus, with this isolation strategy combined with a variety of methods, we were able to characterize two populations of Syb2 organelles, SCG and MCG, the latter containing SMAPs [15, 17].

2

Materials All solutions were prepared with deionized water and analytical grade reagents. Prepare and store all reagents at room temperature unless indicated otherwise.

2.1 Primary Murine T Cells

1. 0.8–1.5 × 108 day 3–4 activated CD8+ T cells isolated from one Syb2 KI mouse (see Note 1). 2. BD FACSAria III analyzer (BD Biosciences) for flow cytometry analysis. 3. Cell surface-specific FITC-, APC-, or PE-conjugated antibodies against CD44, CD62L, and CD25 (BD Pharmingen or eBioscience).

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1. Wash buffer: D-PBS (137.9 mM sodium chloride, 1.47 mM potassium phosphate monobasic, 2.67 mM potassium chloride, 8.09 mM sodium phosphate dibasic) with 0.1% bovine serum albumin (BSA) and 2 mM ethylenediaminetetraacetic acid (EDTA), pH 7.4. 2. Protease inhibitors: 1 mg/mL Pepstatin A (in DMSO), 10 mM E64 (in water), and 400 mM 4-benzenesulfonyl fluoride hydrochloride (Pefabloc SC in water or equivalent). Store separately at -20 °C. 3. Sucrose solubilization buffer: 20 mM HEPES and 5 mM EDTA, pH 7.3 to dissolve sucrose. 4. Homogenization buffer: 300 mM sucrose, 20 mM HEPES, 5 mM EDTA, supplemented with protease inhibitors (2 mM Pefabloc SC, 10 μM E64 and 1 μg/mL Pepstatin A), pH 7.3 (see Note 2). Store at 4 °C. 5. Pre-chilled cell cavitation/disruption bomb (Parr 1019HC T304 SS/Parr Instrument Company) connected to a Nitrogen gas tank. 6. Micromagnetic stir bar. 7. 0.4% Trypan blue solution in 0.8X D-PBS or equivalent.

2.3 Subcellular Fractionation

1. Sucrose solutions for gradient: 0.8, 1.0, 1.2, 1.4, and 1.6 M sucrose in 20 mM HEPES with 5 mM EDTA and protease inhibitors (2 mM Pefabloc SC, 10 μM E64, and 1 μg/mL Pepstatin A), pH 7.3 (see Note 3). 2. Refractometer. 3. Beckmann ultracentrifuge with a SW40Ti rotor (or equivalent) and corresponding 14 mL transparent Ultra Clear centrifuge tubes (Beckmann 344,060, or equivalent). 4. 10 mL syringes and 20-G needles. 5. 1.5 mL low protein binding tubes and pipette tips. 6. Sample buffer for immunoblotting: Lithium dodecyl sulfate (LDS) sample-buffer (4X) and 1 M DTT. 7. Pierce™ 660 nm Protein Assay reagent (see Note 4).

2.4 Immunoisolation of MCG and SCG

1. Dynabeads™ Protein G magnetic beads (Invitrogen). 2. Dynabead wash 20 (pH 7.4).

buffers:

D-PBS

with

0.01%

Tween

3. Anti-Syb2 antibody (clone 69.1, Synaptic Systems or equivalent). 4. Dilution buffer: 320 mM KCl, 10 mM HEPES, 5 mM EDTA solution in distilled water (pH 7.3).

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5. Wash buffer: 160 mM KCl 10 mM HEPES, pH 7.3. 6. Rotator: Intelli mixer RM-2M (neoLab). 7. 1.5 mL low protein binding tubes and pipette tips. 8. Magnet to separate Dynabeads: DynaMag™-2 Magnet (Invitrogen). 9. Sample buffer for immunoblotting: Lithium dodecyl sulfate (LDS) sample-buffer (4X) and 1M DTT. 10. Confocal microscope (LSM 780, Zeiss) or equivalent.

3

Methods

3.1 Cell Homogenization

1. Collect CD8+ cells after expansion to 0.8–1.5 × 108 total CD8+ cells, adequate for granule isolation. 2. Test the quality of the cells and the activation state of the cells by flow cytometry before isolation of cytotoxic granules (see Note 5). 3. Wash the cells in cold wash buffer and resuspend them in 2 mL homogenization buffer. 4. Transfer the cell suspension into the clean and pre-chilled cavitation bomb (4 °C) and add the micro magnetic stir bar. 5. Close the Parr Bomb and place it on top of a magnetic stirrer and connect it to the nitrogen tank. 6. Slowly pressurize to 800 psi with nitrogen gas and stir at 4 °C for 25 min. Open the discharge valve gently and release the pressure to dropwise collect the homogenate in a pre-chilled 15 mL tube (see Note 6; Fig. 1a). 7. Mix 15 μL of the cavitate with trypan blue, add on a microscopy slide, cover with a coverslip, and observe by phasecontrast microscope to judge the effectivity of the cavitation procedure. 8. Centrifuge the cell lysate for 10 min at 1000 × g at 4 °C to pellet unbroken cells, partially disrupted cells, and nuclei. Collect the post nuclear supernatant (PNS, S1) without foam (Fig. 1a). 9. Wash the pellet/foam once in 0.5 mL of homogenization buffer and centrifuge for 10 min at 1000 × g and 4 °C. Add this 0.5 mL supernatant to the PNS from the previous step (~2 mL final volume) and use the pooled supernatant (S1 + S1’) for subcellular fractionation (see Note 7).

3.2 Subcellular Fractionation

1. Prepare 10 mL sucrose solutions of different concentrations (0.8, 1.0, 1.2, 1.4, and 1.6 M) from a 2 M sucrose stock solution in sucrose solubilization buffer and add fresh

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Fig. 1 Immunoisolation of two classes of cytotoxic granules from Syb2 KI mouse CTLs. (a) Flow chart for CG isolation procedure. (b) Confocal images of immune-isolated organelles of Syb2 KI mouse CTLs. Upper and lower panel represent immune-isolated organelles of fraction 6 (IP6) and fraction 8 (IP8), respectively. Scale bar, 1 μm. (c) Representative TEM images of immune-isolated organelles. Electron micrographs from IP6 (upper image) and IP8 (lower image) are shown. The majority of granules in IP6 contain multi-core granules (MCG), while the majority of granules in IP8 have a single core granule (SCG). Sale bar, 0.2 μm

proteinase inhibitors. Keep all solutions on ice and determine the refractive index for each solution using a refractometer (see Note 3). 2. Perform a discontinuous sucrose gradient column from bottom to top in a pre-chilled 14 mL ultra clear centrifuge tube by carefully layering 2 mL of each sucrose solution (see Note 8). 3. Carefully layer the 2 mL of the pooled post-nuclear supernatant (0.3 M sucrose) onto the performed gradient (Fig. 1a). 4. Ultracentrifuge the post-nuclear supernatant at 100,000 × g for 90 min at 4 °C in the SW40Ti rotor with acceleration/ deceleration of 8. 5. Slowly and carefully, transfer 12 × 1 mL aliquot from the top of the gradient into protein low binding 1.5 mL tubes. Avoid mixing the layers during this step. Use for each aliquot a new pipette tip (Fig. 1a).

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6. Keep the aliquots on ice and add again fresh protease inhibitors to avoid protein degradation (see Note 9). 7. Collect 61 μL of each fraction into a new 1.5 mL tube, add 16 μL LDS-buffer and 3 μL DTT, heat at 96 °C for 7 min, and store at -20 °C for SDS-PAGE and immunoblotting. 8. Additionally, collect 100 μL of each fraction to determine the protein concentration with Pierce™ 660 nm Protein Assay reagent for quantitative western blot (see Note 10). 3.3 Immunoisolation of MCG and SCG

1. Resuspend magnetic Dynabeads and transfer 15 μL per fraction to a 1.5 mL tube, place on magnet, and remove supernatant (see Note 11). 2. Wash beads for 20 min with 300 μL D-PBS with 0.01% Tween 20 in the rotator at 20 ± 2 °C (F1, 5 rpm). Repeat two times. 3. Coat 15 μL beads with 5 μg anti-Syb2 antibody in 200 μL D-PBS with 0.01% Tween 20 (pH 7.4). 4. Rotate 40 min at 20 ± 2 °C (F1, 5 rpm). 5. Wash beads three times using 400 μL D-PBS in the rotator (see Note 12). 6. During the wash time, prepare SCG and MCG from fractions 6 and 8 for immune-isolation. Dilute the fractions 6 and 8 in a 1:1 ratio with dilution buffer (see Note 13, Fig. 1a). 7. Add the diluted fractions to the Syb2 antibody-conjugated Dynabeads and incubate by slow rotation (overnight at 20 ± 2 °C, F1, 5 rpm) (see Note 14). 8. After overnight incubation, place the tube in the magnet and remove the supernatant. 9. Wash the beads by adding 300 μL of 160 mM KCl wash buffer and rotate slowly for 10 min. 10. Add the beads to the magnet and carefully remove the supernatant. 11. Repeat the washing steps five times. 12. Use a few microliter of Dynabeads and check the organelle binding efficiency by mRFP fluorescence (561 nm) with confocal microscopy (Fig. 1b). 13. Use organelle-Dynabead Protein G complexes for TEM and/or SEM analysis (Fig. 1c). 14. For mass spectrometry and western blot mix organelleDynabead Protein G complexes add 18 μL LDS-buffer and 2 μL DTT, heat at 96 °C for 10 min, and keep at -80 °C.

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Notes 1. We find it best to use 0.8–1.5 × 108 activated mouse CD8+ cells. For other cell types, the method has to be adapted. 2. We used an isotonic buffer for granule isolation to avoid osmotic stress and to keep the nuclei intact. 5 mM EDTA is included to chelate the cation to inhibit metalloproteases. Prepare the homogenization buffer with protease inhibitors fresh for each experiment. 3. Prepare a 2 M sucrose stock solution. Weigh 68.46 g sucrose into a glass beaker, fill to 90 mL with buffer (20 mM HEPES, 5 mM EDTA, pH 7.3) and stir with a magnetic stir bar. Once completely dissolved (~1 h), fill up to 100 mL. Store the solution at 4 °C until use. We find it best to prepare the stock solution one day before the subcellular fractionation. Always prepare the sucrose gradient solutions fresh on the day of the experiment and keep on ice. 4. Pierce™ 660 nm Protein Assay for total protein quantitation is compatible with a high concentration of most detergents, reducing agents and other commonly used reagents. The maximum compatible concentration for sucrose in these solutions is 50%, which allows determination of the protein concentration of different sucrose fractions after fractionation. 5. Only intact and healthy cells should be used for organelle isolation (>80%). We find it best to determine the viability of the cells based on their size and granularity. Cell surface-specific antibodies against CD25, CD44, and CD62 were used to determine the activation state of the cells. 6. The finale homogenate should appear milky with foam on top. The yield of the granules is mainly affected by the cavitation process. Try to avoid excessive cavitation and minimize foaming. 7. Repeat the centrifugation step until no pellet is visible. The final post nuclear supernatant should be free of nuclei and whole cells. 8. Start with the highest sucrose concentration (1.6 M) and then gradually add the sucrose solutions of lower concentration using a 10 mL syringe with 20xG needle. Submerge the centrifuge tubes with the sucrose gradient in ice until the homogenate is ready. Do not disturb tubes in any way to avoid mixing of the sucrose layers. 9. We find it best to continue immediately after fractionation with the immunoisolation.

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10. Additional aliquots of the fractions can be used for electron microscopy, confocal or structured illumination microscopy to characterize the different fractions and to examine the quality of the isolated organelles. We recommend centrifuging the organelles of the different fractions onto gelatin-coated coverslips for immunostaining and light microscopy. 11. Increase the amount of the magnetic beads depending on the number of fractions that should be analyzed. 12. We suggest to test the binding efficiency of the antibody by western blot. 13. The 1:1 dilution with the 320 mM KCl buffer reduces the sucrose concentration and the viscosity of the solution for better organelle binding. The final 160 mM intracellular KCl concentration stabilizes the organelles. 14. We had less organelle binding efficiency at 4 °C.

Acknowledgments This work was supported by grants from the Deutsche Forschungsgemeinschaft SFB 894 (E.K., U.B., and J.R). Further, this project has received funding from the European Research Council (ERC) under the European Union’s Horizon 2020 research and innovation program (grant agreement No 951329 to J.R.). We thank Ulrike Hahn, Keerthana Ravichandran and Momchil Ninov for their participation in the development of these methods. We thank Margarete Klose, Anja Bergstr€aßer, Nicole Rothgerber, and Anne Weinland for excellent technical assistance. References 1. Peters PJ, Borst J, Oorschot V, Fukuda M, Krahenbuhl O, Tschopp J, Slot JW, Geuze HJ (1991) Cytotoxic T lymphocyte granules are secretory lysosomes, containing both perforin and granzymes. J Exp Med 173(5): 1099–1109. https://doi.org/10.1084/jem. 173.5.1099 2. Page LJ, Darmon AJ, Uellner R, Griffiths GM (1998) L is for lytic granules: lysosomes that kill. Biochim Biophys Acta 1401(2):146–156. https://doi.org/10.1016/s0167-4889(97) 00138-9 3. Griffiths GM, Argon Y (1995) Structure and biogenesis of lytic granules. Curr Top Microbiol Immunol 198:39–58. https://doi.org/ 10.1007/978-3-642-79414-8_3 4. Clark R, Griffiths GM (2003) Lytic granules, secretory lysosomes and disease. Curr Opin

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Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made. The images or other third party material in this chapter are included in the chapter’s Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter’s Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

Chapter 12 Microvillar Cartography: A Super-Resolution Single-Molecule Imaging Method to Map the Positions of Membrane Proteins with Respect to Cellular Surface Topography Shirsendu Ghosh, Andres Alcover, and Gilad Haran Abstract We describe microvillar cartography (MC), a method to map proteins on cellular surfaces with respect to the membrane topography. The surfaces of many cells are not smooth, but are rather covered with various protrusions such as microvilli. These protrusions may play key roles in multiple cellular functions, due to their ability to control the distribution of specific protein assemblies on the cell surface. Thus, for example, we have shown that the T-cell receptor and several of its proximal signaling proteins reside on microvilli, while others are excluded from these projections. These results have indicated that microvilli can function as key signaling hubs for the initiation of the immune response. MC has facilitated our observations of particular surface proteins and their specialized distribution on microvillar and non-microvillar compartments. MC combines membrane topography imaging, using variable-angle total internal microscopy, with stochastic localization nanoscopy, which generates deep sub-diffraction maps of protein distribution. Since the method is based on light microscopy, it avoids some of the pitfalls inherent to electron-microscopybased techniques, such as dehydration, the need for carbon coating, and immunogold clustering, and is amenable to future developments involving, for example, live-cell imaging. This protocol details the procedures we developed for MC, which can be readily adopted to study a broad range of cell-surface molecules and dissect their distribution within distinct surface assemblies under multiple cell activation states. Key words T-cell microvilli, T-cell receptor, Cell-surface topography, Super-resolution microscopy, Variable-angle total internal reflection microscopy

1

Introduction The surfaces of eukaryotic cells are usually rough and may carry multiple protrusions of various types. Some immune cells are covered with protrusions throughout their surface [1, 2]. In particular, neutrophils and lymphocytes, including B and T cells and natural killer cells, are covered with microvilli, which are slender and

Cosima T. Baldari and Michael L. Dustin (eds.), The Immune Synapse: Methods and Protocols, Methods in Molecular Biology, vol. 2654, https://doi.org/10.1007/978-1-0716-3135-5_12, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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flexible actin-rich protrusions, with a typical diameter of 100 nm and a length of a few hundred nanometers [2, 3]. These membrane protrusions do not only create a structural scaffold for the formation of microclusters of specialized receptors and adhesion molecules but also serve as a physical barrier for the diffusion of non-microvillar molecules [4]. Microvilli should be distinguished from other dynamic protrusions that are de novo generated by motile cells. These include thin, sheet-like actin-rich protrusions at the cell’s leading edge called lamellipodia, which play a key role in cellular migration and surface sensing [5, 6]. Additional projections include filopodia, which are critical for cell sensing of cues and directionality of migration [7]. As opposed to these de novo generated protrusions, microvilli are pre-existing structures. Their role in the physiology of lymphocytes has been debated over many years. Lymphocytes travel in blood vessels, and they have to attach to endothelial cells of blood vessel walls before they penetrate into target tissues. It has been shown that adhesion molecules like L-selectin and P-selectin glycoprotein ligand-1 are concentrated on microvilli [8], and therefore, these cellular organelles have been considered as essential for the initial contact between blood-borne lymphocytes and the endothelial surfaces they interact with [8]. A different role for microvilli has been recently suggested following observations that lymphocytes use their microvilli to search for antigenic signals presented by rare antigen-presenting cells (APCs) [3, 9, 10]. Moreover, microvilli have been shown to be involved in force-driven penetration of the dense glycocalyx barrier of APCs, which may otherwise hinder ligand/receptor interaction [11]. This novel understanding was partly based on application of our microvillar cartography (MC) microscopy method (to be described in this protocol), which demonstrated that T-cell receptors (TCRs) are almost exclusively positioned on microvilli in T cells prior to their encounter of antigenic signals [9]. While our experiments were conducted on fixed cells, experiments by Krummel and coworkers [3] using light-sheet microscopy and by Sherman and coworkers [12] using a combination of atomic force and superresolution microscopies demonstrated the dynamic nature of microvillar involvement in contact formation with APCs. Using the MC approach, we further observed that, in addition to TCRs, multiple membrane proteins that are involved in the initiation of the immune response are also enriched on microvilli [10]. We also showed that TCR localization on microvilli is mediated by ERM (ezrin, radixin, and moesin) proteins, which connect membrane proteins to the cortical actin cytoskeleton [10]. Our work has thus established the role of microvilli as specialized activation hubs of T cells. Interestingly, the MC method was also used recently to reflect on the original role attributed to microvilli, that is, the formation of initial contact with endothelial cells

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[13]. Indeed, it was found that the G-protein-coupled receptor CCR7, which is essential for rapid recognition of chemokine signals on endothelial cells, is strongly clustered on the tips of T-cell microvilli, whereas L-selectin is more randomly distributed on these projections. CCR7 likely mediates inside-out signaling to a small subset of LFA-1 integrin molecules that are also positioned on microvilli, thereby allowing sub-second activation and arrest [13]. MC is a novel strategy for mapping the location of membrane molecules with respect to cellular surface topography. It is based on synergistic imaging by variable-angle total internal reflection microscopy (VA-TIRFM) to determine the 3D topography of the cell surface and stochastic localization nanoscopy (SLN) to position of membrane molecules with respect to this topography. A schematic of the MC procedure is shown in the Fig. 1. In this protocol, we describe in detail the implementation of MC, from cell preparation and staining through imaging studies and statistical data analysis. While MC is general and can be applied to any cell, we describe it here based on our studies of T-cell microvilli. The experiment starts with the labeling of specific membrane proteins using fluorescent antibodies, which is done at low temperature to prevent potential cell activation or clustering (Fig. 2A–C) [14]. To increase the resolution, we refrain from using secondary antibodies but rather employ either commercially available or in-house labeled primary antibodies. The cells are then fixed and stained with a membrane dye such as FM143fx (Fig. 2D, E). Importantly, if the antigenic epitope for antibody labeling of the membrane protein of interest is exposed outside the cell membrane, then labeling with antibodies is carried out before fixation (Fig. 2). This is due to the fact that the fixation procedure sometimes leads to partial masking of antigenic epitopes [14]. For detection of intracellular proteins, fixation and permeabilization are performed prior to labeling (Fig. 3). VA-TIRFM [15–18] operates by varying the penetration depth of an evanescent electromagnetic field into a sample, which modulates the illumination pattern of different sample features. The penetration depth of the TIRF illumination (d(θ)) is defined as the distance where the intensity of the evanescent field is 1/e of the incident laser intensity. The simplest way to tune d(θ) is to change the angle of incidence, θ. The distance (δz) of each point in the image from the glass surface can be calculated from the relative intensity of fluorescence at that point, if the value of d(θ) is known. Then, a 3D image of the fluorescently labeled cell membrane can be reconstructed from the set of δz values, leading to a topographical map of the T-cell surface, including the microvillar structure and distribution. To increase the statistical precision of the final topographical membrane map, information from images

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Fig. 1 Schematic of microvillar cartography (MC)

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Fig. 2 Flowchart of T-cell labeling when the membrane protein of interest is exposed outside the cell

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Fig. 3 Flowchart of T-cell labeling when the membrane protein of interest is exposed only within the cell

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obtained at different values of θ is combined. Image analysis, which we carry out in Matlab, leads to what we call “LocTips maps,” namely, detailed maps of the location of individual microvilli and of cell-body areas. The central microvillar region is defined as the region within 20 nm in the z-direction from the pixel with the minimum δz value. This analysis is independent of the diffractionlimited resolution in the lateral (x-y) plane. It rather depends on the very high resolution in the vertical (z) direction of TIRF images. To localize single-membrane protein molecules with respect to the cell-membrane topography with a sub-diffraction resolution [19, 20], we use stochastic localization nanoscopy (SLN). This super-resolution technique relies on the blinking of individual molecules to obtain images with sparse signals that can be localized accurately (Fig. 1). We implemented a simple approach, named dual-plane SLN (Fig. S3 of Ref. [9]), which permits us to differentiate between molecules that are localized on two different z-planes. In this process, we employ a piezo-stage to vertically shift the sample position in the z-direction, and we repeat the SLN imaging on two planes. This eventually leads to the accumulation of signals from blinking molecules, which are situated at distinct z-planes. To achieve depth-based sectioning, the out-offocus signals of single molecules are rejected at the time of data analysis. We superimpose images from VA-TIRFM and SLN, to generate the 3D topographical localization map of membrane proteins with respect to the T-cell microvilli (Fig. 1). We then find the ratio of relative localizations of membrane proteins of interest on cell body and microvillar areas. We also plot the distribution of membrane protein molecules with respect to the distance from central microvillar region. Finally, from simultaneous SLN experiments with two different labeled membrane proteins we can compute the co-localization probability (CP), which allows us to identify pairs of proteins that, due to their interaction, are situated (on the average) closer to each other than expected based on the cellular topography. Here, some caution is in place, because two proteins located on a relatively small object like a microvillus might appear to co-localize even if they do not interact. A careful comparison to pairs of proteins that are not known to interact facilitates this kind of analysis.

2 2.1

Materials Labeling

1. Dulbecco’s phosphate-buffered saline without calcium and magnesium (PBS-/-). 2. Hanks’ balanced salt solution without magnesium or calcium (HBSS-/-).

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3. 0.5 M Ethylenediaminetetraacetic acid (EDTA) aqueous solution, pH 8. 4. 16% Paraformaldehyde aqueous solution, EM grade. 5. 25% Glutaraldehyde aqueous solution, EM grade. 6. FM143fx dye (Invitrogen or equivalent, see Note 1). 2.2

Imaging

1. 0.01% Poly-L-lysine (PLL) in PBS-/-. 2. Glass bottom culture dish (e.g., MatTek, 35 mm petri dish, 14 mm micro-well, No. 1.0 cover-glass, part no. P35G1.0-14C). 3. Fiducial markers, such as fluorescent nanodiamonds.

2.3

Software

1. MATLAB R2020a and onward. 2. OriginPro 2016.

2.4 Buffer Preparation

1. 1 M potassium chloride: Dissolve 74.55 g potassium chloride in 1 L MILLI-Q water. 2. 4 M sodium hydroxide solution: Dissolve 80 g sodium hydroxide in 500 mL of MILLI-Q water. 3. 1 M sodium hydroxide solution: Dissolve 20 g sodium hydroxide in 500 mL of MILLI-Q water. 4. 1 M Trizma® hydrochloride (pH 7.5–7.7): Dissolve 157.60 g Trizma® hydrochloride in 800 mL MILLI-Q water. Adjust pH to 7.5–7.7 with the appropriate volume of 4 M sodium hydroxide. Bring final volume to 1 L with MILLI-Q water. Filter through a 0.22 μm filter and store at room temperature. 5. 1 M Ethylene Glycol-Bis[β-Aminoether]N,N,′N′,N-TetraAcetic Acid (EGTA) Solution: For a 100 mM EGTA stock solution, add 3.8 g to about 20 mL of distilled H2O and bring to pH 11 with 4 M sodium hydroxide and dissolve and then bring to pH 8.0 with hydrochloric acid and add H2O to a final volume of 100 mL. Filter through a 0.22 μM filter. 6. 40% glucose (w/v): Warm (~60 °C) 50 mL of PBS-/- and then add 40 g of glucose. Stir/shake it to dissolve. Bring the final volume to 100 mL with PBS-/-. Filter through a 0.22 μm filter (MILLEXGV) and store at 4 °C (after sealing the container properly with parafilm). 7. Cysteamine solution: Prepare 100 mM cysteamine in 125 mM Trizma® hydrochloride (pH 7.5–7.7). Should be prepared freshly. 8. Blocking solution: PBS-/-, 1% bovine serum albumin (BSA), 5 mM EDTA, 0.05% sodium azide.

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9. Fixation solution: 4% paraformaldehyde, 0.4% glutaraldehyde, 10 mM EGTA (ethylene glycol-bis(β-aminoethyl ether)-N,N, N′,N′-tetra acetic acid), 1 mM EDTA, 2% sucrose in PBS-/-. Should be freshly prepared (see Note 2). 10. Permeabilization buffer: 0.05% saponin+1% BSA in PBS-/-. Should be freshly prepared. 11. TP 50 buffer: Add 100 μL 1 M trizma hydrochloride (pH 7–7.7), 100 μL 1 M potassium chloride, 1 mL glycerol and 800 μL MILLI-Q water and then filter it through 0.22 μm filter (Millex-GV) and store at -20 °C. 12. 50 mg/mL glucose oxidase: Dissolve 50 mg glucose oxidase (from Aspergillus niger) in TP 50 buffer and store at -20 °C. 13. 40 mg/mL catalase: Dissolve 40 mg catalase (from bovine liver) in TP 50 buffer and store at -20 °C. 14. Blinking Buffer: Add 500 μL of 100 mM cysteamine (it should be freshly prepared), 250 μL of 40% glucose, 10 μL of 50 mg/ mL glucose oxidase, 1 μL of 40 mg/mL catalase and 239 μL of 125 mM trizma hydrochloride (pH 7.5–7.7). Should be freshly prepared. 15. Antibody labeling with Alexa Fluor-647/ Alexa Fluor-568: If Alexa Fluor-647/ Alexa Fluor-568 tagged primary antibody is not available commercially, then low endotoxin, azide-free (LEAF) purified commercial unlabeled antibodies may be labeled with Alex-647 in house, using the following protocol (see Note 3). The NHS ester of Alexa Fluor-647/ Alexa Fluor568 is reacted with unlabeled antibody molecules in PBS buffer in a 10:1 molar ratio in the presence of 0.1 M sodium bicarbonate buffer for 1 h at room temperature in the dark. Micro Bio-Spin column with Bio-Gel P-30 is used to remove the unlabeled dye molecules. 2.5

Microscope

1. Super-resolution microscope: We perform the microscopy experiments using a home-built TIRF-based setup. A detailed description of this setup follows (Fig. 4). However, any TIRF microscope that allows varying the excitation angle may be used, in principle. In our microscope, we combine two different laser lines, at 532 nm (50 mW; Cobolt) and 642 nm (150 mW; Toptica), into a polarization-maintaining singlemode fiber. For that, we use a series of dichroic beam splitters (z408bcm, z532bcm; Chroma) and an objective lens (M-20×; N.A., 0.4; Newport). The setup also contains a 405 nm laser, which we keep in our schematic (Fig. 4), though we do not directly refer to it in this protocol. To separately modulate each excitation beam, we connect the optical fiber into an acoustooptic tunable filter (AOTFFnc-VIS-TN-FI; AA OptoElectronic). To maximize coupling efficiency, we use a polarizer

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Fig. 4 Schematic of the microscope setup. See text for details. Abbreviations used: λ/2, half-wave plate, M mirror, DM dichroic mirror, L lenses, λ/4 quarter-wave plate, NF notch filter, EM emission filter

(GT10-A; Thorlabs) for each laser to tune its polarization. Free-space combination of the two laser lines is also possible. A free-space isolator is used to block backscattering into the laser and is followed by a half-wave plate to rectify the polarization rotation caused by the isolator. The power of each laser is controlled by the computer. To expand and collimate the firstorder output beam from the AOTF to a diameter of 6 mm, we use achromatic lenses (01LAO773, 01LAO779, CVI Melles Griot). An achromatic focusing lens ( f = 500 mm; LAO801; CVI Melles Griot) is used to focus the expanded laser beams at the back focal plane of the microscope objective (UAPON 100 × OTIRF; N.A., 1.49; Olympus). We shift the position of the focused beam from the center of the objective to its edge to achieve total internal reflection at the sample. To separate excitation beams from the fluorescence light, we use a multipleedge dichroic beam splitter (Di03-R405/488/532/635t1–25x36; Semrock) in the emission path of the sample, and then the fluorescence is coupled out from the side port of the microscope (Olympus IX71). To block the residual scattered laser light that passes through the dichroic beam splitter, we use notch filters (NF01-405/488/532/635 StopLine Quad-

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notch filter and ZET635NF; Semrock). We also use a selective emission filter (z488-532-647 m; Chroma) within the light path. We further employ a dichroic beam splitter (640dcrx 228869; Chroma) to split the fluorescent image into two parts and then project them onto two areas of a single electron-multiplying charge-coupled device (EMCCD) camera chip (iXonEM +897 back-illuminated; Andor). The upper half of the EMCCD chip is dedicated to record emission below 640 nm, which we refer to as the green channel. The bottom half records emission above 640 nm, which we refer to as the red channel. A single lens ( f = 150 mm; 01LAO551; CVI Melles Griot) is used to refocus each spectrally separated image. The EMCCD camera has a final magnification of 240×, which translates to a pixel size of 66.67 nm.

3

Methods

3.1 Labeling of the Cell Membrane and Membrane Proteins (Timing: 1 Day)

1. Treat the micro-well of the MakTek dish with 400 μL of 1 M sodium hydroxide solution for 40 min. Wash it ten times with MILLI-Q water and five times with PBS-/- and coat with PLL (100 μL of PLL are added for 30 min). Then, wash the well three times with PBS-/-. Note that for experiments with two membrane proteins (in which the membrane channel is not available, see below), add 100 μL of a very diluted fiducialmarker solution to the micro-well of the PLL coated MakTek dish for 15 min and then wash twice with PBS-/-. Coating with fiducial markers is not required for the standard MC procedure. Place 400 μL of PBS-/- within the well to keep it wet until loading the sample. Remove the PBS-/- just before use. 2. Harvest three million T cells from human blood or from a cell culture (e.g., Jurkat cells) and dilute with a 1:1 10 mM EDTA/ PBS-/- solution (so that the final concentration of EDTA in the solution is 5 mM) (see Note 4). 3. Spin the solution at 250 G (~1400 rpm) for 5 min and discard the supernatant. 4. Add 200 μL of the blocking solution to dissolve the pellet (see Note 5). Incubate the solution on ice for 10 min. 5. If the membrane protein of interest has the epitope for antibody labeling exposed outside the plasma membrane, then step 6 should be followed. Otherwise, go directly to step 7. 6. Add Alexa Fluor 647 tagged antibody to the solution such that the final concentration of the antibody is 10–20 μg/mL and incubate on ice for 20–40 min, depending on the antibody (see Note 6).

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7. Add 0.5 mL 5 mM EDTA/ PBS-/- and spin the solution at 300 G, 5 min, 4 °C and then discard the supernatant (if the initial pellet size is small, then a smaller volume of the 5 mM EDTA/PBS-/- solution may be used). 8. Repeat step 7 once again. 9. Add 600 μL of fixation solution to dissolve the pellet and incubate the solution on ice for 2 h (see Note 7). 10. Spin the solution at 800 G for 5 min, 4 °C, and discard the supernatant. 11. Add 0.5 mL PBS-/- to dissolve the pellet, spin the solution at 800 G for 5 min, 4 °C, and discard the supernatant (if the initial pellet size is small, then a smaller volume of the PBS-/solution may be used). 12. Repeat step 11 once again. 13. Add 100 μL of 5 μg/mL FM143fx membrane staining dye solution in HBSS-/- and incubate on ice for 30 min (see Note 8). 14. Add 0.5 mL PBS-/-, spin it at 800 G for 5 min, 4 °C, and discard the supernatant. 15. Add 600 μL fixation solution to dissolve the pellet and incubate it on ice for 30 min. 16. Spin the solution at 800 G for 5 min, 4 °C, and discard the supernatant. 17. Add 0.5 mL PBS-/- to dissolve the pellet, spin it at 800 G for 5 min, 4 °C, and discard the supernatant. (If the initial pellet size is small, then a smaller volume of the PBS-/- solution may be used.) 18. Repeat step 17 once again. Then, suspend the cells in 50 μL PBS-/- buffer and store at 4 °C. Do not freeze the sample. If the membrane protein of interest has the epitope for antibody labeling buried inside the cell membrane, then do not suspend the cells in PBS-/- buffer and store at 4 °C after the washing step but rather follow steps 19–24 (see Note 9). 19. Add 200 μL of permeabilization buffer. 20. Add Alexa Fluor 647 tagged antibody to the solution such that the final concentration of the antibody is 10–20 μg/mL. 21. Incubate the solution at 4 °C for 4 h to overnight, depending on the antibody (see Note 10). 22. Add 0.5 mL PBS-/-, spin at 800 G, 5 min, 4 °C, and discard the supernatant. (If the initial pellet size is small, then a smaller volume of the PBS-/- solution may be used.) 23. Repeat step 22 once again. 24. The cells are suspended in 50 μL PBS-/- buffer and stored at 4 °C. Do not freeze the sample.

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3.2 Labeling of Two Different Membrane Proteins for Colocalization Probability (CP) Analysis (Timing: 1 Day)

3.3 Sample Preparation for Microscopy

3.4 Imaging (Timing ~1 h for Imaging of One Cell) 3.4.1 Variable AngleTotal Internal Reflection Microscopy

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We recommend following a similar protocol as stated above for the labeling of two different membrane proteins, skipping the membrane staining step. Labeling of two different membrane proteins should be done sequentially. We find it convenient to label one of the membrane proteins with alexa-647 tagged antibodies and the other membrane protein with alexa-568 tagged antibodies. If one of the two membrane proteins has an epitope for antibody labeling exposed outside and the other membrane protein has the epitope within the cytoplasm, then labeling may be done in the following way: First, follow steps 1–12. Then directly go to step 19 and follow steps 19–24. However, in step 20, use Alexa Fluor 568 tagged antibodies instead of Alexa Fluor 647 tagged antibodies. Finally, as noted in the analysis section, an important control for CP analysis involves a pair of membrane proteins that do not interact with each other, and one should be ready to label and study such a pair. 1. On the day of imaging, 10 μL of labeled cell solution is mixed in 390 μL of the blinking buffer (see Note 11). 2. Place the solution of labelled cells in blinking buffer into the micro-well of the PLL coated MakTek dish. Imaging should be started after the cells settle down on the glass surface (which happens within ~10 min) (see Note 12). If necessary, for example, for the CP procedure, include fiducial markers when coating with PLL (see discussion below). (see Note 13)

1. Turn on the visible light lamp of the microscope. 2. Select a single cell on the micro-well of the MakTek dish using the eyepiece of the microscope. 3. Capture the bright-field image of the cell. 4. Turn off the visible light lamp of the microscope. 5. Turn on the 532 nm (green) laser, such that the excitation power at the sample will be 10–20 μW (corresponding to 6–12 W/cm2). Set the angle of incidence of the light to 66.8° (see Note 14). 6. Focus the laser light at glass surface. Next, change the angle of incidence of the light to 63°. The focus should not be readjusted when changing the angle of incidence (see Note 15). 7. Record the membrane image movie of the cell using the EMCCD camera (typical camera settings: Exposure time 0.1 s, number of frames 50, camera frame area: 256 pixel × 512 pixel for each channel).

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8. Repeat step 7 for angles of incidence 64.2°, 65.5°, 66.8°, 68.2°, 69.7°, 71.2°, and 73.0°. Critical Step: The focus should not be readjusted between movies. 3.4.2 Stochastic Localization Nanoscopy (SLN)

1. Set the angle of incidence of the light again at 66.8°. 2. Record the membrane image only in the green channel (typical camera settings: Exposure time 0.1 s, number of frames 50, camera frame area: 256 pixel × 256 pixel in the green channel). 3. Block the 532 nm laser with the shutter. 4. Turn on/unblock the 642 nm (red) laser with high power (such that the excitation power will be 60–70 mW, corresponding to 36–44 kW/cm2 at the sample). 5. Record the SLN movie of labelled receptor molecules in the red channel (Camera settings: Exposure time 0.015 s, number of frames 3000, camera frame area: 256 pixel × 256 pixel in the red channel) (see Note 16). 6. Block the 642 nm laser with the shutter. 7. Unblock the 532 nm laser to illuminate the sample with the 532 nm laser light (see Note 17). 8. Repeat steps 2–7 ten times to acquire a total 30,000 frames of SLN images of receptors. 9. Use the piezo stage to shift the imaging plane 400 nm away from the surface of the glass (see Note 18). 10. Record the membrane image only in the green channel (camera settings: exposure time 0.1 s, number of frames 50, camera arame area: 256 pixel × 256 pixel in the green channel). 11. Block the 532 nm laser with the shutter. 12. Unblock the 642 nm (red) laser with high power (such that the excitation power at the sample will be 60–70 mW, corresponding to 36–44 kW/cm2). 13. Record the SLN image movie of labelled receptor molecules only in the red channel (camera setting: exposure time 0.015 s, number of frames 3000, camera frame area: 256 pixel × 256 pixel in the red channel). 14. Block the 642 nm laser with the shutter. 15. Unblock the 532 nm laser to illuminate the sample. 16. Repeat steps 10–15 ten times to acquire a total of 30,000 frames of SLN images of receptors. 17. Bring back the imaging plane to the glass surface using the piezo stage.

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18. Record the membrane image again in the green channel only (camera settings: exposure time 0.1 s, number of frames 50, camera frame area: 256 pixel × 256 pixel in the green channel). For Dual-Color SLN Imaging Only 19. Set the angle of incidence of the light at 66.8°. 20. Choose a cell residing in a region in which you can observe at least one fiducial marker. 21. Block the 532 nm laser with the shutter. 22. Turn on/unblock the 642 nm (red) laser with high power (such that the excitation power will be 60–70 mW, corresponding to 36–44 kW/cm2 at the sample) (see Note 19). 23. Record the SLN movie of labeled receptor molecules in the red channel (camera setting: exposure time 0.015 s, number of frames 4000, camera frame area: 256 pixel × 256 pixel in the red channel). 24. Repeat step 23, seven times to acquire a total of 28,000 frames of SLN images of Alexa Fluro-647 tagged receptors. 25. Block the 642 nm laser with the shutter. 26. Turn on/unblock the 532 nm (red) laser with high power (such that the excitation will be 20–30 mW (corresponding to
12–18 kW/cm2 at the sample). 27. Record the SLN movie of labelled membrane molecules in the green channel (camera settings: exposure time 0.015 s, number of frames 4000, camera frame area: 256 pixel × 256 pixel in the green channel). 28. Repeat step 27, seven times to acquire total 28,000 frames of SLN images of Alexa Fluro-568 tagged receptors (see Note 20). 3.4.3 Analysis (Timing 15–30 min)

(see Note 21) 1. Analyze the SLN movies captured with the Alexa Fluor-647tagged membrane proteins to obtain localization of individual receptor molecules with sub-pixel resolution (see Note 22). 2. Analyze the VA-TIRFM movie files to reconstruct the membrane topography (see Note 23). 3. Correct results for sample drift during the recording of series of the SLN movies (see Note 24). 4. Merge positions of membrane proteins mapped by SLN imaging with the 3D topography of the cell membrane, reconstructed from a VA-TIRFM image after correcting for registration shift and the effect of sample drift (see Note 25).

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5. Determine the distribution of membrane protein with respect to microvilli and cell-body regions. Find out the cumulative fractional increase of the number of molecules of a specific protein on each cell as a function of the distance from the central region of each microvillus (see Note 26). 6. If dual color super-resolution experiments is performed, calculate the distance-dependent co-localization probability (CP) for the protein pair of interest and also for the control protein pair, as described in the experimental design section (see Note 27).

4

Notes 1. Keep in dark at -20 °C. 2. Use of glutaraldehyde is essential to retain the resting state topography of T cells during fixation. 3. The concentration of the antibody should be 1 mg/mL for optimal labeling. 4. This step is essential to keep the membrane topography of T-cells intact. A washing strategy similar to this one may be readily developed for other cell types. 5. To reduce nonspecific staining by antibodies, the cells are incubated in a blocking solution on ice. 6. If the antibody is known not to activate the cells or cluster receptors, then labeling can be done at 37 °C. If the membrane protein of interest has an antigenic epitope exposed outside the plasma membrane, this protein is labeled before cell fixation (Fig. 2c). This is essential since fixation may destroy the exposed antigenic epitope [14]. Further, a low temperature (4 °C) is adopted for the labeling process in order to prevent cellular activation or internalization induced by the labeling antibody [14]. This is particularly important when using antibodies that may affect biological function, such as anti-TCRαβ or anti-CD3, which activate T cells at 37 °C and induce receptor clustering and internalization. We usually verify that low-temperature labeling does not give rise to any artifact by comparing the distribution of two membrane proteins (e.g., L-selectin and CD45) labeled at low temperature as well as under physiological temperature (37 °C). We confirm that the distribution of those two membrane proteins with respect to the membrane topography does not depend on the labelling temperature. We also make sure that we do not observe any artifactual clustering of receptors in those cases. (See the section “Labeling at 4°C captures the bona fide resting state” in Ref. [10].) We recommend performing a similar control for any cell type to be labeled at 4 °C.

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If the number of labeled membrane proteins observed in imaging steps is very small, this might be due to under-labeling. In that case, increase the incubation time of the antibody. Check if the labeling efficiency increases at 37 °C without activating the cells. If the observed number of labeled proteins is still small, then it may be due to low density of membrane proteins on the cell membrane. If the observed number of labeled membrane proteins is suspiciously high, then the antibody used may be nonspecific. Check the specificity of the antibody using flow cytometry. If the antibody is tagged with Alexa Fluor-647 in-house, then check whether the unlabeled antibody is impurity- and azidefree. Try using an antibody from a different source. 7. The fixation procedure is optimized for Jurkat cells and may need to be varied for other cell types. We perform cell fixation with a specific fixation buffer (4% (wt/vol) paraformaldehyde, 0.2–0.5% glutaraldehyde, 2% (wt/vol) sucrose, 10 mM EGTA, and 1 mM EDTA, PBS), which was shown by scanning electron microscopy (SEM) [9, 10] to retain the original membrane topography. We observe that the use of glutaraldehyde (0.2–0.5%) during fixation is essential for this purpose. Fixing the cells in solution prevents changes of membrane topography due to interaction with surface immobilization agents such as poly L-lysine (PLL). Membrane topography and the microvillar density observed in VA-TIRF surface reconstructions of T-cells on PLL surfaces can be compared to scanning electron microscopy (SEM) images of similar cells. Using this approach, we have found that both SEM, which images the cells from the top with respect to the PLL surface, and VA-TIRFM, which images them from the bottom, demonstrated similar topography and microvillar density. (See Fig. 1 and S1–2 of Ref. [9]; Fig. 1 and S1 of Ref. [10].) 8. The membrane staining timing is important as a shorter incubation time leads to under-labeling of cell membrane, whereas a longer incubation time may lead to penetration of membrane dye into the cytoplasm of the cells, causing increased background in imaging. We recommend performing membrane staining following fixation whenever possible. To obtain an unbiased representation of the cellular membrane topography from the TIRFM images, the cell membrane needs to be stained homogenously. Nonhomogeneous staining of the cellular membrane would lead to inherent differences of the registered emission intensity of the dye in different regions of cell membrane. This may mask the intensity differences between dye molecules due to their axial positions, which would lead to errors in reconstruction of the 3D membrane topography. Thus, we choose to use the dye

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FM143fx, as it has been shown to stain cellular membranes homogeneously [21–23]. This can in fact be readily tested. For example, we treated FM143fx stained Jurkat T-cells such that they would spread over a glass surfaces and lose their microvillidominated topography. A homogeneous distribution of the intensity of FM143fx throughout the flattened cell membrane was registered in TIRFM images. (See section “Cellular labeling with antibodies” and Fig. S1V-W in Ref. [10].) 9. If the antigenic epitope of a specific membrane protein is within the cytoplasm, the cell membrane needs to be permeabilized for labeling. In this case, we prefer to invert the order introduced above and label membrane proteins following membrane staining. This order of labeling steps is selected to prevent penetration of FM143fx molecules into the cell, which may occur if permeabilization is done before membrane staining. Penetration of the membrane dye may lead to an enormous background signal during imaging. 10. There may be a chance of only a low level of labeling of membrane proteins whose antigenic epitope is exposed only within the cell. This may due to weak permeabilization. Try using 0.03% Triton x-100 in 1% BSA in PBS as the permeabilization buffer. If the problem persists, increase the incubation time of the antibody. 11. The blinking buffer should be freshly prepared. 12. If cells do not adhere to the glass surface properly, increase the volume of PLL solution used for PLL coating and also increase the incubation time with the PLL solution. 13. We provide a MATLAB code for the analysis of the results (see the section Code Availability below). If you plan to use this MATLAB code, you will need to make sure that files are saved in “.fits” format and that file names are given in the correct format, as explained in Tables S1 and S2 of the supplementary file “step by step guide to run analysis codes.pdf.” 14. Note that to determine the
angle  of incidence, the following equation is used, θ = sin - 1 FDη (Ref. [24]), in which θ is the 1 angle, D is the distance of the illumination beam from the center of the objective (which we measure by reading the scale of a linear stage used to translate the beam), F is the focal length of the objective lens (1.8 mm in our case), and the refractive index of glass and oil (η1) is 1.52. 15. In case the camera saturates with the emission from the membrane stain, decrease the excitation power of the 532 nm laser so that the camera does not saturate and restart the imaging from step 6 under 3.4.1 with the selected power.

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16. The red channel image may sometimes contain a large background signal due to presence of impurities in the blinking buffer. Re-prepare the blinking buffer with caution such that no impurity is present. The background may be also due to over-staining of cell membrane, which causes penetration of membrane dye inside cytoplasm of the cells. Repeat sample preparation with a shorter incubation time for membrane staining. If the 642 nm laser power at the time of imaging of the 0 nm plane is too high, it might cause photobleaching of most Alexa Fluor-647 dye molecules. This might lead to a too-low number of blinking events at the -400 nm plane. In that case, reduce the power of 642 nm laser when recording the 0 nm plane image. 17. To allow us to correct images for sample drift during data collection (see below), we collect membrane dye reference images (50 frames) between the SLN movies under weak illumination of the 532-nm laser (10–20 μW, corresponding to 6–12 W/cm2). The TIRFM images are recorded in the green (532 nm) channel of the EMCCD camera. 18. We use a piezo stage (PI nano Z-Piezo slide scanner; PI) to move the sample up or down and collect the SLN images at the 0 nm and -400 nm focal planes. For each focal plane, we collect 30,000 camera frames, divided into ten movies (3000 frames in each, 15 ms per frame). We achieve the sectioning effect of dual-plane SLN by rejecting the out-of-focus images of single molecules during the localization procedure (see the section “dual-plane SLN” in Note 22 below) [9]. The SLN imaging of membrane protein molecules at the -400 nm focal plane ensures that we are not missing any molecules due to the complex 3D topography of cell membrane. This allows us to rule out the possibility of biased detection of membrane proteins on the microvillar region compared to cell-body regions. Here, we need to clarify the ambiguity between the penetration depth and the detection limit of molecules based on their distance from the glass surface upon illumination by the evanescent field. The penetration depth of the evanescent field signifies the distance at which the field intensity is 1/e (~37%) of the incident light intensity. However, depending on the incident intensity, molecules can be detected much deeper than that distance. We have verified that the intensity of the evanescent field is high enough to detect molecules with equal probability at the 0 nm plane and the -400 nm plane (see details in Ref. [10], particularly the “TIRF setup” section and Figs. S1M–S1O).

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19. We prefer to measure with the 642-nm excitation first because we observe that Alexa-568 is more photo-stable, and reversal of order may cause photobleaching of a fraction of the Alexa647 dye due to direct excitation. This needs to be gauged for every pair of dyes used. 20. From simultaneous SLN experiments with two different labeled membrane proteins, we can compute the co-localization probability (CP), which allows us to identify pairs of proteins that, due to their interaction, are situated (on the average) closer to each other than expected based on the cellular topography. Here, some caution is in place, because two proteins located on a relatively small object like a microvillus might appear to co-localize even if they do not interact. A careful comparison to pairs of proteins that are not known to interact facilitates this kind of analysis. Thus, for using the CP method, one needs to also measure dual color SLN images for an appropriate control sample, following the same procedure as above (steps 19–28) under 3.4.2. 21. These analysis steps can be performed using our Matlab code, for which operation instructions are provided in the supplementary file “step by step guide to run analysis codes.pdf” (see the section Code Availability below). 22. Analysis of SLN movies to generate super-resolved maps of membrane proteins: Two common methods for localization of single molecules are 2D Gaussian fitting (2D-Fit) and center-of-mass (CM) analysis [20, 25, 26] based on a point spread function model. 2D-Fit provides better precision than CM analysis. However, the speed of 2D-Fit is much reduced due to the need for multiple iterations, especially while attempting to fit a “bad” molecule (i.e., an impurity, a cluster of multiple molecules, an out-of-focus molecule, etc.). On the other hand, CM analysis is less precise but much faster, since it requires a one-step calculation per molecule. We developed a simple algorithm, CFSTORM (Center of mass and 2d-Guassian Fit STORM, Fig. 5) that combines the two methods: individual emitters are identified in each frame by steps of thresholding. More specifically, a binary image is created from each frame by setting a threshold intensity level. Then the “bwlabel” function in MATLAB is used to segment the binary image to define individual emitters. Next, the data in a square of 11 × 11 pixels around the local maximum of each individual emitter is used in Eqs. 1 and 2 below for calculating the center of mass coordinates (cx,cy) and the size of the pattern (sx, sy) [26]. We use a fixed threshold for the size of molecules (sx, sy) in the CM analysis. If a molecule has a larger size than the threshold, it is discarded. We then perform 2D-Fit (Eq. 3) on those molecules that pass CM fitting and again discard bad molecules based on

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Fig. 5 Flowchart for detection of location of single molecules from SLN movies

standard deviation (SD) values obtained from the fit (σ x, σ y). The emitters that pass the 2D-Fit step are true single molecules whose sub-pixel localization is determined from the parameters of the fit with maximum precision. P P I ij x ij I ij x ij cx = P ; cy = P ð1Þ I ij I ij  P    P  c y I    ij y ij I ij c x - x ij P P sx = ; sx = ð2Þ I ij I ij 0 !2 1  2 y - y0 A x - x0 þ bdg ð3Þ þ f ðx, y Þ = amp × exp@2σ 2x 2σ 2y

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In these equations, the i, j indices are the x-axis, and y-axis indices for each pixel xi,j and yi,j are the coordinates of the pixel in nanometers, and Ii,j is the intensity of the pixel. The coefficient “amp” is an amplitude, “x0,” “y0” is the center position, and “bgd” is a background parameter. To reconstruct an image of all protein molecules in the cell membrane, we combine the locations of single emitters determined from all SLN movies (Fig. 5). Determination of localization precision and spatial resolution: To this end, we image fluorescent beads (~110 nm for our case) and adjust the intensity of laser in such a way that emission intensity of the beads matches the range of photon numbers emitted from a single blinking event of an Alexa Fluor 647 molecule. We measure the x and y coordinates and number of photons of each bead for several frames. Based on these measurements, we estimate that the uncertainty in average localization in x–y is ~11 nm. However, the localization precision is NOT the spatial resolution obtained in the final reconstructed image, as the latter also depends on the density of single-molecule detection events. To calculate the true image resolution, we suggest using the Fourier ring correlation analysis of images [27], which takes into account of both localization uncertainty and labeling density. Dual-Plane SLN: We achieve a sectioning effect at the two measured planes (0 nm and -400 nm) by discarding images of out-of-focus single molecules at the time of data analysis. The recorded pattern of an out-of-focus molecule has a lower overall signal intensity and a higher spatial width. We use these features to reject out-of-focus molecules. To assess the defocusing of single molecules, we use images of fluorescent beads (
110 nm in our case) coated on a glass surface. We employ the piezo-stage to vertically move the sample in the z-direction with a 100 nm step-size and image the fluorescent beads. The relative change of the SD of 2D Gaussian fits of individual fluorescent bead is determined in order to quantify the defocusing effect. We observe that the SD is substantially different when the images are acquired 400 nm away from glass surface (i.e., the difference between the glass plane and focal plane is 400 nm) than when acquired at the glass surface (Fig. S3 in Ref. [9]). This shows that one may selectively capture signals from “in-focus” molecules that are localized at either the 0 nm or the 400 nm plane by applying a suitable threshold on the SD. 23. Analysis of VA-TIRFM images of cell membrane to reconstruct the 3-D topography of cell membrane: As noted in the introduction, the intensity of fluorescence emission of a point on a uniformly stained object following excitation by an evanescent

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field depends on the distance of that point from the glass surface on which the object sits [15–18]. This property is used to reconstruct the 3D surface topography (Fig. 6A–E). Thus, we assume that the pixel that is nearest to the glass surface should have the maximum intensity of emission, dependent on the specific angle of incidence θ [Imax(θ)]. To compute the relative z-height from the glass (δz) of each pixel of the cellular membrane image [with intensity I(θ)], we use the following equation: δz = ln (I max (θ)/I(θ))/d(θ). The penetration depth of the TIRF illumination is represented by d(θ), which is calculated with the following equation,  λ  2 2  - 1=2 d ðθÞ = 4π , where λ represents the waven1 sin θ - n22 length (532 nm), n1 is the refractive index of the glass coverslip and immersion oil (1.52), and n2 is the refractive index of the buffer (1.35). We use a refractometer to determine n2 at 23 °C. The TIRF image taken at each angle of incidence leads to a separate topographical map. To define the regions of individual microvilli (MV) and the regions of the cell body (CB) on the topographical maps obtained with the above procedure, we use an algorithm implemented in MATLAB (see flowchart of the algorithm in Fig. 7). We describe below the detailed implementation, which is of course subject to modifications by each user. Initially, we create a two dimensional “Laplacian of a Gaussian” (LoG) filter (Matlab; image processing tool kit) with the following parameters: Gaussian σ = 0.5; kernel size = 10 × 10 (Fig. 6F) using the function “fspecial.” Then, we process the VA-TIRFM images with the created LoG filter using “imfilter” with the option “replicate.” This leads to flattening of cell body regions while keeping the tip positions of microvilli appearing as peaks. We then create binary images from the LoG-filter processed images by setting a threshold intensity level (we use a value of 5) to capture the most distinguishable microvilli regions. We now need to separate individual microvilli from larger microvilli-containing regions. For that, we perform an erosion operation on the binary image with proper morphological structural elements. To this end, we create a diamond shaped morphological structuring element using the “strel” function in MATLAB. We then erode the binary image created for the VA-TIRFM image taken at the minimum angle using the diamond-shaped morphological structuring element and the function “imerode.” To segment the eroded binary images to individual microvillar areas, we use the function “bwlabel.” The structures that appear to have a size greater than 10 pixels after the segmentation mostly represent the merging of two individual microvillar areas. To segment them, we first multiply the binary value of the structure with the corresponding values

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Fig. 6 The analysis procedure. (A–C) A series of 3D surface reconstruction maps of a Jurkat T cell from TIRFM images, based on calculated δz values from TIRFM measurements at angles of incidence (A) 65.5°, (C) 66.8°,

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from the image obtained at the maximum angle to create a new binary map and then try to segment them again with “bwlabel.” If the structure is not segmented, we create two orthogonal “line”-shaped morphological structuring elements using the “strel” function in MATLAB. We then erode the newly created binary map with the combination of these two elements using “imerode.” We try to segment the eroded binary image again with “bwlabel.” If it is still not separable, we repeat the above steps one more time. As a last step, in case we do not achieve segmentation yet, we create another diamond-shaped morphological element using the “strel” function and erode the eroded binary image of the object twice again. In our experience, after these four steps of erosion all the structures with size greater than 10 pixels can be segmented by “bwlabel.” In the above procedure, we intentionally segment the VA-TIRFM image recorded at the smallest angle of incidence first and then combine with information from the image taken at the largest angle of incidence to separate individual microvilli. This guarantees the inclusion of the maximum number of microvilli. After segmentation of all individual microvilli, we recover the eroded binary image using the function “imdilate.” Finally, we combine the segmented areas generated from the images obtained at angles 65.5°, 66.8°, and 68.2°, and the mean of the δz of each pixel is calculated and used to generate a map of the location of microvilli, which we call the “LocTips map” (Fig. 6H). The tip of each microvillus is defined as the pixel within that microvillus with the minimum δz value. We tested that our method is capable of identifying true microvilli. For that, we used localization of L-selectin, which is well known as a microvilli marker [8]. We showed that L-selectin ä Fig. 6 (continued) and (C) 68.2° under weak illumination of a 532-nm laser. The δz map is shown from a direction perpendicular to the y–z plane. (D) A representative 3D surface reconstruction map of a T cell, calculated as the mean of the δz values of B-D. The δz values are represented by different hues with a step size of 6.25 nm. (E) A bottom-to-top 2D projection of D. (F) An example of a TIRFM image (Left) and its Laplacian of Gaussian (LoG) filter image (Right). (G) The regions above a threshold value of 5 in LoG filter images (white), calculated from a series of images acquired at variable angles of incidence (indicated above each image). (H) A LocTips map. The segmentation map of distinguishable protruding areas is determined by combining a series of segmentation analyses of VA-TIRFM measurements at incident angles from 63° to 73°. The coordinates of the pixel of minimum δz in each individual microvillus are marked (black cross). (I–J) Positions of protein molecules obtained from SLN (white dots) at the 0 nm plane (I) and at the -400 nm plane (J) are superimposed on membrane topography maps obtained from VA-TIRFM. (K) Cumulative increase of the fraction of total molecules on the cell as a function of the distance from the central microvilli region, normalized by the cumulative increase in the fraction of area (δCount/δArea) as a function of distance from microvilli. (L) Percentage of molecules on the microvillar (MV) region and the cell-body (CB) region of the membrane

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Fig. 7 Flowchart for membrane topography analysis from VA-TIRFM Images

molecules are localized selectively on those areas of the T-cell membrane that are identified as microvilli by our analysis method (see Ref. [10], in the section “microvilli can be identified using L-selectin localization” and in Figs. S1J-L). We also compared the microvillar density on T cells detected using our method with that detected by SEM imaging and found them to be similar [9, 10].

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24. Determination of time-dependent sample drift: The SLN experiment involves recording of a large number of separate movies. There are multiple ways to minimize sample drift occurring at the time of imaging and correct for it during analysis. We adopted the following movie-by-movie procedure, which relies on the TIRFM membrane images recorded between two successive SLN movies. We calculate the 2D cross-correlation function of each such TIRFM image with the TIRFM image acquired at the end of the sequence. A lateral drift of the sample leads to a peak in the 2D cross-correlation image that is shifted from the origin in both the x and y directions. The position of this peak, calculated from a 2D Gaussian fit, can be used for drift correction. For dual-color SLN experiments, in which we do not have the membrane TIRF channel for drift correction, we use fiducial markers (e.g., nanodiamonds) and correct using a similar procedure as above. 25. Determination of channel shift: The two recorded SLN channels on the camera, green (532 nm) and red (647 nm), are typically slightly shifted from each other, and correction is required in order to superimpose them. This correction may involve both a shift in the x or y direction and a more complex distortion (e.g., a stretch or a rotation). A careful design and alignment of the emission image splitter (image-splitting device) can eliminate the need to perform any operation on the images other than x-y translation in order to match them. To perform the latter, we observe that a fraction of emission of the 532 nm channel leaks into the 647 nm channel when we record membrane images (the 532 nm laser is off when SLN images are recorded). Thus, we cross-correlate the leaked membrane image in the 647 nm channel with the original membrane image in 532 nm channel. The shift is estimated from a 2D cross-correlation image and is used to adjust the images accordingly. Our Matlab code (see the section Code Availability below) includes a procedure that corrects for both sample drift and channel shift self-consistently. 26. Analysis of the distribution of membrane proteins with respect to microvilli: To obtain the distribution of each studied membrane protein, we merge the super-resolved map of membrane protein obtained from SLN movies acquired in the red channel with the membrane topography obtained from the TIRFM image acquired in the green channel after correcting for the channel shift and membrane drift effect in the red channel. We segment the membrane of each cell into the microvillar (MV) area and the non-microvillar or cell-body (CB) area using the “LocTips map” procedure described above.

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We also calculate the “cumulative fractional increase” of the number of molecules of a specific protein on each cell as a function of the distance from the central region of each microvillus (Fig. 6K, see also Figs. S6E–S6G of Ref. [10]). To this end, we define the central region of each microvillus as the region that is not more than 20 nm from the microvillar tip, which is the pixel of the minimum δz value. The “boundary” function of MATLAB is used to encapsulate this central region (see Fig. S6F of Ref. [10]). This leads to a different shape of the central region of each microvillus, depending on the shape and orientation of the specific microvillus with respect to the surface. Concentric closed curves of a similar shape and increasing size (Fig. S6G of Ref. [10]) are plotted, and the number of molecules in each concentric sector that forms between two closed curves is calculated. From these values, we determine the cumulative fractional increase, which is then normalized by the cumulative fractional increase of the area, to obtain the δCount/δArea plot, where δCount represents the fraction of total counts recorded within a concentric sector at a particular distance from the central microvillar region, whereas δArea represents the fractional area of that sector. The more a protein is localized to the microvillar region, the steeper is the slope of the δCount/δArea plot as a function of distance from the microvillar central region. To decipher the distribution of a specific membrane protein in a specific cell type, we advise to image at least ten cells. The actual pattern of distribution of a membrane protein may only be recognized from the average percentage of membrane protein molecules localized in different regions of the cells and the δCount/δArea plot averaged over all cells. Finally, we calculate the percentage of molecules on MV regions and on CB regions of each cell (Fig. 6L, see also Figs. S6A–S6D of Ref. [10]). 27. Co-localization Probability (CP) analysis: As noted in the overview section, two proteins that are situated on the microvillar regions may appear to be spuriously localized together due to the tight topography. However, a careful comparison can differentiate between protein pairs that are truly interacting and those that appear to be close just due to the membrane topography. For a detailed analysis of the proximity of a pair of proteins, we developed the CP analysis. CP is the distancedependent probability of having a partner protein molecule within a specific distance from a protein molecule of the other type. Mathematically, the CP for molecules i and j within a distance R may be simply calculated by Nii(R)/Ni, where the total number of detected points of molecule i is represented by Ni, and Nij(R) represents the number of points of molecule i that have at least one point of molecule j within R. The CP for

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any protein pair increases with distance and eventually approaches 1 at a distance comparable to the size of microvilli. It is essential to compare the distance-dependent CP plot of a certain protein pair with that of a pair of proteins that are known not to interact with each other even, while they both reside on the same microvillus. From our experience, the CP of an interacting pair rises significantly “faster” with distance compared to the CP of a noninteracting pair. Other methods for co-localization may be found in the literature.

Acknowledgments We thank Drs. Sara W. Feigelson and Ronen Alon of the Weizmann Institute of Science, Israel, and Dr. Yunmin Jung of the Institute for Basic Science Center for Nanomedicine, Seoul, Korea, for their kind involvement in this project and for their advice. G.H. is the incumbent of the Hilda Pomeraniec Memorial Professorial Chair. Data Availability Sample datasets for “microvillar cartography” and “co-localization probability” analysis can be downloaded from the “BioImage archive” using the links given below: Microvillar Cartography data: https://www.ebi.ac.uk/biostudies/ studies/S-BSST520 (Accession: S-BSST520) Co-localization Probability Data: https://www.ebi.ac.uk/ biostudies/studies/S-BSST521 (Accession: S-BSST521) All other data generated during and/or analyzed during studies similar to those detailed above are available from the corresponding author upon request. Code Availability The custom MATLAB code described in this study can be found in the following link in the Github repository: https://github.com/shirsendughosh/Micorvillar-Catographyand-Colocalization-Probability-Code. The code is accompanied by operation instructions in the file “step by step guide to run analysis codes.pdf.” It can be accessed and used by readers without restriction. References 1. Sauvanet C, Wayt J, Pelaseyed T, Bretscher A (2015) Structure, regulation, and functional diversity of microvilli on the apical domain of epithelial cells. Annu Rev Cell Dev Biol 31: 593–621. https://doi.org/10.1146/annurevcellbio-100814-125234 2. Majstoravich S, Zhang J, Nicholson-Dykstra S, Linder S, Friedrich W, Siminovitch KA, Higgs

HN (2004) Lymphocyte microvilli are dynamic, actin-dependent structures that do not require Wiskott-Aldrich syndrome protein (WASp) for their morphology. Blood 104(5): 1396–1403. https://doi.org/10.1182/ blood-2004-02-0437 3. Cai E, Marchuk K, Beemiller P, Beppler C, Rubashkin MG, Weaver VM, Gerard A, Liu

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TL, Chen BC, Betzig E, Bartumeus F, Krummel MF (2017) Visualizing dynamic microvillar search and stabilization during ligand detection by T cells. Science 356(6338). https://doi.org/10.1126/science.aal3118 4. Yi JC, Samelson LE (2016) Microvilli set the stage for T-cell activation. Proc Natl Acad Sci U S A 113(40):11061–11062. https://doi.org/ 10.1073/pnas.1613832113 5. Mogilner A, Keren K (2009) The shape of motile cells. Curr Biol 19(17):R762–R771. https://doi.org/10.1016/j.cub.2009.06.053 6. Keren K, Pincus Z, Allen GM, Barnhart EL, Marriott G, Mogilner A, Theriot JA (2008) Mechanism of shape determination in motile cells. Nature 453(7194):475–480. https:// doi.org/10.1038/nature06952 7. Mattila PK, Lappalainen P (2008) Filopodia: molecular architecture and cellular functions. Nat Rev Mol Cell Biol 9(6):446–454. https:// doi.org/10.1038/nrm2406 8. von Andrian UH, Hasslen SR, Nelson RD, Erlandsen SL, Butcher EC (1995) A central role for microvillous receptor presentation in leukocyte adhesion under flow. Cell 82(6): 989–999. https://doi.org/10.1016/00928674(95)90278-3 9. Jung Y, Riven I, Feigelson SW, Kartvelishvily E, Tohya K, Miyasaka M, Alon R, Haran G (2016) Three-dimensional localization of T-cell receptors in relation to microvilli using a combination of superresolution microscopies. Proc Natl Acad Sci U S A 113(40): E5916–E5924. https://doi.org/10.1073/ pnas.1605399113 10. Ghosh S, Di Bartolo V, Tubul L, Shimoni E, Kartvelishvily E, Dadosh T, Feigelson SW, Alon R, Alcover A, Haran G (2020) ERM-dependent assembly of T cell receptor signaling and Co-stimulatory molecules on microvilli prior to activation. Cell Rep 30(10): 3434–3447 e3436. https://doi.org/10. 1016/j.celrep.2020.02.069 11. Sage PT, Varghese LM, Martinelli R, Sciuto TE, Kamei M, Dvorak AM, Springer TA, Sharpe AH, Carman CV (2012) Antigen recognition is facilitated by invadosome-like protrusions formed by memory/effector T cells. J Immunol 188(8):3686–3699. https://doi. org/10.4049/jimmunol.1102594 12. Razvag Y, Neve-Oz Y, Sajman J, Reches M, Sherman E (2018) Nanoscale kinetic segregation of TCR and CD45 in engaged microvilli facilitates early T cell activation. Nat Commun 9(1):732. https://doi.org/10.1038/s41467018-03127-w

13. Ghosh S, Feigelson SW, Montresor A, Shimoni E, Roncato F, Legler DF, Laudanna C, Haran G, Alon R (2021) CCR7 signalosomes are preassembled on tips of lymphocyte microvilli in proximity to LFA-1. Biophys J 120(18):4002–4012 14. McCarthy DA, Macey MG, Cahill MR, Newland AC (1994) Effect of fixation on quantification of the expression of leucocyte functionassociated surface antigens. Cytometry 17(1): 39–49. https://doi.org/10.1002/cyto. 990170106 15. Sundd P, Gutierrez E, Pospieszalska MK, Zhang H, Groisman A, Ley K (2010) Quantitative dynamic footprinting microscopy reveals mechanisms of neutrophil rolling. Nat Methods 7(10):821–824. https://doi.org/10. 1038/nmeth.1508 16. Axelrod D (2003) Total internal reflection fluorescence microscopy in cell biology. Methods Enzymol 361:1–33. https://doi.org/10. 1016/s0076-6879(03)61003-7 17. Stock K, Sailer R, Strauss WSL, Lyttek M, Steiner R, Schneckenburger H (2003) Variable-angle total internal reflection fluorescence microscopy (VA-TIRFM): realization and application of a compact illumination device. J Microsc (Oxford) 211:19–29. https://doi.org/10.1046/j.1365-2818.2003. 01200.x 18. Truskey GA, Burmeister JS, Grapa E, Reichert WM (1992) Total Internal-Reflection Fluorescence Microscopy (TIRFM). 2. Topographical mapping of relative cell substratum separation distances. J Cell Sci 103:491–499. https://doi. org/10.1529/biophysj.105.066738 19. van de Linde S, Loschberger A, Klein T, Heidbreder M, Wolter S, Heilemann M, Sauer M (2011) Direct stochastic optical reconstruction microscopy with standard fluorescent probes. Nat Protoc 6(7):991–1009. https:// doi.org/10.1038/nprot.2011.336 20. Rust MJ, Bates M, Zhuang X (2006) Subdiffraction-limit imaging by stochastic optical reconstruction microscopy (STORM). Nat Methods 3(10):793–795. https://doi.org/ 10.1038/nmeth929 21. Jensen KH, Berg RW (2016) CLARITYcompatible lipophilic dyes for electrode marking and neuronal tracing. Sci Rep 6:32674. https://doi.org/10.1038/srep32674 22. Rea R, Li J, Dharia A, Levitan ES, Sterling P, Kramer RH (2004) Streamlined synaptic vesicle cycle in cone photoreceptor terminals. Neuron 41(5):755–766. https://doi.org/10. 1016/s0896-6273(04)00088-1

Mapping Proteins with Respect to Membrane Topography 23. Sharp MD, Pogliano K (1999) An in vivo membrane fusion assay implicates SpoIIIE in the final stages of engulfment during Bacillus subtilis sporulation. Proc Natl Acad Sci U S A 96(25):14553–14558. https://doi.org/10. 1073/pnas.96.25.14553 24. Ajo-Franklin CM, Ganesan PV, Boxer SG (2005) Variable incidence angle fluorescence interference contrast microscopy for z-imaging single objects. Biophys J 89(4): 2759–2769. https://doi.org/10.1529/ biophysj.105.066738 25. Wolter S, Loschberger A, Holm T, Aufmkolk S, Dabauvalle MC, van de Linde S, Sauer M (2012) rapidSTORM: accurate, fast open-

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Chapter 13 T Cell Immunological Synaptosomes: Definition and Isolation Hye-Ran Kim, Jeong-Su Park, Na-Young Kim, and Chang-Duk Jun Abstract In addition to microvilli’s role as structural scaffold for TCR clustering, we recently discovered a novel function as message senders. We found that microvilli are separated from the T cell body shortly upon TCR stimulation and vesiculated to form T cell microvilli particles (TMPs), a new type of membrane vesicles. TMPs and synaptic ectosomes, which bud from the synaptic cleft, constitute “T cell immunological synaptosomes (TISs)” and act as conveyors of T cell messages or traits to cognate antigen-presenting cells. In practice, it is almost impossible to distinguish between TMPs and synaptic ectosomes. Here, we describe a newly developed protocol to isolate TISs from activated T cells using antibody-immobilized agarose beads and density gradient ultracentrifugation. We further describe the methods for TIS quantification with flow cytometry and to evaluate TIS efficacy on dendritic cells. Key words Microvilli, Extracellular vesicles, T cell microvilli particles (TMPs), T cell immunological synaptosomes (TISs), Antibody-immobilized agarose beads

1

Introduction The message transfer between T cells and antigen-presenting cells (APCs) determines the nature and extent of immune responses. Thus, understanding the mechanisms of the information exchange between two cells has received great interest. The immunological synapse (IS) has been most actively studied as a communication architecture between T cells and APCs, discovered, and named approximately two decades ago [1, 2]. Many important molecules involved in T cell activation or deactivation are clustered in the IS and can be exchanged [3–9]. However, recent evidence identified the same T cell molecules clustered in the IS in the microvilli tips of naı¨ve and effector T cells, indicating the importance of these surface projections as structural platform for clustering T cell molecules including TCRs [10–13]. Microvilli are fingerlike membrane structures on the cell surface enriched with actin bundles. T cells contain

Cosima T. Baldari and Michael L. Dustin (eds.), The Immune Synapse: Methods and Protocols, Methods in Molecular Biology, vol. 2654, https://doi.org/10.1007/978-1-0716-3135-5_13, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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abundant microvilli on their surface, but little is known about the physiological function of microvilli in relation to T cell life and fate. We and others recently reported that T cell contact to APCs occurs through microvillar extensions [10, 14, 15]. Extended microvilli appear to serve as spatiotemporal platforms for scaffolding immunologically relevant molecules, including TCR complexes, costimulatory and adhesion molecules, and various cytokines [11, 15]. Surprisingly, we observed that microvilli are rapidly separated from the T cell body by the combined action of two independent mechanisms (TCR activation and adhesion-dependent trogocytosis) and deposited on the surface of antigen-bearing APCs, potentially serving as a means of delivering T cell messages to activate APCs. Interestingly, the nano-sized small particles originating from the T cell surface could activate DCs regardless of the presence of antigens. Mounting evidence indicates that activated T cells release exosomes and ectosomes during IS formation [4, 8, 16]. However, we found that activation itself does not increase the release of extracellular vesicles except when in contact with the adherent substrate, be it cognate APCs, p-MHC/ICAM-1-embedded lipid bilayer, or plate-coated anti-CD3/28 antibodies. Interestingly, we further observed accumulation of microvilli-specific makers and TCRs in the central region of supramolecular activation clusters (cSMAC) during IS maturation, implying that the accumulation of TCR clusters into cSMACs is related to the microvilli’s centripetal movement [15]. Despite the possibility of different types of microvesicles being released from activated T cells, we suggest an overlapping origin of TCR-enriched microvesicles [4], synaptic ectosomes [8], and exosomes [16] with T cell-derived microvilli particles (TMPs). We therefore called the membrane particles released from the activated T cells on adhesion substrates T cell immunological synaptosomes (TISs). In these protocols, we used immobilized anti-CD3/28 antibodies CNBr-activated Sepharose beads to purify TISs. The strong covalent binding between antibodies and Sepharose beads prevents antibody co-elution with TISs, which can avoid antibody contamination in crude TISs. To isolate TISs, mouse T blasts were incubated with immobilized anti-CD3/28 antibodies, CNBr-activated Sepharose beads, and TISs were collected by sucrose gradient ultracentrifugation (Fig. 1). T cell adhesion on immobilized antiCD3/28-sepharose beads was confirmed by confocal microscopy (Fig. 2). Mouse CD4+ T cells expressing Vstm5_GFP, a maker protein localized in the stem region of microvilli [15] (Fig. 2a), were incubated with immobilized anti-CD3/28 antibodies Sepharose beads for 2 h and observed under confocal microscopy (Fig. 2b). Vstm5_GFP+ particles shed from activated T cells were scattered around the beads (Fig. 2b) or released in culture supernatants. The purified TISs according to the procedure described in

T Cell Immunological Synaptosomes: Definition and Isolation

s ad ion Be stitut n o c e

TISs

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(2) Vortex and spin down in the presence of EDTA and harvest supernatant

100,000 x g

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(5) Collect 4&5 fraction & ultracentrifugation

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1 0.2 M 2 3 4 5 6 P 2M

00

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(3) Repeat centrifugation to remove remaining cells & beads

Save supernatants 100,000 x g

(4) Collect TISs via sucrose gradient ultracentrifugation

Fig. 1 Schematic illustration of T cell immunological synaptosome (TIS) purification. Mouse CD4+ T blasts were incubated with anti-CD3/28 antibody-immobilized, CNBr-activated Sepharose beads in exo-free mouse T cell media for 3 h and allow TISs to dissociate from beads and cell mixture by vigorous vortex in the presence of 10 mM EDTA. All supernatants were subjected to two successive centrifugations at 2000
g for 10 min to completely remove cells and debris. TISs were centrifuged for 1 h at 100,000
g, and pellets were resuspended in PBS and reloaded onto a sucrose gradient of 12 different sucrose concentrations from top to bottom and centrifuged at 100,000
g for 16 h. Fractions were then carefully collected at 2 mL each from the bottom of the tube. Fractionated samples at levels 4–5 from the top were diluted with PBS and centrifuged at 100,000
g for 90 min to pellet the vesicles. While beads were reconstituted by separation of cells from beads using a 20 μm pore-sized strainer and adding triethanolamine (pH 11.5) followed by a neutralization buffer to remove residual particles after use

Fig. 1 were further quantified by CytoFLEX flow cytometry (Beckman Coulter, Brea, CA) (Fig. 3a). CytoFLEX could successfully detect extracellular vesicles (EVs) down to 150 nm offering the possibility of detecting particles 15,000 g for >30 min at 4 °C. Collect the supernatant, filter through 0.45 μm syringe filters and add 0.1 mM PMSF. 2. Load supernatant onto a custom-prepared 14.4.4S monoclonal antibody (mAb) affinity column (2 mL bed volume) with a flowrate of ~1 mL/min. Wash column with 10 column volumes of 1× PBS and elute I-Ek/ANP in 1 mL fractions with CAPS elution buffer. Prior to elution, prepare 0.5 mL of neutralization buffer in 1.5 mL tubes to immediately neutralize the high pH of the CAPS elution buffer. After elution, wash the 14.4.4S mAb column with 1 × PBS and store in 1× PBS supplemented with 0.05% NaN3 at 4 °C. Optionally, load the eluted and refolded I-Ek a second time onto the 14.4.4S mAb affinity column. 3. Subject the eluted protein to S200 gel filtration (Superdex 200 10/300) to remove aggregates and concentrate fractions containing monomeric I-Ek/ANP with Amicon®Ultra:4 centrifugal filters (10 kDa) to 1–2 mg/mL. Snap freeze aliquots in liquid N2 for storage at -80 °C or directly proceed with biotinylation.

3.5 Site-Specific Biotinylation of I-Ek/ ANP Using the BirA Biotin Ligase

For site-specific biotinylation of I-Ek/ANP with the GST-tagged BirA biotin ligase, we refer to the protocol described by Avidity (Avitag™ Technology). 1. Prepare 1–2 mg I-Ek/ANP at a concentration of 20–40 μM in 1× PBS. Prepare 10× stocks of Biomix A, Biomix B and additional biotin (0.5 mM d-biotin). 2. Dilute I-Ek/ANP (in 1× PBS) at least two- to threefold with ddH20 to reduce the NaCl concentration in the biotinylation reaction as NaCl strongly reduces the activity of the BirA biotin ligase. For example, mix 100 μL Biomix A (10×), 100 μL Biomix B (10×), 100 μL biotin (10×), 300 μL ddH20, and 300 μL of I-Ek/ANP protein solution. To ensure rapid biotinylation, increase the substrate concentration again to 20–40 μM using Amicon®Ultra:4 centrifugal filters (10 kDa). Finally, add at least 1 μg BirA ligase per 4 nmol substrate and incubate at 30 °C for 40 min. To reach quantitative biotinylation, adjust the amount of BirA biotin ligase to the quantity and concentration of the substrate as described by the manufacturer Avidity. In case of incomplete biotinylation of the substrate, increase the amount of BirA biotin ligase or the concentration of the substrate. If low protein yields after biotinylation occur, substitute NaCl for 0.2 M potassium glutamate and biotinylate overnight at 4 °C.

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3. After biotinylation, supplement the reaction mix with 100 mM NaCl and remove the BirA ligase using glutathione agarose according to the manufacturer’s instructions. Incubate the sample at 4 °C for 1 h, centrifuge at 1000 g for 1 min to remove the resin and filter the supernatant through a 0.22 μm centrifuge tube filter (spin at 10,000 g for 2 min). 4. Purify the biotinylated I-Ek/ANP via S200 gel filtration (Superdex 200 10/300), collect monomeric fractions and concentrate the protein to ~1 mg/mL using Amicon®Ultra:4 centrifugal filters (10 kDa). Snap freeze biotinylated I-Ek/ ANP with liquid N2 for storage at -80 °C or directly proceed with peptide exchange. 5. Verify quantitative biotinylation of I-Ek/ANP with a streptavidin-based gel shift assay and SDS-PAGE analysis. 3.6 Site-Specific Labeling of a Peptide with MaleimideConjugated Dyes

For site-specific labeling of peptides presented by I-Ek, we typically extend suitable peptides (e.g., moth cytochrome c peptide, ANERADLIAYLKQATK) with a GGSC-linker at the C-terminus (MCC-GGSC), which allows for maleimide-based conjugation to fluorophores as described in (Huppa et al. 2010). 1. Solubilize peptides in ddH20 or any other appropriate buffer and purify via reversed-phase HPLC (e.g., Pursuit XRs C18 5 μm 250 × 21.2 mm column) using an 0–100% linear gradient from an ionic solvent (0.1% trifluoroacetic acid in ddH20) to an organic solvent mix (0.1% trifluoroacetic acid, 10% ddH20, 89.9% acetonitrile) over 100 min with a flowrate of 5 mL/ min. Collect peak fractions. 2. Verify peak fractions via mass spectrometry (e.g., MALDI-TOF or nanoHPLC-nanoESI Orbitrap) and lyophilize peptide fractions with the correct molecular mass. Store lyophilized peptides at ≤ -20 °C. 3. Solubilize peptides at a concentration of 1–5 mg/mL (0.5–2.5 mM) in 1× PBS or any other buffer at pH 6.5–7.5 recommended for maleimide-based conjugation (e.g., 10–100 mM Tris–HCl or HEPES). 4. Reduce oxidized sulfhydryl groups with immobilized TCEP (Tris[2-carboxyethyl] phosphine hydrochloride) disulfide reducing gel according to the manufacturer’s instructions. Add one volume of TCEP resin (washed once with 1× PBS) to one or two volumes of peptide solution and incubate for 1 h at room temperature. After incubation, centrifuge the sample 1000 g for 1 min and filter the supernatant through a 0.22 μm centrifuge tube filter (spin at 10,000 g for 2 min). 5. Dissolve maleimide-conjugated dyes (e.g., Alexa Fluor™ 555 C2 Maleimide) at a concentration of 10–20 mM in DMSO or DMF immediately before use. Add sufficient

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maleimide-conjugated dyes from the stock solution to give approximately 2 moles of dye for each mole of peptide and incubate for 2 h at room temperature or overnight at 4 °C. We recommend mixing at least 300 μg MCC-GGSC with 356 μg Alexa Fluor™ 555 C2 Maleimide (1:2 molar ratio) to obtain sufficient amounts of dye-conjugated peptide for a quantitative I-Ek peptide exchange. The reaction can be stopped with an excess of glutathione. 6. Purify the reaction mix via reversed-phase HPLC to separate dye-conjugated peptide from unconjugated peptide or dye as described above. Verify peak fractions with mass spectrometry and lyophilize the site-specifically labeled peptide for storage at -80 °C. 3.7 Exchange of the I-Ek-Associated ANP Placeholder Peptide with Site-Specifically Labeled Peptides

The ANP placeholder peptide can be substituted with any peptide that binds into the peptide-binding cleft of I-Ek under slightly acidic conditions (pH 5.1) as described in [3, 4]. 1. Dilute 0.25 mg of I-Ek/ANP (~1 mg/mL) in 1× PBS to a final concentration 20 μM. Dissolve the lyophilized and sitespecifically labeled peptide (e.g., MCC-AF555) in 25–50 μL 1× PBS and add the solubilized peptide to I-Ek/ANP. Mix carefully. 2. For quantitative peptide exchange, add citric acid buffer pH 4.9 at a final concentration of 200 mM to the peptide exchange reaction to reach an exact pH of 5.1. Incubate for 1 h at room temperature and subsequently centrifuge for 2 min at 15,000 g to pellet and remove denatured protein. Collect supernatant and incubate for 1–3 days at room temperature. 3. Subject the protein-peptide solution to an S75 or S200 gel filtration step (e.g., Superdex 75 10/300 or Superdex 200 10/300) to remove protein aggregates and unbound peptide. Concentrate fractions containing monomeric I-Ek/ MCC-AF555 with Amicon®Ultra:4 centrifugal filters (10 kDa) and store the protein at a concentration of 0.2–1 mg/mL in 1× PBS supplemented with 50% glycerol at -20 °C. 4. Determine the protein-to-dye ratio and consequently the efficiency of peptide exchange by measuring the absorption at 280 nm and 555 nm with a spectrophotometer. A protein-todye ratio of 1 reflects quantitative peptide loading.

3.8 DNA Origami Preparation

DNA origami structures are typically assembled from a long, singlestranded scaffold DNA and short, specifically designed staple strands in a one-pot folding reaction using a thermal cycler. We have characterized in detail three rectangular, single-layer DNA origami tiles of three different sizes (30 × 20 nm, 65 × 54 nm,

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100 × 70 nm) [5] based on the M13mp18 scaffold [6]. In these core structures, specific staple strands can be elongated for functionalization with ligands (on the top side of the tile) or for anchorage to the SLB via hybridization to cholesterol-modified DNA oligonucleotides (on the bottom side). For functionalization, we either elongate one centrally located staple strand or two strands at 10, 20, or 30 nm distance; at the bottom side, the DNA origami feature six to ten elongations for the attachment of cholesterolDNA (Fig. 1). Layouts can easily be adapted for individual purposes using the open-source software caDNAno [7] (see Note 1). 1. Mix DNA (prewarmed to 24 °C), MgCl2, and folding buffer FoB 10× (50 mM Tris–HCl (pH = 8.0), 500 mM NaCl, 10 mM EDTA) in a DNA LoBind® tube (see Note 2). A representative folding protocol for 100 × 70 nm DNA origami tiles is shown in table below. Modified (i.e., biotinylated and/or fluorescently labeled) oligonucleotides are designed to be complementary in sequence to elongated staples for ligand attachment on the top side (“V sequence” 5′-ACATGACACTACTCCAC-3′), and cholesterol- DNA is complementary in sequence to elongations on the bottom side (“Z sequence” (5′-GGCTAAATATGCTAGGACTCT3′)) (see [8]).

Component

DNA strands Concentration [n]

M13mp18 scaffold

100 nM

–

10.0

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10×

–

10.0

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MgCl2

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–

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50 μM

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50 μM

1

0.2

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Biotin-DNA (V-sequence)

50 μM

1

1.0

500 nM/ staple

H2O, ultrapure Total volume

Volume Final [μL] concentration

31.5 100.0

2. Distribute the folding mix to ten PCR tubes and place them into a thermal cycler (CFX Connect Real-Time PCR Detection System) following a thermal protocol optimized for the

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structure to be folded. At this stage, DNA origami can be stored in the freezer at -20 °C for up to 3 months. The optimized thermal protocol for the described DNA origami layouts is (24 °C–90 °C, 10 °C min-1; 90 °C, 15 min; 90 °C– 4 °C, 1 °C min-1; 4 °C, 6 h). The critical temperature for DNA duplex formation is between 55 °C and 65 °C depending on the GC content. 3.9 DNA Origami Purification

Purification of DNA origami structures can be achieved via several methods [9], with yields and purity depending on the specific structure. For simple structures such as the described tiles, spin column purification (with, e.g., Amicon®Ultra centrifugal filters) constitutes an adequate and user-friendly option. For more complex DNA nanoarchitectures (e.g., 3D tripods), we recommend purification via agarose gel electrophoresis or FPLC to remove larger DNA aggregates. 1. Prepare 5 mL aliquots of the PuB. The lower Mg2+ content (5 mM) of the PuB compared to the folding mix improves recovery yields of correctly folded DNA origami structures from filter membranes. Always use freshly prepared PuB. 2. Pre-rinse Amicon®Ultra:0.5 (100 kDa cutoff membrane) with PuB according to the manufacturer’s instructions by evenly distributing 500 μL of PuB on the filter (do not touch the filter) followed by centrifugation at 5000 g for 5 min at 24 °C. Discard the flow-through and insert the filter into a clean tube. The molecular weight of staple strand DNA is ≤10 kDa, but in our hands, 100 kDa membranes yield better separation. 3. Mix 100 μL DNA origami solution with 400 μL PuB and evenly distribute the solution on the filter. Centrifuge at 7000 g for 5 min at 24 °C and discard the flow-through. 4. Add PuB to a volume of 500 μL to the filter followed by spinning at 7000 g for 5 min at 24 °C. Discard the flowthrough and repeat the procedure one more time. If the volume in the filter exceeds 50 μL, spin five to ten more minutes at 7000 g at 24 °C. 5. Add 50 μL PuB to a new clean collection tube, invert the filter and put it into the collection tube for recovering purified DNA origami structures. Spin at 5000 g for 4 min at 24 °C. Adjust the concentration of DNA origami to 10 nM by adding appropriate amounts of PuB. At this stage, DNA origami structures can be stored in Biopur® tubes for up to 4 weeks at -20 °C.

3.10 DNA Origami Functionalization Strategy Using Divalent Streptavidin

We have recently assessed multiple different strategies for the sitespecific decoration of DNA origami structures with regard to achieving optimal yields and full functionality of attached proteins [10]. We here describe a functionalization strategy based on dSA.

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In principle, also commercially available tetravalent streptavidin can be used, but this will likely lead to structures featuring up to three pMHC molecules per functionalization site [10]. 1. Add 50 μL of a 10 nM DNA origami solution into a DNA LoBind® tube, prewarm to 24 °C and add a 10× molar excess of dSAv. Incubate for 30 min at 24 °C. 2. Pre-incubate an Amicon®Ultra:2.0 centrifugal filter (100 kDa) with 2 mL PuB. Spin at 4000 g for 10 min at 4 °C. Discard the flow-through and reinsert the filter into the tube. 3. Mix the DNA origami solution with PuB to a final volume of 1.5 mL and evenly distribute the solution on the filter. Centrifuge at 4000 g for 15 min at 4 °C and discard the flow-through. Add 1.5 mL PuB to the filter followed by spinning at 4000 g for 15 min at 4 °C. Discard the flow-through. If the volume in the filter exceeds 50 μL, spin 5–10 min at 4000 g at 4 °C. 4. Invert the filter, add 50 μL PuB to the conical collection tube and spin at 2000 g for 4 min at 4 °C. Adjust the concentration of DNA origami to 5 nM with PuB. 5. Incubate the desired amount of DNA origami (pre-warmed to 24 °C) at 10× molar excess of AF555-conjugated and sitespecifically biotinylated pMHC in a DNA LoBind® tube for 60 min at 24 °C. 6. Repeat steps 2–4. 7. Functionalized DNA origami structures can be stored at this step (or after step 4) in Biopur® tubes up to 1 week at 4 °C. 3.11 DNA Origami Quality Control: Gel Electrophoresis

We recommend verifying the integrity of DNA origami structures as well as the successful functionalization applying several different methods including gel electrophoresis, high-speed atomic force microscopy (hs-AFM), electron microscopy, DNA Points Accumulation for Imaging in Nanoscale Topography (DNA-PAINT), and others. Representative images are shown in Fig. 3. We here describe the quality control via gel electrophoresis as a rapid initial quality feedback, which is readily available in most biochemical labs. We recommend performing gel electrophoresis after folding, after initial purification, and after dSAv attachment. hs-AFM imaging and DNA-PAINT imaging are recommended after dSAv functionalization but before addition of biotinylated pMHC. 1. Prepare a 1% w/v agarose gel with 1× Tris-acetate-EDTA (TAE) buffer pH 8; stain with, for example, Sybr™-Gold. For larger or smaller DNA origami structures, agarose content can be varied between 0.5% and 2.0%. Load ~5 ng DNA origami, the M13mp18 scaffold and a 1kB DNA ladder on the gel and run the gel for 75 min at 100 V and 24 °C in 1× TAE pH 8 supplemented with 10 mM MgCl2. Avoid currents

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Fig. 3 DNA origami quality control. (a) Agarose gel electrophoresis of DNA origami platforms. Schematic sketches above the individual lanes indicate the different DNA origami layouts functionalized with dSAv. from left to right: M13mp18 scaffold, S, M, L DNA origami. (b) Schematics of a 100 × 70 nm (L) DNA origami tile. Circles indicate available modification sites on the top side of the DNA origami tile; full circles indicate sites used for functionalization with two ligands at 20 nm distance (Ldiv 20 nm). The distances are approximated with 0.34 nm per base pair along the helical axis and with 2 nm per helix perpendicular to the helical axis [6]. Interhelical gaps were assigned 1 nm, based on the spacing of crossovers between helices. Distances are given in nm. (c) hs-AFM image of an L DNA origami platform featuring two dSAvs spaced 20 nm apart. Scale bar, 25 nm. (d) Mapping of dSAv positions on DNA origami platforms via DNA-PAINT. (i) Biotinylated ligands are replaced with biotinylated DNA-PAINT docking strands. These are detected via transient binding of fluorescently labeled imager strands. (ii) Representative pseudo-color DNA-PAINT super-resolution image of the large DNA origami platform featuring two ligand attachment sites at 20 nm distance. Ligand (cyan) and platform (red) positions were imaged consecutively by Exchange-PAINT [19]. Scale bar, 50 nm. (iii), Crosssectional histogram of ligand positions from DNA-PAINT localizations summed up from100 individual DNA origami platforms

above 150 mA. The presence of MgCl2 is necessary to maintain DNA origami integrity but may result in temperature peaks (>>40 °C), which may damage the DNA origami as well as attached proteins. We recommend to regularly monitor the temperature and, if necessary, run the gel on ice. 2. Image the gel with an UV-light source. Correctly folded DNA origami will appear as a discrete band shifted to a larger apparent size when compared to the M13mp18 scaffold. 3.12

SLB Preparation

Planar supported lipid bilayers (SLB) form spontaneously on a hydrophilic glass surface upon addition of small unilamellar vesicles (SUVs, ; = 20–100 nm), which can be easily generated from dried lipid mixtures via bath sonication. We use a lipid mixture containing 98% 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine (POPC)

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and 2% 1,2-dioleoyl-sn-glycero-3-[(N-(5-amino-1-carboxypentyl) iminodiacetic acid)succinyl] (nickel salt) (DGS Ni-NTA) to create a mobile SLB that allows for protein attachment via poly-histidine tags. 1. Add 48.6 μL of POPC (10 mg/mL) and 13.8 μL DGS Ni-NTA (1 mg/mL) to a Fiolax® borosilicate test tube to achieve a 49:1 molar ratio and a total of 500 μg of lipid. 2. Fully evaporate the chloroform under a gentle stream of inert gas (e.g., nitrogen) inside a chemical hood until a lipid film has formed. 3. For preparing a 5× stock suspension of lipid vesicles (0.5 mg/ mL), add 1 mL of PBS 10× to the dried lipid film and gently resuspend the lipids until the suspension assumes a milky appearance. Firmly seal the test tube (with, e.g., Parafilm) and sonicate the lipid suspension in the test tube for at least 15 min at 20 °C until the lipid suspension has cleared up. Best results are achieved by placing the test tube into the center of a whirl with the surface of the lipid suspension ~3 mm below the water-air interface. The temperature of the water bath should be above the phase transition temperature of the lipid mixture. 4. The 5× lipid vesicle stock suspension can be stored for 2–4 days at 4 °C. Dilute 1:5 with PBS 10× for further use. 5. Ensure glass coverslips are completely dry. Place coverslips into a plasma cleaner, evacuate for at least 10 min to a pressure of 0.2–0.3 mbar before igniting the plasma (pale purple color). Clean for 3 min at a radio frequency power of maximally 30 W. If treatment with an O2 plasma does not yield satisfactory results, immerse coverslips in Piranha solution prior plasma cleaning (1:1 mixture of concentrated sulfuric acid and 30% hydrogen peroxide) for 30 min followed by thorough rinsing with ddH2O). 6. Remove the tape of a sticky-slide 8-well chamber and gently press the cleaned coverslip onto the chamber. 7. Flip the chamber, add 220 μL of the vesicle suspension per well and incubate for 10 min at 24 °C. 8. Carefully rinse the SLB with 20 mL PBS 1× using a 25 mL serological pipette. From a full well, remove 330 μL to leave 350 μL PBS 1× in the well. SLBs can be stored overnight at 4 °C. 3.13 SLB Functionalization

1. Thaw cholesterol-DNA and incubate it on the SLB at a final concentration of 0.1 μM for 60 min at 24 °C. 2. Remove excessive cholesterol-DNA by gently washing the SLB with 10 mL PBS 1× supplemented with 1% BSA. This washing step is crucial for proper attachment of DNA origami to the

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SLB. Residual cholesterol-DNA in solution will promote formation of DNA origami aggregates. 3. To achieve a surface density of ~1 pMHC per μm2 on the SLB, dilute 5 nM DNA origami solution 1:200 in PBS 1× and add 3 μL to the SLB. Allow to hybridize the elongated staple strands at the bottom side of the DNA origami to complementary cholesterol-DNA in the SLB for 60 min at 24 °C. The surface density of DNA origami structures on the SLB increases nonlinearly with increasing their concentration in solution. The actual density of fluorescently labeled pMHC should always be determined following the protocol given in Subheading 3.16. As a rough guideline, for ~10 pMHC per μm2, dilute 5 nM DNA origami 1:10 in PBS 1× and add 5 μL to the SLB; for ~100 pMHC per μm2, use 20 μL of a 25 nM DNA origami solution. 4. Remove unbound DNA origami by rinsing the SLB with 10 mL PBS 1×. 5. Add His10-tag ICAM-1 and His10-tag B7–1 to a concentration of 5 nM each to the SLB and incubate for 75 min at 24 °C to achieve a surface density of 100 molecules per μm2 for each protein. Rinse with 10 mL PBS 1×. Replace PBS 1× with imaging buffer (HBSS) before cell imaging experiments. Following the precise order of incubation steps on the SLB is critical for achieving efficient and homogeneous decoration of the SLB with all components. Changing the order will decrease DNA origami attachment yields. 3.14 Diffusion Analysis of DNA Origami Structures

1. Set up TIR illumination and focus onto the DNA origamidecorated SLB (see Note 3). 2. Record multiple time-lapsed movies (≥10 movies) at different locations of the SLB for adequate sampling. We typically use an illumination time (till) of 3 ms and a delay of 7 ms between images, yielding tlag = 10 ms based on an expected mobility of DNA origami of 98% ethanol, design side down, and sonicate for 5 min. 6. Recover the microfluidic chip using the tweezers, dry it using compressed dry air, and clean it with tape. If possible, check the integrity of the designs and the placement of the injection holes using a benchtop magnifier (see Note 24). 7. Place the microfluidic chip and the fluorodish, without its cover, in the plasma cleaner for 1 min (follow manufacturer’s instructions for manipulating the atmospheric plasma cleaner, we use it at maximum power, i.e., 18 W), and then take the chip and add it on the glass bottom of the fluorodish, design facing down. Ensure both sides are in contact by looking at the homogeneity of the color of the contact zone. Heat for at least 1 h at 60 °C to reinforce bonding. 8. Activate the bonded chip in the plasma cleaner for 1 min before injecting gently with the 0.2% w/v PVP-K90 solution in MilliQ water using a syringe with a filter, until some solution exited and visual control confirms the whole design is filled (see Note 25). Then, add more solution on top to prevent drying and store the dish at 4 °C (see Note 26).

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This is one method of droplet formulation and functionalization that allows to obtain droplets of a diameter comparable to that of a cell, and mobility of the ligand at the surface of the droplet to follow antigen clustering during immune synapse formation, but other techniques allow to obtain very different droplet characteristics (see Note 27). 1. Prepare the oil phase by resuspending DSPE-PEG(2000)-biotin at 10 mg/mL in chloroform. Add 150 μL of this solution to 30 g of soybean oil, stir gently, and leave in a vacuum bell jar (low vacuum, ~100 mBar) for over 3 h 30 m at room temperature (see Note 28). 2. Prepare the continuous aqueous phase by adding 2.5 g 4%w/v sodium alginate with a spatula in a small beaker, then completing to 5 g with deionized water. Add 5 g of 30% w/v Pluronic F68 solution. Stir gently with the spatula, avoiding bubbles. Remove bubbles using a vacuum bell (low vacuum, ~100 mBar) for 1 h once the solution is prepared. 3. Slowly add the oil phase to the aqueous phase: add two to three drops, stir gently with a spatula until the oil is incorporated before adding more drops, and repeat this process. Progressively, the oil phase gets incorporated more easily and can be added faster. During this whole step, avoid introducing air bubbles. A rough, white emulsion should be obtained. 4. Add the emulsion to the Couette cell, and shear at 150 rpm (e = 100 μm) (see Note 29). Recover the emulsion as it gets out from the Couette cell, and collect it in a tared beaker. 5. Determine the mass of the obtained emulsion, which is at 25% aqueous phase, with 15%w/v Pluronic F68 in it. 6. Add 14 times the estimated aqueous volume of deionized water to obtain a final emulsion at 5% in the oil phase and 1% in Pluronic-F68. Place the emulsion in a separating funnel, protected from light, for 24 h at 1%w/v Pluronic F68, 5%w/v oil phase, then remove the clearer part under the cream of droplets. Wash the emulsion in freshly prepared 1%w/v Pluronic F68 like this at least two times, or until the lower part of the solution is clear (see Note 30). 7. After the last wash, remove the clear lower phase and discard it. Recover the final emulsion and store it in glass vials at 12 °C, protected from light (see Notes 31 and 32). This emulsion is used as stock emulsion and can be used for up to a year, or until the appearance of the emulsion has changed, i.e., oil and water are two separated phases. 8. Before starting to do experiments with droplets, take some emulsion, dilute it 100 times and verify their size distribution

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Fig. 2 Functionalization of oil droplets. The stock emulsion is prepared as described in Subheading 3.3 Droplet formulation. Oil droplets presenting biotinylated phospholipids at their surface are sequentially functionalized with a fluorescent streptavidin and a biotinylated ligand, here a biotinylated BCR ligand acting as antigen. Images of diluted, functionalized droplet suspension by spinning disk confocal microscopy below. Scale bar 20 μm

(by light microscopy or by flow cytometry). Using this protocol, the droplet diameter should be around 9 μm on average, with ~10% dispersion in size. 3.4 Droplet Functionalization

Droplet functionalization should be performed on the day of the experiment (Fig. 2). Throughout the protocol of droplet functionalization, use only low-binding microtubes. See Note 28 for an alternative functionalization. 1. Take 2 μL of stock droplet emulsion using a cut pipet tip to not damage the emulsion, and put it into a microtube, before adding 198 μL of PB + Tween 20 buffer. 2. Wash the emulsion: centrifuge the emulsion at 600 g for 30 s in a minifuge. Remove 180 μL of the aqueous phase (see Note 33). 3. Add 180 μL PB + Tween 20. Repeat the washing step (step 2) three times (see Note 34). 4. After the last wash, leave 20 μL of emulsion in the microtube. In another low-binding microtube, prepare 180 μL of PB + Tween 20 buffer with 2.5 μg of streptavidin. Add this solution to the 20 μL of droplet emulsion and homogenize well by gently pipetting up and down a few times (no vortex).

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5. Incubate for 30 min on a rotating wheel at RT, protected from light. 6. Wash droplets as detailed in steps 2–3 to remove the excess of streptavidin. 7. After the last wash, leave 20 μL of emulsion in the microtube. In another low-binding microtube, prepare 180 μL of PB + Tween 20 buffer with 5 μg of biotinylated ligand. 8. Incubate for 30 min on a rotating wheel at RT, protected from light. 9. Wash droplets as detailed in steps 2–3 to remove the excess of ligand, and leave in 200 μL total (see Note 35). 3.5 Immune Synapse Reconstitution in the Microfluidic Chip

1. On the morning of the experiment, take the microfluidic chips out of the fridge and to room temperature. 2. Remove the PVP solution from the fluorodish, and inject the microfluidic chip with cell culture media (with a drug if the experiment involves a treatment, see Note 36) until a large drop has formed on the exit side. Add some media (~1 mL) on top of the microfluidic chip to prevent drying. Put the microfluidic chip in an incubator (37 °C, 5% CO2) until the time of experiment (see Note 37). 3. Cut three pieces of tubing or similar length, long enough to join the reservoir of the pressure controller and the stage of the microscope. Insert a metal injector at the extremity of each piece of tubing, and install two of these tubes on the two pressure controllers. 4. Take a microfluidic chip, remove some of the media on top, and install it on the microscope stage (see Note 38). Orient it so that the entry injection point is on the side of the pressure controller and reservoirs. 5. Prepare one microcentrifuge tube with the well-resuspended target suspension (see Note 39), and one microcentrifuge tube with cell culture media +25 mM HEPES (± drug). Install them on the reservoir holders. 6. Gradually increase the pressure for the target suspension, until a drop is getting out of the metal injector. Lower the pressure to decrease the flow and prevent losing the content of the reservoir, and deposit the drop on the entry point of the microfluidic chip before inserting the injector in the entry point hole. Only then, increase the pressure to inject the targets (see Note 40). Lower the pressure when enough targets have been trapped to avoid more targets getting into the microfluidic chip (Fig. 3) (see Note 41). 7. Gradually increase the pressure for the cell culture media, until a drop is getting out of the metal injector. Lower the pressure

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Fig. 3 Steps for immune synapse reconstitution in the microfluidic chip

to decrease the flow and prevent losing the content of the reservoir, and deposit the drop on the entry point of the microfluidic chip. Hold the fluorodish and the cell culture media tubing with one hand, remove the target suspension tubing and injector with the other one, and replace it by the cell culture media tubing and injector (see Note 42). Increase the injection pressure to wash the microfluidic chip from all non-trapped targets (Fig. 3). Cut the pressure for the target suspension, attach the metal injector at a high point to prevent the remaining liquid in the tubing from flowing out (see Note 43). Then, disconnect its tubing and reservoir. 8. While the microfluidic chip is being washed with cell culture media, set up the microscopy settings (see Note 44). Look at the fluorescence of the target and adjust the settings, identify the fields of view to image, and add some of the lymphocyte suspension on the side of the fluorodish, next to the microfluidic chip, to adjust settings for fluorescent imaging if they are stained. 9. Install the third tubing + metal injector on the free pressure controller. Prepare a microcentrifuge tube with the lymphocyte suspension, and set it up on the pressure controller. As in step 7, replace the cell culture media inlet with the lymphocyte suspension inlet. When the injector is in place, keep the pressure low while selecting the fields of view for acquisition. 10. Launch the acquisition, let one image be acquired before increasing the injection pressure of the lymphocyte suspension to have the initial state of the target, and the first time of contact between the target and the lymphocyte (Fig. 4). When there are enough target-lymphocyte doublets (see Note 41), decrease the injection pressure to prevent more lymphocytes from accumulating. 3.6

Final Remarks

The 4D (+ color if necessary) data collected with a confocal or epifluorescence microscope can be reconstructed in a single movie to obtain a multi-tiff file (see Fig. 4 for examples of images

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Fig. 4 Examples of images obtained using this system, on a spinning disk confocal microscope: (a) B lymphocyte cell line (IIA1.6) stained with Hoechst to visualize the nucleus, in contact with an antigencoated droplet, (b) naive primary murine B lymphocyte expressing LifeAct-GFP, stained with Hoechst to visualize the nucleus, in contact with an antigen-coated droplet, (c) primary CD8+ T cell from OT-I mouse, stained with Hoechst to visualize the nucleus, in contact with an antigen-coated droplet (antigen: pMHC OVA SIINFEKL/H2-Kb/monomer). Scale bar 5 μm. Illustrations are z-projections over a few planes at the center of the cell, from z-stacks covering the full depth of the cell

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obtained). To analyze polarization phenomena we recommend to crop, on each field of view, the traps where both a target and a lymphocyte are present, and to remove the time points that precede the arrival of the cell. Image analysis tools (Imaris, FIJI, etc.) can then be applied to track, for example, organelles and cytoskeleton dynamics and/or the morphodynamics of the nucleus and the cell.

4

Notes 1. This is to remove humidity/water that might prevent photoresist adhesion. 2. Pre-heat the hot plates to have a stable temperature, and make sure to keep the humidity in the room as low as possible. 3. Various designs have been described over the years [9, 10, 14, 15, 19–21] and can be used for that protocol. Here, we describe the protocol for the design used in [9] and described in [14], with a B lymphocyte cell line (they can be found in the Github repository https://github.com/FattaccioliLab/Phen otypingBCellsWithLipidDroplets). One can also use smaller micro-traps for primary lymphocytes, as shown in Figs. 1 and 4 [9]. Photomasks can be ordered via several online services (e.g., https://www.selba.ch, https://www.jd-photodata.co. uk/ https://www.microlitho.co.uk/) 4. We find that the other commonly-used PDMS, Sylgard, has other mechanical properties and tends to break more easily than RTV during the repeated injections with needles. Of note, RTV and Sylgard may differ in adsorbing drugs during experiments. 5. The PDMS components can be mixed in a plastic cup, or a falcon tube, mixed using a disposable pipette. The PDMS mix can also be prepared in advance, in excess, and stored at -20 °C in a falcon tube. In that case, bubbles will disappear over time, and the tube should be put back to room temperature a little before using, but the PDMS-filled molds will not need to be left very long in the vacuum bell. 6. One can also use a 0.75 mm diameter microfluidic puncher, and replace the metal injectors and needles by PTFE tubing by making a Tygon-PTFE junction. 7. This can also be replaced by DOPE, or other functionalization of DSPE-PEG. 8. One can also use other oils, such as mineral oil, to achieve different droplet properties.

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9. Keep a stock of PB buffer, pH = 7, 20 mM, and a stock of PB + Tween 20 20% v/v, to prepare the 0.2% solution on the day of the experiment. 10. We tried different providers, fluorophores, and stock concentrations for the streptavidin, and all were compatible with our system. 11. This ligand can be changed depending on the experiment. We have also used biotinylated BSA as a negative control, F(ab′)2 anti-mouse IgM or biotinylated HEL for primary B cells, biotinylated ICAM-1, biotin anti-CD3 or pMHC OVA SIINFEKL/H2-Kb/ monomer for primary T lymphocytes (see Fig. 4). These ligands can also be mixed to have several functions on the droplets, although their proportions would have to be calibrated. 12. Metal injectors can also be custom-made, but we find that this is the most cost-effective protocol. Metal injectors can be re-used between experimental days but need to be flushed with MilliQ water after an experiment, kept in ethanol, and sonicated before being reused. This avoids the presence of dried cells or medium components, and therefore debris or clogging. 13. During the experiment, the chip is constantly connected to the pressure controller via the metal injector and tubing, preventing us from adding the CO2 control cover on top. We find that adding 25 mM HEPES and keeping the whole microscope at 37 °C keeps the cells healthy for the duration of the experiment (1 h maximum in our case). 14. We find that the concentration of the lymphocyte cell suspension depends on the cell type and version of the microfluidic design used. Typically, for the lymphocyte cell line, we use 1.5.106 cells/mL, and for primary murine lymphocytes, we use 3.106 cells/mL. 15. We use one well/chip. We find that it is better to separate the cells in wells so as to prevent repetitive resuspension and manipulation of the same cells and facilitate addition of drugs for pre-incubation in a systematic manner during the day of the experiment. Overall, this improves reproducibility. 16. Here, we use antigen-coated oil droplets to activate B lymphocytes. However, any object of similar dimension such as beads or other cells can be used. The concentration of this solution should be tuned to the size of the object. In experiments using beads or droplets, and involving drug treatment, we advise to resuspend the beads/droplets with cell culture media +25 mM HEPES + drugs. Of note, under a certain concentration of droplets, we found that almost nothing was trapped.

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In our case, we use antigen-coated droplets on which the antigen is mobile, in order to visualize antigen recruitment and follow immune synapse formation. 17. This protocol should be adapted for smaller cells, e.g., primary cells, where not only the trap design is modified, but also the height of the microfluidic chip should be close to 8 μm to avoid cells going above or below droplets, or being stacked on top of each other. 18. Viscosity of the SU-8 varies between lots and with aging of the resin, so this protocol, and in particular spin-coating (time, speed) of the SU-8 photoresist in step 2, should be adapted to the SU-8 batch to control final thickness of mold. It is good practice to aliquot the stock resin solution in 50–100 mL bottles to limit aging. 19. For a source having a power of 20 mW/cm2, the exposure time is calculated by dividing the dose by the lamp power, which gives an exposure time of 7.0 s. 20. If the wafer takes on a whitish aspect when put in contact with isopropanol, that means that the development is not complete, and you should increase the time or refresh the developer solution. 21. If at this stage, the patterns are not well resolved, or you see detachment, oblique sidewalls, etc., you have to consider optimizing the exposure and development steps. 22. This step improves long-term use and storage of wafers and facilitates the detachment of cured PDMS from the wafer. We advise frequent users to prepare epoxy replicates of the microfluidic chips when possible (depending on size and design resolution), thus as to avoid using the initial wafers on a dayto-day basis as they are very fragile. An example of well adapted protocol for it can be found in Refs. [22, 23]. Of note for chips with small patterns the procedure might not work. 23. Visualizing the injection sites can be made easier by putting ethanol on the chip, which will be retained on the designs. 24. The cleaning procedures of the microfluidic chip can vary depending on the environment they are prepared in. Here, we propose a protocol for chips prepared in a cleanroom-free facility, and some steps could be eliminated or simplified in a microfluidics-dedicated space, with less dust. 25. PDMS is initially very hydrophobic, making it difficult to inject liquid in it and avoid bubbles. Filling the chip with the PVP solution allows a coating of the PDMS, and its permanent contact with an aqueous solution, keeping it more hydrophilic until the time of the experiment.

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26. Microfluidic chips can be kept at 4 °C in the 0.2% w/v PVP K90-filled fluorodish for up to a week before the experiment. 27. Of note we tested two types of functionalization: one where the biotinylated phospholipids are mixed with the oil (bulk) and one where they are incorporated in the droplet after droplet fabrication (surface) [10]. We present here only the bulk one, as it is the one that permits the accumulation of the antigen at the synapse. Surface-functionalized droplets do not show antigen accumulation but allow, nevertheless, the activation of the cells. This functionalization technique allows very precise quantification of the ligands present at the interface and is less demanding in terms of materials. The reader could be also interested in changing the mechanical properties of the droplets, which is doable modifying the oil mixture. Details on alternative functionalization methods and surface tension can be found in [10, 24]. 28. It is crucial that the chloroform has well evaporated, otherwise this changes the viscosity of the solution and will affect the final droplet size. 29. The rough emulsion is sheared in the Couette cell to obtain more homogeneous and smaller droplets. Changing the speed/settings of the Couette cell or the viscosity of the water phase will change the average size of the droplets. 30. These steps allow to remove some of the smallest droplets, which will not get decanted to the surface of the funnel, and obtain a more homogeneous emulsion. 31. Separate the stock into several glass vials, as repeated openings of the vial will degrade the emulsion faster. This way, the emulsion stock can be used for longer. 32. To store products at 12 °C, we find that the most cost-effective item is a small wine cellar for service, where the door can be covered with aluminum foil to block the light. 33. We encourage you to leave the tube in the centrifuge for at least 30 s after the spin so that the emulsion is well concentrated at the surface, on one side. To remove the aqueous phase, use a gel-loading pipet tip and insert it on the side of the microtube with the least emulsion, then slowly aspirate the aqueous phase sitting below. 34. During these washes, the Pluronic F68 surfactant in continuous phase is removed. At the oil/water interface, Tween 20 surfactant is progressively added to ensure a homogeneous protein coating. 35. For experiments using drugs, we separate this solution into several microcentrifuge tubes and resuspend the droplets in media+drug to ensure the good equilibrium of drug concentration.

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36. PDMS is very permeable and can adsorb drugs. It is therefore important to pre-incubate the microfluidic chip with cell culture media + drug to obtain a stable concentration of drug treatment during the experiment. 37. Equilibrating the microfluidic chip in an incubator allows it to be at the right temperature and gas (CO2) content for the experiment and therefore maintain good conditions for the cell as well as for imaging, as temperature differences increase the risk of drifting during time-lapse microscopy. 38. Do not dry the surface of the chip, and do not let it with little media for a long time. The surface should not dry because this increases the risk of introducing air bubbles into the microfluidic chip when injecting and of dry media debris entering and damaging the microfluidic chip. 39. We find that it is better to not put all target suspension in the microcentrifuge tube at once, but rather add some more for each microfluidic experiment during the day. Indeed, in case of a problem with the pressure controller, one could lose the full content of the microcentrifuge tube. In addition, this allows the use of a suspension from the same fresh and clean stock for each acquisition. 40. With our design, our pressure controller, and when putting the reservoir on the side of the microscope stage at a height close to the one of the microfluidic chip, we find that setting the pressure around 12 mbar until liquid gets out of the injector, lowering the pressure to 8–9 mbar while inserting the metal injector into the hole, then injecting targets/lymphocytes with a pressure around 20 mbars, and setting to a resting pressure (very low flow to prevent arrival of more targets/lymphocytes) around 6–8 mbars (this can vary a lot depending on the design and the length of the tube) is a good starting point. 41. A good rule of thumb is that if the microscope settings allow to image 30 traps, then having 10–15 traps with a target/lymphocyte in it is enough (keeping in mind the field of view of the microscope). One should keep in mind that more targets/ lymphocytes will certainly arrive and be trapped in the next few minutes, during the wash with cell culture media, and that if there are traps containing several targets/lymphocytes, analysis will be compromised. 42. This manipulation is critical and will be repeated in the next step. It requires some training before being able to avoid injecting air bubbles, which can be mostly prevented by ensuring that the incoming metal injector has media going out of it. Another risk is to trigger backflow and have targets flowing back to the entry.

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43. We find that taping it to the ceiling of the 37 °C box of the microscope is the most convenient. 44. As an example of imaging settings, with a spinning disk confocal microscope, we could do 40 min acquisitions with 1 image every 30 s, at 3 positions, in 3 colors (Transmission +2 fluorescent channels), and 21 z slices, using a spinning disk confocal microscope and streaming each channel. For the cell line microfluidic chip design, the Nikon perfect focus system does not work, but one can use the software autofocus on the transmission channel at each time point, and an offset on the z-position for the fluorescent channels.

Acknowledgments JP was supported by the Ecole Doctorale FIRE-Programme Bettencourt, Universite´ Paris Cite´ and by funding from the Agence Nationale de la Recherche (ANR-21-CE30-0062-01 IMPerIS). LP was supported by the IPV scholarship program (Sorbonne Universite´). JF and PP acknowledge funding from CNRS; PP and JP acknowledge funding from the Agence Nationale de la Recherche (ANR-10-IDEX-0001-02 PSL*); and JF acknowledges funding from the Agence Nationale de la Recherche (ANR Jeune Chercheur PHAGODROP, ANR-15- CE18-0014-01). References 1. Yuseff M-I, Pierobon P, Reversat A, LennonDume´nil A-M (2013) How B cells capture, process and present antigens: a crucial role for cell polarity. Nat Rev Immunol 13:475–486. https://doi.org/10.1038/nri3469 2. Dustin ML (2014) The immunological synapse. Cancer Immunol Res 2:1023–1033. https://doi.org/10.1158/2326-6066.CIR14-0161 3. Dustin ML, Long EO (2010) Cytotoxic immunological synapses. Immunol Rev 235: 2 4 – 3 4 . h t t p s : // d o i . o r g / 1 0 . 1 1 1 1 / j . 0105-2896.2010.00904.x 4. Batista FD, Iber D, Neuberger MS (2001) B cells acquire antigen from target cells after synapse formation. Nature 411:489–494. https:// doi.org/10.1038/35078099 5. Ritter AT, Asano Y, Stinchcombe JC, Dieckmann NM, Chen BC, Gawden-Bone C, van Engelenburg S, Legant W, Gao L, Davidson MW, Betzig E, Lippincott-Schwartz J, Griffiths GM (2015) Actin depletion initiates events leading to granule secretion at the immunological

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10. Pinon L, Ruyssen N, Pineau J, Mesdjian O, Cuvelier D, Chipont A, Allena R, Guerin CL, Asnacios S, Asnacios A, Pierobon P, Fattaccioli J (2022) Phenotyping Polarization Dynamics of Immune Cells Using a Lipid Droplet – Cell Pairing Microfluidic Platform. Cell Reports Methods 2, 11, 100335. https://doi.org/10. 1016/j.crmeth.2022.100335 11. Bourouina N, Husson J, Hivroz C, Henry N (2012) Biomimetic droplets for artificial engagement of living cell surface receptors: the specific case of the T-cell. Langmuir 28: 6106–6113. https://doi.org/10.1021/ la300398a 12. Barek KBM (2015) Adhesion et phagocytose de gouttes d’emulsions fonctionnalisees. Doctoral dissertation, Ecole Normale Supe´rieure, Paris 13. Ben M’Barek K, Molino D, Quignard S, Plamont MA, Chen Y, Chavrier P, Fattaccioli J (2015) Phagocytosis of immunoglobulincoated emulsion droplets. Biomaterials 51: 270–277. https://doi.org/10.1016/j. biomaterials.2015.02.030 14. Mesdjian O, Ruyssen N, Jullien MC, Allena R, Fattaccioli J (2021) Enhancing the capture efficiency and homogeneity of single-layer flowthrough trapping microfluidic devices using oblique hydrodynamic streams. Microfluid Nanofluid 25(11):91. https://doi.org/10. 1007/s10404-021-02492-1 15. Dura B, Dougan SK, Barisa M, Hoehl MM, Lo CT, Ploegh HL, Voldman J (2015) Profiling lymphocyte interactions at the single-cell level by microfluidic cell pairing. Nat Commun 6: 5 9 4 0 . h t t p s : // d o i . o r g / 1 0 . 1 0 3 8 / ncomms6940 16. Jang JH, Huang Y, Zheng P, Jo MC, Bertolet G, Zhu MX, Qin L, Liu D (2015) Imaging of cell-cell communication in a vertical orientation reveals high-resolution structure of immunological synapse and novel PD-1 dynamics. J Immunol 195(3): 1320–1330. https://doi.org/10.4049/ jimmunol.1403143

17. Mesdjian O (2018) Etude des phe´nome`nes d’adhe´sion entre des cellules B et des gouttes d’huile fonctionnalise´es par des anticorps a` l’aide de pie`ges microfluidiques, Doctoral dissertation, Ecole Normale Supe´rieure 18. Microchem PROCESSING GUIDELINES FOR:SU-8 2000.5, SU-8 2002, SU-8 2005, SU-8 2007, SU-8 2010 and SU-8 2015. https://kayakuam.com/su-8-series/ 19. Skelley AM, Kirak O, Suh H, Jaenisch R, Voldman J (2009) Microfluidic control of cell pairing and fusion. Nat Methods 6(2):147–152. https://doi.org/10.1038/nmeth.1290 20. Sinha N, Subedi N, Tel J (2018) Integrating immunology and microfluidics for single immune cell analysis. Front Immunol 9:2373. https://doi.org/10.3389/fimmu.2018. 02373 21. Hoehl MM, Dougan SK, Ploegh HL, Voldman J (2011) Massively parallel microfluidic cellpairing platform for the statistical study of immunological cell-cell interactions. In: 15th International Conference on Miniaturized Systems for Chemistry and Life Sciences 2011, MicroTAS 2011. MicroTAS, pp 1508–1510 22. Heuze ML, Collin O, Terriac E, LennonDumenil AM, Piel M (2011) Cell migration in confinement: a micro-channel-based assay. Methods Mol Biol 769:415–434. https://doi. org/10.1007/978-1-61779-207-6_28 23. An inexpensive and durable epoxy mould for PDMS – Chips and Tips. https://blogs.rsc. org/chipsandtips/2009/04/22/an-inexpen sive-and-durable-epoxy-mould-for-pdms/? doing_wp_cron=1662134774.70260810 85205078125000 24. Pinon L, Montel L, Mesdjian O, Bernard M, Michel A, Menager C, Fattaccioli J (2018) Kinetically enhanced fabrication of homogeneous biomimetic and functional emulsion droplets. Langmuir 34(50):15319–15326. https://doi.org/10.1021/acs.langmuir. 8b02721

Chapter 23 Quantifying Immune Cell Force Generation Using Traction Force Microscopy Marcel Issler, Huw Colin-York, and Marco Fritzsche Abstract Immune cells rely on the generation of mechanical force to carry out their function. Consequently, there is a pressing need for quantitative methodologies that permit the probing of the spatio-temporal distribution of mechanical forces generated by immune cells. In this chapter, we provide a guide to quantify immune cell force generation using traction force microscopy (TFM), with a specific focus on its application to the study of the T-cell immunological synapse. Key words Mechanical forces, Traction force microscopy, Cytoskeleton, Immunological synapse, T cell activation, Mechanobiology, Biophysics

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Introduction The dynamics of the actin-myosin cytoskeleton and signaling events at the T-cell membrane during immunological synapse formation lead to mechanical force generation [1–4]. Whilst the precise role that mechanical force generation plays during early T-cell signaling and during the formation and continuation of signaling at the immunological synapse remains under investigation, there is an increasing reliance on methodologies that allow the quantification of force generation during immunological events such as synapse formation [2, 5]. Traction force microscopy (TFM) represents a powerful tool to quantify the spatio-temporal distribution of such forces and is rapidly becoming a standard method in immunological research [6, 7]. As such, we provide a traction force microscopy protocol specifically designed to study the mechanical forces generated by T-cells during immunological synapse formation. By observing in real-time the deformation of a planar elastic gel in contact with a single cell, or monolayer, TFM permits the spatiotemporal mapping of the forces induced by the cellular contact. Typically, a thin layer of elastic gel, either a hydrogel such as

Cosima T. Baldari and Michael L. Dustin (eds.), The Immune Synapse: Methods and Protocols, Methods in Molecular Biology, vol. 2654, https://doi.org/10.1007/978-1-0716-3135-5_23, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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polyacrylamide (PAA) or an elastomer such as polydimethylsiloxane (PDMS) is formed on a glass coverslip. Fluorescent beads are copolymerized within the gel, serving as fiducial markers and the top surface of the gel is functionalized with molecules that facilitate cell attachment. When a cell in contact with the gel applies a mechanical force, the gel surface is displaced. Imaging the displacement of the fluorescent beads within the gel leads to a spatial map of the distribution of applied forces. Because the mechanical properties of the elastic material are well known, the 2D distribution of gel displacements can be reconstructed into a two-dimensional (2D) map of cellular mechanical force generation [6, 7]. The quality of the force measurement using TFM depends on several factors, including substrate stiffness, bead distribution, surface functionalization, fluorescent imaging quality, image analysis, and force reconstruction. In this protocol, we detail the necessary conditions to achieve high-quality quantification of the mechanical forces generated by T-cells during immune synapse formation. We will focus on the specific details of experiments using immune cells, which present a challenge to TFM, and how these can be overcome.

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Materials Prepare all solutions using ultrapure water (Milli-Q or equivalent) and analytical grade reagents. Prepare and store all reagents at room temperature (RT) unless indicated otherwise. Diligently follow all waste disposal regulations when disposing waste materials.

2.1 Cover Glass Preparation

1. Sharp tweezers. 2. Top cover glass: 12 mm #1.5 borosilicate glass. 3. Bottom glass: 18 mm #1.5 borosilicate glass coverslips suitable for high-resolution microscopy. 4. Plasma cleaner. 5. 0.5% vol/vol 3-Aminopropyltrimethoxysilane (APTES) (Sigma 281778-5ML, or equivalent) solution in Milli-Q water. 6. 0.5% vol/vol glutaraldehyde solution in water.

2.2 Gel Preparation and Polymerization

1. 40% acrylamide stock (Sigma A4058-100ML, or equivalent). 2. 2% bis-acrylamide stock (Sigma M1533-25ML, or equivalent). 3. Acrylic acid (Sigma 147230-5G, or equivalent). 4. N,N,N,N0 -tetramethyl-ethylenediamine T9281-25ML, or equivalent).

(TEMED)

(Sigma

5. Ammonium persulfate (APS) (Sigma A3678, or equivalent).

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6. Fluorescent beads, e.g., Dark Red FluoSpheres™ CarboxylateModified Microspheres 0.2 μm, (Thermo F8807, or equivalent). 7. N2 gas. 8. Phosphate-buffered saline (PBS): 1 mM Potassium Phosphate monobasic, 155 mM sodium chloride, 3 mM sodium phosphate dibasic. 9. 10 M NaOH in water. 2.3 Gel Functionalization

1. 100 mM Tris–HCl-buffer, pH 8. 2. PBS adjusted to pH 8 with NaOH. 3. 1-Ethyl-3-(3-dimethylaminopropyl)carbodiimide (EDC) powder (Sigma 341006-5GM, or equivalent). 4. N-Hydroxysuccinimide (NHS) powder (Sigma 130672-5G, or equivalent). 5. 100 mM N-morpholinoethanesulfonic acid (MES) buffer, pH 6. 6. 0.1% poly-L-lysine solution. 7. Anti-CD3 (OKT3)—e.g., Biolegend Ultra Leaf 317347 or equivalent. 8. Anti-CD28—e.g., Biolegend Ultra Leaf 302943 or equivalent.

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TFM Acquisition

1. T cells, Jurkat Clone E-6.1. 2. Leibovitz’s L-15 Medium, no phenol red (Thermo 21083027 or equivalent). 3. Culture media: RPMI-1640 with 10% fetal bovine serum (FBS), 1% penicillin-streptomycin, 1% sodium pyruvate, 1% 1 M 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES), 1% L-Glutamine.AQ1

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Methods In the following, we present the steps necessary to produce a protein functionalized soft PAA hydrogel capable of quantifying forces generated by T-cells during immune synapse formation. In addition, we present a brief guide to the acquisition and analysis of TFM data generated by Jurkat T-cells on an anti-CD3/CD28 functionalized 672  95 Pa PAA gel. PAA hydrogels represent a good choice to measure immune cell force generation for several reasons, (i) they can be produced with low elastic moduli (1 min and collect the supernatant (see Note 20). 3. Resuspend conjugates in 1 mL of CSK buffer and agitate gently up and down (see Note 21). 4. Place the tubes back in the magnetic separation rack, remove supernatant, and repeat the wash as in steps 2 and 3 above. 5. Resuspend the samples in 1 mL of CSK buffer +0.5% Triton X-100 supplemented with protease inhibitor tablets according to the manufacturer’s recommendation to permeabilize cell membrane and help preserving the crosslinked proteins. 6. Put aside a sample of 5 μL of the suspension into a 96 well plate containing 95 μL of PBS. When there is a suitable break in the protocol, to visualize conjugate formation in washed samples, image the sample in the 96 well plate using an inverted brightfield microscope (such as EVOS) (Fig. 2b). 7. In order to remove the cell bodies from the beads, sonicate the samples in a sonicator using a program optimized beforehand to the samples (see Note 22). 8. After sonication keep samples on ice. 9. Put aside a sample of 5 μL of the suspension into a 96 well plate containing 95 μL of PBS. When there is a suitable break in the protocol, to visualize cell body detachment from beads, image the sample in the 96 well plate using an inverted brightfield microscope (such as EVOS) (Fig. 2c). 10. Place the tubes in the magnetic separation rack for >1 min and remove the supernatant (see Note 23). 11. Add 1 mL of CSK buffer +0.5% Triton X-100 supplemented with protease inhibitor tablets and agitate gently up and down. 12. Repeat the steps 10 and 11 to wash 5 more times. 13. Put aside a sample of 5 μL of the suspension into a 96 well plate containing 95 μL of PBS. When there is a suitable break in the protocol, to visualize successful washing of the beads and better evaluate the success of sonication, and image the sample in the 96 well plate using an inverted brightfield microscope (such as EVOS) (Fig. 2d).

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Fig. 2 Monitoring of the conjugate formation and sonication efficiency Conjugation using 25 × 106 mouse primary B cells and 25 × 106 magnetic beads coated with activatory or non-activatory antibodies was performed as described. Inverted brightfield microscopy was used to image (a) conjugates after adding the DTBP crosslinker at 15 min after the start of the conjugation, (b) after washing away the unbound cells, (c) after sonication, and (d) after washing off the cell bodies and debris. The results are representative of a set of 12 experiments

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14. Remove the supernatant using the magnet. 15. Add 20 μL of 2× Laemmli buffer with 5% β-mercaptoethanol and incubate the sample tubes for 30 min at 70 °C thermoshaker (1000 rpm) to denature the proteins and elute the from the beads. 16. Cleave the DTBP crosslinker by incubating the sample further for 10 min at 95 °C on a thermo-shaker (1000 rpm) (see Note 24). 17. Separate the beads using the magnet and gently transfer the supernatants, containing the extracted immune synapse or control adhesion proteins, into new Eppendorf tubes. These eluted samples can be stored at -20 °C for several days (see Note 25). 3.5

In-Gel Digestion

1. Run the eluted samples on a gradient polyacrylamide gel suitable for a wide range of polypeptides for 1 cm (approximately 20 min). 2. Stain the gel with reversible zinc staining, according to the manufacturer’s indications, for protein quantification (see Note 26). 3. Excise the zinc-stained gel bands with a clean scalpel, split in three pieces and place them in three tubes, Eppendorf® Protein LoBind, or equivalent. 4. Chop each gel piece further into smaller pieces inside the Eppendorf (see Note 27). 5. Destain the gel pieces with 1 mL of Tris-glycine pH 8 until Zinc Reversible Stain disappears (see Note 26). 6. Wash the gel pieces with 1 mL of Milli-Q water for 10 min with gentle rotation and discard supernatant. 7. Repeat the wash two more times as described in step 6. 8. Shrink and dehydrate the gel pieces by covering the gel pieces with 150 μL of ACN. 9. Wait until the gel pieces become white (about 5–10 min) and remove all ACN. 10. To reduce disulphide bonds, rehydrate the gel pieces in 150 μL of 20 mM DTT for 30 min at 56 °C. Remove the excess liquid if needed (see Note 28). 11. Repeat shrinking and dehydration of the gel pieces as described in step 8 of this section. 12. Rehydrate the gel pieces by adding 150 μL of 55 mM IAA for 20 min in the dark RT to block reoxidation of disulphide bonds. Remove the excess liquid if needed (see Note 28).

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13. Wash the gel pieces two times with 100 μL of 100 mM ammonium bicarbonate. 14. Repeat shrinking and dehydration of the gel pieces as described in step 8 of this section. 15. To start enzymatic digestion, add 75 μL of 0.02 μg/μL trypsin solution and allow it to absorb to the gel pieces for 20 min on ice (see Note 29). 16. After 20 min, add about 150 μL solution containing 40 mM NH4HCO3/10% ACN to completely cover the gel pieces and keep them wet during enzymatic cleavage (see Note 30). 17. Incubate the samples for up to 18 h at 37 °C. The incubation with trypsin generates the peptides for mass spectrometry analysis. 18. Add about 225 μL ACN (equal volume with the digestion mixture used in steps 15 and 16), vortex and incubate 15 min at 37 °C. 19. Collect the supernatant to another Eppendorf tube and repeat the extraction with 150 μL of solution containing 50% ACN / 5% Formic Acid. 20. Combine the supernatants containing the eluted peptides and dry the samples in a vacuum concentrator (see Note 31). 21. Store the dried peptides at -20 °C. 22. Immediately prior to mass spectrometry analysis, dissolve the peptides in 10 μL 2% Formic Acid by vortexing, incubate at 37 °C for 15 min and vortex again (see Note 32).

4

Notes 1. Ideally, the buffers should be prepared fresh but can also be stored at +4 °C for up to 1 month. 2. We use mice at age of 2–3 months. 3. CSK buffer can be stored at -20 °C for a few months. Store CSK without Triton and add Triton every time before use. 4. To avoid plasticizer contamination, which can be detrimental to mass spectrometry, tubes made of high-quality polypropylene must be used. 5. For good coating efficiency, we have determined that a maximum of 20 × 106 magnetic beads should be coated per one 1.5 mL Eppendorf tube. 6. During the coating of the beads, there are many washing steps where some beads are lost. To account for that, in our experimental setup, 40 × 106 beads were initially prepared per

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condition, in two different Eppendorf tubes. This ensures the recovery of a minimum of 25 × 106 coated beads for conjugation with cells. 7. BSA is added to aid the orientation and presentation of ligands. 8. Based on the manual provided for tosyl-activated Dynabeads by the manufacturer, maximal chemical binding is achieved after 16–24 h at 37 °C. Coupling at 18–25 °C (room temperature) would require an incubation time period longer than 48 h to obtain the same degree of chemical binding. Consider if your ligand is temperature-sensitive in order to determine the right temperature and time. 9. BS3 is moisture sensitive. Equilibrate at RT for 30 min before opening the vial to avoid condensation. 10. BS3 is an amine-to-amine crosslinker. The addition of BS3 helps to avoid leakage of the bead-bound antibodies to the final adhesion protein elutes, subject to mass spectrometry. 11. Tris in the Washing Buffer 3 deactivates remaining free tosyl groups, as well as quenches the BS3 crosslinker. 12. Antibody-coated beads may typically be stored in Washing Buffer 2 (optionally containing 0.02% sodium azide) at 2–8 ° C for months. The stability of coated beads with other ligands should be determined individually. For example, beads coated with fibronectin do not store for long. 13. During the coating steps, beads may be lost by suction or other errors, so an accurate count of the number of final beads is recommended. 14. The purity (for example, surface staining for B220 and IgM) and viability (for example, Zombie Violet) of the isolated B cells should be checked by flow cytometry routinely after each isolation. 15. The volume and amount of cells were optimized to ensure the optimal formation of single conjugates (single cell:bead contacts). It may vary depending on the experiment setup, cells, coating ligand, and magnetic beads used. 16. The ratio of cells to beads used is 1:1. This ratio provided a balanced output of single conjugate formation compared to other ratios tested, but again, this may vary depending on the experiment setup, cells, coating ligand, and magnetic beads used. 17. For the experimental procedure described, the activation timepoint used was 15 min. This can be changed depending on the experimental setup. 18. DTBP crosslinker is a membrane-permeable reversible crosslinker. DTBP has been considered a powerful tool to stabilize

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protein-protein interactions in different biological settings. The chosen DTBP conditions (2.5 mM, 5 min) were determined by a minimal concentration and time that yielded sufficient protein retrieval. Depending on the given experimental application, incubation time may be adjusted to fit the reader’s criteria. 19. Conjugation with activatory ligand was determined to be more effective than conjugation with non-activatory ligand. 20. The supernatant collected is stored in a new refrigerated Eppendorf. This one contains the cells not bound to any beads and can be used to give a rough estimate of the number of conjugates formed. In our experiments, the activatory ligand beads have approximately 70% conjugation rate, while the non-activatory ligand beads have approximately a 50% conjugation rate. 21. The magnetic separation rack used has two components: a magnetic band and a rack for test tubes. When agitating gently to homogenize conjugates in the CSK buffer, the tube rack is separated from the magnet and rotated slowly up and down. Pipetting conjugates to homogenize was deemed too harmful for the samples. 22. Sonication is required to remove the cell bodies from the beads, leaving only the crosslinked proteins at the adhesion structures for elution. This is a critical step that is sensitive to various parameters including the concentration of conjugates and efficiency of the crosslinking. Different sonication programs (time and intensity) bring different results and the best settings for each experimental procedure should be optimized beforehand. Our example protocol with our sonicator had settings of HIGH, 30″ on/30″ off, for 5 cycles. 23. Washing the beads should be performed with extra care to avoid any loss of weakly crosslinked proteins from the crosslinked immune synapse complexes. 24. For effective DTBP crosslinker cleavage, an incubation with 2× Laemmli buffer with 5% β-mercaptoethanol for 10 min with a temperature higher than 90 °C is required. 25. When preparing solutions and handling your samples keep in mind that the most common keratin contaminations, causing problems in mass spectrometry, arise from dust, hair, or skin. 26. Zinc staining is a sensitive and fast method for protein detection in polyacrylamide gels immediately after electrophoresis. Here, it is used also to quantify the loaded amount of eluted proteins from the different ligand-coated beads using whole cell lysates (WCL) with known protein concentration as standards (in our experimental setup, 1 and 3 μg of WCL was

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used). Ideally, matching amounts of protein should be submitted for mass spectrometry analysis for each sample. Additionally, the dye can be removed easily for subsequent sample preparation and is compatible with mass spectrometry analysis. 27. The volumes detailed here are calculated for our experimental setup. Gel lanes were run for a length of 1 cm and cut into three pieces. Each gel piece was then placed in a different Eppendorf and further chopped into smaller pieces. Too big gel pieces lead into reduced peptide recovery, while smaller pieces can be discarded during pipetting steps, thus, the user should consider the aimed final size of the gel pieces beforehand and perform chopping similarly in all the samples. 28. On the one hand, DTT acts as a reducing agent able to convert cysteine disulphide bonds into free sulfhydryl groups. On the other hand, IAA is an alkylating agent that reacts with free sulfhydryl groups of cysteine to form S-carboxyamidomethylcysteine, which cannot be reoxidized to form disulphide bonds. In this conformation, trypsin has maximum access to the cleavage sites within the proteins. 29. Gel pieces should be completely saturated with trypsin solution (when saturated, the gel pieces will be transparent). More trypsin solution should be added if gel pieces are not initially covered. Trypsin solution diffuses into the gel and cleaves the proteins into peptides. After 20 min on ice, gel pieces should be transparent. If not, more trypsin solution should be added. 30. The presence of low amounts of ACN (10% v/v) enhances trypsin activity by accelerating and improving the cleavage of the proteins during tryptic digestion. 31. We have used Medium Capacity Integrated Vacuum Concentrator System (#SPD1030A-115, Thermo Fisher Scientific) with 45 °C heating for 20 min and maximum vacuum pressure of 5.1, using manual run. Other vacuum centrifuges with comparable settings can be used. 32. The complexity of the protocol correlates with the fine-tuning of the conjugation (cells + beads). Conjugation efficiency with the non-activatory controls was found significantly lower than the conjugation with activatory ligand coated beads, which generated almost 50% more conjugates. To enable a direct comparison between conditions, the same amount of protein should be analyzed by mass spectrometry. Inherently, we recommend to assess the amount of eluted protein obtained from the tested conditions and compensate for that difference in future conjugation assays. Moreover, sonication settings have to be actively tested for efficient sonication without much protein loss, which presents a glaring challenge since only limited quantities of protein were retrieved for the mass

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spectrometry analysis (in the range of 2–4 μg per sample). Hence, if higher protein quantities are warranted, optimization of the protocol for higher numbers of conjugates should be performed.

Acknowledgments We thank Dr. Alexandre Carisey (St Jude’s Research Hospital, Memphis, USA) for his generous help with setting up the protocol. We also acknowledge Prof. Jordan Orange and Dr. Anna Meyer for their pioneering efforts in the immune synapse isolation in NK cells (Department of Pediatrics, Vagelos College of Physicians and Surgeons, Columbia University Irving Medical Center, New York, New York, USA). Turku Proteomics Facility of Turku Bioscience and Biocenter Finland are acknowledged for providing the research infrastructures. This work was supported by the Academy of Finland (decision number: 339810 to PKM), Sigrid Juselius foundation (to PKM), Erasmus+ program (to DMC), the Finnish Cultural Foundation (Suomen Kulttuurirahasto; to SHP), and by InFLAMES Flagship Programme of the Academy of Finland (decision number: 337530). References 1. Marshall JS, Warrington R, Watson W, Kim HL (2018) An introduction to immunology and immunopathology. Allergy Asthma Clin Immunol 14:49. https://doi.org/10.1186/ s13223-018-0278-1 2. Nicholson LB (2016) The immune system. Essays Biochem 60:275–301. https://doi. org/10.1042/EBC20160017 3. Batista FD, Iber D, Neuberger MS (2001) B cells acquire antigen from target cells after synapse formation. Nature 411:489–494. https:// doi.org/10.1038/35078099 4. Kuokkanen E, Sˇusˇtar V, Mattila PK (2015) Molecular control of B cell activation and immunological synapse formation. Traffic 16: 311–326. https://doi.org/10.1111/tra. 12257 5. Batista FD, Harwood NE (2009) The who, how and where of antigen presentation to B cells. Nat Rev Immunol 9:15–27. https://doi. org/10.1038/nri2454 6. Carrasco YR, Batista FD (2007) B cells acquire particulate antigen in a macrophage-rich area at the boundary between the follicle and the subcapsular sinus of the lymph node. Immunity 27:160–171. https://doi.org/10.1016/j. immuni.2007.06.007

7. Avalos AM, Ploegh H (2014) Early BCR events and antigen capture, processing, and loading on MHC class II on B cells. Front Immunol 5:92. https://doi.org/10.3389/ FIMMU.2014.00092 8. Hou P, Araujo E, Zhao T, Zhang M, Massenburg D, Veselits M, Doyle C, Dinner AR, Clark MR (2006) B cell antigen receptor signaling and internalization are mutually exclusive events. PLoS Biol 4:1147–1158. https://doi.org/10.1371/journal.pbio. 0040200 9. Yuseff MI, Reversat A, Lankar D, Diaz J, Fanget I, Pierobon P, Randrian V, Larochette N, Vascotto F, Desdouets C, Jauffred B, Bellaiche Y, Gasman S, Darchen F, Desnos C, Lennon-Dume´nil AM (2011) Polarized secretion of lysosomes at the B cell synapse couples antigen extraction to processing and presentation. Immunity 35:361–374. https://doi.org/10.1016/J.IMMUNI.2011. 07.008 10. del Valle Batalla F, Lennon-Dumenil AM, Yuseff MI (2018) Tuning B cell responses to antigens by cell polarity and membrane trafficking. Mol Immunol 101:140–145. https://doi. org/10.1016/j.molimm.2018.06.013

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11. Tolar P (2017) Cytoskeletal control of B cell responses to antigens. Nat Rev Immunol 1710(17):621–634. https://doi.org/10. 1038/nri.2017.67 12. Mattila PK, Batista FD, Treanor B (2016) Dynamics of the actin cytoskeleton mediates receptor cross talk: an emerging concept in tuning receptor signaling. J Cell Biol 212:267– 280. https://doi.org/10.1083/jcb.201504137 13. Porat-Shliom N, Milberg O, Masedunskas A, Weigert R (2013) Multiple roles for the actin cytoskeleton during regulated exocytosis. Cell Mol Life Sci 70:2099. https://doi.org/10. 1007/S00018-012-1156-5 14. Mattila PK, Feest C, Depoil D, Treanor B, Montaner B, Otipoby KL, Carter R, Justement LB, Bruckbauer A, Batista FD (2013) The actin and Tetraspanin networks organize receptor nanoclusters to regulate B cell receptormediated signaling. Immunity 38:461–474. https://doi.org/10.1016/j.immuni.2012. 11.019 15. Liu C (2018) B Cell receptor signaling. Humana Press, p 1707. https://doi.org/10. 1007/978-1-4939-7474-0 16. Wang JC, Bolger-Munro M, Gold MR (2018) Imaging the interactions between B cells and antigen-presenting cells. Methods Mol Biol 1707:131–161. https://doi.org/10.1007/ 978-1-4939-7474-0_10/COVER/ 17. Raybould MIJ, Rees AR, Deane CM (2021) Current strategies for detecting functional convergence across B-cell receptor repertoires. https://doi.org/10.1080/19420862.2021. 1996732 18. Breden F, Luning Prak ET, Peters B, Rubelt F, Schramm CA, Busse CE, Vander Heiden JA,

Christley S, Bukhari SAC, Thorogood A, Matsen FA, Wine Y, Laserson U, Klatzmann D, Douek DC, Lefranc MP, Collins AM, Bubela T, Kleinstein SH, Watson CT, Cowell LG, Scott JK, Kepler TB (2017) Reproducibility and reuse of adaptive immune receptor repertoire data. Front Immunol 8:1418. https:// doi.org/10.3389/FIMMU.2017.01418/ BIBTEX 19. Awoniyi LO, Sˇusˇtar V, Herna´ndez-Pe´rez S, Vainio M, Sarapulov AV, Petrov P, Mattila PK (2020) APEX2 proximity biotinylation reveals protein dynamics triggered by B cell receptor activation. bioRxiv 2020.09.29.318766. https://doi.org/10.1101/ 2020.09.29.318766 20. Satpathy S, Wagner SA, Beli P, Gupta R, Kristiansen TA, Malinova D, Francavilla C, Tolar P, Bishop GA, Hostager BS, Choudhary C (2015) Systems-wide analysis of BCR signalosomes and downstream phosphorylation and ubiquitylation. Mol Syst Biol 11:810. https:// doi.org/10.15252/msb.20145880 21. Cunha DM, Sara Hernandez-Perez S, Awoniyi LO, Carisey AF, Jacquemet G, Mattila PK (2023) Proteomic profiling of isolated immune synapses from primary mouse B cells. bioRxiv https://doi.org/10.1101/2023.02.23. 529674 22. Jones MC, Humphries JD, Byron A, MillonFre´millon A, Robertson J, Paul NR, Ng DHJ, Askari JA, Humphries MJ (2015) Isolation of integrin-based adhesion complexes. Curr Protoc Cell Biol 2015:9.8.1–9.8.15. https://doi. org/10.1002/0471143030.cb0908s66

Chapter 26 Analyzing Single Cell Secretions by “Shadow Imaging” Ashley R. Ambrose, Khodor S. Hazime, and Daniel M. Davis Abstract Here, we describe a method, which we term “shadow imaging,” to analyze the secretions of individual cells at immune synapses or other cell contacts. Following immune synapse formation and cellular activation on ligand-rich slides, the position of each cell is recorded using a pulsed immunofluorescence stain against the proteins on the ligand-rich slide surface. The pulsed stain does not penetrate the synaptic cleft, resulting in an unlabeled region or “shadow” beneath cells that is retained following cellular detachment. The secreted components, such as perforin, exosomes, or other types of extracellular vesicles, are retained on the slide and can be analyzed on a single-cell basis using immunofluorescence. The ability to identify single cells secreting different combinations of particles, proteins, and vesicles enables us to better understand the heterogeneity in immune cell secretions and can be used as a novel approach for phenotyping cell populations. Key words Immune synapse, Secretion, Extracellular vesicles, Cytotoxicity, Single-cell analysis, Bioimaging, Immunofluorescence

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Introduction Lymphocytes, such as cytotoxic T lymphocytes (CTLs) and natural killer (NK) cells, form highly organized contacts when encountering other cells such as antigen-presenting cells, virally infected cells, or cancerous cells. This area of contact is known as the immune synapse and is a tightly regulated platform for intercellular communication which is initially defined and organized by receptor-ligand interactions [1–4]. A variety of effector molecules are exchanged between conjugated cells in a context dependent manner. For example CTL and NK cells that form an immune synapse with cancerous or virally infected cells, polarize toward their target, and release cytotoxic granules [5–7]. These granules contain perforin, a pore-forming protein, and granzyme B that induces apoptosis in target cells [8, 9]. Perforin and granzyme B can be secreted

Ashley R. Ambrose and Khodor S. Hazime contributed equally. Cosima T. Baldari and Michael L. Dustin (eds.), The Immune Synapse: Methods and Protocols, Methods in Molecular Biology, vol. 2654, https://doi.org/10.1007/978-1-0716-3135-5_26, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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as a complex with thrombospondin-1 (TSP-1) in specialized structures termed supramolecular attack particles (SMAPs) [10–12]. In addition to these cytotoxic molecules, lymphocytes have been shown to secrete cytokines [13, 14], extracellular vesicles [15–17], and other proteins across the synapse [18]. Even viral particles can traffic across synapses, too [19, 20]. Methods have been established that can analyze the secretions from a bulk population of cells by looking at individual molecules using techniques such as ELISA or mass spectrometry. However, analyzing the secretions of single cells has been challenging, and this is of particular importance at the immune synapse. Microfluidic devices have been developed that can measure the extracellular vesicles released by single cells, but this poorly recapitulates the immune synapse and provides low-resolution information [21, 22]. Here, we demonstrate a method that enables the identification and capture of synaptic secretions from single cells, enabling a greater understanding of the information transferred at individual immune synapses. In this example, we apply it to observing perforin and extracellular vesicle secretion from NK cells activated on glass slides coated with ICAM-1 alongside activating ligands. Understanding the composition of the diverse milieu of effector molecules and how this varies from cell to cell can provide insights into the nuanced intercellular communication at the immune synapse.

2 2.1

Materials Disposables

1. T75 flask; Cat # CC7682-4175; CytoOne or equivalent. 2. Pasteur pipette; Cat # E1414-0311; Starlab or equivalent. 3. 24-well Plates with Lid, flat bottom; Cat # CC7682-7524; CytoOne or equivalent. 4. Automatic pipette and serological pipettes (5 mL, Cat # 86.1253.001; 10 mL, Cat # 86.1254.001 and 25 mL, Cat # 86.1685.001; all from Sarstedt or equivalent). 5. LS column; Cat # 130-042-401; Miltenyi Biotec or equivalent. 6. 8-chambered LAB-TEK II glass slides, Coverglass #1.5; Cat #155409; Nunc or equivalent (see Note 1). 7. 0.22 μm Syringe Filter, sterile; Starlab Cat # E4780-1223 or equivalent. 8. 50 and 15 mL sterile screw cap conical centrifuge tubes (Falcon or equivalent).

2.2

Equipment

1. Cell culture incubator with humidified atmosphere (37
C, 5% CO2). 2. Sterile cell culture laminar flow hood safety level II.

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3. Haemocytometer, Neubauer, 0.1 mm, 0.0025 mm2. 4. GT2 Benchtop Centrifuge; Fisherbrand; 15818722 or equivalent. 5. Sliding door drying cabinet—model # 322/0103/00; serial # R1586713 or equivalent. 6. Eclipse Ti inverted Microscope with total internal reflection fluorescence illuminator and objective—NIKON or equivalent. 7. DMIL light Microscope—Leica; 090-135-001 or equivalent. 2.3

Reagents

1. Phenol red free RPMI; Cat # R7509; Sigma or equivalent. 2. Ficoll-Paque Plus; Cat # 17144003; Cytiva or equivalent. 3. Red Blood cell lysis buffer; Cat # R7757-100 mL; Sigma or equivalent. 4. 0.4% Trypan Blue Stain, Cat# 17-942E; BioWhittaker or equivalent. 5. Poly-L-Lysine; Cat # P8920; Sigma or equivalent. 6. Clone media: Dulbecco’s Modified Eagle Medium [DMEM] medium; Cat # D6429, containing 30% F12 Ham; Cat # V6658, 10% human serum; Cat #, M7145, 1% Non-essential amino acids; Cat # S8636, 2 mM L-glutamine; Cat # G7513, 50 U/mL penicillin/streptomycin; Cat # P4333, all from Sigma-Aldrich or equivalent, 50 μM 2-mercaptoethanol; Cat # 31350-010; 1 mM sodium pyruvate; Cat # 11360070; Gibco or equivalent. 7. Bovine serum albumin (BSA); Cat # A9418, Sigma-Aldrich or equivalent. 8. Non-enzymatic cell-dissociation solution, composed of EDTA, glycerol, and sodium citrate (the precise formulation of this reagent is proprietary); Cat # C5789, Sigma-Aldrich or equivalent. 9. Human serum; Cat # H4522; Sigma-Aldrich or equivalent. 10. Phosphate-buffered saline (PBS): containing 0.2 mg/mL KCl, 0.2 mg/mL KH2PO4, 8 mg/mL NaCl, and 1.15 mg/mL Na2PO4 (anhydrous) pH 7.4; Cat # D8537, Sigma-Aldrich or equivalent. 11. Blocking solution: PBS/3% BSA/1% Human Serum (ThermoFisher or equivalent), filter sterilized using a 0.22 μM filter. 12. MACS buffer: 0.5% BSA, 2 mM EDTA (Cat # E5134; Sigma) in PBS. 13. NK cell negative isolation kit; Cat # 130-092; Miltenyi Biotec. 14. Recombinant human IL-2; Roche or equivalent. 15. His-ICAM-1, Sino Biological or equivalent.

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16. His-MICA, Peak Proteins Ltd. or equivalent. 17. Anti-ICAM-1 antibody, purchased already conjugated to Bv421 (Clone HCD54), BioLegend of equivalent. 18. Anti-CD63, purchased already conjugated to Alexa Fluor 647 (Clone H5C6), BioLegend or equivalent. 19. Anti-Perforin, purchased already conjugated to Alexa Fluor 488 (clone dG9), Biolegend or equivalent. 2.4 Biological Materials

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1. Plateletpheresis biproducts (leucocyte cones) from healthy donors were acquired from the National Blood Service under ethics license REC 05/Q0401/108 (University of Manchester, Manchester, UK). If these are not available whole blood can be substituted with some reduction in the yield of NK cells.

Methods

3.1 Primary NK Cell Isolation

1. Mix the contents of the leucocyte cone with 50 mL of phenol red free RPMI pre-heated to 37
C in a T75 flask. 2. Add 15 mL of Ficoll-Paque Plus to each of two 50 mL centrifuge tubes. 3. Add 2–3 mL of the leucocyte mixture gently and dropwise to each 50 mL centrifuge tube while holding it at a 45
angle using a Pasteur pipette. 4. Divide the remaining leucocyte mixture equally between the two tubes using 25 mL serological pipettes. 5. Centrifuge the tubes at 535  g for 40 min at room temperature with the brake on the centrifuge disabled to ensure the interface remains distinct during deceleration. 6. Carefully collect the layer of peripheral blood mononuclear cells (PBMCs) from above the Ficoll Paque cushion from each tube and transfer into a fresh 50 mL centrifuge tube using a Pasteur pipette. Collect all the cells with minimal Ficoll Paque solution. 7. Fill the tube to a total volume of 50 mL with phenol red free RPMI and centrifuge at 350  g for 10 min at room temperature. 8. Aspirate the supernatants taking care not to disrupt the PBMC pellets and resuspend the PBMC pellets in 50 mL phenol red free RPMI for each tube before a second centrifugation step of 350  g for 10 min at room temperature. 9. Aspirate the supernatant and resuspend the PBMC pellet in 10 mL of red blood cell lysis buffer and then mix with the second pellet before incubating for 5 min at room temperature.

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10. Stop the lysis reaction by adding 40 mL phenol red free RPMI and then pellet the PBMCs by centrifugation at 350  g for 10 min. 11. Aspirate the supernatant and resuspend the pellet in 10 mL phenol red free RPMI. 12. Count the PBMCs using a haemocytometer on a light microscope using a 20 objective. Mix 10 μL of cells with 40 μL phenol red free RPMI and 50 μL 0.4% trypan blue for counting. 13. Add 1.0  108 PBMCs to a 15 mL centrifuge tube and centrifuge at 350  g for 10 min at 4
C. 14. Aspirate the supernatant and resuspend the cells in 400 μL MACS buffer and 100 μL of the biotin-antibody cocktail from the NK cell negative selection kit and incubate at 4
C for 5 min. 15. Add 200 μL anti-biotin microbeads and 300 μL MACS buffer, mix well, and incubate at 4
C for 10 min. 16. Centrifuge at 350  g for 10 min at 4
C. 17. While centrifuging the cells, prepare an LS column in a magnetic stand and rinse with 3 mL MACS buffer, collecting flow through in a 15 mL centrifuge tube. 18. Aspirate the supernatant from the centrifuged cells and resuspend in 500 μL MACS buffer. 19. Place a fresh 15 mL centrifuge tube below the LS column and add the cell suspension to the prepared LS column. 20. Wash the column three times with 3 mL MACS buffer; the flow through in the 15 mL centrifuge tube contains the NK cells. 21. Centrifuge the NK cells at 350  g for 10 min at 4
C. 22. Aspirate supernatant and resuspend in 1 mL of 37
C NK cell clone media. 23. Count the NK cells with a haemocytometer on a light microscope using a 20 objective; mix 10 μL of cells with 10 μL 0.4% trypan blue for counting. Cells were then diluted to 1  106/mL with NK cell clone media. 24. Add IL-2 to a final concentration of 200 Units/mL. 25. Plate the NK cells in 24-well tissue culture plates, adding 1 mL of NK cells per well. 26. Incubate NK cells for 6 days at 37
C and 5% CO2 (see Note 2). 3.2 Sample Preparation

1. Incubate each well of an eight chambered glass slide with 200 μL of 0.01% PLL in ddH2O for 15 min at room temperature.

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2. Aspirate the PLL solution and wash each well with 500 μL ddH2O three times. 3. Place the slides in a drying cabinet and dry at 60
C for 1 h. 4. Incubate slides with 200 μL of PBS comprising 2.5 μg/mL His-ICAM-1, 2.5 μg/mL His-ICAM-1 plus 2.5 μg/mL His-MICA, or 2.5 μg/mL His-ICAM-1 plus 10 μg/mL antiNKp30 overnight at 4
C (see Notes 3 and 4). 5. Wash the slides three times with 500 μL PBS. 6. Incubate 1.0  105 NK cells in 200 μL NK cell clone media on coated slides for 1 h at 37
C in a 5% CO2 incubator (see Note 5). 7. Aspirate media and gently wash each well three times with 500 μL room temperature PBS so that NK cell attachment to the surface is not disrupted. 8. Stain slides with 200 μL of 5 μg/mL anti-ICAM-1-Bv421 in PBS for 1 min, and then immediately aspirate and wash three times with room temperature PBS. 9. Detach cells by addition of 200 μL of non-enzymatic celldissociation solution per well and incubation for 20 min at 37
C (see Note 6). 10. Aspirate the cell dissociation solution and wash each well three times with PBS to remove all cells. 11. Incubate the slides with 200 μL of blocking solution for 1 h at room temperature to block non-specific antibody binding site. 12. Stain the slides for 1 h at room temperature with 10 μg/mL anti-CD63-AF647 and 2.5 μg/mL anti-Perforin-AF488 in blocking solution (steps 6–12 are illustrated in Fig. 1) (see Note 7). 13. Wash the slide three times with PBS and image immediately. 3.3

Imaging Setup

1. Apply immersion oil to the 100 1.49NA objective and place the slide onto the inverted microscope and focus at the coverslip-media interface. 2. Check the alignment at the center of the well to ensure TIRF illumination. 3. Image the samples sequentially using 405, 488, and 647 nm laser lines in TIRF mode. 4. Acquire images using the same laser power and exposure settings for all conditions. A preliminary survey of the dimmest and brightest samples may be necessary to choose the best settings to avoid saturation. The precise settings will depend upon your system.

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Fig. 1 Shadow imaging protocol schematic. (a) NK cells were incubated on glass slides coated with 0.01% PLL followed by 2.5 μg/mL ICAM-1 plus 2.5 μg/mL MICA for 1 h at 37
C and 5% CO2 to trigger immune synapse formation, cellular activation and secretion of extracellular vesicles and proteins such as perforin and granzyme B. (b) Slides were pulse-stained with anti-ICAM-1 conjugated to Bv421 for 1 min, before 3 washes with PBS. (c) Cells were removed by incubation with non-enzymatic cell dissociation solution for 20 min at 37
C and 5% CO2. (d) Slides were blocked and then stained with fluorescent antibodies for proteins of interest

5. Acquire ~30–40 images per condition for each biological replicate. 6. Save datasets for analysis. 3.4

Data Analysis

1. Open datasets in ImageJ [23] equipped with the latest version of the Bio Formats plugin. 2. Create a composite image showing all three channels simultaneously.

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3. Identify the positions of cells using the shadow (405 nm) channel. Count the number of cells per field of view based on the number of shadows. Shadows which were only partially within the field of view were excluded from further analysis (see Notes 8 and 9). 4. Identify the cells secreting perforin or CD63 by measuring the fluorescence intensity of multiple dark (empty) shadows in each channel (AF488 and AF647), and calculate the mean value to use as a threshold. Those with a fluorescence intensity value higher than the threshold are considered to be cells that secrete the labelled protein (see Note 10). Example data is shown in Fig. 2.

4

Notes 1. It is possible to use coverslips instead of chamber slides for this assay. However, we recommend chambered slides as it is essential that staining with the anti-ICAM-1 antibody and the subsequent washing steps are fast. We find that this is easier to achieve in chambered slides. 2. For the experiments described here, NK cells were cultured for 6 days at 37
C with 5% CO2 following addition of 200 U/mL IL-2; this increases NK cell numbers, and after 6 days, they are in a resting state and ready for use in experiments. Protocols for expanding NK cells can vary the cytokines used or the time NK cells are rested for, and many labs have developed different protocols. 3. This assay can be readily adapted to achieve adherence and activation or inhibition of cells to trigger cellular secretions with a variety of ligands. In addition to the ligands demonstrated here, we have also utilized; B7-H6-Fc (Cat # 7144-B7050; R&D Systems), αNKp30 stimulating antibody (Cat # MAB18491; R&D Systems or 325204; Biolegend), and αNKG2A (Cat # MAB1059; R&D Systems) all in combination with ICAM-1 [11]. Theoretically, any of the surface ligands can be targeted by fluorescent antibodies to generate the shadows, but the concentrations and timing will require optimization. 4. To increase the physiological relevance of this assay we have previously used it in combination with planar lipid bilayers instead of glass slides [24]. This was successful, however, these bilayers were not mobile and further investigation would be required to determine the feasibility of generating shadows on mobile planar lipid bilayers. It is possible that ligand crosslinking with fluorescent antibodies could retain the shadows in place, for example.

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Fig. 2 Secretions from activated NK cells. (a) TIRF images showing cell shadows (cyan), secreted perforin (green), and secreted exosomes (magenta) following cell detachment. NK cells were incubated on slides coated with 2.5 μg/mL ICAM-1 alone (upper panel), 2.5 μg/mL ICAM-1 plus 10 μg/mL αNKp30 (middle panel) or 2.5 μg/mL ICAM-1 plus 2.5 μg/mL MICA (lower panel) for 1 h at 37
C / 5% CO2. (b–d) Quantification of the percentage of cells that secrete perforin (b), the intensity of secreted perforin (c), and the intensity of the extracellular vesicle marker, CD63 (d), from individual cells. Scale bar: 10 μm for all images; 5 μm for zoomed images. Statistical analysis was performed using Prism (GraphPad Software; v8.4.2) and significance was calculated using one-way Anova. Data are presented as Mean  SD. *** ¼ P  0.001, **** ¼ P  0.0001

5. This assay can be used to create shadows for a variety of cell types. The only requirements are that the cells adhere during the assay, thereby preventing ingress of the shadow-generating antibody, and that the cells can be readily detached. In addition to NK cells, we have also used shadow imaging to analyze the secretions of human macrophages [24].

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6. To our knowledge, the capture of proteins on PLL coated slides occurs through non-specific electrostatic interactions, which results in the capture of a broad range of secretions. While we have been able to detect a variety of different proteins, we cannot exclude the possibility that some proteins are unable to attach and therefore remain undetected with this assay. 7. Due to the non-specific protein capture of the PLL coated surface, it is possible to analyze a wide variety of proteins within shadows. We have previously detected membrane proteins of extracellular vesicles and secreted proteins. This assay can easily be adapted to stain for any protein of interest. In addition, it is possible to use the setup without shadow staining to isolate proteins for analysis by mass spectrometry to help identify potential targets of interest. 8. The quality of cellular adherence is a critical component of this assay. With both attachment that is too strong and too weak causing issues. If the cell is difficult to remove without the application of significant force, then it is difficult to know whether the proteins left behind are truly secreted or whether the shear forces from the washing tore the cells from the surface. In the case of weak adherence, this can be problematic for generating shadows in the staining that clearly define the locations of individual cells. A potential solution for both too weak and too strong attachments could be to optimize the ligand coating, particularly proteins that modulate binding via integrins. 9. When analyzing the shadow data, cells that share a border can cause issues. It is not always possible to determine whether there are two shadows or one when there is no gap between the shadows. It is best to incubate cells on a slide at a sufficiently low density that most cells attach to the slide as single cells; however, the optimal density for this will vary depending on the cell type used. When it is ambiguous whether there are one or two cells, it is best to exclude these cells from the analysis to ensure consistency. 10. Shadows for individual cells should be distinct with an absence of fluorescence and clear borders. However, there are several factors that can influence the quality of the shadows observed: (a) Cell motility—Not all cells form stable immune synapses and may instead form motile kinapses. This can cause problems whereby the detected secretions are not all localized within the shadows. This could be resolved by optimizing the ligand coating to ensure that cells form a more stable synapse or by using shorter incubations so that cells do not move on from the point at which they secrete.

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(b) Cell attachment—As mentioned in Note 7, cellular adherence is essential for the generation of clear shadows. If the difference between signal beneath cells and around cells is small or many cells are still attached, consider optimizing the ligand concentrations used, reducing the length of the incubation or changing the cell detachment strategy. (c) Fluorescent staining of ICAM-1—if the staining of ICAM-1, or your chosen ligand does not leave distinct shadows then optimizing the staining protocol may provide better results. Modifying the antibody concentration, the fluorophore used, for example, using quantum dots or adjusting the length of time the antibody incubates on the slide.

Acknowledgements This work was supported by a Wellcome Trust Investigator Award (110091/Z/15/Z), Bristol Myers Squibb, and the Manchester Collaborative Centre for Inflammation Research (funded by a Precompetitive Open-Innovation Award from GSK, AstraZeneca, and The University of Manchester). References 1. Davis DM, Chiu I, Fassett M, Cohen GB, Mandelboim O, Strominger JL (1999) The human natural killer cell immune synapse. Proc Natl Acad Sci U S A 96:15062–15067. https://doi.org/10.1073/pnas.96.26.15062 2. Grakoui A, Bromley SK, Sumen C, Davis MM, Shaw AS, Allen PM, Dustin ML (1999) The immunological synapse: a molecular machine controlling T cell activation. Science (80- ) 285:221–227. https://doi.org/10.1126/sci ence.285.5425.221 3. Monks CR, Freiberg BA, Kupfer H, Sciaky N, Kupfer A (1998) Three-dimensional segregation of supramolecular activation clusters in T cells. Nature 395:82–86. https://doi.org/10. 1038/25764 4. Stinchcombe JC, Bossi G, Booth S, Griffiths GM (2006) Centrosome polarization delivers secretory granules to the immunological synapse. Nature 443:462–465. https://doi.org/ 10.1038/nature05071 5. Krzewski K, Coligan JE (2012) Human NK cell lytic granules and regulation of their exocytosis. Front Immunol 3:1–16. https://doi. org/10.3389/fimmu.2012.00335

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Chapter 27 Exploiting the RUSH System to Study Lytic Granule Biogenesis in Cytotoxic T Lymphocytes Nagaja Capitani, Chiara Cassioli, Keerthana Ravichandran, and Cosima T. Baldari Abstract The Retention Using Selective Hooks (RUSH) system allows for the synchronized release of one or more proteins of interest from a donor endomembrane compartment, usually the endoplasmic reticulum, and the subsequent monitoring of their traffic toward acceptor compartments. Here we describe the RUSH system applied to cytotoxic T cells to characterize the biogenesis of lytic granules, using as a proof-of-concept granzyme B trafficking to this specialized compartment. Key words Intracellular trafficking, RUSH system, Cytotoxic granules, Granzyme B, Lytic granule biogenesis, Confocal microscopy, Spinning disk microscopy

1

Introduction Cytotoxic T lymphocytes (CTLs) are the T cell effectors specialized for the elimination of cells infected by intracellular pathogens as well as cancerous cells. CTLs exert their cytotoxic function through two well-characterized cytotoxic mechanisms: release of cytotoxic granules (CG) and Fas/FasL mediated apoptosis [1]. The first mechanism involves the release of cytotoxic molecules from CGs via exocytosis at the immunological synapse (IS), a specialized contact area that forms at the interface of the T cell with its cognate target cell [2, 3]. Upon TCR-mediated recognition of a cell presenting specific peptide associated with MHCI, CTLs rapidly polarize and reorganize their cytoskeleton to translocate the microtubule-organizing center (MTOC) toward the synaptic interface [4, 5]. MTOC docking beneath the IS is a key step in the formation of a lytic synapse as it ensures the microtubule-assisted directional transport of CGs that contain soluble cytolytic proteins, such as granzymes and perforin, that are eventually released into the synaptic cleft [6, 7]. The other mechanism by which CTLs exert

Cosima T. Baldari and Michael L. Dustin (eds.), The Immune Synapse: Methods and Protocols, Methods in Molecular Biology, vol. 2654, https://doi.org/10.1007/978-1-0716-3135-5_27, © The Author(s) 2023

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their killing activity is based on the FasL-dependent pathway. This mechanism involves cross-linking of the surface death receptor Fas on target cells induced by cell surface FasL expressed on CTLs. Fas/FasL interaction rapidly induces activation of the caspase cascade to promote target cell apoptosis [8]. A third mechanism of CTL-mediated killing has been recently reported by Balint and colleagues [9]. This is based on the synaptic release of a non-membranous type of cytotoxic particles, known as supramolecular attack particles (SMAPs) [9]. Proteomic and structural analysis of SMAPs revealed that together with canonical cytolytic components previously observed in CGs, such as granzyme B (GZMB) and perforin (Prf), this new class of particles is enriched in glycoproteins such as thrombospondin-1 (TSP-1) and galectin-1 (Gal1) that form a shell surrounding the cytotoxic core [9]. The recent in-depth characterization of CGs by Chang and colleagues has identified two populations of CGs, the “classical” single-core CGs (SCG) and the multicore CGs (MCG), which are the source of SMAPs [10]. The co-existence of SCGs and MCGs in CTLs and the fact that they are released independently supports the notion that their biogenesis and trafficking are governed by specific pathways. The development of a method to unravel these pathways will allow to achieve insight into the relationship between SCGs and MCGs and their interplay in CTL-mediated killing. In 2012, Boncompain and colleagues developed an assay known as Retention Using Selective Hooks (RUSH) to analyze the traffic of secretory proteins in eukaryotic cells [11]. This system is based on the simultaneous expression of two proteins: the hook used as “anchor” protein, and the reporter, the protein of interest whose trafficking will be analyzed. The hook is stably expressed in a donor compartment, usually the endoplasmic reticulum (ER), and the Golgi compartment. The hook is fused at its N- or C-terminal end with core streptavidin, which can be oriented toward the lumen of compartment or at the cytoplasmic face. The reporter is fused to the streptavidin-binding peptide (SBP) to allow its interaction with streptavidin on the hook protein and with a fluorescent protein (e.g., EGFP, mCherry) to visualize it within the cell. Once the RUSH construct is expressed by the cell, the reporter interacts with the hook in the donor compartment due to the streptavidin-SBP interaction and is thus retained in the donor compartment. When biotin is added to the culture medium, it efficiently binds streptavidin, displacing the SBP, and the reporter protein is released and can traffic toward its destination compartment, also known as acceptor compartment (Fig. 1). This system can be applied to more than one reporter, allowing to track the synchronized trafficking of two proteins fused to different fluorescent tags. Here we describe the application of the

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Fig. 1 RUSH system. The RUSH (Retention Using Selective Hooks) system is a two-phase assay. In the initial phase, known also as “retention phase,” the reporter protein is anchored in the donor compartment (ER in the example) by the hook due to the streptavidin-SBP (streptavidin-binding peptide) interaction. Addition of biotin in the culture medium introduces the second phase of the assay, known as “release phase.” The reporter protein is released from the hook as the result of displacement by biotin and it traffics, together with the associated fluorescent protein, to its acceptor compartment (lysosomes in the example)

RUSH system to the study of the biogenesis and intracellular trafficking of the CG components. As a proof-of-concept, we used the RUSH system to synchronize the traffic of granzyme B (GZMB), which is known to exploit the cation-independent mannose-6-phosphate pathway for specific targeting to CGs [12]. As a hook, we used the KDEL sequence, which retains the fused protein in the ER lumen [13]. To verify the correct localization to CGs, which are specialized lysosomes, we co-stained CTLs with the lysosomal marker LAMP-1 at the experiment endpoint. The results provide proof-of-concept that the RUSH system can be exploited to track the traffic of CG-associated proteins in CTLs. This paves the way to dissect the interplay of the pathways that regulate the biogenesis of SCGs and MCGs by synchronizing the trafficking of different CG components in CTLs co-transfected with RUSH constructs expressing the respective reporters.

2

Materials

2.1 Cloning of GZMB in a RUSH Construct

1. Commercial Addgene plasmid mCherry, Plasmid #65279).

(Str-KDEL_TNF-SBP-

2. Phusion DNA Polymerase. 3. 5× Phusion HF Buffer (ThermoFisher scientific or equivalent). 4. dNTP Mix (10 mM each).

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5. Primers containing restriction enzymes for amplification of the insert to be cloned in Addgene vector: Primer forward: TTGGCGCGCCATGCAACCAATCCTGC TTCTGCTGG Primer reverse: GAATTCCGGTAGCGTTTCATGG TTTTCTTTATCC 6. UltraPure DNase/RNase-Free Distilled Water. 7. Thermal cycler. 8. Restriction enzymes: FastDigest SgsI (AscI) and EcoRI (ThermoFisher scientific). 9. Agarose gel. 10. NucleoSpin Gel and PCR Clean-up Kit (Macherey-Nagel or similar). 11. UV-Vis spectrophotometer for μL volumes of nucleic acids (QIAxpert system (QIAGEN) and QIAxpert slides (QIAGEN) or similar). 12. Ligation: T4 DNA ligase 10× buffer and T4 DNA ligase. 13. E. coli thermocompetent cells (e.g., DH5α Competent Cells). 14. LB (Luria-Bertani) liquid medium (for each 950 mL of MilliQ H2O, add 10 g Tryptone, 10 g NaCl, and 5 g Yeast Extract) (see Note 1). 15. LB agar plates with 100 μg/mL ampicillin (see Note 2). 16. Petri dishes. 17. NucleoSpin Plasmid, Mini kit for plasmid DNA (MachereyNagel or similar). 18. NucleoBond Xtra Midi Plus EF, Midi kit for endotoxin-free plasmid DNA (Macherey-Nagel or similar). 2.2 CD8+ Cell Purification from Peripheral Blood and CTL Differentiation

1. RosetteSep Human CD8+ T cell Enrichment Cocktail (STEMCELL Technologies or similar methods of purification of untouched CD8+ T cells). 2. Lympholyte Cell Separation Media or equivalent. 3. Phosphate buffered saline (PBS) (Medicago AB), 1 tablet in 1000 mL of deionized water (0.14 M NaCl, 0.0027 M KCl, 0.010 M Phosphate buffer, pH 7.4), sterile filtered. 4. Bovine Calf Serum (BCS)—0.1 μm sterile filtered and low endotoxin. 5. PBS with 0.2% (v/v) BCS: 2 mL of BCS and 98 mL of PBS (Medicago AB, see point 3), keep sterile. 6. High speed temperature. 7. Transfer pipette.

centrifuge,

swing-bucket

rotors,

room

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8. R10 Medium: RPMI-1640 medium (Sigma-Aldrich) with 2 mM L-Glutamine and 20 mM HEPES, supplemented with 10% (v/v) BCS, 2 mg/mL penicillin, 1% (v/v) MEM Non-essential amino acids. 9. Recombinant human IL-2 IS, premium grade (Miltenyi Biotec) (see Note 3). 10. Dynabeads Human T-activator CD3/CD28 for T cell Expansion and Activation (ThermoFisher Scientific or equivalent) (see Note 4). 11. Magnetic particle concentrator MCP-6 (Dynal or equivalent). 12. PBS 0.1% (v/v) BCS: 1 mL of BCS and 99 mL of PBS, keep sterile. 13. 12-well cell culture plate with flat bottom. 2.3 CTLs Nucleofection with the RUSH-GZMB Construct

1. Amaxa T cell nucleofector kit (Lonza).

2.4 Evaluation of RUSH on Fixed Cells by Confocal Microscopy

1. CTLs transfected with RUSH-GZMB construct.

2. Nucleofector 2b Device (Lonza).

2. Cell culture plate, 96 well. 3. 4 mM biotin (stock solution) (see Note 5). 4. 10-well diagnostic microscope slides. 5. Absolute ethanol. 6. 0.1% (w/v) Poly-L-Lysine (PLL) stock solution in sterile water (Sigma-Aldrich or similar). 7. Hellendahl-type dish. 8. 3MM paper. 9. Fixation solution: 4% (v/v) paraformaldehyde (PFA) in PBS (see Note 6). 10. Permeabilization solution: PBS with 1% (w/v) BSA and 0.01% (v/v) Triton X-100]. 11. Primary antibodies: anti-RFP, rabbit polyclonal and anti-RFP mouse monoclonal (Rockland); anti-human CD107a (LAMP-1), mouse monoclonal (BioLegend), anti-CanX, rabbit polyclonal (Sigma-Aldrich). 12. Secondary fluorochrome-conjugated antibodies. 13. Slide mounting medium: 90% (v/v) glycerol in PBS. Coverslips 24 × 60 mm and conventional nail polish. 14. Confocal microscopy, Zeiss LSM700.

2.5 Evaluation of RUSH in Live Cells

1. CTLs transfected with RUSH-GZMB construct. 2. LysoTracker Blue DND-22 (ThermoFisher Scientific).

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3. 4 mM biotin (stock solution). 4. μ-Slide 8 Well Chamber Slide. 5. Medium RPMI 1640 no phenol red. 6. Spinning disk microscope with an inverted objective and a thermostated incubation chamber (CSU-W1-SoRa, Nikon, or equivalent).

3

Methods

3.1 Generation of the RUSH-GZMB Construct 3.1.1 GZMB Amplification by PCR

PCR amplification of GZMB is performed using as template a GZMB-mCherry home-made vector. PCR reaction mix (final volume 100 μL) (see Note 7): 20 μL 5× Phusion HF Buffer 2 μL 10 mM dNTPs 0.5 μM (final concentration) forward primer 0.5 μM (final concentration) reverse primer 50 ng template DNA 1 μL Phusion High–Fidelity DNA Polymerase Add H20 to 100 μL PCR protocol: Initial Denaturation

98 °C, 30 sec

Denaturation

98 °C, 10 sec

Annealing

68 °C, 30 sec

Extension

72 °C 15–30 s/kb

Final extension

72 °C, 7 min

9 > = > ;

35 cycles

Hold 4°C 5 μL of the PCR product is run on an agarose gel (see Note 8) to verify the quality of PCR amplification. The remaining part is purified using NucleoSpin Gel and PCR Clean-up Kit and quantified using QIAxpert system (see Note 9). 3.1.2 Insert and Vector Restriction Enzyme Digestion

Purified GZMB PCR product obtained in paragraph 3.1.1 and Addgene Plasmid #65279 are digested using the following protocol (see Note 10): 6 μL 10× FastDigest Buffer 2 μL FastDigest SgsI 2 μL FastDigest EcoRI

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template DNA (0.4 μg purified PCR product or 2 μg vector) Add H20 to 60 μL Digestion protocol: 30 min at 37 °C and 5 min at 80 °C The digested products are loaded onto agarose gels, the DNA fragments are excised from the gel and purified using NucleoSpin Gel and PCR Clean-up Kit and quantified using QIAxpert system. 3.1.3 DNA Ligation and Transformation

Digested inserts and vectors obtained in Subheading 3.1.2 are ligated using the following proportion ng of vector × kb size of insert insert × molar ratio = ng insert kb size of plasmid plasmid where the amount of vector for a ligation reaction is usually 50–100 ng and the molar ratio vector:insert varies depending on vector and insert size. In our ligation reactions we used 100 ng vector and a molar ratio of 3 (insert): 1 (vector) to obtain Str-KDEL_GZMB-SBP-mCherry (see Note 11). Assemble the following reaction in a sterile microcentrifuge tube (see Note 12): 100 ng vector DNA 50–100 ng insert DNA 1 μL 10× Buffer 1 μL T4 DNA Ligase Nuclease-free water to 10 μL Incubate the reaction at room temperature for 3 h, or 4 °C overnight, and inactivate the reaction by heating at 70 °C for 10 min. The ligation reaction obtained is transformed in E. coli DH5α Competent Cells following this procedure: 1. Thaw on ice one tube of DH5α cells. 2. Gently mix cells with the pipette tip and aliquot 50 μL cells for each transformation reaction in a 1.5 mL microcentrifuge tube. 3. Add the ligated DNA (10 μL) to the cells and mix gently. 4. Incubate tubes on ice for 30 min. 5. Heat shock cells for 30 s in a 42 °C water bath. 6. Place back tubes on ice for 2 min. 7. Add 850 μL pre-warmed LB medium. 8. Incubate tubes in shaking incubator at 37 °C for 1 h at 225 rpm. 9. Spread 20 μL to 200 μL from each transformation on pre-warmed selective plates (see Notes 13 and 14). 10. Incubate plates overnight at 37 °C.

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The next day pick single colonies and inoculate into LB medium. Incubate at 37 °C in shaking incubator at 225 rpm for 18–20 h. 3.1.4 Plasmid DNA Preparation and Colony Screening

3.2 CD8+ Cell Purification from Peripheral Blood and CTL Differentiation

Bacterial cells obtained from single colonies are used to extract the plasmid using the NucleoSpin Plasmid Mini kit for plasmid DNA (Macherey-Nagel or similar). Plasmid DNA obtained (~200–300 ng) is digested with restriction enzymes as described in Subheading 3.1.2 and loaded onto agarose gels to verify the quality of ligated DNA. Plasmid DNA containing the correct insert is selected and a Midiprep is prepared from it using the NucleoBond Xtra Midi Plus EF, Midi kit for endotoxin-free plasmid DNA (Macherey-Nagel or similar). 1. Isolate peripheral blood CD8+ T cells from buffy coats of healthy donors by negative selection (>95%) using the RosetteSep Human CD8+ T cell Enrichment Cocktail and subsequent centrifugation over Lympholyte Cell Separation Media. Briefly, add 50 μL of RosetteSep Human CD8+ T cell Enrichment Cocktail per mL of whole blood sample, mix, and incubate at room temperature for 20 min. Dilute the sample with PBS 2% (v/v) BCS and layer the diluted sample on a density gradient medium, being careful to minimize their mixing. 2. Centrifuge without brake. Carefully harvest the enriched cell layer with a transfer pipette and transfer to a new centrifuge tube. Wash enriched cells 2× with PBS 2% (v/v) BCS and proceed with the differentiation protocol. 3. Activate and expand resting CD8+ T cells (day 0) at a cell density of 0.5–2.5 × 106 cells/mL in complete R10 medium with 50 U/mL rIL-2 (see Note 15). 4. Resuspend the Dynabeads Human T-activator CD3/CD28 in the vial, transfer the desired volume (12.5 μL of beads per 1 × 106 of CD8+ T cells) to a tube, add at least 1 mL of PBS 0.1% (v/v) BCS, and mix. Place the tube on a magnet for 1 min and discard the supernatant. Remove the tube from the magnet and resuspend the washed Dynabeads in the same volume of complete R10 medium as the initial volume of Dynabeads taken from the vial. 5. Add washed Dynabeads to CD8+ T cells (cell-to-bead ratio = 1: 0.5). 6. Seed the cell/bead suspension at a cell density of 1 × 106 cells/ mL into a flat-bottom 12-well cell culture plate (2 mL/well) and incubate in a humidified CO2 incubator at 37 °C (see Note 16). 48 h after activation remove the beads (day 2) (see Note 17) and expand the cells in complete R10 medium with 50 U/ mL rIL-2 for further 3 days (day 5).

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Fig. 2 Schematic representation of a method to obtain differentiated CTLs starting from human naı¨ve/resting CD8+ cells purified from buffy coats of healthy donors. Freshly isolated naı¨ve/resting CD8+ cells are activated with commercial Dynabeads (without the need of antigen or antigen presenting cells) and expanded for 5–7 days to generate CTLs acquiring cytotoxic activity. CTLs were transfected 6 days after activation and analyzed at day 7. (The figure was partially generated with BioRender.com)

7. Split the cells back to a density of 1×106 cells/mL in complete R10 medium with 30 U/mL rIL-2 the day before nucleofection (Fig. 2). 3.3 CTL Nucleofection with the RUSH-GzmB Construct

At day 6, count the CTLs obtained and determine cell density. CTL nucleofection with the RUSH-GZMB construct is performed using the Amaxa T cell nucleofector kit, with the following protocol: 1. Centrifuge 2×106 cells per sample at 200×g for 10 min at room temperature. 2. Discard supernatant completely and resuspend the cell pellet carefully in 100 μL room temperature Human T Cell Nucleofector Solution (see Note 18). 3. Combine 100 μL of cell suspension with 1.5 μg DNA (Str-KDEL_GZMB-SBP-mCherry). 4. Transfer cell/DNA suspension Amaxa certified cuvette (see Note 19). 5. Insert the cuvette with the cell/DNA suspension into the Nucleofector Cuvette Holder and apply the T-023 Nucleofector Program. 6. Add ~500 μL of the pre-equilibrated culture media to the cuvette and gently transfer the sample into a 12-well plate (final volume 2 mL media/well/sample) (see Note 20).

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3.4 Evaluation of RUSH on Fixed Cells by Confocal Microscopy

1. Cell preparation: collect the nucleofected CTLs and seed 105 cells in a 96-well cell culture plate in a volume of 200 μL R10 medium/sample and incubate at 37 °C, 5% CO2. 2. Perform a time-course of release with biotin: plan a time-course for the reporter release, depending on the reporter that you are analyzing. In this protocol, we apply the following time-course: 0–15 min–30 min–60 min where the sample “0” is without biotin addition while the other time points indicate the time of biotin treatment. Add 40 μM biotin (from biotin 4 mM stock solution) to the treated samples. At the end of the time course, wash the samples with PBS and resuspend each sample in 30 μL PBS. 3. In the meantime, wash 10-well diagnostic microscope slides with absolute ethanol and allow them to dry. Coat each well with 300 μL of 0.01% PLL (1:10 dilution of stock solution in sterile water; see Note 21) and keep at RT for 20 min in the dark, then discard the solution and carefully wash with MilliQ. Air-dry the slides and keep in the dark. 4. Transfer all samples to the PLL-coated slides, one sample/well, and allow samples to adhere for 15 min at RT. 5. Remove the supernatant using a pipette (see Note 22), add 4% (v/v) PFA and keep for 20 min at RT in the dark. 6. Wash slides by immersion into a Hellendahl-type dish containing PBS. Dry the spaces between wells using 3 MM paper, being careful to not completely remove PBS from the wells to avoid cell disruption. 7. Add permeabilization solution [PBS with 0.1% (w/v) BSA and 0.01% (v/v) Triton X-100] dropwise to the wells and incubate for 20 min at RT. 8. Wash slides (as described in step 5). 9. Add 15 μL/well of anti-CanX antibody diluted 1:50 or antiLAMP1 antibody diluted 1:400 mixed with anti-RFP antibody (see Note 23) diluted 1:500 in PBS and keep the slide in a humidified chamber at 4 °C overnight. 10. The next day wash slides (as described in step 5). 11. Add 15 μL/well of a mix of AlexaFluor secondary antibodies, diluted 1:80 each in PBS and keep the slide in a humidified chamber at RT for 45 min (see Note 24). 12. Wash slides (as described in step 5). 13. Add 90% (v/v) glycerol in PBS, one drop/well (see Note 25), and then overlay the coverslip, remove the excess of slide mounting solution by very carefully pushing onto the coverslip, then fix it using conventional nail polish. 14. Store the slides at 4 °C in the dark.

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15. Analyze fluorescence on a confocal microscope to determine the localization of the protein of interest fused to the fluorescent tag. In our setting, where the hook is specific for the ER, the protein of interest should be localized in the ER in the absence of biotin (Fig. 3) and reach its destination compartment when biotin is added to the medium, depending on the kinetics of trafficking of the protein itself (Fig. 4). An example of the results for CTLs transfected with the Str-KDEL_GZMB-SBP-mCherry RUSH constructs is shown in Figs. 3 and 4. 3.5 Evaluation of RUSH on Live Cells

1. Preparation of μ-Slide 8 Well Chamber Slide: coat each required well with 200 μL of PLL and maintain at RT for 20 min in the dark, then discard the solution and carefully wash with MilliQ. Air-dry the slide and lodge it in the thermostated incubation chamber of the microscope. 2. Cell preparation: collect the nucleofected CTLs and seed 2×105 cells in 200 μL of Medium RPMI 1640 no phenol red per well in PLL-coated μ-Slide 8 Well Chamber Slide. 3. Add immersion oil to the 60× objective and wait for transfected cells to adhere to the PLL-coated well. 4. Choose the appropriate configuration of your microscope for the fluorescent protein to be imaged. Adjust the focus and mark positions of interest. 5. Add 40 μM biotin (from biotin 4 mM stock solution) to the sample. 6. Acquire images with a time interval of 5 min for a total time of 60 min at 37 °C. 7. During long-term acquisition, having the following precautions: use the Perfect Focus System and prevent photobleaching by using low laser intensity and short exposure time whenever possible.

4

Notes 1. LB liquid medium composition is described in paragraph 2.1. Combine the reagents and shake until the solutes have dissolved. Adjust the pH to 7.0 and the final volume of the solution to 1 L with H2O. Sterilize by autoclaving. 2. LB agar plates with antibiotic selection are obtained as described for the LB liquid medium with the 15 g bacteriological agar per 1 L of water. Mix well by inverting the bottle several times until powder is dissolved and sterilize by autoclaving. Following autoclaving (while media is still liquid but cool enough to safely handle the bottle), add the desired

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Fig. 3 Reporter release from the “donor” compartment. (a) Schematic representation of a RUSH plasmid with the hook placed upstream of the IRES (internal ribosome entry site) and the reporter is downstream. (b, c)

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antibiotic at the final concentration of 100 μg/mL and pour LB agar into Petri dishes. When agar has set, replace lid, invert plates, and store at 4 °C. 3. Resuspend lyophilized human IL-2 IS with deionized sterilefiltered water to a final concentration of 500 U/μL (corresponding to 0.1 mg/mL), and store aliquots at -20 °C or below. Avoid repeated freeze-thaw cycles. 4. Resuspend the Dynabeads carefully before use by vortexing the vial for >30 s. 5. Prepare a 4 mM biotin stock solution by dissolving 48 mg biotin in 50 mL culture medium. Filter using a 0.22 μm membrane to sterilize the solution and store up to 3 months at 4 °C. 6. 4% (v/v) PFA solution is prepared diluting a stock solution (16% PFA) in PBS1×, in sterile conditions, in the dark. Freshly 4% (v/v) PFA is prepared for each experiment. 7. Thaw on ice all the PCR reagents. Prepare a 10 μM primer solution in sterile water from a 100 μM stock. Be careful drawing the DNA Polymerase, because of its viscosity. 8. For gels, agarose is commonly used at concentrations of 0.7–2% depending on the size of bands to be separated. For example, to prepare a 1% (w/v) agarose gel 0.5 g of agarose in 50 mL 1× TAE (50× TAE buffer: 242.28 g Tris, 18.61 g EDTA Disodium, 57.1 mL Glacial Acetic Acid, dH2O up to 1 L). Mix agarose powder with 100 mL 1×TAE in a beaker. Microwave for 1–3 min until the agarose is completely dissolved, then let agarose solution cool down to about 50 °C and add 5 μL GelRed Nucleic Acid Gel Stain. 9. Pipet 2 μL of each sample for up to 16 samples onto a QIAxpert Slide avoiding bubbles. 10. Thaw on ice the FastDigest Buffer and keep the enzymes on ice throughout the experiment. Be careful drawing the enzymes, because of their viscosity. 11. Insert:vector ratio is calculated on the base of insert and vector length (kb) and mass. The ideal ratio is usually 3:1, because a larger amount of insert increases the chances of successful cloning reaction.

ä Fig. 3 (continued) Human primary CTLs expressing Str-KDEL_GZMB-SBP-mCherry (red) and stained for the ER marker CanX Ab (green), with and without biotin treatment. Scale bar, 5 μm (c) Co-localization analyses of Str-KDEL_GZMB-SBP-mCherry with CanX were obtained using Manders’ coefficient (mean ± SD; 10 cells/ sample) (b). The results show that before biotin addition GZMB strongly co-localizes with the ER, while after biotin addition the co-localization decreases in a time-dependent fashion, indicating release from the ER

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Fig. 4 Reporter trafficking toward the “acceptor” compartment. (a) Human primary CTLs expressing Str-KDEL_GZMB-SBP-mCherry (red) and stained with anti-LAMP-1 Ab (green) with and without biotin

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12. Thaw on ice all the ligation reagents. Be careful drawing the T4 DNA ligase, because of its viscosity. 13. Plates should be removed from 4 °C at least 1 h before use and placed in 30 °C incubator. 14. Plating two different volumes of bacterial culture is recommended to ensure that at least one plate will have well-spaced colonies. 15. Split the cells when cell density exceeds 2.5 × 106 cells/mL or when the medium turns yellow. 16. Examine cell culture daily, noting changes in cell size and shape. 17. Upon activation, some cells will bind strongly to the beads. Resuspend and transfer the bead/cell suspension to a suitable tube by thoroughly pipetting before cell separation from the beads on a magnet. 18. Avoid storing the cell suspension longer than 20 min, as this reduces cell viability and gene transfer efficiency. 19. Sample must cover the bottom of the cuvette without air bubbles. 20. Use the supplied pipettes and avoid repeated aspiration of the sample. 21. 0.1% (w/v) PLL stock solution is diluted 1:10 in distilled water and used fresh. 22. Do not completely dry the wells to avoid cell disruption. 23. The use of anti-RFP antibody in immunofluorescence experiments performed on cells transfected with Str-KDEL_TNFSBP-mCherry construct permits to amplify the mCherry signal, if required. 24. The fluorescently labelled secondary Ab must be kept safe from the light. For this reason, prepare dilutions and perform incubations in the dark. 25. Dispense slide mounting medium dropwise onto the slide wells using a Pasteur pipette.

ä Fig. 4 (continued) treatment. Scale bar, 5 μm. (b) Co-localization analyses of Str-KDEL_GZMB-SBP-mCherry with LAMP-1 were obtained using Manders’ coefficient (mean ± SD; 10 cells/sample). The results show that before biotin addition GZMB has a low co-localization with lytic granules (marked by LAMP-1), while after biotin addition the co-localization increases in a time-dependent fashion, indicating that after release from the ER GZMB is correctly transported to its destination compartment

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Acknowledgments This project has received funding from the European Research Council (ERC) under the European Union’s Horizon 2020 research and innovation program (grant agreement No 951329). References 1. Cassioli C, Baldari CT (2022) The expanding arsenal of cytotoxic T cells. Front Immunol 13: 883010. https://doi.org/10.3389/fimmu. 2022.883010 2. Griffiths GM, Tsun A, Stinchcombe JC (2010) The immunological synapse: a focal point for endocytosis and exocytosis. J Cell Biol 189: 399–406. https://doi.org/10.1083/jcb. 201002027 3. Kabanova A, Zurli V, Baldari CT (2018) Signals controlling lytic granule polarization at the cytotoxic immune synapse. Front Immunol 9: 307. https://doi.org/10.3389/fimmu.2018. 00307 4. Huse M (2012) Microtubule-organizing center polarity and the immunological synapse: protein kinase C and beyond. Front Immunol 3:1–11. https://doi.org/10.3389/fimmu. 2012.00235 5. Ritter AT, Asano Y, Stinchcombe JC, Dieckmann NMG, Chen B, Gawden-Bone C et al (2015) Actin depletion initiates events leading to granule secretion at the immunological synapse. Immunity 42:864–876. https://doi.org/ 10.1016/j.immuni.2015.04.013 6. Stinchcombe JC, Majorovits E, Bossi G, Fuller S, Griffiths GM (2006) Centrosome polarization delivers secretory granules to the immunological synapse. Nature 443:462–465. https://doi.org/10.1038/nature05071 7. Stinchcombe JC, Griffiths GM (2007) Secretory mechanisms in cell-mediated cytotoxicity. Annu Rev Cell Dev Biol 23:495–517. https://

doi.org/10.1146/annurev.cellbio.23.090506. 123521 8. Bossi G, Griffiths GM (1999) Degranulation plays an essential part in regulating cell surface expression of Fas ligand in T cells and natural killer cells. Nat Med 5:90–96. https://doi. org/10.1038/4779 9. Ba´lint Sˇ, Mu¨ller S, Fischer R, Kessler BM, Harkiolaki M, Valitutti S et al (2020) Supramolecular attack particles are autonomous killing entities released from cytotoxic T cells. Science 368:897–901. https://doi.org/10. 1126/science.aay9207 10. Chang H, Schirra C, Ninov M, Hahn U, Ravichandran K, Krause E et al (2022) Identification of distinct cytotoxic granules as the origin of supramolecular attack particles in T lymphocytes. Nat Commun 13:1029. https:// doi.org/10.1038/s41467-022-28596-y 11. Boncompain G, Perez F (2012) Synchronizing protein transport in the secretory pathway. Curr Protoc Cell Biol 15:unit 15.19. https:// doi.org/10.1002/0471143030.cb1519s57 12. Griffiths GM, Isaaz S (1993) Granzymes a and B are targeted to the lytic granules of lymphocytes by the mannose-6-phosphate receptor. J Cell Biol 120:885–896. https://doi.org/10. 1083/jcb.120.4.885 13. Boncompain G, Divoux S, Gareil N, de Forges H, Lescure A, Latreche L et al (2012) Synchronization of secretory protein traffic in populations of cells. Nat Methods 9:493–498. https://doi.org/10.1038/nmeth.1928

Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made. The images or other third party material in this chapter are included in the chapter’s Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter’s Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

Chapter 28 Interactions of Tissue-Resident T Cells Rut Mora-Buch, Hasan Akbaba, and Shannon K. Bromley Abstract Resident memory T cells (TRM) are non-circulating cells that play a critical role in protection from local infections and cancers. Flow cytometric and transcriptional analyses of these cells have defined their distinct phenotypes; imaging allows study of their morphology, localization, and interactions within tissues. Here, we describe commonly used methods to generate cutaneous CD8+ TRM and to prepare skin samples for analysis, including staining of cryostat sections, epidermal sheets, and tissue whole mounts. Key words Resident memory CD8+ T cells, TRM, Skin, Immunofluorescence microscopy, Epidermal sheets, Whole mount

1

Introduction CD8+ TRM lodge long term in both barrier and non-barrier tissues of mice and humans [1]. Mouse models that induce cutaneous CD8+ TRM are widely used to study CD8+ TRM regulation and function. Receptors expressed by skin CD8+ TRM are well defined [2], and so these cells are easily identified by microscopy and flow cytometric analyses. Following local cutaneous infection with viruses, including herpes simplex virus (HSV) [3] and vaccinia virus (VV) [4], large numbers of virus-specific CD8+ TRM persist long-term within the skin, without return to the circulation. Similarly tattoo vaccination with DNA encoding cognate antigen induces CD8+ T cell recruitment into the skin and CD8+ TRM formation [5, 6]. Additionally, although antigen recognition within the tissue boosts numbers of CD8+ TRM [7, 8], antigen non-specific inflammation can be used to induce the recruitment of early effector CD8+ T cells into the skin [9]. These cells persist for months and can deliver robust protection from infectious challenge.

Cosima T. Baldari and Michael L. Dustin (eds.), The Immune Synapse: Methods and Protocols, Methods in Molecular Biology, vol. 2654, https://doi.org/10.1007/978-1-0716-3135-5_28, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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Ex vivo imaging analysis provides considerable information about TRM numbers, localization, and morphology within tissues, and different skin preparations have allowed characterization of cutaneous CD8+ TRM. For example, imaging of frozen cryosections has revealed that TRM localize within distinct tissue niches. In mouse skin, CD4+ T cells interact with CD11b+ antigen-presenting cells (APCs) and are retained within clusters around hair follicles within the dermis [10], while CD8+ TRM localize to the basal layer of the epidermis [11]. On the other hand, imaging of skin whole mounts has captured the characteristic dendritic morphology of CD8+ TRM [12, 13]. Finally, separation of epidermal and dermal sheets followed by flow cytometric and/or microscopic analysis can also be used to quantitate TRM numbers and localization [14]. The techniques detailed in this protocol are relatively simple and do not require specialized equipment. Nonetheless, they are valuable tools to investigate how TRM numbers, localization, and morphology are affected by changes in their gene expression or surrounding environment.

2

Materials All work with mice should be performed according to approved ethical guidelines governing the use of animals in your laboratory. Additionally, follow institutional safety guidelines when handling and disposing hazardous chemicals, including DNFB and paraformaldehyde, as well as biohazardous materials such as recombinant HSV-OVA virus. T cell donor mice and recipient mice should be sex-matched, and all reagents, tubes, and equipment used for T cell preparation and adoptive transfer into mice should be sterile. The following mouse models generate epidermal CD8+ TRM, and nearly all the cutaneous CD8+ TRM express the receptors CD69, CD103, and CD49a.

2.1

Mice

1. C57Bl/6 recipient mice expressing the congenic marker, CD45.1 (see Note 1). 2. C57Bl/6 mouse for source of antigen-presenting cells (APCs). 3. OT-I mouse expressing a transgenic T cell receptor that recognizes the antigen, ovalbumin 257–264 (see Note 2) to be used as a source of T cells.

2.2 Preparation of T Cells for Adoptive Transfer

1. Tools for dissection (dissection board, pins, scissors, and forceps). 2. Spray bottle containing 70% ethanol. 3. 50 mL conical tube. 4. 70 μm nylon mesh strainer.

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5. Plunger with rubber tip from 3 cc syringe. 6. Fetal bovine serum—lot tested for T cell growth. 7. 0.5 M EDTA solution, pH 8 with NaOH. 8. Dulbecco’s Phosphate Buffered Salt Solution (DPBS): 137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 1.8 mM KH2PO4. 9. T cell purification buffer: DPBS without calcium or magnesium, 2% fetal bovine serum, 2 mM EDTA. 10. Immunomagnetic negative selection naı¨ve CD8+ T cell isolation kit (StemCell Technologies). 11. 5 mL polystyrene round-bottom tube. 12. 15 mL conical tube. 13. Magnet (StemCell equivalent).

Technologies

EasySep

magnet

or

14. 0.4% Trypan blue in 0.85% saline and hemacytometer. 15. Red blood cell lysis buffer: 155 mM NH4Cl, 12 mM NaHCO3, 0.1 mM EDTA. 16. Cell irradiator (see Note 3). 17. 1 mg/mL Ovalbumin (257–264) SIINFEKL peptide (stock in water). 18. T75 flasks. 19. RPMI-1640 media. 20. 1 M Hepes pH 7.4 in water. 21. 200 mM L-glutamine in water. 22. 100× Non-essential amino acid solution. 23. 55 mM β-mercaptoethanol stock in DPBS stored at 4 °C. 24. 10 mg/mL streptomycin, 10,000 U/mL penicillin in water. 25. T cell growth media: RPMI-1640, 10% fetal bovine serum, 10 mM HEPES, 2 mM L-glutamine, 1× non-essential amino acids, 55 μM β-mercaptoethanol, 100 μg/mL streptomycin, 100 U/mL penicillin. 26. 100 μg/mL Murine IL-2 (stock in water). 2.3 Models for Generation of Cutaneous CD8+ TRM

1. Syringes with attached needles (Monoject 28 g × ½ inch). 2. Ketamine (80 mg/kg) and xylazine (12 mg/kg). 3. Ophthalmic ointment (Puralube or equivalent). 4. Depilatory cream. 5. Electric shaver. 6. Ovalbumin-expressing herpes simplex virus-1 (HSV-1) KOS strain [2].

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7. Hanks balanced salt solution (HBSS). 8. Sandpaper (150 grit) glued to the blunt end of a pencil. 9. Opsite FlexiGrid tape (Smith & Nephew or equivalent). 10. Micropore tape (3 M or equivalent). 11. Transpore tape (3 M or equivalent). 12. Ethanol wipe. 13. Plasmid encoding model antigen, ovalbumin [6]. 14. Rotary tattoo machine (Biotouch or equivalent). 15. 7 prong tattoo needles (Biotouch or equivalent). 16. 0.3% 1-Fluoro-2,4-dinitrobenzene (DNFB) prepared in 4:1 acetone:olive oil (see Note 4). 2.4 Skin Preparation and Staining Methods for Analysis by Microscopy

1. Positively charged glass microscope slides. 2. Dissecting scissors and forceps. 3. Tissue embedding molds. 4. Pyrex container. 5. Dry ice pellets. 6. 2-methyl-butane. 7. Cryostat and blades. 8. Slide container. 9. 37 °C lab oven. 10. Parafilm. 11. Ziploc bag. 12. Pap pen. 13. Acetone. 14. Blocking buffer: PBS, 5% BSA, 1% TruStain FcX (Biolegend or equivalent) (see Note 5). 15. Streptavidin/Biotin Blocking Kits (Vector Labs). 16. PBS 0.05% Tween 20. 17. 0.5 mg/mL Anti-CD45.2-biotin. 18. 0.2 mg/mL Anti-CD8β-PE. 19. 0.5 mg/mL Streptavidin-Alexa Fluor 647. 20. 1 mg/mL Hoechst in water. 21. Prolong Diamond antifade mountant, equilibrated to room temperature. 22. Nail polish. 23. Humidified chamber (see Note 6). 24. Dissecting scissors and forceps.

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25. 5 mL polypropylene tubes. 26. Periodate-lysine-paraformaldehyde (PLP) Buffer: 0.1 M sodium phosphate buffer, 0.2 M L-lysine pH 7.4, 0.01 M sodium periodate, and 2% paraformaldehyde (see Note 7). 27. Blocking buffer + serum block: PBS, 2% BSA, 0.5% normal serum block (Biolegend or equivalent), 0.2% TruStain FcX (Biolegend or equivalent). 28. 0.2 mg/mL Anti-CD8β-PE. 29. PBS 0.1% Tween 20. 30. Hoechst (1 mg/mL stock concentration). 31. Microscope slides. 32. Nail polish. 33. Crystal Clear Gorilla tape. 34. 3 mg/mL Dispase in Dulbecco’s modified eagle medium. 35. 6-well plate.

3

Methods

3.1 Preparation of CD8+ T Cells for Adoptive TransferNaı¨ve CD8+ T Cell Purification

1. Euthanize a 6–8 week old OT-I mouse. 2. Rinse the mouse by spraying with 70% ethanol. 3. Position the mouse in the supine position and fix its limbs in place using sterile needles. 4. Cut the skin from the belly to neck, taking care not to cut the peritoneum. Then, make perpendicular incisions in the skin from the midline down each limb. 5. Gently peel back the skin on each side of the animal and pin it down. 6. Locate the axillary, brachial, and inguinal lymph nodes (Fig. 1), remove them using fine forceps, and place them into a 15 mL conical tube containing 5 mL CD8+ T cell purification buffer. Keep the tube on ice. 7. Make an incision on the left side of the mouse’s peritoneum. Locate the spleen, gently remove it using blunt forceps, and place it into the 15 mL tube on ice. 8. In a tissue culture hood using sterile technique, place the 70 μm strainer onto a 50 mL conical tube. Wet the filter with 1 mL T cell purification buffer. Then, transfer the spleen and lymph nodes into the strainer. 9. Mash the spleen and lymph nodes using the rubber tip of a 3-cc syringe plunger. Rinse the strainer with 10 mL T cell purification buffer. Repeat.

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A

B axillary brachial inguinal

spleen

Fig. 1 Dissection of mice for isolation of spleen and lymph nodes: (a) Mice are pinned in the supine position, and incisions are made from the belly to neck. Then, perpendicular cuts are made down the arms and legs of the mice, and the skin on each side of the animal is peeled back and pinned down. (b) Location of the spleen, and the axillary, brachial and inguinal lymph nodes

10. Remove an aliquot of cells, dilute them with trypan blue and determine the number of live cells by counting on a hemacytometer. 11. Centrifuge the cells at 300 × g for 10 min. 12. Aspirate the supernatant, resuspend the cells at 108 cells/mL in T cell purification buffer and isolate the cells according to the manufacturer’s directions (see Note 8). 13. After purification, wash the cells with DPBS and resuspend at 106 cells/mL. Keep the cells on ice until intravenous (i.v.) adoptive transfer. An aliquot of the purified cells can be stained with antibodies directed against CD8, CD44, and CD62L to ensure that they have a naı¨ve, CD8+ CD44low CD62Lhi phenotype. 3.2 Preparation of CD8+ T Cells for Adoptive TransferGeneration of Early Effector CD8+ T Cells

1. Follow Subheading 3.1 steps 1–7 to isolate the spleen from 1 C57Bl/6 mouse and transfer it to a 15 mL conical tube containing 5 mL PBS. 2. In a tissue culture hood using sterile technique, place a 70 μm strainer onto a 50 mL conical tube. Wet the filter with 1 mL PBS. Then, transfer the spleen into the strainer. 3. Mash the spleen using a 3-cc syringe plunger. Rinse the strainer with 10 mL PBS. Repeat. 4. Centrifuge the cells at 400 × g for 5 min.

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5. Aspirate the supernatant and resuspend the cells in 1 mL red blood cell lysis buffer. Incubate at room temperature for 3 min. 6. Add 20 mL T cell growth medium to stop the cell lysis and then centrifuge the cells at 400 × g for 5 min. 7. Aspirate the supernatant and resuspend the cells in 20 mL T cell growth medium. 8. Irradiate the splenocytes (3000 rad). 9. Add 20 μL Ovalbumin (257–264) peptide to the cells and invert to mix. Loosen the cap on the 50 mL tube to allow air exchange and place the cells into a 37 °C incubator for 1 h (see Note 9). 10. After 1 h, spin down the splenocytes at 400 × g for 5 min, aspirate the supernatant, and wash the cells with 20 mL T cell growth media. Repeat. 11. Determine the total number of live splenocytes by diluting an aliquot of the cells with trypan blue and counting on a hemocytometer. Finally, resuspend the splenocytes at 5 × 107 cells in 20 mL T cell growth media and transfer the cells into a T75 flask. 12. Isolate the axillary, brachial, and inguinal lymph nodes and spleen from one OT-I mouse and place the tissues into 5 mL DPBS in a 15 mL conical tube on ice. 13. In a tissue culture hood using sterile technique, place a 70 μm strainer onto a 50 mL conical tube. Wet the filter with 1 mL DPBS. Then, transfer the spleen and lymph nodes into the strainer. 14. Mash the tissues using a 3-cc syringe plunger. Rinse the strainer with 10 mL DPBS. Repeat. 15. Centrifuge the cells at 400 × g for 5 min. 16. Aspirate the supernatant and resuspend the cells in 1 mL red blood cell lysis buffer. Incubate at room temperature for 3 min. 17. Add 20 mL T cell growth medium to stop the cell lysis and then centrifuge the cells at 400 × g for 5 min. 18. Aspirate the supernatant and resuspend the cells at 5 × 107 cells in 20 mL T cell growth media. 19. Transfer the OT-I cells into the T75 flask containing the OVA-pulsed splenocytes. 20. Culture the flask standing upright (to maximize interactions with antigen-presenting cells) in a 37 °C incubator. 21. After 48 h, check the T cells under a microscope. If the T cells have been activated, they should form spherical clumps of cells visible under the microscope. The activated T cells are also larger than naı¨ve CD8+ T cells.

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22. Pipet the cells up and down, and transfer 20 mL T cells to a new T75 flask. Add 20 mL T cell growth media containing 10 ng/ mL murine IL-2 to each flask. Place the flask on its side in the 37 °C incubator. 23. On day 3, split the cells 1:1 again with T cell growth media containing 10 ng/mL IL-2 and incubate at 37 °C. 24. On day 4.5, recover cells from the flasks, remove dead cells by density gradient centrifugation following the manufacturer’s instructions (for example, using Lympholyte or Ficoll Premium). Then, wash the cells with DPBS and resuspend them at 107 cells/mL in DPBS for adoptive transfer. The cells can be stained with antibodies directed against CD8, CD69, and CD44 to confirm CD8+ T cell activation. 3.3 Models for Generation of Cutaneous CD8+ TRMZosteriform Model of HSV Infection

1. Anesthetize 7- to 8-week-old female (see Note 10) congenic recipient mice by intraperitoneal (i.p.) injection of 80 mg/kg ketamine and 12 mg/kg xylazine (see Note 11). 2. Adoptively transfer 105 naı¨ve CD8+ OT-I T cells in 100 μL DPBS by i.v. injection, either via lateral tail vein or retro-orbital injection. 3. Shave and depilate the mouse’s left flank. Do not leave depilating cream on skin for more than 1 min, or it can cause skin redness and inflammation. Rinse the skin with sterile PBS and dry with sterile gauze. 4. Abrade the skin near the top of the spleen with 20 strokes of 150 grit sandpaper (see Note 12). 5. Apply 10 μL HBSS containing 106 plaque forming units (pfu) HSV-OVA virus. Rub the virus into the site of abrasion using 20 strokes of a pipet tip. 6. Cover the infection site with a 1 × 2 cm piece of Opsite FlexiGrid, then wrap the flank with micropore tape and Transpore tape (see Note 13). 7. Remove bandages on day 2. The primary infection site should be visible. 8. Monitor the mice daily for formation of a zosteriform band of vesiculating lesions and progression of disease. The virus infects the innervating dorsal root ganglia, re-emerges to infect distal sites along the dermatome, and within ~5–6 days, a band of lesions develops from the primary site of infection at the tip of the spleen spreading ventrally toward the belly [15].

3.4 Models for Generation of Cutaneous CD8+ TRMOVA Plasmid Tattoo

1. Anesthetize 7- to 8-week-old female congenic recipient mice by i.p. injection of 80 mg/kg ketamine and 12 mg/kg xylazine. 2. Adoptively transfer 105 naı¨ve CD8+ OTI T cells in 100 μL PBS by i.v. injection.

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3. Draw a 1 cm2 square centered on the flank skin overlaying the top of the spleen. 4. Pipette 15 μL PBS containing 3 μg OVA plasmid in the center of the square. 5. Using a rotary tattoo device with a sterile 7 prong circle needle, tattoo the plasmid into the marked area of skin for 30 s. OT-I T cells can be recovered from the skin by day 8. The geometric mean fluorescence intensity (gMFI) of CD103 expression by cutaneous OT-I T cells increases over time until by day 30, nearly all cutaneous OT-I T cells are CD103+. 3.5 Models for Generation of Cutaneous CD8+ TRMDNFB-Induced Skin Inflammation

1. Anesthetize 7- to 8-week old female recipient mice by i.p. injection of 80 mg/kg ketamine and 12 mg/kg xylazine. 2. Adoptively transfer 106 d4.5 CD8+ OT-I T cells i.v. into congenic recipient C57Bl/6 mice. 3. Shave the left flank of the recipient mice, remove remaining hair with depilatory cream, and rinse the flank with sterile PBS. 4. Dry the flank with sterile gauze and then outline a 1 cm2 square on the skin overlaying the top of the spleen. 5. Pipette 10 μL of 0.3% DNFB in 4:1 acetone:olive oil (3 μL DNFB in a mix of 800 μL acetone and 200 μL olive oil) within the square. In our hands, by day 30, ~80% of cutaneous OT-I T cells express CD103 in this model.

3.6 Skin Preparation and Staining Methods for Analysis by MicroscopyCryosectioning

1. In a fume hood, precool 2-methyl-butane in a Pyrex beaker surrounded by dry ice pellets. 2. Euthanize a recipient mouse. 3. Isolate the 1 cm2 area of skin overlaying the spleen and place it into a petri dish containing PBS on ice. 4. Hold down the edge of the skin with blunt forceps and scrape off any fat using the curved shafts of forceps. Then, dry off the skin to remove excess PBS. 5. Cut the skin into ~3 mm wide strips. Then, submerge the skin into a tissue embedding mold containing optimum cutting temperature (OCT) medium at room temperature for 10 min. Remove bubbles from the OCT using fine forceps. Position the skin on edge in the embedding mold so that the epidermis will be perpendicular to the cryostat blade when the skin is sectioned (see Note 14). 6. Carefully add dry ice pellets to the 2-methyl-butane, a couple at a time. Wait for any bubbling to stop. 7. Using forceps or hemostats, carefully lower the embedded tissue molds onto the surface of the 2-methyl-butane. The OCT should rapidly turn white as it freezes (see Note 15).

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8. Remove the mold and temporarily store it on dry ice until all samples have been frozen. 9. Wrap the samples in aluminum foil and store in a Bitran bag in a -80 °C freezer for long-term storage. 10. Cool cryostat to -20 °C. Then, place samples in the cryostat for ~30 min to equilibrate to the cryostat temperature. 11. Remove the OCT tissue block from the mold, and adhere it to the cryostat metal grid using OCT. 12. Section the samples at desired thickness. We routinely cut 10–12 μm skin sections. Sections can be moved using a paintbrush if necessary. 13. Quickly transfer the section to a room temperature slide by placing the slide onto the tissue. Using a gloved finger, rub the underside of the slide to help the section adhere. 14. Dry the sections at 37 °C for 1 h to maximize adherence to the slides. 15. Slides can be stored in a slide box at -20 °C. Wrap the slide box with parafilm and place it into a Ziploc bag to minimize humidity before storage. 16. For staining, first fix sections in ice cold acetone for 10 min at room temperature. 17. Wash the slides by immersing them into PBS for 5 min at room temperature. Repeat for a total of three 5-min washes. 18. Wick excess moisture from slides, taking care not to disturb the tissue section. 19. Circle the tissue using a Pap pen. This provides a hydrophobic barrier to contain solutions during the staining procedure. 20. Pipette Blocking Buffer containing Streptavidin solution to cover the area circled with the pap pen and incubate for 1 h at room temperature. Perform incubations with the slides in a humidified chamber so that the tissue sections do not dry out. 21. Wash the sections by immersing the slides in PBS, 0.05% Tween 20 for 5 min at room temperature. Repeat. 22. Pipette Biotin solution (4 drops/mL in PBS) to cover the sections. Incubate for 15 min at room temperature. 23. Wash the sections by immersing the slides in PBS 0.05% Tween 20 for 5 min at room temperature. Repeat. 24. Pipette anti-CD45.2-biotin (1:50) in PBS 5% BSA to cover the sections and incubate 1 h at room temperature. 25. Wash the sections with PBS 0.05% Tween 20 at room temperature for 5 min, three times.

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26. Wash the sections with PBS at room temperature for 5 min, three times. 27. Pipette anti-CD8β-PE (1:100) and SA-AF647 (1:250) in PBS 5% BSA onto the sections and incubate for 1 h at room temperature. From this point on, the humidified chamber should be covered with aluminum foil to protect the samples from light. 28. Wash the sections with PBS 0.05% Tween at room temperature for 5 min, three times. 29. Wash the sections with PBS at room temperature for 5 min, three times. 30. Immerse the slides in Hoechst Solution (1 μL/10 mL PBS) and incubate for 10 min at room temperature. 31. Wash the slides with PBS for 5 min at room temperature. 32. Wick excess PBS from the slides, taking care not to touch the tissue. 33. Pipette Prolong Diamond mounting media onto the tissue. Try to avoid pipetting any bubbles onto the section and remove any using a pipette tip. Then, cover the section with a coverslip by placing the edge of the coverslip on the slide and slowly lowering it over the tissue, again avoiding trapping of air bubbles. 34. Allow the mounting media to cure at room temperature for 24 h. 35. Nail polish the edges of the coverslip to seal and allow the nail polish to dry before imaging. Slides can be stored at 4 °C. This method can be used to quantitate numbers of CD8+ TRM in the epidermis and dermis (Fig. 2a).

Fig. 2 Immunofluorescence microscopy images. (a) A histological cross-section of flank skin stained with antibodies directed against CD8 (red), CD45.2 (cyan), and Hoecsht (blue). Yellow asterisks indicate CD8+ CD45.2+ T cells localized within the epidermis. (b) Skin whole mounts stained with anti-CD8β

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3.7 Skin Preparation and Staining Methods for Analysis by Microscopy- Skin Whole Mount

All incubations and washes in this protocol are performed with the skin in a 5 mL polypropylene tube rotating on a Nutating mixer. 1. Euthanize a recipient mouse. 2. Isolate a 1 cm2 area of skin overlaying the spleen and place it into a petri dish containing PBS on ice. 3. Hold down the edge of the skin with blunt forceps and scrape off the fat using the curved shafts of forceps. Then, dry off the skin to remove excess PBS. 4. Fix the skin in 1 mL PLP buffer for 1 h at 4 °C. 5. Wash the skin in 2 mL PBS 0.1% Tween 20 for 20 min at 4 °C, three times. 6. Block the skin in blocking buffer + serum block for 2 h at room temperature. 7. Stain the skin overnight with a 1:100 dilution of CD8β-PE in blocking buffer + serum block at 4 °C. 8. Wash the tissue with 2 mL PBS, 0.1% Tween 20 for 20 min at 4 °C, three times. 9. Incubate the tissue in Hoechst for 2 h at room temperature. 10. Wash the tissue with 2 mL PBS for 20 min at 4 °C, three times. 11. Remove excess liquid from the tissue and then place the skin on a slide with the epidermis facing up. 12. Pipette Prolong diamond antifade mountant to cover the tissue. 13. Slowly place a coverslip over the skin, avoiding trapping of air bubbles. Allow the mounting media to cure for 24 h at room temperature. Finally, nail polish the edges of the coverslip to seal, and allow to dry before imaging by confocal microscopy. We have used this method to visualize the morphology of CD8+ TRM (Fig. 2b).

3.8 Skin Preparation and Staining Methods for Analysis by MicroscopyEpidermal Sheet

1. Euthanize mouse, isolate a 1 cm2 piece of skin, and place it in a petri dish containing PBS on ice. 2. Hold down the edge of the skin with blunt forceps and scrape off the fat using the curved shafts of forceps. Then, remove excess PBS from the skin, ensuring that the epidermis is dry. 3. Affix the epidermis side of the skin to Gorilla tape. Rub the tape onto the skin to ensure that all the epidermis adheres to the tape. 4. Float the tape with the epidermis side up in a 6-well plate containing 5 mL DMEM, 3 mg/mL dispase for 1 h at 37 °C. 5. Gently use forceps to pull the dermis away from the epidermis (see Note 16).

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6. Fix the epidermis in 1 mL PLP Buffer for 15 min at room temperature. Incubations can be performed in a 24-well plate. 7. Stain the epidermis using the same protocol as detailed for staining cryosections. Staining can be performed with the epidermis in a 24-well plate using 300 μL of solutions. 8. After staining, cut away excess tape around the epidermis. 9. Place the sample tape side down onto a slide. 10. Add Prolong Diamond Mounting Media to cover the epidermis, remove bubbles, and carefully place a coverslip to cover the epidermal sheet. Allow the mounting media to cure at room temperature overnight, nail polish the edges of the coverslip to seal, and allow to dry before imaging.

4

Notes 1. Adoptive transfer of T cells into congenic recipient mice allows identification of host and donor cells by staining for CD45.1 and CD45.2. Alternatively, investigators can use Thy1 (CD90) congenic mice. However, we have observed that in some models, adoptively transferred CD90.1 cells obtained from Jackson Laboratories B6.PL-Thy1a/CyJ mice do not persist in C57Bl/ 6 CD90.2 hosts long-term; donor mice may require additional backcrossing to the C57Bl/6 strain. 2. Use of OT-I transgenic mice expressing a T cell receptor recognizing ovalbumin 257–264 allows T cell activation and expansion following introduction of cognate OVA antigen. Alternative TCR transgenic mice and cognate antigen systems can also be used. 3. If an irradiator is not available, feeder splenocytes can be treated with mitomycin C to inhibit proliferation [16]. Mitomycin C is toxic, and users should follow the manufacturer’s precautions. 4. We routinely treat female mice with 0.3% DNFB in 4:1 acetone:olive oil. In order to recover similar numbers of CD8+ TRM from the skin of male mice, we apply 0.5% DNFB. 5. When streptavidin/biotin blocking is required, streptavidin can be added to the blocking buffer step. 6. A humidified chamber can be home-made by placing wet paper towels or a thin layer of water on the base of a plastic container. Pipettes can be broken to fit the length of the container and taped together at a width to carry the glass slides, avoiding any contact of the slides with the water. 7. Prepare 0.2 M L-lysine in 0.1 M phosphate buffer, pH 7.4 and store at 4 °C. Prepare 4% paraformaldehyde by diluting 16%

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paraformaldehyde (Electron Microscopy Sciences or similar) in 0.1 M phosphate buffer. Immediately before use, mix l-lysine buffer and paraformaldehyde 1:1, and add sodium metaperiodate to 0.01 M. 8. We routinely recover 6–10 × 106 naı¨ve CD8+ T cells from the spleen and lymph nodes of one OT-I mouse. Depending on the number of cells required for adoptive transfer, the number of starting cells can be scaled down to save on purification reagents per the manufacturer’s instructions. 9. We usually purify CD8+ T cells during the incubation of irradiated splenocytes with cognate antigen. 10. We prefer to use female mice. Male mice may fight, leading to inflammation of the skin and less consistent results. It is also important to use mice that are not undergoing hair growth. When mice are in anagen phase of the hair cycle, HSV infection tends to be less successful, and results are more variable. 11. The blink reflex is suppressed in mice anesthetized with ketamine and xylazine. To prevent corneal damage, apply ophthalmic ointment to each eye. 12. The skin can be abraded with sandpaper or with a handheld motorized pedicure/manicure nail drill to remove the keratinized layer of skin and expose the epidermal layer. The area of abraded skin should appear shiny after abrasion. Caution should be used not to abrade the skin too deeply into the dermal layer or cause bleeding to avoid variability in the kinetics and symptoms of disease [17]. Abrasion of skin at the top of the spleen provides a landmark to locate the site of infection at memory timepoints. 13. Wrapping the flank with tape prevents removal of the Opsite FlexiGrid and disruption of infection. Depending on the size of the mice, the Transpore tape may need to be cut in half to allow normal ambulation of the mice. 14. Fine forceps can be used to position the skin and remove bends in the skin before freezing. If we have difficulty positioning the skin on end in the OCT without twisting or bending it, we have also placed the skin flat at the bottom of the embedding mold. Then, to obtain transverse sections, we attach the skin to the cryostat metal grid so that the epidermis is perpendicular to the blade. Additionally, if the skin is infected, it should be fixed before freezing in OCT and sectioning (2 h in 4% paraformaldehyde with 10% sucrose in PBS on ice) [3]. 15. 2-methyl-butane can be reused. After freezing tissues, pour any remaining 2-methyl-butane back into a container. Allow it to come to room temperature before tightening the lid to prevent pressure buildup.

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16. If the dermis does not easily separate from the epidermis, return the skin to the dispase solution at 37 °C and periodically check whether the epidermis can be separated from the dermis. In our hands, prolonging exposure of the skin to dispase can result in loss of some cell surface receptor expression; analysis of expression of receptors of interest can be checked by flow cytometric analysis. Additionally, we have observed that the morphology of CD8+ TRM within the epidermis is changed by incubation in dispase. Skin whole mounts may be more amenable to investigation of CD8+ TRM morphology.

Acknowledgements This work was supported by NIH grant 1R01AI163517 (to S.K. B.) and National Eczema Association Catalyst Research Grant NEA19-CRG121 (to R.M-B). Drawings were created with Biorender.com References 1. Enamorado M, Khouili SC, Iborra S, Sancho D (2018) Genealogy, dendritic cell priming, and differentiation of tissue-resident memory CD8 (+) T cells. Front Immunol 9:1751. https:// doi.org/10.3389/fimmu.2018.01751 2. Mackay LK, Rahimpour A, Ma JZ, Collins N, Stock AT, Hafon ML, Vega-Ramos J, Lauzurica P, Mueller SN, Stefanovic T, Tscharke DC, Heath WR, Inouye M, Carbone FR, Gebhardt T (2013) The developmental pathway for CD103(+)CD8+ tissue-resident memory T cells of skin. Nat Immunol 14(12): 1294–1301. https://doi.org/10.1038/ni. 2744 3. Gebhardt T, Wakim LM, Eidsmo L, Reading PC, Heath WR, Carbone FR (2009) Memory T cells in nonlymphoid tissue that provide enhanced local immunity during infection with herpes simplex virus. Nat Immunol 10(5):524–530. https://doi.org/10.1038/ni. 1718 4. Jiang X, Clark RA, Liu L, Wagers AJ, Fuhlbrigge RC, Kupper TS (2012) Skin infection generates non-migratory memory CD8+ T (RM) cells providing global skin immunity. Nature 483(7388):227–231. https://doi. org/10.1038/nature10851 5. Ariotti S, Hogenbirk MA, Dijkgraaf FE, Visser LL, Hoekstra ME, Song JY, Jacobs H, Haanen JB, Schumacher TN (2014) T cell memory. Skin-resident memory CD8(+) T cells trigger a state of tissue-wide pathogen alert. Science

346(6205):101–105. https://doi.org/10. 1126/science.1254803 6. Mani V, Bromley SK, Aijo T, Mora-Buch R, Carrizosa E, Warner RD, Hamze M, Sen DR, Chasse AY, Lorant A, Griffith JW, Rahimi RA, McEntee CP, Jeffrey KL, Marangoni F, Travis MA, Lacy-Hulbert A, Luster AD, Mempel TR (2019) Migratory DCs activate TGF-beta to precondition naive CD8(+) T cells for tissueresident memory fate. Science 366(6462): eaav5728. https://doi.org/10.1126/science. aav5728 7. Hirai T, Yang Y, Zenke Y, Li H, Chaudhri VK, De La Cruz Diaz JS, Zhou PY, Nguyen BA, Bartholin L, Workman CJ, Griggs DW, Vignali DAA, Singh H, Masopust D, Kaplan DH (2021) Competition for active TGFbeta cytokine allows for selective retention of antigenspecific tissue- resident memory T cells in the epidermal niche. Immunity 54(1):84–98.e85. https://doi.org/10.1016/j.immuni.2020. 10.022 8. Khan TN, Mooster JL, Kilgore AM, Osborn JF, Nolz JC (2016) Local antigen in nonlymphoid tissue promotes resident memory CD8+ T cell formation during viral infection. J Exp Med 213(6):951–966. https://doi.org/10. 1084/jem.20151855 9. Mackay LK, Stock AT, Ma JZ, Jones CM, Kent SJ, Mueller SN, Heath WR, Carbone FR, Gebhardt T (2012) Long-lived epithelial immunity by tissue-resident memory T (TRM) cells in

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the absence of persisting local antigen presentation. Proc Natl Acad Sci U S A 109(18): 7037–7042. https://doi.org/10.1073/pnas. 1202288109 10. Collins N, Jiang X, Zaid A, Macleod BL, Li J, Park CO, Haque A, Bedoui S, Heath WR, Mueller SN, Kupper TS, Gebhardt T, Carbone FR (2016) Skin CD4(+) memory T cells exhibit combined cluster-mediated retention and equilibration with the circulation. Nat Commun 7:11514. https://doi.org/10. 1038/ncomms11514 11. Gebhardt T, Whitney PG, Zaid A, Mackay LK, Brooks AG, Heath WR, Carbone FR, Mueller SN (2011) Different patterns of peripheral migration by memory CD4+ and CD8+ T cells. Nature 477(7363):216–219. https:// doi.org/10.1038/nature10339 12. Ariotti S, Beltman JB, Chodaczek G, Hoekstra ME, van Beek AE, Gomez-Eerland R, Ritsma L, van Rheenen J, Maree AF, Zal T, de Boer RJ, Haanen JB, Schumacher TN (2012) Tissue-resident memory CD8+ T cells continuously patrol skin epithelia to quickly recognize local antigen. Proc Natl Acad Sci U S A 109(48):19739–19744. https://doi.org/10. 1073/pnas.1208927109 13. Bromley SK, Akbaba H, Mani V, MoraBuch R, Chasse AY, Sama A, Luster AD (2020) CD49a regulates cutaneous resident memory CD8(+) T cell persistence and

response. Cell Rep 32(9):108085. https:// doi.org/10.1016/j.celrep.2020.108085 14. Mohammed J, Beura LK, Bobr A, Astry B, Chicoine B, Kashem SW, Welty NE, Igyarto BZ, Wijeyesinghe S, Thompson EA, Matte C, Bartholin L, Kaplan A, Sheppard D, Bridges AG, Shlomchik WD, Masopust D, Kaplan DH (2016) Stromal cells control the epithelial residence of DCs and memory T cells by regulated activation of TGF-beta. Nat Immunol 17(4):414–421. https://doi.org/10.1038/ni. 3396 15. van Lint A, Ayers M, Brooks AG, Coles RM, Heath WR, Carbone FR (2004) Herpes simplex virus-specific CD8+ T cells can clear established lytic infections from skin and nerves and can partially limit the early spread of virus after cutaneous inoculation. J Immunol 172(1): 3 9 2 – 3 9 7 . h t t p s : // d o i . o r g / 1 0 . 4 0 4 9 / jimmunol.172.1.392 16. Kruisbeek AM, Shevach E, Thornton AM (2004) Proliferative assays for T cell function. Curr Protoc Immunol Chapter 3:Unit 3.12. https://doi.org/10.1002/0471142735. im0312s60 17. Goel N, Docherty JJ, Fu MM, Zimmerman DH, Rosenthal KS (2002) A modification of the epidermal scarification model of herpes simplex virus infection to achieve a reproducible and uniform progression of disease. J Virol Methods 106(2):153–158. https://doi.org/ 10.1016/s0166-0934(02)00160-x

Chapter 29 Live Imaging of CAR T Cell Ca2+ Signals in Tumor Slices Using Confocal Microscopy David Espie, Sarah Barrin, Irena Rajnpreht, Lene Vimeux, and Emmanuel Donnadieu Abstract The immune synapse is a key structure organizing T-cell activation against foreign entities, such as cancer cells expressing neoantigens. One crucial step in this activation cascade is the intracellular Ca2+ ([Ca2+]i) response that shapes T cells for proliferation, differentiation, and cytotoxicity. The development of calcium probes coupled to real-time fluorescence microscopy has allowed a close study of this phenomenon in vitro. Such systems have provided valuable insights on the consequences of Ca2+ responses on T cells, including cytotoxicity and cytoskeletal remodeling. However, in vitro models do not recapitulate the tissue architecture that T cells come in contact with in vivo. Thus, there is a growing necessity for better understanding the factors influencing Ca2+ response in T cells including in genetically modified T cells (e.g., CAR T cells). In this methodology chapter, we describe an experimental system to measure [Ca2+]i signals of CAR T cells loaded with the calcium probe Fluo-4 on fresh tumor slices. Combined with confocal fluorescent imaging, this model offers an approach to image early T-cell activation in a three-dimensional (3D) tissue environment. Key words T cells, Intracellular calcium, Tissue slice, Confocal microscopy

1

Introduction The intracellular Ca2+ ([Ca2+]i) is a crucial second messenger of T-cell activation following TCR engagement in the context of an immune synapse. Increase of T-cell [Ca2+]i levels activate intracellular signaling cascades regulating T-cell effector functions, including mobility, proliferation, cytokine secretion and cytotoxicity [1]. Calcium probes and dyes can be used to measure this phenomenon in real-time using live fluorescence microscopy. These probes bind calcium and exhibit increased fluorescence from a resting background signal (non-ratiometric dyes) or a shift in their

David Espie and Sarah Barrin contributed equally. Cosima T. Baldari and Michael L. Dustin (eds.), The Immune Synapse: Methods and Protocols, Methods in Molecular Biology, vol. 2654, https://doi.org/10.1007/978-1-0716-3135-5_29, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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excitation-emission spectrum (ratiometric dyes) [2]. Hence, they represent useful tools to assess the early activation of both endogenous and genetically modified T cells. Chimeric antigen receptor (CAR)-modified T cells have gained much interest in the past decades due to their remarkable efficacy in haematological malignancies [3]. In a nutshell, the CAR functions as a synthetic receptor comprising of an extracellular antigen-recognition domain with intracellular signaling and co-stimulatory domains based on CD3ζ and CD28/4-1BB. This novel construct permits recognition of extracellular antigens on cancer cells in a non MHC-restricted manner [4]. The presence of a CD3-based signaling domain allows the CAR to trigger signalling events leading to T cell activation. Hence, measure of [Ca2+]i signals in CAR-T can be useful to validate the specificity of CAR-T against their target antigen and study the factors required for proper activation. Our laboratory has previously developed a novel preclinical model to study T-cell mobility and function in ex vivo fresh human tumours and lymphoid tissue slices using confocal microscopy [5–8]. This model has been expanded to assess the behavior of CAR-T in the tumor slice system through measurement of [Ca2+]i signals [9]. We describe, in this methodology chapter, a detailed protocol of this technique. The different steps consist of loading CAR-T and control T cells with the fluorescent calcium indicator Fluo-4, embedding the tumor tissue into an agarose gel, cutting it into thick slices, and adding the Fluo-4-loaded cells on the tissue. Spinning disk confocal microscopy is then used to monitor calcium fluxes over time.

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2.1 Machines and Equipment

1. Spinning disk upright confocal microscope with heating chamber (37 °C). 2. Vibratome: Leica VT1200S (other models are suitable).

2.2 Buffers and Powders

1. Phosphate-buffered saline (PBS) without calcium and magnesium: (2.67 mM KCl, 1.47 KH2PO4, 137.93 mM NaCl, 8.06 mM Na2HPO4-7H2O). 2. Hanks balanced salt solution (HBSS) with calcium and magnesium (no phenol red). 3. Complete medium: RPMI 1640 (no phenol red) supplemented with 3% human serum blood type AB and 1% penicillin/ streptomycin (0.2 μM filtered). 4. Low-gelling temperature agarose, type VII-A or equivalent. 5. Fluo-4 AM in 1 mM aliquots in DMSO stored at -20 °C. 6. Perfusion medium: RPMI 1640 (no phenol red).

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1. Fine forceps. 2. Double edged razor blade. 3. Scalpel, Swann-Morton. 4. Butyl cyanoacrylate glue, Vetbond (3M). 5. 35 mm plastic petri dishes. 6. 10 cm plastic tissue culture dishes. 7. 30 mm culture hydrophilic PTFE).

inserts,

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8. Stainless steel washers, 4 mm inner diameter. 9. 95% O2, 5% CO2 gas cylinder, regulator with needle valve and flexible tubing. 10. Perfusion system with adjustable flow rate and flexible tubing. 2.4

Antibodies

Below is a list of directly coupled antibodies that have been validated in tumor slices: 1. Brilliant Violet 421-conjugated anti-human EpCAM (CD326) (clone 9C4). 2. Alexa Fluor 647-conjugated anti-human and mouse fibronectin (clone HFN7.1). 3. APC-conjugated REA266).

anti-human

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4. Brilliant Violet 421-conjugated anti-mouse podoplanin (gp38) (clone 8.1.1).

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3.1 CAR-T Production and Culture

1. CAR-T cells are obtained from transduction of activated T cells with lentivirus encoding for the CAR transgene. The resulting CAR-T are tested for CAR expression levels, phenotype and functionality (e.g. degranulation, cytotoxicity and cytokine production). For protocols covering the CAR-T generation process including lentivirus production, T cell isolation and viral transduction, please refer to [10].

3.2 Labeling CAR-T and Non-transduced T Cells with Fluo-4

1. Use a culture of CAR-T and non-transduced T cells (or T cells transduced with an irrelevant CAR) at 10 to 14 days postactivation with a concentration between 1–2 × 106 cells/mL (see Note 1). 2. Isolate the desired number of cells and wash them twice in pre-warmed HBSS. 3. Put cells at 1 × 106 cells/mL in HBSS with 1 μM Fluo-4.

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4. Incubate for 30 min at 37 °C in the dark to allow the dye to be loaded inside the cells (see Note 2). 5. Wash cells twice in complete medium and resuspend to final concentration of 1 × 106 cells/mL in complete medium. 6. The labeled cells can be kept for several hours at 4 °C in the dark to allow time for tissue slice preparation. 3.3 Obtaining Human Tumor Samples

1. Obtain fresh human tumor sample from patients undergoing tumorectomy or another tumor sample from a xenograft mouse model. 2. Transport the fresh sample in ice-cold complete medium for immediate processing (see Note 3).

3.4 Tissue Processing

1. Place the tumor sample in a 10 cm plastic tissue culture dish with some complete medium to keep the tissue moist. 2. Cut the sample with a razor blade into small cube fragments between 3–5 mm in on each side. 3. Transfer the tissue cubes onto absorbent paper to remove excess medium (see Note 4).

3.5 Preparation of Agarose Gel

1. Add 0.5 g low gelling agarose to a glass beaker containing 10 mL PBS. 2. Microwave the above solution until agarose is dissolved (boils), swirl gently, and pour in a pre-warmed 35 mm petri dish (placed inside an incubator). 3. Place the petri dish inside an incubator at 37 °C to allow the agarose to cool and the bubbles to disappear (approx. 5 min) (see Notes 5 and 6).

3.6 Embedding of Tumor Samples in Agarose Gel

1. Take out the petri dish from the incubator and quickly place the tumor samples inside the agarose gel using fine forceps (see Notes 7 and 8). 2. Let the agarose gel solidify on ice for 5–10 min. 3. Remove the agarose gel from the petri dish using a scalpel. 4. Trim the excess agarose surrounding the tumor tissue with a razor blade leaving border 3–5 mm around the tissue.

3.7 Vibratome Slicing of Human Tumor Samples

1. Mount each agarose block on the specimen disk of the vibratome with a drop of butyl cyanoacrylate glue (avoid air bubbles if possible). 2. Place the specimen disk of ice for 2–3 min to let the glue solidify under the agarose block (the glue solidifies quickly). 3. Place the specimen disk inside the buffer tray and fill with ice-cold PBS until the agarose blocks are immersed.

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4. Cut the agarose blocks into 300–400 μM slices using the vibratome (depending on the consistency or stiffness of the tissue), at a low cutting speed (0.2–0.3 mm/s), and with a cutting amplitude of 1.55 mm. 5. Collect the tumor slices using fine forceps and place them in a 6-well plate filled with 2–3 mL of pre-chilled complete medium per well (see Note 9). 3.8 Immunostaining of Tumor Slices

1. Place 1 mL complete medium per well of a 6-well plate and place a culture insert making sure to avoid air bubbles. 2. Transfer up to two slices per insert using fine forceps and place a stainless-steel washer onto the tissue. 3. Dilute antibodies to final concentration 10 μg/mL in a 40 μL final volume of complete medium per slice. 4. Using a pipette, add 40 μL antibody mix per slice (see Note 10). 5. Incubate the plate at 37 °C for 15 mins to allow antibody staining. 6. With fine forceps, remove the washers and place each tissue slice in a 6-well plate filled 2–3 mL pre-chilled complete medium to wash excess antibody. 7. Maintain the plate at 4 °C in the dark before imaging.

3.9 Microscope Preparation

The confocal microscope used in this protocol is upright spinning disk equipped with a 25× water immersion objective (25× /0.95 NA): 1. Set the temperature of the microscope heat chamber at 37 °C a few hours before starting the imaging session. 2. Set the perfusion system of a pre-warmed solution of RPMI without phenol red (perfusion medium), bubbled with 95% O2 and 5% CO2 to 0.3 mL/min. 3. Using a pipette tip, place some butyl cyanoacrylate glue at the bottom of a pre-warmed 37 °C 35 mm petri dish. 4. Using fine forceps, place a tissue slice onto the glue taking care to avoid air bubbles. 5. Remove excess humidity around the agarose and tissue using absorbent paper. 6. Place a stainless-steel washer around the tissue and place on the imaging stage of the microscope. 7. Put the input and output perfusion tips of the perfusion system inside the petri dish. Please note that both tips should be placed close to the bottom of the petri dish to avoid creating a medium flux that might cause drift during acquisition.

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8. Using a pipette tip, create a meniscus inside stainless-steel washer using complete medium (approx. 200 μL) (see Note 11). 9. Bring the microscope objective slowly into the meniscus and close to the tissue sample. Avoid touching or damaging the tissue. 10. Using brightfield light, adjust the objective inside the meniscus to the appropriate z position in order to focus at the cut surface of the slice. 11. Set an imaging session with the following parameters: (a) Set a time lapse with one image every 10 s (1 time point) for 10–15 min. (b) Set the acquisition for the channel recording the Ca2+ signal every time point and the other channels used to image the TME every 5–10 time points to avoid photobleaching. (c) Depending on the intensity of the fluorescent signal of the fluorophores, set the exposure time between 50 and 800 ms and the laser intensity between 10% and 40%. (d) Set a z-stack acquisition with approximately 10 z-steps with a z-stack thickness between 5–10 μM. 12. Turn on the camera to visualize the slice in live. 13. Use appropriate fluorescent lights to select a region of interest containing tumor cells and other elements of the tumor microenvironment (TME) and adjust the z focus if needed. This live visualization step should be done quickly to avoid photobleaching. 3.10 Imaging of Ca2+ Levels of CAR-T

1. Resuspend 0.1–0.2 × 106 CAR-T or T cells in 10 μL pre-warmed complete medium. 2. Slowly add the cells inside the meniscus using a 10 μL pipette, you may raise the microscope objective slightly to have more space if needed. 3. Turn on the camera of the microscope and quickly readjust the z-stack if necessary. 4. Quickly check the labeled CAR-T or T cells are progressively landing on the tissue slice. 5. As soon as the first cells land on the slice, start recording the timelapse and turn on the perfusion system set at 3 mL/min briefly. 6. As soon as the media touches the agarose border, lower the perfusion between 0.2 and 0.3 mL/min to avoid any drift during acquisition.

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Fig. 1 Diagram of the experimental setup for the calcium assay. The agarose slice containing the tissue of interest (orange) is glued to the bottom of a petri dish using the VetBond glue (blue). A stainless-steel ring (black) is placed around the tissue slice. It is very important that the ring is placed flat on the agarose surface with no gaps underneath to create a complete seal. The petri dish is brought under the pre-warmed microscope and a meniscus is created inside the metal ring. The microscope objective can be lowered gently inside the meniscus. There should be enough space to add the Fluo-4-loaded cells (green) and observe their calcium response (red) upon contact with the target cells of interest. A perfusion system can be set up inside the microscope for long-term recording. Created with BioRender.com

7. The perfused media should gently and slowly cover the stainless ring without disturbing the T cells or CAR-T on the tissue. 8. It is possible to realize this experiment without perfusion system, but it is recommended to use it to be closer to physiological conditions and avoid stress induced by hypoxia. 9. A schematic presentation of the experimental setup is shown in Fig. 1. 3.11 Analyzing CART Activation

1. Import the data to an automated tracking software (e.g. Image J or Imaris) and locate single CAR-T or non-transduced T cells by adjusting cell diameter and detection threshold according to the imaged fluorescent cells. 2. Analyze fluorescence intensity of individual cells over time. The Ca2+ response can be measured as the ratio of the mean fluorescence intensity (F) occurring during the course of the experiment from the fluorescence intensity determined at the beginning of the movie (F0) before the cells activate. 3. Compare the Ca2+ levels between CAR-T and non-transduced T cells. A representative example is shown in Fig. 2.

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Fig. 2 Ca2+ responses of CAR-T vs. non-transduced T cells in tumor slices expressing the target antigen. The tumour arises from an immunodeficient mouse xenotransplanted with a human tumour cell line. The tumor

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Notes 1. If using cryopreserved T cells and CAR-T, defrost them for 2–3 days before the experiment to allow sufficient time for recovery. 2. Fluo-4 cell loading step can lead to cell loss of up to 50% due to successive washings and presence of DMSO with T cells. Carefully calculate the final number of Fluo-4-loaded cells required before the experiment. 3. If the tumor sample is received late in the day, it is possible to keep it overnight at 4 °C in MACS Tissue Storage solution (Miltenyi) for processing the next day. However, optimal results are achieved when the sample is processed immediately. 4. This step is important since excess medium can weaken the agarose structure around the tissue and affect tissue slice quality. 5. Having a clear agarose is key. Presence of bubbles can weaken the agarose and diminish tissue slice quality. 6. One petri dish can accommodate up to 1–4 tumor samples, scale up the number of petri dishes if more tumor samples are to be studied. 7. Pick tumor samples from below using the forceps to avoid damaging the tissue. 8. Wipe excess agarose on the forceps between each sample using tissue paper; not doing this can weaken the agarose and affect the final tissue slice quality. 9. Use great care when handling slices with fine forceps as they can be easily damaged. This critical step needs training. A cut in the agarose will compromise the seal made by the stainless-steel washer resulting in a leakage of the antibody containing solutions and consequently a poor staining. 10. Make sure washers are well positioned on the agarose surrounding the tumor tissue. Washers are used to concentrate the antibodies on the vibratome-cut slice. 11. A cut in the agarose will compromise the seal made by the stainless-steel and can disturb the stability of the meniscus and make imaging more complicated.

ä Fig. 2 (continued) cells (green) are labeled by an antibody against human ICAM-1 and the stroma (blue) labeled by an antibody against mouse podoplanin: (a) Confocal pictures of a tumor slice showing the Ca2+ level of CAR-T (left) and non-transduced T cells (right); (b) tracks of a representative Ca2+ responses for a single CAR-T (left) vs. non-transduced T cell (right); (c) average Ca2+ responses of CAR-T (n = 3) vs. non-transduced T cells (n = 20) plotted against time

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Acknowledgments We wish to thank Marie Escande (Invectys, Paris, France) for critical reading of the manuscript. DE and ED are members of the T2Evolve network (Innovative Medicines Initiative). The T2Evolve network received funding from the Innovative Medicines Initiative 2 Joint Undertaking (grant agreement number 116026). This Joint Undertaking receives support from the European Union’s Horizon 2020 Research and Innovation program and European Federation of Pharmaceutical Industries and Associations (EFPIA). IR and ED are members of the OptiCAN network (ERAPERMED2020-222). DE PhD is co-funded between the academic lab led by ED as PhD supervisor and the industrial partner Invectys. We would like to thank the staff of the IMAG’IC facility of the Cochin Institute for their advice during this study. IMAG’IC facility is supported by the National Infrastructure France BioImaging (grant ANR-10-INBS-04) and IBISA consortium. References 1. Trebak M, Kinet JP (2019) Calcium signalling in T cells. Nat Rev Immunol 19(3):154–169. https://doi.org/10.1038/s41577-0180110-7 2. Grynkiewicz G, Poenie M, Tsien RY (1985) A new generation of Ca2+ indicators with greatly improved fluorescence properties. J Biol Chem 260(6):3440–3450. https://doi.org/10.1016/ S0021-9258(19)83641-4 3. Gill S, June CH (2015) Going viral: chimeric antigen receptor T-cell therapy for hematological malignancies. Immunol Rev 263(1): 68–89. https://doi.org/10.1111/imr.12243 4. June CH, Sadelain M (2018) Chimeric antigen receptor therapy. N Engl J Med 379(1):64–73. https://doi.org/10.1056/NEJMra1706169 5. Peranzoni E, Bougherara H, Barrin S, Mansuet-Lupo A, Alifano M, Damotte D, Donnadieu E (2017) Ex vivo imaging of resident CD8 T lymphocytes in human lung tumor slices using confocal microscopy. J Vis Exp 130. https://doi.org/10.3791/55709 6. Nicolas-Boluda A, Donnadieu E (2019) Obstacles to T cell migration in the tumor microenvironment. Comp Immunol Microbiol Infect Dis 63:22–30. https://doi.org/10. 1016/j.cimid.2018.12.006 7. Salmon H, Franciszkiewicz K, Damotte D, Dieu-Nosjean MC, Validire P, Trautmann A, Mami-Chouaib F, Donnadieu E (2012) Matrix

architecture defines the preferential localization and migration of T cells into the stroma of human lung tumors. J Clin Invest 122(3): 8 9 9 – 9 1 0 . h t t p s : // d o i . o r g / 1 0 . 1 1 7 2 / JCI45817 8. Peranzoni E, Lemoine J, Vimeux L, Feuillet V, Barrin S, Kantari-Mimoun C, Bercovici N, Guerin M, Biton J, Ouakrim H, Regnier F, Lupo A, Alifano M, Damotte D, Donnadieu E (2018) Macrophages impede CD8 T cells from reaching tumor cells and limit the efficacy of anti-PD-1 treatment. Proc Natl Acad Sci U S A 115(17):E4041–E4050. https://doi.org/ 10.1073/pnas.1720948115 9. Kantari-Mimoun C, Barrin S, Vimeux L, Haghiri S, Gervais C, Joaquina S, Mittelstaet J, Mockel-Tenbrinck N, Kinkhabwala A, Damotte D, Lupo A, Sibony M, Alifano M, Dondi E, Bercovici N, Trautmann A, Kaiser AD, Donnadieu E (2021) CAR T-cell entry into tumor islets is a two-step process dependent on IFNgamma and ICAM-1. Cancer Immunol Res 9(12): 1425–1438. https://doi.org/10.1158/ 2326-6066.CIR-20-0837 10. Mo F, Mamonkin M (2020) Generation of chimeric antigen receptor T cells using Gammaretroviral vectors. Methods Mol Biol 2086: 119–130. https://doi.org/10.1007/978-10716-0146-4_8

Chapter 30 Measuring CTL Lytic Granule Secretion and Target Cell Membrane Repair by Fluorescent Lipophilic Dye Uptake at the Lytic Synapse Sabina Mu¨ller, Liza Filali, Marie-Pierre Puissegur, and Salvatore Valitutti Abstract CD8+ cytotoxic T lymphocytes (CTL) play a key role in anti-tumor immune response. They are therefore at the heart of current immunotherapy protocols against cancer. Despite current strategies to potentiate CTL responses, cancer cells can resist CTL attack, thus limiting the efficacy of immunotherapies. To optimize immunotherapy, it is urgent to develop rapid assays allowing to assess CTL-cancer cell confrontation at the lytic synapse. In this chapter, we describe a flow cytometry-based method to simultaneously assess the extent of CTL activation and of tumor cell reparative membrane turnover in CTL/target cell conjugates. Such a method can be performed using a limited number of cells. It can therefore be employed in clinical settings when only a few patient-derived cells might be available. Key words Cytotoxic T lymphocytes, Lytic synapse, Fluorescent lipophilic dyes, Plasma membrane turnover, Flow cytometry, Time lapse video microscopy

1

Introduction Perforin-mediated cytotoxicity is a key pathway used by human CTL to annihilate their target cells. Within minutes or seconds after productive TCR engagement, the pore-forming protein perforin, granzyme A and B and other enzymes stored in CTL cytoplasmic granules (named lytic granules) are secreted at the CTL/target cell lytic synapse [1–6]. Perforin-mediated penetration of granzyme A and B into target cells triggers a complex cascade of apoptotic and pyroptotic pathways leading to target cell death [4, 7].

Supplementary Information The online version contains supplementary material available at https://doi.org/ 10.1007/978-1-0716-3135-5_30. Cosima T. Baldari and Michael L. Dustin (eds.), The Immune Synapse: Methods and Protocols, Methods in Molecular Biology, vol. 2654, https://doi.org/10.1007/978-1-0716-3135-5_30, © The Author(s) 2023

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Recent observations revealed that target cells do not passively receive CTL-derived lytic components at the cell-cell contact site; on the contrary, they deploy rapid and efficient synaptic defense mechanisms to counteract CTL attack. We showed that melanoma cells are resistant to CTL-mediated cytotoxicity when compared to conventional cytotoxicity-sensitive target cells, thanks to a process of membrane reparation based on rapid lysosome exocytosis [2, 8]. The reparation process is triggered by perforin pore formation into target cell membrane and Ca2+ entry at the lytic synapse [3]. Lysosome exposure limits the efficacy of perforin-mediated cytotoxicity by inducing perforin degradation and rapid plasma membrane reparative turnover [2, 3]. Additional lines of evidence strengthened the notion of target cell resistance at the lytic synapse. Synaptic actin cytoskeleton polymerization in human breast cancer cells has been shown to limit NK cell-mediated cytotoxicity [9, 10]. The ESCRT-dependent repair mechanism has been identified as a key molecular machinery involved in membrane reparation in mouse cancer cells [11]. Lipophilic dies, such as FM1-43 and FM4-64, have been thoroughly used to monitor membrane turnover associated with cellular endo/exocytosis processes [12, 13] (Fig. 1). We showed that they can be used to visualize by time-lapse-microscopy, ongoing membrane recycling occurring at the lytic synapse during CTL-mediated cytotoxicity [3] and to quantify in a high throughput manner CTL and target cell membrane turnover by flow cytometry ([3]; L. Filali et al. filed patent WO2020109355). In the present chapter, we describe this method and its use to monitor, at the same time: (i) the membrane re-modelling occurring in CTL upon TCR engagement; (ii) the reparative turnover occurring in target cells upon CTL attack.

Fig. 1 Schematic representation of FM lipophilic dye uptake in plasma membrane upon secretive/reparation turnover. Non-fluorescent lipophilic dye (pink) stably intercalates into the lipid bilayer increasing in fluorescence intensity (red)

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Materials 1. HLA-A2-restricted human CD8+ CTL clones specific for the NLVPMVATV peptide of the cytomegalovirus protein pp65. CAUTION-human cell lines should always be treated as a biohazard following local regulations in approved facilities and with all associated precautions. 2. Clone medium: RPMI 1640 GlutaMAX medium supplemented with heat inactivated 5% human AB serum, 50 μM 2-mercaptoethanol, 10 mM Hepes, 1× MEM-Non-Essential Amino Acids Solution (Gibco or equivalent), 1× sodium pyruvate (Sigma-Aldrich or equivalent), 10 μg/mL ciprofloxacin (AppliChem or equivalent), 100 IU/mL human recombinant interleukin-2, and 50 ng/mL human recombinant interleukin15 (Miltenyi Biotec or equivalent). 3. Human Peripheral Blood obtained and processed following ethical procedures and with approval of local regulatory authorities. CAUTION- human blood products should always be treated as a biohazard following local regulations in approved facilities and with all associated precautions. 4. Ficoll®Paque Plus, Cytiva 17-1440-03 (GE17-1440-03, Sigma or equivalent). 5. Phosphate buffered saline without calcium and magnesium (PBS), (GIBCO or equivalent). 6. Sterile 15 mL or 50 mL polypropylene centrifuge tubes with screw caps. 7. Centrifuge with a swinging bucket rotor. 8. JY EBV-transformed B cell line (ATCC 77441 or equivalent). 9. D10 cell line (isolated from metastatic melanoma patients, kindly provided by Dr. G. Spagnoli, Basel, Switzerland, [2]). 10. Complete medium: RPMI 1640 GlutaMAX medium supplemented with heat inactivated 10% fetal calf serum and 50 μM 2-mercaptoethanol, 10 mM Hepes, 1× MEM-NEAA (Gibco or equivalent), 1× sodium pyruvate (Sigma-Aldrich or equivalent), and 10 μg/mL ciprofloxacin (AppliChem or equivalent). 11. MycoAlert mycoplasma detection kit (Lonza or equivalent). 12. Oregon Green™ 488 Taxol, Bis-Acetate (Tubulin Tracker™ Green, Invitrogen T34075 or equivalent). 13. 1 mM PKH67 in ethanol (Sigma PKH67GL or equivalent). 14. N-(3-Triethylammoniumpropyl)-4-(6-(4-(Diethylamino) Phenyl) Hexatrienyl) Pyridinium Dibromide, (FM™4-64 Dye, Invitrogen, T13320 or equivalent). 15. 15-well chambered μ-slide (Ibidi, Biovalley, 81506 or equivalent).

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16. 0.5 mg/mL Poly-D-Lysine hydrobromide in sterile tissue culture grade water (Sigma P6407 or equivalent). 17. Antigenic peptide: NLVPMVATV (CMV peptide p65 NV-9, GeneCust, Luxembourg or equivalent). 18. R5: RPMI-1640, 5% FCS, 1 mM HEPES. 19. Automated inverted microscope adapted for live cell time-lapse fluorescence microscopy with a temperature-controlled chamber maintained at 37 °C at constant 5% CO2 (see Note 1). 20. Computer workstation running ImageJ software. 21. Ethylenediaminetetraacetic disodium salt solution (EDTA), (Sigma 2854 or equivalent). 22. Sodium azide (Sigma-Aldrich, S2002 or equivalent). 23. N-(3-Triethylammoniumpropyl)-4-(4-(Dibutylamino) Styryl) Pyridinium Dibromide, (FM™ 1-43 Dye Invitrogen, T3163 or equivalent). 24. eBioscience™ Fixable Viability Dye eFluor™ 780, (Invitrogen, 65-0865-14 or equivalent). 25. CellTrace™ Violet Cell Proliferation Kit, for flow cytometry, (Invitrogen C34557 or equivalent). 26. 96-u-bottom tissue culture plates, (FALCON/CORNING, 353077 or equivalent) 24 well tissue culture plates (FALCON/CORNING, 353047 or equivalent). 27. FACS-Buffer: PBS, 1% heat inactivated foetal calf serum, 1% heat inactivated human serum, 0.1% sodium azide, 0.5 mM EDTA. 28. Flow cytometer (see Note 2).

3 3.1

Methods Cell Culture

1. Dilute heparin anticoagulated human blood from healthy donors 1:1 with RPMI (see Note 3). 2. Put 15 mL of Ficoll®Paque Plus solution into a 50 mL centrifuge tube and carefully overlay with up to 30 mL of diluted blood without disturbing the interface. 3. Centrifuge at 800 × g in a well-balanced swinging bucket rotor for 30 min with no acceleration and no break. 4. Carefully collect the cloudy layer of cells at the interface between the plasma and the Ficoll-Paque with a 5 mL pipette (= peripheral blood mononuclear cells or PBMC) and transfer to a fresh 50 mL centrifuge tube. Fill with RPMI and pellet cells at 400 × g for 15 min. 5. Discard supernatant, resuspend pelleted PBMC and fill tube with fresh sterile RPMI.

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6. Centrifuge at 400 × g 10 min. 7. Resuspend PBMC in 30 mL RPMI, take a sample for counting, centrifuge 180 × g for 10 min, discard supernatant and resuspend pellet to 1 × 106/mL in Clone medium. 8. Irradiate the PBMC at 35 Gy. (CAUTION- Use X-ray or γ-radiation with appropriate regulatory approval and safety procedures, see Note 4). 9. Stimulate CD8+ T cell clones Clone medium containing 1 μg/mL phytohemagglutinin with 1 × 106/mL irradiated PBMC. Use 24 well culture plates (2 mL final volume). 10. Do not touch the cultures for the first 3 days at least. 11. The irradiated PBMC will die after 2–3 days. 12. Maintain the viable CD8+ T cell clone cells at 1–2 × 106 per mL in Clone medium. Always use 24 well plates (or smaller if needed). 13. Restimulation of clones is routinely performed every 2–3 weeks. OK 3.2 Procedure for Time-Lapse Microscopy

1. Pulse target cells with 10 μM antigenic peptide for 2 h at 37 °C/5% CO2 in R5 and vortex tubes every 30 min (see Notes 5–7). 2. Wash the Ibidi chambers twice with sterile PBS. Add 40 μL of 0.05 mg/mL Poly-D-lysine (freshly diluted 1:10 v/v with sterile PBS) per well and leave for 10 min at 37 °C/5% CO2. 3. Flip the slide over and let it air dry for at least 10 min in the sterile tissue culture hood. 4. Rinse coated wells twice with sterile PBS, once with RPMI 1640 and once with R5 prior to use (see Note 8). 5. Stain CTL with 1 μM Tubulin Tracker™ Green (1:500, from stock solution in 10% Pluronic/90% DMSO) at 37 °C/ 5% CO2 in R5 for 30 min and vortex for a few seconds every 10 min. Alternatively, stain CTL with 2 μM PKH67 in R5 for 1–2 min at room temperature (see Notes 9–12). 6. Wash target cells and CTL in R5 3× at room temperature: spin 5 min at 400 × g twice. Wash once in R5 for 10 min at 180 × g (see Note 13). 7. Resuspend CTL at 80.000 cells/5 μL in R5. 8. Resuspend target cells at 20.000 cells/5 μL in R5. 9. At least 5 min before starting the imaging, mount the coated and washed ibidi chamber on the heated stage within the temperature-controlled chamber at 37 °C and constant 5% CO2 of the microscope and add 40 μL of R5. 10. Add the 5 μL of target cells (20.000 cells/well) into the preheated well (see Note 14).

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Fig. 2 Visualization of membrane turnover at the lytic synapse. Snapshots show FM4-64 (red) uptake at the contact sites between a melanoma cell and two cognate CTLs (loaded with PKH67, green) (see Note 15)

11. Set acquisition conditions on your microscope: adjust laser powers and time of recording (usually 30 min) and choose 1–3 s interval between images. 12. Check by visual inspection, that target cells have sedimented on the bottom of the slide. 13. Delicately open the chambered slide and add FM4-64 at a final concentration of 1 μg/mL into the well with the target cells (note that the maximum volume per well is 50 μL). 14. Add 5 μL of the stained CTL (80.000 cells/well) to the well (see Note 14). 15. Adjust focus and start acquisition. Figure 2 and Electronic Supplementary Movie 1 show typical experiments of FM uptake as monitored by time-lapse live cell imaging. 3.3 Procedure for Flow Cytometry

1. Pulse target cells with 10 μM antigenic peptide, or leave unpulsed, for 2 h at 37 °C/5% CO2 in R5. Vortex for a few seconds tubes every 30 min (see Notes 15–18). 2. Stain CTL with Cell Trace Violet (CTV, 1:1000, dilution according to manufacturer’s instructions, final concentration 10 μM) for 20 min at 37 °C/5% CO2 in R5. This staining allows to discriminate T cells from target cells in the gating strategy. 3. Wash target cells and CTL three times in R5: spin 5 min at 400 × g three times.

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4. Resuspend both pulsed and unplused target cells at 100.000 cells/25 μL in R5. 5. Distribute 25 μL of target cells (20.000 cells) into the wells of a 96-u-bottom plate for conjugation with the T cells (use one plate per time point: 2, 5, or 15 min). 6. Prepare a solution of FM1-43 at 20 μg/mL in R5 (see Note 19). 7. Resuspend CTL at 40.000 cells/25 μL in R5 and add them to the target cells in the 96-u bottom wells (2 CTL:1 target cell ratio). 8. Add 50 μL of the FM1-43 solution to each well containing the target and the T cells in a total volume of 50 μL (25 μL + 25 μL) to obtain a final concentration of 10 μg/mL of FM1-43. Keep unstained cells as control for flow cytometry. 9. Pellet cells by centrifuging the 96-u-bottom plates for 1 min at 400 × g and incubate at 37 °C/5% CO2 for 2, 5, or 15 min (see Note 20). 10. At the end of the different incubation times, put the plates on ice. 11. Add 100 μL of ice-cold FACS buffer to each well and disrupt conjugates by pipetting cell pellets in each well up and down (use a multichannel pipet). 12. Spin plates for 2 min at 400 × g at 4 °C and discard supernatant. 13. Wash plates twice by adding 200 μL of ice cold FACS-Buffer/well: spin plates at 400 × g for 2 min at 4 °C (see Note 21). 14. Dilute the Fixable Viability Dye eFluor™ 780 1:1000 in ice cold FACS-Buffer (50 μL/well) (see Note 22). 15. After washing, add 50 μL of the diluted Fixable viability dye eFluor780 per well and incubate 20 min on ice and in the dark. 16. Wash plates twice with 200 μL ice cold FACS-Buffer/well: spin at 400 × g for 2 min at 4 °C. 17. Resuspend the cells in 100 μL ice cold FACS-Buffer (see Note 23). 18. Acquire samples on a Flow Cytometer (see Note 24). See Fig. 3 for the gating strategy. 19. Once the two cell populations are identified as target cells and T cells, it is possible to study the uptake of the fluorescent dye on each cell population individually by applying the analysis on the differently gated populations. The geometric mean of FM fluorescence intensity is measured and presented as histogram plots (Figs. 4a, c and 5a, c) (see Note 25).

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Fig. 3 Gating strategy used for flow cytometry analysis. Left panels: Live cells were initially selected on side scatter/forward scatter criteria. Low staining fixable viability dye efluor-780, corresponding to live cells, were then selected. CTL were distinguished from target cells on the basis of their positivity for Cell Trace violet (CTV). Central panels: On each population we applied gates that allow excluding cell doublets. Right panels show histogram of FM1-43 fluorescence in the two cell populations

20. Pooled data from different experiments are expressed as geometric means ± SEM (Figs. 4b, d and 5b, d) (see Note 25). A dose–response curve of FM1-43 uptake in CTL interacting for 2, 5, or 15 min with JY cells pulsed with increasing concentrations of the antigenic peptide is show in Fig. 6. Data show the high sensitivity of the method (see Note 26).

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ll ll

ll ll

Fig. 4 FM1-43 internalization is enhanced after CTL attack in resistant and sensitive target cells. Time kinetics of FM1-43 fluorescence intensity in D10 or JY cells unpulsed or pulsed with 10 μM antigenic peptide. Analysis was performed on target cells either alone or following conjugation with CTL during 2, 5, or 15 min. (a, c) Typical flow cytometry plots of FM1-43 fluorescence intensity on melanoma cells (a) or JY cells (c). (b, d) Plots show time dependent FM1-43 uptake in melanoma cells (b) and in JY cells (d). Geometric mean fluorescence intensities of samples are indicated. Data are from four independent experiments realized in duplicate. Two-way ANOVA test using GraphPad Prism software was used to determine the statistical significance after 15 min of conjugation. **P < 0.01; ***P < 0.001. Reproduced after modification from L. Filali et al. filed patent; WO2020109355, 2020 with permission from INSERMTransfert

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l ll ll

l ll ll

Fig. 5 FM1-43 internalization is equally enhanced in CTL after activation by resistant or sensitive target cells (a, c) Typical flow cytometry plots of FM1-43 fluorescence intensity on CTLs interacting with melanoma cells (a) or JY cells (c). (b, d) Time kinetics of FM1-43 fluorescence intensity in CTL either alone or following conjugation with D10 (b) or JY cells (d) during 2, 5, or 15 min. Geometric mean fluorescence intensities of samples are indicated. Data are from four independent experiments. Results are from the same experiments shown in Fig. 4. Two-way ANOVA test using GraphPad Prism software was used to determine the statistical significance after 15 min of conjugation. **P < 0.01; ***P < 0.001. (Reproduced after modification from L. Filali et al. filed patent; WO2020109355, 2020 with permission from INSERMTransfert)

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Fig. 6 Typical flow cytometry plots showing dose response curves of FM1-43 uptake in CTL. FM1-43 uptake in CTL interacting for 2, 5, or 15 min with JY cells pulsed with increasing concentrations of the antigenic peptide is shown. The percentage of activated CTL is indicated. (Reproduced after modification from L. Filali et al. filed patent; WO2020109355, 2020 with permission from INSERMTransfert)

4

Notes 1. For example, we use an automated inverted microscope (Nikon) with a spinning-disk confocal scan head (Yokogawa) with a sCMOS Hamamatsu ORCA-Flash 4.0 V3 camera operated with MetaMorph software or a laser scanning confocal microscope (LSM780, Zeiss). On either system, we use a 63× objective (1.4 NA, oil immersion). 2. For example, we use a FACS MACSQuant 10 (Miltenyi Biotec) and analyze the resulting files using the FlowJo 10 software. 3. Ideally, the allogeneic PBMC should come from two or more donors to optimize performance of the feeder cells to promote growth of the CD8+ T cell clone. 4. The objective of the irradiation is to prevent T cells in the PBMC from dividing. If an appropriate radiation source is not available, 10 μg/mL mitomycin C for 1 h at 37 °C in complete medium followed by 5× washes can also be used to prevent division of feeder cells. 5. The combination of human HLA-A2–restricted clonal antigen specific CD8+ T cell clones with different HLA-A2+ antigenpresenting target cells, used in the above-described methods, is technically challenging and does not need to be the method of choice. It requires specialized cell culture and cloning skills which have been setup and adjusted over years of practice. It

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is possible to use polyclonal CD8+ T cell lines, purified from healthy donor blood samples using negative selection approaches and expanded with CD3/CD28 beads or CD3/CD28 antibodies coupled to tissue culture plates. Polyclonal T cells can be stimulated by conjugation with target cells pulsed with a cocktail of Bacterial Super Antigens (SAgs). SAgs bind directly to the T-cell receptor Vβ region and to the α1 domain of the MHC Class II molecules, bypassing the need for antigen processing and presentation [14]. More in general, this method can be applied to all cytotoxic lymphocytes (including anti-CD3 redirected target cell lysis, NK cell, CAR-T cells, etc.). 6. For successful live cell imaging it is important to make sure that cells are in optimal conditions (split target cells the day before the experiment, change medium, be gentle when suspending, avoid bubbles, etc.). Also, periodically check cell lines for mycoplasma using a sensitive test kit. 7. Perform experiments under sterile conditions. This will avoid bacterial contamination coming up during the time of imaging. 8. The choice of the chambered slides (shape, dimension, etc.) depends mainly on the type of mounted insert, on the heated stage of the microscope and on the number of available cells. 9. Staining with dyes, probes or antibodies depends a lot on the cell types (mouse, human, lines, primary cells, etc.). It is therefore important to assess the optimal concentrations, times of incubation, temperature, etc., in preparative experiments, to obtain the best results. 10. For experiments requiring cell fixation, it is possible to use a fixable FM analog (FM™ 1-43FX, fixable analog of FM™ 1-43 membrane stain, Invitrogen™ F35355). 11. Prepare all solutions and dye dilutions the same day of the experiment. The coated chambered slides can be kept at 4 °C for a few days. 12. During staining and prior to imaging, keep all the solutions and cells at 37 °C/5% CO2. 13. Cells should never be put on ice. This will slow down any cellular process involved in the interaction between T cells and target cells and signal transduction. It can bias the results of the experiment and also alter the localization of the loaded dyes. 14. The ready to use cells, loaded with the dyes, will metabolize dyes and lose staining over time (1–2 h if left at RT, best), quicker if put at 37 °C) and will form aggregates in the tube (not suitable for investigating single cell interactions). In the case several time-lapse acquisitions are planned, it is important to prepare sequential lots of freshly loaded cells.

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15. Here we describe the method to visualize FM day uptake at the synapse using time-lapse imaging. The membrane turnover occurring at both sides of the lytic synapse can be quantified by manual or automated image analysis as described in Filali et al. [3]. 16. Design your experiment in advance and calculate the number of cells needed to perform it. Make sure that cells are in good conditions. 17. Prepare cells for experiments under sterile conditions. 18. Keep R5 at 37 °C/5% CO2. 19. Prepare the right amount of FM1-43 solution in advance. 20. Form conjugates with cells that have been kept at 37 °C. Keep centrifuge at room temperature. Never spin at 4 °C before the co-culture. 21. Keep FACS-Buffer at 4 ° C (on ice). 22. Prepare the right amount of diluted Fixable Viability Dye eFluor™ 780, and keep it on ice. Any other way to distinguish dead cells from alive cells can be used (additional Viability Dye eFluor, 7-AAD, PI, etc.). 23. Stop incubation on ice and rapidly disrupt conjugates using ice-cold FACS-Buffer. 24. FACS data are analyzed using FlowJo 10 software, but any other software will do. 25. The results are represented here as the geomean values, but the percentage of activated CTLs can also be presented. 26. FM1-43 uptake exhibits an antigen dose response comparable to that exhibited by CD107a surface upregulation, a gold standard method for quantifying CTL lytic granule secretion [3]. Interestingly, FM uptake is a more rapid procedure, does not require staining with antibodies, and allows to detect in parallel target cell membrane turnover. References 1. Bertrand F, Muller S, Roh KH, Laurent C, Dupre L, Valitutti S (2013) An initial and rapid step of lytic granule secretion precedes microtubule organizing center polarization at the cytotoxic T lymphocyte/target cell synapse. Proc Natl Acad Sci U S A 110(15): 6073–6078. https://doi.org/10.1073/pnas. 1218640110 2. Khazen R, Muller S, Gaudenzio N, Espinosa E, Puissegur MP, Valitutti S (2016) Melanoma cell lysosome secretory burst neutralizes the CTL-mediated cytotoxicity at the lytic synapse. Nat Commun 7:10823. https://doi.org/10. 1038/ncomms10823

3. Filali L, Puissegur MP, Cortacero K, CussatBlanc S, Khazen R, Van Acker N, Frenois FX, Abreu A, Lamant L, Meyer N, Vergier B, Muller S, McKenzie B, Valitutti S (2022) Ultrarapid lytic granule release from CTLs activates Ca(2+)-dependent synaptic resistance pathways in melanoma cells. Sci Adv 8(7): eabk3234. https://doi.org/10.1126/sciadv. abk3234 4. McKenzie B, Khazen R, Valitutti S (2022) Greek fire, poison arrows, and scorpion bombs: how tumor cells defend against the siege weapons of cytotoxic T lymphocytes.

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Front Immunol 13:894306. https://doi.org/ 10.3389/fimmu.2022.894306 5. Cassioli C, Baldari CT (2022) The expanding arsenal of cytotoxic T cells. Front Immunol 13: 883010. https://doi.org/10.3389/fimmu. 2022.883010 6. Stinchcombe JC, Bossi G, Booth S, Griffiths GM (2001) The immunological synapse of CTL contains a secretory domain and membrane bridges. Immunity 15(5):751–761. https://doi.org/10.1016/s1074-7613(01) 00234-5 7. Tuomela K, Ambrose AR, Davis DM (2022) Escaping death: how cancer cells and infected cells resist cell-mediated cytotoxicity. Front Immunol 13:867098. https://doi.org/10. 3389/fimmu.2022.867098 8. Caramalho I, Faroudi M, Padovan E, Muller S, Valitutti S (2009) Visualizing CTL/melanoma cell interactions: multiple hits must be delivered for tumour cell annihilation. J Cell Mol Med 13(9B):3834–3846. https://doi.org/10. 1111/j.1582-4934.2008.00586.x 9. Al Absi A, Wurzer H, Guerin C, Hoffmann C, Moreau F, Mao X, Brown-Clay J, Petrolli R, Casellas CP, Dieterle M, Thiery JP, Chouaib S, Berchem G, Janji B, Thomas C (2018) Actin cytoskeleton remodeling drives breast cancer cell escape from natural killer-mediated cytotoxicity. Cancer Res 78(19):5631–5643. https://doi.org/10.1158/0008-5472.CAN18-0441 10. Wurzer H, Filali L, Hoffmann C, Krecke M, Biolato AM, Mastio J, De Wilde S, Francois

JH, Largeot A, Berchem G, Paggetti J, Moussay E, Thomas C (2021) Intrinsic resistance of chronic lymphocytic leukemia cells to NK cell-mediated lysis can be overcome in vitro by pharmacological inhibition of Cdc42induced actin cytoskeleton remodeling. Front Immunol 12:619069. https://doi.org/10. 3389/fimmu.2021.619069 11. Ritter AT, Shtengel G, Xu CS, Weigel A, Hoffman DP, Freeman M, Iyer N, Alivodej N, Ackerman D, Voskoboinik I, Trapani J, Hess HF, Mellman I (2022) ESCRT-mediated membrane repair protects tumor-derived cells against T cell attack. Science 376(6591): 377–382. https://doi.org/10.1126/science. abl3855 12. Brumback AC, Lieber JL, Angleson JK, Betz WJ (2004) Using FM1-43 to study neuropeptide granule dynamics and exocytosis. Methods 33(4):287–294. https://doi.org/10.1016/j. ymeth.2004.01.002 13. Gaffield MA, Betz WJ (2006) Imaging synaptic vesicle exocytosis and endocytosis with FM dyes. Nat Protoc 1(6):2916–2921. https:// doi.org/10.1038/nprot.2006.476 14. Esquerre M, Tauzin B, Guiraud M, Muller S, Saoudi A, Valitutti S (2008) Human regulatory T cells inhibit polarization of T helper cells toward antigen-presenting cells via a TGFbeta-dependent mechanism. Proc Natl Acad Sci U S A 105(7):2550–2555. https://doi. org/10.1073/pnas.0708350105

Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made. The images or other third party material in this chapter are included in the chapter’s Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter’s Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

Chapter 31 In Vitro Generation of Human Tolerogenic Monocyte-Derived Dendritic Cells Catharien M. U. Hilkens, Julie Diboll, Fiona Cooke, and Amy E. Anderson Abstract Human monocyte-derived dendritic cells (moDC) are commonly used as a research tool to investigate interactions between antigen-presenting cells and T cells. Generation of these cells involves the isolation of CD14 positive monocytes from peripheral blood and their in vitro differentiation into immature moDC by the cytokines GM-CSF and IL-4. Their functional characteristics can then be manipulated by maturing these cells with a cocktail of agents, which can be tailored to induce either immune activating or tolerogenic properties. Here, we describe a protocol for the generation of moDC with stable tolerogenic function, referred to as tolerogenic dendritic cells. These cells have been developed as an immunotherapeutic tool for the treatment of autoimmune disease but have also proven useful to dissect mechanisms of T cell tolerance induction in vitro. Key words Human, Monocytes, Dendritic cells, Tolerogenic dendritic cells, Peripheral blood mononuclear cells, Interleukin-4, Granulocyte-macrophage colony stimulating factor, Dexamethasone, 1,25-dihydroxy-vitamin D3, Toll-like receptor 4 ligand

1

Introduction Dendritic cells (DC) are antigen-presenting cells that play a central role in the activation and regulation of T cells. They reside in healthy tissues as immature cells that are non-immunogenic. In response to environmental signals, they mature and acquire ability to efficiently activate and expand antigen-specific T cells. DC were discovered in the 1970s, but the field only rapidly expanded in the 1990s when in vitro protocols for culturing and expanding mouse and human DC from progenitor cells became available [1–3]. For human DC, a widely used protocol is the in vitro culture of peripheral blood monocytes in the presence of GM-CSF and IL-4 to induce differentiation of DC [3]. Nowadays, these DC are commonly referred to as monocyte-derived DC (moDC) to distinguish them from the classical DC (cDC) types that arise from specific pre-DC precursors in vivo. Nevertheless, despite their

Cosima T. Baldari and Michael L. Dustin (eds.), The Immune Synapse: Methods and Protocols, Methods in Molecular Biology, vol. 2654, https://doi.org/10.1007/978-1-0716-3135-5_31, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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different ontogenies, moDC share morphological, phenotypical, and functional features with cDC. These include the presence of dendrites, expression of typical DC markers (e.g., CD1c, CD11c, HLA-DR), and, depending on their maturation status, potent ability to activate effector T cells and/or induce T-cell tolerance. As a result, they have become a useful in vitro model to understand the mechanisms by which DC activate or regulate antigen-specific T-cell responses. Early studies mainly focused on the immunogenic properties of moDC, for example, the accumulation of MHCII-peptide complexes [4], and the production of bioactive IL-12p70 [5]. But as it became clear that moDC could be manipulated to gain immunosuppressive and tolerogenic properties [6], they became a target for the development of moDC with stable tolerogenic function (coined ‘tolerogenic DC’ in general, but for clarity we will refer to these cells as tolerogenic moDC). These cells have been extensively used as research tools to study mechanisms of T-cell tolerance, for example, the activation/expansion of regulatory T cells or the induction of T-cell anergy. They have also been developed as cell therapies for the treatment of autoimmune diseases and for the prevention of transplant rejection [7]. A variety of methods have been described by which stable tolerogenic function can be induced in moDC [8, 9]. These include genetic engineering to either silence or enforce the expression of key molecules of interest, but a more popular and straightforward method is to treat moDC with immunosuppressive agents. There is an ever-growing list of agents that can induce tolerogenic properties in moDC; the most commonly used are glucocorticoids, 1,25dihydroxy-vitamin D3, rapamycin, and IL-10. Many of these agents act to some extent by inhibiting or modulating NFκB signalling, a key transcriptional pathway for maturation of DC into immunogenic cells. However, although there are some commonalities (e.g., reduced expression of co-stimulatory molecules, antiinflammatory cytokine profile), a common tolerogenic transcriptomic “signature” cannot be defined among the various tolerogenic moDC types. In fact, transcriptomic profiles differ greatly between these cell types [9], and it is therefore likely that they exert their immunoregulatory actions through both overlapping and distinct pathways. Here, we describe a protocol to generate human tolerogenic moDC. The protocol involves extraction of peripheral blood mononuclear cells (PBMC) through density centrifugation, followed by the isolation of CD14 positive monocytes using magnetic CD14 microbeads. Cells are subsequently cultured for a period of 7 days in the presence of GM-CSF and IL-4 to induce differentiation of moDC. To generate tolerogenic moDC, the immunosuppressive drugs dexamethasone and 1,25-dihydroxy-vitamin D3 (active form of vitamin D3) are added to the culture at day 3 (dexamethasone) and day 6 (dexamethasone and 1,25-dihydroxy-

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vitamin D3). In addition, on day 6 the cells are treated with a Tolllike receptor (TLR)-4 ligand to enhance their antigen-processing and migratory capacity [10]. As a control, mature moDC are generated by treatment with a TLR4 ligand in the absence of the immunosuppressive drugs. This protocol of tolerogenic moDC generation has been used to investigate regulatory effects on naı¨ve and memory CD4+ T cells [10–13] and has been translated into a Good Manufacturing Practice (GMP)-compliant therapeutic cell product that has undergone safety testing in a clinical trial in inflammatory arthritis patients [14, 15]. The main characteristics of these tolerogenic moDC are summarized in Table 1.

Table 1 Characteristic cell surface and cytokine profiles of human mature and tolerogenic moDC cultured according to the protocol described in this Chapter. 1Cell surface markers were determined by flow cytometry and 2cytokine production by ELISA, except for TGFβ1 which was determined on the basis of mRNA expression (Taqman low density array). Markers and cytokines in bold are recommended for minimum quality control of tolDC products. Markers and cytokines may vary between individual donors and culture medium used. matDC: mature moDC; tolDC: tolerogenic moDC. ++ high expression; + moderate expression; -/+ low or non-detectable expression; - non-detectable expression Cell surface markers1

Cytokine production2

Marker

matDC

tolDC

Cytokine

matDC

tolDC

CD1c

++

+

IL-1β

++

-/+

CD11c

++

++

IL-6

++

+

CD14

-/+

-/+

IL-10

+

++

CD32

+

++

IL-12p70

++

–

CD40

++

+

IL-23

++

-/+

CD80

++

++

TNF

++

+

CD83

++

-/+

TGFβ1

+

++

CD86

++

+

CD206

+

++

CCR7

+

-/+

CXCR4

+

+

HLA-DR

++

++

PDL-1

++

++

MerTK

-/+

++

TLR-2

-/+

+

TGFβ-LAP

-/+

++

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Materials All reagents and plastics are sterile, tissue culture-grade. Store reagents at 4–8 °C (unless indicated otherwise).

2.1 Peripheral Blood Mononuclear Cell (PBMC) Isolation

1. Blood diluent: Hanks Balanced Salt Solution (HBSS) containing 2 mM EDTA. Add 2 mL endotoxin-free 0.5 M EDTA to a 500 mL bottle of calcium/magnesium-free HBSS. Store and use at room temperature. 2. Density gradient medium: 9.1% w/v sodium diatrizoate, 5.7% w/v Ficoll® 400 or equivalent 400,000 g/mol polysucrose to a final density of 1.077 g/mL, e.g., Ficoll-Paque™ or Lymphoprep™. Store at room temperature, protected from light (see Note 1). 3. Washing medium: HBSS containing 1% foetal calf serum (FCS). Add 5 mL heat-inactivated FCS (see Note 2) to a bottle of calcium/magnesium-free HBSS. 4. Trypan blue viability dye: 0.4% solution in 0.85% NaCl. 5. 50 mL polypropylene centrifuge tubes (see Note 3). 6. 3 mL plastic Pasteur pipettes. 7. 70 μm nylon cell strainer. 8. Haemocytometer and glass coverslip (see Note 4).

2.2 Monocyte Isolation

1. 25–30 mL polystyrene universal tubes with a v-bottom (see Note 5). 2. MACS buffer: Dulbecco’s Phosphate Buffered Saline (PBS; 8.0 g/L NaCl, 0.2 g/L KCl, 0.2 g/L KH2PO4), pH 7.2, containing 0.5% FCS and 2 mM EDTA. Add 2.5 mL FCS and 2 mL 0.5 M endotoxin-free EDTA to a 500 mL bottle of calcium/magnesium-free PBS and filter through a 0.2 μm filter unit (see Note 6). Store at 4 °C and chill on ice before use. 3. Miltenyi MACS™ human CD14 MicroBeads. 4. Miltenyi MACS™ magnetic separator for use with LS columns (see Note 7). 5. Miltenyi LS Column. 6. Ice bucket and ice. 7. RF10: RPMI-1640 medium (see Note 8) supplemented with 10% FCS, 2 mM L-glutamine (see Note 9), 100 U/mL penicillin, and 100 μg/mL streptomycin.

2.3 Generation and Culture of MonocyteDerived Dendritic Cells (moDC)

1. RF1: RPMI-1640 medium, 1% FCS, 2 mM L-glutamine, 100 U/mL penicillin, and 100 μg/mL streptomycin. 2. Recombinant human interleukin-4 (IL-4) and granulocytemacrophage colony stimulating factor (GM-CSF).

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Reconstitute cytokines according to manufacturer’s recommendations and then dilute to 5 μg/mL using RF1, aliquoted and stored at -20 °C (see Notes 10 and 11). 3. 1 × 10-4 M dexamethasone aliquoted and stored at -20 °C (see Note 12). 4. 1 × 10-4 M 1,25-dihydroxy-vitamin D3 aliquoted and stored at -20 °C (see Note 13). 5. Lipopolysaccharide (LPS): Reconstitute LPS to 1 mg/mL in RF1. Store suitable sized aliquots at -20 °C (see Note 14). 6. 24 well, flat bottom, polystyrene tissue culture treated plate (s) (see Note 15).

3

Methods

3.1 Peripheral Blood Mononuclear Cell (PBMC) Isolation

1. Collect blood by venepuncture into sterile, evacuated blood tubes containing EDTA as an anticoagulant, ensuring the blood is thoroughly mixed with the anticoagulant by inverting the vacuum tube several times—do not shake. 2. Dilute blood at a 1:1 ratio with room temperature blood diluent (see Note 16). 3. Transfer 15 mL density gradient medium into a 50 mL centrifuge tube and gently layer 15–25 mL of diluted blood, without mixing the two phases (see Note 17). Use more than one tube if the volume of diluted blood is >25 mL. 4. Centrifuge at 900 × g, room temperature for 30 min, with the brake off or reduced to a low setting (see Note 18). 5. Using a 3 mL plastic Pasteur pipette harvest the PBMC layer from the interface (see Note 19) and transfer to a new 50 mL centrifuge tube. Cells from two tubes may be pooled into one tube at this point to carry out the washing steps. 6. Top up the tube(s) to 50 mL with cold washing medium and centrifuge at 600 × g, 4 °C for 8 min with the brake on (see Note 20). 7. Check the bottom of the tube for a cell pellet (see Note 21). 8. Aspirate the supernatant using a pipette or by pouring into a waste container (see Note 22) and resuspend the cell pellet thoroughly (see Note 23). If you have more than one tube, pool cells into one tube at this point. 9. Top up the tube to 50 mL with cold washing medium and centrifuge at 300 × g, 4 °C for 8 min with the brake on (see Note 24). 10. Discard the supernatant and resuspend the cell pellet as described previously.

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11. Top up the cell suspension to 50 mL with cold washing medium. Return the cells to ice. 12. Using a 25 mL serological pipette, gently pipette the cell suspension through a 70 μm nylon cell strainer into a new 50 mL centrifuge tube to remove any debris or cell clumps. 13. Count the cells using a haemocytometer counting chamber (see Note 4). 3.2 Monocyte Isolation

1. Transfer the required number of PBMC (see Note 25) to a suitably sized centrifuge tube (see Note 26), top up with cold washing medium, and centrifuge at 400 × g, 4 °C for 8 min, brake on. 2. Discard the supernatant and resuspend the cell pellet as described previously. 3. Resuspend PBMC in ice-cold MACS buffer (90 μL MACS buffer per 10 × 106 PBMC) (see Note 27). 4. Add 10 μL Miltenyi MACS™ human CD14 MicroBeads per 10 × 106 cells (see Note 28). Gently swirl the tube to mix. 5. Incubate at 4 °C for 15 min (see Note 29). Gently agitate the tube every 5 min. 6. Wash away the unbound MicroBeads by adding 25–50 mL ice-cold MACS buffer and centrifuge at 400 × g, 4 °C for 8 min, brake on. 7. While the cells are centrifuging place a LS column into the MACS separator with a 25–30 mL universal tube directly below the column nozzle. 8. Pre-rinse the column by adding 3 mL ice-cold MACS buffer to the column and allow to flow through into the tube below. 9. Once the cells have finished centrifuging, check for the presence of a cell pellet and discard the supernatant and resuspend the cell pellet as described previously. 10. Add 2 mL ice-cold MACS buffer to the pellet and resuspend the cells gently but thoroughly by pipetting up and down with a P1000 pipette. 11. Transfer the cell suspension to the LS column, avoiding air bubbles, and allow the unlabelled cells to pass through into the tube below. 12. Wash the column three times by adding 3 mL ice-cold MACS buffer, allowing it to flow through completely each time to ensure complete removal of the unlabelled cells. This tube contains CD14 negative cells (see Note 30). 13. Remove the LS column from the magnetic separator and place into a 25–30 mL universal tube. Add 5 mL MACS buffer to the

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column and push through using the supplied plunger to flush the labelled cells from the column. This tube contains the CD14 positive monocytes. 14. Top up the tube containing the CD14 positive monocytes with MACS buffer and centrifuge at 400 × g, 4 °C for 8 min with the brake on. 15. Discard the supernatant and resuspend the cell pellet as described previously. 16. Resuspend the cells in 5 mL cold RF10 (see Note 31) and count the cells (see Note 32). 3.3 Generation and Culture of tolDC

Day 0—Seed Monocytes

1. Resuspend the required number of monocytes in cold RF10 (see Note 31) at a concentration of 0.5 × 106/mL. 2. Thaw freezer stock aliquots (5 μg/mL) of IL-4 and GM-CSF and add the required volume to the CD14 positive cell suspension to give the final concentrations of 50 ng/mL of each cytokine. 3. Seed 1 mL cells per well into a 24 well tissue culture plate and incubate at 37 °C in 5% CO2 (see Note 33). Day 3—Refresh Culture Medium

4. Remove the plate of cells from the incubator and examine the cells under a microscope to check cell health and for signs of contamination. Return cells to incubator whilst preparing reagents. 5. Warm the RF10 culture medium to 37 °C in a water bath (see Note 34). 6. Thaw freezer stock aliquots of IL-4 and GM-CSF (both 5 μg/ mL) and dexamethasone (1 × 10-4 M). 7. Prepare refresh medium: RF10 culture medium containing IL-4 and GM-CSF at 100 ng/mL, and dexamethasone at 2 × 10-6 M (see Note 35). 8. Carefully remove 450 μL medium from each well and discard into a waste container containing disinfectant (see Note 36). 9. Add 500 μL of refresh medium to each well (see Note 37) and return the plate to incubator (see Note 33). Day 6— Add Further Tolerizing Agent and Maturation Stimulus

10. Remove the plate of cells from the incubator and examine the cells under a microscope to check cell health and for signs of contamination. Return cells to the incubator whilst preparing reagents.

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11. Warm the RF10 culture medium to 37 °C in a water bath (see Note 34). 12. Thaw freezer stock aliquots of dexamethasone (1 × 10-4 M), 1,25-dihydroxy-vitamin D3 (1 × 10-4 M), and LPS (1 mg/ mL). 13. Prepare a 1 × 10-8 M working stock of 1,25-dihydroxy-vitamin D3 as follows: (a) Dilute the 1 × 10-4 M freezer stock to 1 × 10-6 M in 100% ethanol (intermediate stock) (see Note 38). (b) Dilute the 1 × 10-6 M intermediate stock to 1 × 10-8 M in RF10 culture medium (working stock). 14. Prepare a 10 μg/mL working stock of LPS using RF1 as the diluent (see Notes 14 and 39). 15. Prepare tolerizing/maturation medium: RF10 containing 1 × 10-5 M dexamethasone (use the 1 × 10-4 M freezer stock), 1 × 10-9 M 1,25-dihydroxy-vitamin D3 at (use the 1 × 10-8M working stock) and 1 μg/mL LPS (use the 10 μg/mL working stock) (see Notes 40 and 41). 16. Add 100 μL of the tolerizing/maturation medium to each well (see Note 42) and return the plate to the incubator for 16–24 h. Day 7— Harvest Cells for Downstream Use

17. Remove the plate of cells from the incubator and examine the cells under a microscope to check cell health, cell morphology (see Note 43), and for signs of contamination. 18. Incubate the plates on ice for 1 h to loosen the cells from the plastic (see Notes 44 and 45). 19. Harvest the cells from each well into an appropriately sized centrifuge tube (see Note 26) by gently pipetting and scraping the bottom of the well (see Notes 14 and 46). 20. Top up the tube with ice cold washing medium and centrifuge at 400 × g, 4 °C for 8 min, brake on. 21. Once the cells have finished centrifuging, check for the presence of a cell pellet and discard the supernatant and resuspend the cell pellet as described previously. 22. Top up the tube with ice cold washing medium and centrifuge as before. 23. Repeat the wash steps a further two times (four washes in total) (see Note 47). 24. Resuspend the cells in a small volume of cold RF10 and count with trypan blue.

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Notes 1. Prolonged exposure to direct sunlight leads to release of iodine from the sodium diatrizoate molecule. 2. Select FCS (also called fetal bovine serum (FBS)) with low endotoxin levels and/or test different FCS batches to ensure optimal cell viability, yield, and a low level of “spontaneous” maturation of the moDC. All FCS must be heat-inactivated at 56 °C for 1 h to inactivate complement. 3. Polypropylene centrifuge tubes reduce the loss of cells due to adhesion to the tube wall. 4. A suitable haemocytometer and associated counting method should be used, such as a Neubauer or Bu¨rker counting chamber. Alternatively, an automated cell counting method may be used. 5. We recommend using Starlab 30 mL universal tubes or a similar tube with a v-bottom. Do not use more than 25 mL volumes with these tubes. 6. We recommend filtration through a 0.2 μm filtration unit to remove any particulate matter, e.g., FCS precipitation, prior to use. Excess gas in the buffer may form bubbles in the column matrix during separation, which can decrease the efficiency of separation. If possible, the buffer should be de-gassed by stirring with a magnetic stir bar under a low vacuum until bubbles stop forming (this can take over an hour). 7. Any suitable Miltenyi MACS magnet system, for example, a MidiMACS or VarioMACS with LS adapter. If multiple isolations are being performed in parallel, e.g., from multiple donors, we recommend using the QuadroMACS system for efficiency. 8. We recommend that you use RPMI-1640, which contains phenol red as an indicator of pH. At near neutral pH the medium will have a pink/orange color. A build-up of waste products from metabolizing cells or growth of contaminants will lead to acidification of the medium, indicated by a color change to orange then yellow. Above pH 8, phenol red turns a bright pink (fuchsia) color and is usually indicative of medium that is “old,” i.e., a breakdown of the glutamine and/or loss of CO2 due to air exchange, and should therefore be discarded. 9. L-glutamine is required to support the growth of cells in culture. It is unstable at physiological pH in liquid culture medium and degrades over time, breaking down to form ammonia and pyroglutamate. We recommend adding fresh glutamine to RF10 culture medium after 1 month if you wish to continue using the medium. Note that it is only possible to

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do this on one occasion and medium should be discarded if not used up within a 2-month period as the pH may increase to an unsuitable level due to the ammonia breakdown products. 10. We reconstitute cytokines (from various commercial sources) as follows: bring the vial containing the lyophilized cytokine to room temperature and centrifuge briefly to ensure no powder remains in the cap. Reconstitute to 5 μg/mL in RF1, by pipetting a small amount of RF1 (e.g., 200 μL) into the vial, mixing by pipetting up and down gently, but thoroughly, and transferring the content of the vial into a centrifuge tube with the appropriate volume of RF1. Repeat this procedure to ensure that all of the content is transferred to the centrifuge tube. Once reconstituted, sterile filter if necessary, and freeze at -20 °C in appropriate size aliquots. 11. Once thawed, cytokines in RF1 are stable at 4–8 °C for 1 week only. Discard after this time, do not re-freeze. So appropriate aliquot size is amount that is likely to be used in 1 week. 12. Follow the manufacturer’s recommendation for the initial reconstitution of dexamethasone. We mostly use dexamethasone powder from Sigma (D4902) and dissolve it in absolute ethanol to 1 mg/mL (2.55 × 10-3 M), then dilute it 25.5 times in RPMI-1640 medium to 1 × 10-4M, and freeze at 20 °C in appropriate size aliquots. Alternatively, water-soluble dexamethasone sodium phosphate is suitable as well and is available as a clinical-grade solution (dexamethasone 3.3 mg/ mL for injection). 13. We recommend reconstituting 1,25-dihydroxy-vitamin D3 in 100% ethanol and storing at -20 °C for up to 1 month only. 1,25-dihydroxy-vitamin D3 is light-sensitive and should be protected from exposure to light. 14. When working with LPS, use filter tips to prevent contamination of pipettes. This includes the harvesting stage of the protocol on Day 7. 15. We recommend using Corning Costar™ 24 well cell culture plates. We have noted changes in cell morphology when using products from alternative manufacturers. 16. This technique is intended to be used for isolation of PBMC from whole blood (healthy and patient samples) but may be used to harvest PBMC from buffy coats or Leucocyte Reduction System (LRS) cones with a minor modification to the dilution step. Both buffy coats and LRS cones contain highly concentrated numbers of leucocytes and therefore require a greater dilution in order to reduce the occurrence of aggregates and obtain good yields of pure PBMC. We suggest diluting buffy coats 1:2 or 1:3, and leucocyte cones 1:4 prior to layering. PBMC yields are optimal from blood drawn

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Fig. 1 Layering diluted blood onto the density gradient medium. (a) Hold the centrifuge tube as close to a horizontal position as possible (without pouring out the tube contents) to increase the surface area of the density gradient medium and reduce mixing when layering. Place the pipette containing the diluted blood at the top of the tube at an approximate 90° angle to the tube. (b) Slowly pipette the blood into the tube. (c) As the blood covers the density gradient medium the tube can be tilted to more of a vertical position and the pipette angle decreased. (d) The diluted blood should sit on top of the density gradient medium without mixing

between 30 min and 6 h prior to separation. If used immediately after being drawn the blood may be too warm resulting in reduced PBMC yields. A useful guide for the expected PBMC yield from each source is 1–2 × 106 PBMC per mL of whole blood, 400–1000 × 106 PBMC per buffy coat or 400–1500 × 106 PBMC per LRS cone. 17. For efficient separation, it is important that the blood does not mix with the density gradient medium. For a description of how to layer the blood, see Fig. 1. 18. Temperature is an extremely important factor when carrying out this method. Samples, density gradient medium, and the initial centrifugation step must all be at room temperature. We find that during the winter months in the north of England the density centrifugation medium needs to be warmed slightly to avoid variations in density which ultimately would lead to granulocyte contamination of the PBMC population. Take care not to heat the medium too much, e.g., by leaving in a 37 °C water bath for any length of time as this will also alter the density of the medium with detrimental effects such as reduced PBMC yield or platelet contamination. It is important to use a slow deceleration for this centrifugation step to prevent mixing of the separated layers. 19. The blood will separate into a red blood cell/granulocyte pellet, density gradient medium layer, PBMC interphase layer and plasma/blood diluent layer (Fig. 2). Hold the tube upright and insert the Pasteur pipette through the plasma layer until you reach the “fluffy” cell layer sitting on top of the density gradient medium. Gently move the pipette around the perimeter of the tube to ensure collection of the entire cell layer. 20. When left in contact with cells for extended periods, density gradient media are toxic to cells. This first wash step removes any contaminating medium and should be performed immediately after harvesting the cells.

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Plasma/blood diluent layer

PBMC interphase layer Density gradient medium

Red blood cell/granulocyte pellet

Fig. 2 Appearance of blood separated over a density gradient medium

21. The cell pellet should be visible to the naked eye and will usually be off-white in color. Occasionally, the presence of contaminating red blood cells can be seen around the edge of the pellet. These red cells are usually low in number when working with blood from “healthy” donors. Factors which may affect this and cause increased numbers of contaminating red cells include blood which is >24 h old, blood obtained from individuals with inflammatory diseases, and/or those on certain drug treatments, or where there were inconsistencies with the venipuncture procedure. 22. Ensure all waste is discarded into a disinfectant solution prior to discarding. Check your local waste disposal regulations for further information. 23. Resuspend cells gently but thoroughly. We recommend that once the supernatant has been removed, and only a small volume ( 0). 3.3 Generation of Primary CAR T Cells

1. Prepare T cell media as in Materials 2.19. Store the medium at 4 °C and warm it to 37 °C before usage. 2. (Day 1) Count donor PBMC number. Plate 7.5 × 106 cells per 24-wells well in 2 mL T cell media also containing 300 IU/mL IL2 and 50 ng/mL OKT3 (see Note 4). 3. (Day 2) Coat non-tissue culture treated 6-well plates with retronectin at a final concentration of 20 μg/mL in 1.5 mL DPBS per well. Wrap the plate with plastic wrap and incubate plate overnight at 4 °C (see Note 5). 4. (Day 3) Warm-up centrifuge to 32 °C. 5. Remove retronectin solution from the 6-wells plate then block the plate with 2 mL 2% BSA in DPBS for 30 min at room temperature. 6. Thaw a lentivirus stock on rack at room temperature and place the stock on ice. 7. Wash each well with 2 mL DPBS, leave the wash in well. 8. Prepare viral medium (MOI = 50) in T cell culture media (see Note 6). 9. Remove the DPBS and add 2 mL/well viral medium. 10. Wrap the plate with plastic wrap and spin the plate at 2000 × g for 2 h at 32 °C. 11. During the spin, harvest, wash, and count T cells to be transduced (see Note 7). 12. When the centrifugation of plates is finished, remove the viral medium. 13. Add 2 × 106 of T cells/well in the T cell culture media (300 IU/mL IL2) at the concentration of 1 × 106 cells/well.

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14. Wrap the plate, and spin for 10 min at 500 × g with the acceleration and brake set at lowest setting. 15. Gently remove the plate from the centrifuge and leave it in the incubator overnight. 16. (Day 4) Transfer transduced cells onto a new tissue culturetreated 6-well plate to get rid of retronectin. Cells from the identically treated wells can be combined into a T25 or T75 Flask at concentration 0.5 × 106 cells/well. 17. Verify CAR-GFP expression using fluorescence microscopy if available. 18. (Day 6) Flow cytometry analysis of CAR-GFP expression 3 days post-transduction (see Note 8). 19. Split cells if they are growing well and reaching a density of 0.5 ~ 1 × 106 cell/mL. 3.4 Culturing Primary CAR T Cell and Raji B-mCherry-CAAX Cells

1. Thaw and culture Raji B-mCherry-CAAX cells (Raji B expressing the membrane marker mCherry-CAAX) in RPMI1640, 10% heat-inactivated FBS, and 1% PSG for 3 ~ 5 days before performing the conjugation assay. 2. Count cell number of Raji B-mCherry-CAAX and primary CAR T cells. 3. Wash cells with DPBS once and re-suspend in RPMI1640 medium supplemented with 20 mM HEPES pH 7.4 at a cell density of 2 × 107 cells/mL. 4. Co-culture Raji B- mCherry-CAAX and primary CAR T cells at a 1:1 ratio in RPMI1640 medium supplemented with 20 mM HEPES pH 7.4 for 0.5 h at 37 °C (Fig. 1a) (see Note 9). 5. Fix cells in 4% paraformaldehyde for 15 min at room temperature. 6. Wash cells with DPBS and stain with an anti-CD45-APC antibody in FACS staining buffer for 30 min on ice. 7. Wash cells with DPBS, re-suspend in RPMI1640 medium supplemented with 20 mM HEPES, and image by confocal microscopy.

3.5 Imaging CAR T Cell-Tumor Cell Conjugation and CD45 Exclusion in the Synapse

1. Perform confocal microscopy. For the images shown in Fig. 1, this was performed on a Nikon Ti2-E inverted motorized microscope stand equipped with a motorized stage with Yokogawa CSU-X1 spinning disk confocal, Agilent laser combiner with four lines, 405, 488, 561, and 640 nm, and scientific CMOS camera Photometrics Prime 95B. 2. Perform imaging on a 96-well glass-bottom plate or equivalent. 3. Add ~0.1 × 106 cells to the well in 200 μL imaging buffer and wait for 10 min to allow cells to settle down (see Note 10).

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Fig. 1 (a) Schematics of co-culture of CAR-T cells with Raji B cells. (b) Confocal microscopy revealing the synapse formed between a T cell expressing CAR-GFP and a Raji B cell expressing mCherry-CAAX. CD45 was stained with an APC-conjugated anti-CD45 antibody. (c) Fluorescence intensities of CAR and CD45 in the synapse

4. Acquire images. The images shown in Fig. 1 were acquired using Nikon Elements and analyzed in Fiji (ImageJ). The same brightness and contrast were applied to images within the same panels (Fig. 1b). 5. To analyze cell conjugation rate, score CAR-T cells with or without binding to Raji-B cells by the Image J plugin Cell Count. Conjugation percentage = number of CAR-T binding to Raji cells/total CAR-T number) ×100%. 6. To measure the level of CD45 exclusion from the synapse draw a segmented line over the synapse formed between the CAR-T and Raji B cell. Use a line scan function to measure the pixel intensities of CAR-GFP and CD45-APC (Fig. 1c). The exclusion percentage = (1 – Intensity of CD45 in CAR zone/Intensity of CD45 out CAR zone) ×100%.

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Notes 1. Raji B-mCherry-CAAX cells: expressing a membrane marker mCherry-CAAX, which is established by lentiviral infection and mCherry flow sorting (plasmid: pHR-mCherry-CAAX). 2. This is a lentiviral vector encoding the third generation of CD19 CAR with a GFP tag on the C-terminus. 3. Depending on the number of viruses to be generated, the seeding cell number may vary: e.g., 0.8 × 106 cells per well in a 6-well plate, 4 × 106 cells in a 100 mm plate, and 10 × 106 cells in a 150 mm plate. 4. OKT-3 antibody could be replaced by Dynabeads™ Human T-Activator CD3/CD28 (ThermoFisher, #11161D or equivalent). Prepare the beads according to the user manual. 5. Retronectin incubation could be done at 37 °C in a 5% CO2 incubator for 4–6 h. 6. MOI could be adjusted (~5–100) based on your virus titer and total amount. Assuming the virus titer is 1 × 108 TU/mL (or 1 × 105 TU/μL), 1 mL of the viral suspension needs to be added to 1 × 107 cells to achieve an MOI of 10. 7. Include a negative control of un-transduced wells and a positive control (e.g., GFP)-transduced wells. 8. If needed, stain for cell surface markers (such as CD3, CD4, CD8, or FMC63) first and analyze cells on a flow cytometer. 9. Co-culture volume may affect cell conjugation efficiency. We recommend using 25 μL of CAR-T cells and 25 μL of tumor cells. 10. Evaluate the cell density in the bright field channel. A cell density between 40% and ~70% will allow individual cell pairs to be clearly resolved. Adjust the cell number if needed.

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replication mediated by CD8 T cells expressing a re-engineered CD4-based chimeric antigen receptor. PLoS Pathog 13(10):e1006613. https://doi.org/10.1371/journal.ppat. 1006613 4. Hale M, Mesojednik T, Romano Ibarra GS, Sahni J, Bernard A, Sommer K, Scharenberg AM, Rawlings DJ, Wagner TA (2017) Engineering HIV-resistant, anti-HIV chimeric antigen receptor T cells. Mol Ther 25(3):570–579. https://doi.org/10.1016/j.ymthe.2016. 12.023

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5. Ellebrecht CT, Bhoj VG, Nace A, Choi EJ, Mao X, Cho MJ, Di Zenzo G, Lanzavecchia A, Seykora JT, Cotsarelis G, Milone MC, Payne AS (2016) Reengineering chimeric antigen receptor T cells for targeted therapy of autoimmune disease. Science 353(6295):179–184. https://doi.org/10. 1126/science.aaf6756 6. Elinav E, Waks T, Eshhar Z (2008) Redirection of regulatory T cells with predetermined specificity for the treatment of experimental colitis in mice. Gastroenterology 134(7):2014–2024. https://doi.org/10.1053/j.gastro.2008. 02.060 7. Aghajanian H, Kimura T, Rurik JG, Hancock AS, Leibowitz MS, Li L, Scholler J, Monslow J, Lo A, Han W, Wang T, Bedi K, Morley MP, Linares Saldana RA, Bolar NA, McDaid K, Assenmacher CA, Smith CL, Wirth D, June CH, Margulies KB, Jain R, Pure E, Albelda SM, Epstein JA (2019) Targeting cardiac fibrosis with engineered T cells. Nature 573(7774): 430–433. https://doi.org/10.1038/s41586019-1546-z 8. Majzner RG, Rietberg SP, Sotillo E, Dong R, Vachharajani VT, Labanieh L, Myklebust JH, Kadapakkam M, Weber EW, Tousley AM, Richards RM, Heitzeneder S, Nguyen SM, Wiebking V, Theruvath J, Lynn RC, Xu P, Dunn AR, Vale RD, Mackall CL (2020) Tuning the antigen density requirement for CAR T cell activity. Cancer Discov 10:702. https:// doi.org/10.1158/2159-8290.CD-19-0945 9. Dustin ML (2009) Modular design of immunological synapses and kinapses. Cold Spring Harb Perspect Biol 1(1):a002873. https://doi. org/10.1101/cshperspect.a002873 10. Dustin ML (2014) The immunological synapse. Cancer Immunol Res 2(11):1023–1033.

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INDEX A Adhesion .................................... 4, 7, 123, 124, 127–129, 131, 152, 170, 202, 278, 314, 328, 333, 338, 343, 356, 396, 397, 402, 404, 405, 485 Antibody epitopes ......................116, 171, 178, 180, 181 Antiflex........................................124, 125, 127, 130, 132 Antigen recognition ................................... 218, 277–301, 393, 437, 454 Artificial neural networks (ANN)................................. 496

B B cell receptor signaling..................................... 2, 91–110 B cells ............................................2–4, 9, 13–16, 91–110, 345, 346, 357, 393–407, 465, 503, 504, 510 Biofunctionalization................................... 264, 265, 269, 271, 317, 320, 323, 324 Bioimaging .................................................................... 462 Biointerfaces ......................................................... 277–301 Biophysics ...................................................................... 379 Bottom-up synthetic biology .............................. 263–274

C CAR-T .......................453–462, 474, 493, 494, 503–511 Catch bond..........................................376–378, 381, 389 CD45 ...........................................52, 156, 184, 218, 220, 222, 225, 438, 447, 449, 504, 506, 509, 510 Cell adhesion ................................. 52, 56, 123–132, 202, 314, 328, 331, 332, 336, 337, 340 Cell-cell contacts ......................................... 138, 246, 464 Cell conjugation ................................................... 509–511 Cell surface proteins............................................... 4, 8, 41 Cell-surface topography....................................... 169–197 Cell to cell interactions ................................137, 149–157 Chimeric antigen receptor (CAR)............. 308, 454, 455, 493–501, 503–511 Co-localization ......................................... 1–20, 175, 181, 184, 188, 196, 218, 226, 261, 280, 296, 300, 301, 313, 334, 433, 435 Confocal microscopy .......................................... 114, 145, 163, 202, 218, 265, 309, 352, 425, 430, 431, 448, 453–461, 509, 510 Cytoskeleton.........................................71, 160, 170, 324, 346, 356, 363, 394, 421, 464

Cytotoxic granules (CGs).......................... 159, 160, 162, 163, 409, 421, 423, 494 Cytotoxicity ................................................ 272, 453, 455, 463, 464, 493, 504 Cytotoxic T lymphocytes (CTLs) ...................... 159, 160, 163, 409, 421–435, 463–465, 467–473, 475

D Dendritic cells (DCs) ................................. 137, 139, 144, 146, 206–208, 212, 214, 218, 219, 223, 393, 477–491 Dexamethasone .......................................... 478, 481, 483, 484, 486, 490, 491 Diffraction-limit ................................................... 150, 278 Diffusion analysis ................................................... 11, 295 Diffusion dynamics ...................................................61–86 1,25-dihydroxy-vitamin D3 ...................... 478, 481, 484, 486, 490, 491 Dissociation constant (Kd) ............................................. 25 DNA origami...................................... 278–280, 289–297, 299, 300, 303–311 Drug delivery ................................................................ 263 Dynamics ................................................ 1–20, 37, 41–49, 51–57, 61–63, 76, 77, 80, 85, 124, 137, 138, 146, 149–151, 170, 217, 218, 223, 228, 231, 232, 248, 265, 278, 342, 356, 363, 394, 504

E Epidermal sheets .................................................. 448, 449 Exosomes..................................................... 202, 213, 417 Extracellular vesicles (EVs)........................ 202, 203, 212, 263–274, 410, 415, 417, 418

F Fc Receptor .......................................................... 304, 308 Flow cytometry .......................................... 160, 162, 185, 203, 205, 206, 208, 212, 257, 265, 352, 404, 464, 466, 468–473, 479, 508, 509 Fluctuations ................................... 62–67, 72, 80, 83, 84, 124, 125, 130–132, 151, 489 Fluorescence lifetime microscopy .................................. 92 Fluorescence localization microscopy .............................. 1

Cosima T. Baldari and Michael L. Dustin (eds.), The Immune Synapse: Methods and Protocols, Methods in Molecular Biology, vol. 2654, https://doi.org/10.1007/978-1-0716-3135-5, © 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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514 Index

Fluorescence microscopy ................................26, 41, 151, 279, 368, 372, 453, 466, 508, 509 Fluorescent lipophilic dyes ............................71, 463–475 Force measurement.................... 330–332, 334, 364, 377 4D live cell imaging ............................................. 221, 354

Interleukin-4 (IL-4)................................... 222, 223, 253, 260, 477, 478, 480, 483, 490 Intracellular calcium...................................................... 453 Intracellular trafficking ................................................. 423 Intravital microscopy ........................................... 105, 106

G

L

Germinal center (GC).................. 91–110, 291, 380, 384 Glycocalyx............................................................. 131, 170 Glycoproteins ....................................................... 170, 422 Granulocyte-macrophage colony stimulating factor (GM-CSF) ...................................... 207, 210, 222, 223, 258, 260, 477, 478, 480, 483, 490 Granzyme B (GZMB)................................ 160, 409, 415, 422–424, 426, 427, 433, 435

Large unilamellar vesicles (LUVs) ..................... 265, 266, 268–274 Lattice light-sheet microscopy (LLSM) ........................... 137, 218–223, 225–228 Ligand-receptor binding................................................. 25 Live imaging .....................................................33, 98, 348 Lymphocyte.................................................. 42, 152, 169, 170, 217, 251, 252, 313, 345, 346, 348, 349, 354–357, 360, 399, 409, 410, 474 Lytic granule ..............................159, 435, 463, 475, 494 Lytic granule biogenesis ...................................... 421–435 Lytic synapse ................................................421, 463–475

H Human............................................. 4, 7, 28, 66, 80, 159, 178, 221, 252, 254, 260, 309, 317, 329, 338, 411, 417, 424, 425, 428, 429, 432–434, 454, 456, 463–466, 473, 474, 477–491, 505, 511

I IgG........................................................4, 7, 73, 258, 304, 311, 317, 322, 331, 348, 368, 372 Imaging................................................. 1, 2, 4–10, 12, 16, 18–20, 33–35, 42, 44–46, 49, 68, 72, 73, 80, 82, 92, 95–99, 107, 113, 114, 116–119, 121, 124, 125, 127, 132, 137–146, 149–157, 171, 175, 176, 181–187, 194, 195, 217–219, 221–223, 225–228, 235, 241–245, 247, 248, 258, 260, 265, 278, 283, 292, 295, 297, 300, 305, 306, 308–311, 315, 320, 331, 339, 354, 360, 361, 364, 368, 372, 382, 394, 395, 399, 409–419, 438, 447–449, 453–461, 467, 468, 474, 475, 496, 497, 504, 506, 509, 510 Immune synapse.....................................61, 62, 149, 151, 252, 313, 345, 351, 353, 354, 358, 364, 365, 393–407, 409, 410, 415, 418, 453 Immunofluorescence .................257, 258, 279, 435, 497 Immunofluorescence microscopy ................................ 447 Immuno-isolation ................................................ 159–166 Immunological memory ................................91, 251, 252 Immunological synapse (IS) .............................25, 33, 61, 62, 72, 78, 137, 151, 201, 202, 251–261, 278, 309, 363, 393, 394, 396, 397, 421, 425, 433, 493–501, 503, 504 Inhibition..................................................... 313, 314, 416 Innate immunity ......................................... 231, 232, 303 Interactions............................................... 1, 2, 12, 25–27, 38, 51, 61–86, 120, 121, 123, 137, 138, 149–154, 157, 170, 175, 185, 188, 218, 232, 264, 300, 305, 314, 365, 375–378, 387, 393, 405, 409, 418, 422, 423, 443, 474, 494, 497, 504

M Macrophage................................................ 101–103, 108, 109, 303–305, 308, 309, 311, 345, 393, 417 Magnetic beads .......................................... 160, 161, 166, 394, 396–399, 401, 403, 404 Mass spectrometry (MS)............................ 160, 163, 281, 288, 289, 394, 397, 403–406, 410, 418 Mechanical forces ................................................. 363, 364 Mechanobiology ........................................................... 327 Mechanosensor.............................................................. 376 Membrane protein structure .......................................... 52 Membranes ......................................................8, 9, 12, 13, 15, 19, 25–39, 51–56, 65, 70, 71, 73–75, 80, 82, 113, 114, 116–121, 123–132, 150, 156, 169–197, 201, 202, 204, 217, 218, 225, 232, 235, 236, 238, 241–244, 253, 255, 260, 266, 267, 271, 272, 282, 284, 291, 314, 324, 363, 378, 400, 418, 433, 463–475, 488, 504, 509, 511 Microfluidics...................... 330, 339–342, 345–361, 410 Micropillar arrays........................ 145, 329, 334–339, 343 Microscopy ........................................ 1, 2, 4, 8, 9, 18, 26, 29, 41, 42, 46, 47, 73, 92, 105, 106, 114–115, 123–132, 137, 143, 145, 150, 151, 153, 159, 160, 162, 164, 166, 170, 171, 177, 181–182, 185, 202, 217–220, 222, 234, 235, 239, 241–244, 260, 265, 278, 279, 282, 292, 307, 309, 317, 319, 321, 335–339, 346, 352, 354, 360, 363–372, 376, 401, 425, 430–431, 437, 440–441, 445–450, 453–461, 466–468, 508–510 Microvilli.................................... 169–171, 175, 184, 191, 193–195, 197, 201, 202, 217–228 Monocytes ....... 309, 477, 478, 480, 482, 483, 488, 489 Myddosome.......................................................... 231–249

THE IMMUNE SYNAPSE: METHODS N NADPH oxidase (NOX) .................................92, 99, 107 Nanoparticle tracking analysis (NTA)..............43, 45, 48, 235, 265, 266, 268, 270, 271, 274, 315, 317, 323 Natural killer cells.......................................................... 169 Nitrogen cavitation ....................................................... 160

O Oligomerization ..................................................... 63, 504 Optical tweezers (OTs)....................................... 375, 376, 378, 379, 381, 382, 391

P Pair distribution .............................................................. 12 Peripheral blood mononuclear cells (PBMCs) ......................................... 252, 254, 260, 412, 413, 466, 467, 473, 478, 480–482, 486, 487, 489, 505, 508 Phagocytosis ......................................................... 303–311 Plasma membrane (PM) .................................4, 8, 18, 65, 79, 115, 121, 150, 155, 156, 160, 178, 184, 203, 208, 300, 303, 304, 342, 375, 464 Plasma membrane turnover.......................................... 464 Polarization ....................................... 121, 127, 132, 178, 345, 346, 356, 394, 494 Polydimethylsiloxane (PDMS) .......................44–47, 138, 140–146, 329, 336–338, 343, 346, 347, 350, 356, 358, 360, 364 Protein height ............................................................... 114 Proteomics.................................. 160, 265, 393–407, 422

R Resident memory CD8+ T cells ................ 345, 437–445, 449–451, 473, 474 RUSH system ....................................................... 421–435

S Scanning fluorescence correlation spectroscopy (sFCS) ............................................................61–86 Secretion ....................322, 324, 409–419, 453, 463–475 Senescence ............................................................ 246, 251 Signaling ...........................................1–20, 25, 73, 79, 80, 91, 92, 149, 150, 171, 217, 227, 231, 232, 243, 246, 248, 252, 260, 263, 278, 303–305, 308, 313–324, 345, 377, 394, 453, 454, 493, 494, 503, 504, 506 Signalosome.........................................231–249, 260, 394 Single-cell analysis .............................................. 29, 31–33 Single molecule ........................................ 1, 2, 4, 5, 8–12, 20, 35, 38, 39, 47, 49, 63, 114, 151, 175, 187–190, 235, 242, 243, 246–248, 278–280, 295–297, 299, 300, 328, 330, 332, 376–379, 382, 385–389

AND

PROTOCOLS Index 515

Single molecule imaging................ 4, 169–197, 245, 300 Single molecule tracking................................26, 296, 300 Skin ..................... 34, 405, 437, 438, 440–443, 445–451 Small unilamellar vesicles (SUVs) .......................... 30, 45, 117, 240, 241, 246, 247, 264–268, 270, 272–274, 293, 305, 307, 310 Spatial control of receptors.................................. 313, 314 Spinning disk microscopy ................................... 114, 121, 348, 352, 355, 361, 426, 454, 473 Subcellular fractionation...................................... 159–166 Sucrose density gradient centrifugation ...................... 160 Super-resolution........................................... 1, 4, 5, 9, 18, 20, 113, 114, 149–157, 169–197, 293, 368 Super-resolution microscopy .................................. 1, 125, 150, 151, 170 Supported lipid bilayers (SLBs)........................26, 27, 29, 31–39, 72, 73, 83, 86, 116, 119, 124, 131, 218, 234–235, 240–244, 246–248, 265, 278, 279, 290, 293–297, 299, 304, 307–310 Surface proteins..............................................61, 156, 264 Surfaces .......................................... 6, 7, 9, 10, 26, 29, 31, 32, 35, 36, 38, 39, 41–47, 49, 51, 62, 67, 71, 73, 80, 83, 101–103, 109, 113–125, 127, 131, 132, 143, 150, 151, 156, 157, 169–197, 201, 202, 208, 217, 218, 220, 225, 227, 228, 240, 245, 246, 248, 255, 264, 265, 271, 277, 290, 293–295, 297, 299, 300, 303, 305, 314, 318–320, 323, 327, 328, 332, 333, 336, 337, 340, 342, 343, 346, 349, 351, 352, 359, 360, 364–371, 375, 378, 380, 381, 384, 387, 389, 390, 393, 404, 414, 416, 418, 422, 445, 451, 458, 459, 475, 479, 487, 504, 511 Synapse ................................................. 61, 137–146, 149, 150, 252, 346, 348, 359, 363, 393, 396, 400, 402, 410, 418, 475, 494, 498, 500, 503–511

T T-cell activation ....................................52, 201, 218, 279, 280, 313, 368, 376, 377, 379, 444, 449, 453, 454 T cell antigen search...................................................... 218 T cell immunological synaptosomes (TISs) .............202–204, 206, 208, 209, 211–214 T-cell microvilli...........................171, 175, 217, 218, 223 T cell microvilli particles (TMPs) ................................. 202 T cell receptor (TCR) .......................................41, 42, 44, 51–53, 79, 80, 143, 170, 201, 202, 217, 218, 220–222, 225, 226, 248, 277, 313, 345, 375–379, 381, 385–389, 438, 449, 453, 463, 464, 474, 494, 503, 504 T-cells.........................................25, 41, 51, 61, 137, 149, 159, 169, 201, 217, 243, 277, 313, 336, 345, 363, 375, 421, 437, 453, 477, 503 Telomeres ............................................................. 251–261 Tension gauge tether (TGT) ..............328–334, 342, 380 Time lapse video microscopy...................... 360, 464, 467

THE IMMUNE SYNAPSE: METHODS AND PROTOCOLS

516 Index

TIRF microscopy .....................42, 46, 47, 242, 246, 307 Tissue-resident memory T (TRM) ...................... 437–440, 447–449, 451 Tissue slice ....................................................456–459, 461 Tolerogenic........................................................... 477–491 Toll-like receptor (TLR) 4 ligand ................................ 479 Total internal reflection fluorescence (TIRF) microscopy .................................................. 42, 218 Tracking ........................................... 12, 41, 42, 101, 102, 218, 243, 278, 338, 339, 459 Traction force microscopy (TFM) ..................... 334–339, 363–372 Trans-endocytosis................................................. 327–343

V Variable-angle total internal reflection microscopy (VA-TIRFM) .................................. 171, 175, 183, 185, 190, 191, 193, 194 Vesicle functionalization ............264, 265, 267, 268, 271

W Whole mount ......................................438, 447, 448, 451

Z Zhu-Golan analysis ............................................ 26, 36, 37