Handbook of ELISPOT: Methods and Protocols (Methods in Molecular Biology, 2768) [4 ed.] 1071636898, 9781071636893

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Methods in Molecular Biology 2768

Alexander E. Kalyuzhny  Editor

Handbook of ELISPOT Methods and Protocols Fourth 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.

Handbook of ELISPOT Methods and Protocols Fourth Edition

Edited by

Alexander E. Kalyuzhny Biotechne, Minneapolis, MN, USA

Editor Alexander E. Kalyuzhny Biotechne Minneapolis, MN, USA

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-3689-3 ISBN 978-1-0716-3690-9 (eBook) https://doi.org/10.1007/978-1-0716-3690-9 © The Editor(s) (if applicable) and The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024 Chapters 5, 7, 11, 13, and 15 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 chapter. This work is subject to copyright. All rights are solely and exclusively licensed by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, expressed or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. This Humana imprint is published by the registered company Springer Science+Business Media, LLC, part of Springer Nature. The registered company address is: 1 New York Plaza, New York, NY 10004, U.S.A. Paper in this product is recyclable.

Preface In 2023, we celebrate the 40th anniversary of the inception of ELISpot. In 1983, Sedgewick and Holt published a paper (J. Immunol. Methods 57, 301–309) describing a novel technique for the enumeration of antibody-secreting cells. Later that same year, another article describing a similar antibody detection technique was published in the same journal by Czerkinsky and colleagues (J. Immunol. Methods 65, 109–121) who coined the name we have been using for four decades: Enzyme-Linked ImmunoSpot, or ELISpot. The first three additions of this book received strong positive feedbacks, which indicate the popularity of ELISpot and encouragement for continuation with additional volumes. This assay remains a mainstream application for enumeration of both cytokines and IgGs as an easy to set and perform high-sensitivity analytical technique. The amazing flexibility of an ELISpot assay allows for frequent improvisions of its specification and adaptation to research and diagnostic needs. The 4th edition of the Handbook of ELISpot expands upon the 1st, 2nd, and 3rd editions keeping our goals the same: provide researchers with protocols that can be applied in different laboratory landscapes spanning from purely research to clinical diagnostic environments. Our current volume focuses on ELISpot validation in addition to the evaluation of antibodies used in this assay. We included the chapters elaborating on peptides in T-cell assays and ELISpot application for monitoring infection diseases, such as tuberculosis and SARS-CoV2 and for immunotherapy clinical trials. Due to a growing trend in utilizing multi-omics approach, we included several chapters on ELISpot’s sibling, FluoroSpot, to enarm researchers with setting and utilizing multi-color assays to analyze T- and B-cell function. While FluoroSpot has been known for quite some time, its younger, but not less potent, sibling FOLISPOT has recently entered the multiplex arena, so we decided to include an interesting chapter for researchers curious about high-level multiplexing. Additionally, Artificial Intelligence (AI) is actively entering experimental biology and ELISpot is not an exception to such a trend. To address the AI challenges, we included the chapter on applying AI for ELISpot studies. Furthermore, other chapters focus on comparing ELISpot with flow cytometry, feline coronavirus (FCoV2), and memory B-cells. As with the 1st, 2nd, and 3rd editions, the goal of compiling this volume was to create a comprehensive technical reference and a troubleshooting guide for researchers with different training backgrounds. All the chapters were written by known ELISpot experts who were eager to share their “how-to” knowledge and experience with novices and other experts around the globe. Contributing chapters to an existing book is not an easy endeavor, and I wish to express my deep gratitude to our contributing authors who committed their time to writing, revising, and polishing their chapters. It was a great experience for me as the book’s editor. I met outstanding scholars, learned a lot, and enjoyed working on this project. I hope this book will help to advance science and technology and inspire scientists in their determination and perseverance for new discoveries. Minneapolis, MN, USA

Alexander E. Kalyuzhny

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

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1 Important Considerations for ELISpot Validation. . . . . . . . . . . . . . . . . . . . . . . . . . . 1 Sylvia Janetzki 2 Current Trends in Validating Antibody Specificities for ELISpot by Western Blotting . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 15 Biji T. Kurien and R. Hal Scofield 3 An Overview of Peptides and Peptide Pools for Antigen-Specific Stimulation in T-Cell Assays . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 29 Karsten Schnatbaum, Pavlo Holenya, Sebastian Pfeil, Michael Drosch, Maren Eckey, Ulf Reimer, Holger Wenschuh, and Florian Kern 4 The Applications of ELISpot in the Identification and Treatment of Various Forms of Tuberculosis and in the Cancer Immunotherapies . . . . . . . . 51 Hemant K. Mishra 5 Artificial Intelligence-Based Counting Algorithm Enables Accurate and Detailed Analysis of the Broad Spectrum of Spot Morphologies Observed in Antigen-Specific B-Cell ELISPOT and FluoroSpot Assays . . . . . . . . 59 Alexey Y. Karulin, Melinda Katona, Zolta´n Megyesi, Greg A. Kirchenbaum, and Paul V. Lehmann 6 Comparing Flow Cytometry and ELISpot for Detection of IL-10, IL-6, and TNF Alpha on Human PBMCs. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 87 Kristina Boss, Jodi Hagen, Megan Constans, Christine Goetz, and Alexander E. Kalyuzhny 7 Reagent Tracker™ Platform Verifies and Provides Audit Trails for the Error-Free Implementation of T-Cell ImmunoSpot® Assays . . . . . . . . . . . 105 Alexander A. Lehmann, Diana R. Roen, Zolta´n Megyesi, and Paul V. Lehmann 8 Detection of SARS-CoV-2-Specific Cells Utilizing Whole Proteins and/or Peptides in Human PBMCs Using IFN-ƴ ELISPOT Assay . . . . . . . . . . . 117 Madeleine M. Rasche, Ella C. Kaufmann, Tamar Ratishvili, Ilya M. Swanson, Inna G. Ovsyannikova, and Richard B. Kennedy 9 Interferon-γ/IL-2 ELISpot and mRNA Responses to the SARS-CoV2, Feline Coronavirus Serotypes 1 (FCoV1), and FCoV2 Receptor Binding Domains by the T Cells from COVID-19-Vaccinated Humans and FCoV1-Infected Cats . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 135 Sabarinath Nair, Bikash Sahay, Ananta P. Arukha, Lekshmi K. Edison, Chiquitha D. Crews, John G. Morris Jr, Subhashinie Kariyawasam, and Janet K. Yamamoto

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Frequencies of SARS-CoV-2 Spike Protein-Specific Memory B Cells in Human PBMCs, Quantified by ELISPOT Assay. . . . . . . . . . . . . . . . . . . . . . . . . . Ilya M. Swanson, Iana H. Haralambieva, Madeleine M. Rasche, Inna G. Ovsyannikova, and Richard B. Kennedy Monitoring Memory B Cells by Next-Generation ImmunoSpot® Provides Insights into Humoral Immunity that Measurements of Circulating Antibodies Do Not Reveal . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Paul V. Lehmann, Zhigang Liu, Noe´mi Becza, Alexis V. Valente, Junbo Wang, and Greg A. Kirchenbaum Tracking Circulating HLA-Specific IgG-Producing Memory B Cells with the B-Cell ImmunoSpot Assay . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Delphine Kervella, Sebastiaan Heidt, Robert Fairchild, Stephen Todryk, and Oriol Bestard Assessing the Affinity Spectrum of the Antigen-Specific B Cell Repertoire via ImmunoSpot® . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Noe´mi Becza, Zhigang Liu, Jack Chepke, Xing-Huang Gao, Paul V. Lehmann, and Greg A. Kirchenbaum Using PBMCs in a Multiplex FluoroSpot Assay for Detection of Innate Immune Response-Modulating Impurities (IIRMIs) . . . . . . . . . . . . . . . Bartek Makower and Niklas Ahlborg Four-Color ImmunoSpot® Assays Requiring Only 1–3 mL of Blood Permit Precise Frequency Measurements of Antigen-Specific B Cells-Secreting Immunoglobulins of All Four Classes and Subclasses . . . . . . . . Lingling Yao, Noe´mi Becza, Andrea Maul-Pavicic, Jack Chepke, Greg A. Kirchenbaum, and Paul V. Lehmann High-Plex ELISPOT: FOLISPOT Based on Fluorescence Detection and DNA Complementary Pairing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Mi Liu Triple-Color FluoroSpot Analysis of Polyfunctional Antigen-Specific T Cells by Quantification of Spot-Forming Units and Relative Spot Volumes . . . . . . . . . Niklas Ahlborg, Christian Smedman, and Bartek Makower Performance and Stability of New Class of Fetal Bovine Sera (FBS) and Its Lyophilized Form in ELISpot and FluoroSpot Assays: Applications for Monitoring the Immune Response in Vaccine, and Cell and Gene Immunotherapy in Clinical Trials . . . . . . . . . . . . . . . . . . . . . . . . Zhinous Hosseini, Christopher J. Groves, Penny Anders, Kristen Cave, Madelyn Krunkosky, Brandi Chappell, Sofie Pattyn, Devin Davis, Sylvia Janetzki, and Elizabeth Reap

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

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Contributors NIKLAS AHLBORG • Mabtech AB, Nacka Strand, Sweden; Department of Molecular Biosciences, The Wenner-Gren Institute, Stockholm University, Stockholm, Sweden PENNY ANDERS • Translational Science and Innovation Laboratory (TSAIL), Q Solutions, Durham, NC, USA ANANTA P. ARUKHA • Department of Comparative, Diagnostic, and Population Medicine (CDPM), College of Veterinary Medicine, Gainesville, FL, USA NOE´MI BECZA • Research and Development Department, Cellular Technology Limited, Shaker Heights, OH, USA ORIOL BESTARD • Nephrology and Kidney Transplantation Department, Vall d’Hebron Research Institute (VHIR), Barcelona, Spain; Nephrology and Kidney Transplantation Department, Vall d’Hebron University Hospital, Barcelona, Spain KRISTINA BOSS • Bio-Techne, Minneapolis, MN, USA KRISTEN CAVE • Translational Science and Innovation Laboratory (TSAIL), Q Solutions, Durham, NC, USA BRANDI CHAPPELL • Translational Science and Innovation Laboratory (TSAIL), Q Solutions, Durham, NC, USA JACK CHEPKE • Research & Development Department, Cellular Technology Limited, Shaker Heights, OH, USA MEGAN CONSTANS • Bio-Techne, Minneapolis, MN, USA CHIQUITHA D. CREWS • Department of Small Animal Clinical Science, College of Veterinary Medicine, University of Florida, Gainesville, FL, USA DEVIN DAVIS • Seraworx, Providence, UT, USA MICHAEL DROSCH • JPT Peptide Technologies, Berlin, Germany MAREN ECKEY • JPT Peptide Technologies, Berlin, Germany LEKSHMI K. EDISON • Department of Comparative, Diagnostic, and Population Medicine (CDPM), College of Veterinary Medicine, University of Florida, Gainesville, FL, USA; Laboratories of Comparative Immunology & Virology for Companion Animals, CDPM, College of Veterinary Medicine, University of Florida, Gainesville, FL, USA ROBERT FAIRCHILD • Department of Inflammation & Immunity, Lerner Research Institute, Cleveland Clinic, Cleveland, OH, USA XING-HUANG GAO • Research & Development Department, Cellular Technology Limited, Shaker Heights, OH, USA CHRISTINE GOETZ • Bio-Techne, Minneapolis, MN, USA CHRISTOPHER J. GROVES • Translational Science and Innovation Laboratory (TSAIL), Q Solutions, Durham, NC, USA JODI HAGEN • Bio-Techne, Minneapolis, MN, USA IANA H. HARALAMBIEVA • Mayo Clinic Vaccine Research Group, Mayo Clinic, Rochester, MN, USA SEBASTIAAN HEIDT • Department of Immunology, Leiden University Medical Center, Leiden, The Netherlands PAVLO HOLENYA • JPT Peptide Technologies, Berlin, Germany ZHINOUS HOSSEINI • Translational Science and Innovation Laboratory (TSAIL), Q Solutions, Durham, NC, USA

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SYLVIA JANETZKI • ZellNet Consulting, Fort Lee, NJ, USA ALEXANDER E. KALYUZHNY • Bio-Techne, Minneapolis, MN, USA SUBHASHINIE KARIYAWASAM • Department of Comparative, Diagnostic, and Population Medicine (CDPM), College of Veterinary Medicine, University of Florida, Gainesville, FL, USA; Laboratories of Comparative Immunology & Virology for Companion Animals, CDPM, College of Veterinary Medicine, University of Florida, Gainesville, FL, USA ALEXEY Y. KARULIN • Cellular Technology Limited, Shaker Heights, OH, USA MELINDA KATONA • Cellular Technology Limited, Shaker Heights, OH, USA ELLA C. KAUFMANN • Mayo Clinic Vaccine Research Group, Mayo Clinic, Rochester, MN, USA RICHARD B. KENNEDY • Mayo Clinic Vaccine Research Group, Mayo Clinic, Rochester, MN, USA FLORIAN KERN • JPT Peptide Technologies, Berlin, Germany; Brighton and Sussex Medical School, Brighton, UK DELPHINE KERVELLA • Nephrology and Transplantation Laboratory, Vall d’Hebron Research Institute (VHIR), Barcelona, Spain; Nephrology and Kidney Transplantation Department, Vall d’Hebron University Hospital, Barcelona, Spain GREG A. KIRCHENBAUM • Research & Development Department, Cellular Technology Limited, Shaker Heights, OH, USA MADELYN KRUNKOSKY • Translational Science and Innovation Laboratory (TSAIL), Q Solutions, Durham, NC, USA BIJI T. KURIEN • Arthritis and Clinical Immunology Program, Oklahoma Medical Research Foundation, Oklahoma, OK, USA; Department of Veterans Affairs Medical Center, Oklahoma City, Oklahoma, OK, USA ALEXANDER A. LEHMANN • Department of Research & Development, Cellular Technology Limited, Shaker Heights, OH, USA PAUL V. LEHMANN • Department of Research & Development, Cellular Technology Limited, Shaker Heights, OH, USA MI LIU • College of Pharmaceutical Sciences, Soochow University, Suzhou, People’s Republic of China; Suzhou Ersheng Biopharmaceutical Co., Ltd, Suzhou, People’s Republic of China ZHIGANG LIU • Research and Development Department, Cellular Technology Limited, Shaker Heights, OH, USA BARTEK MAKOWER • Mabtech AB, Nacka Strand, Sweden ANDREA MAUL-PAVICIC • Research and Development Department, Cellular Technology Limited, Shaker Heights, OH, USA ZOLTA´N MEGYESI • Department of Research & Development, Cellular Technology Limited, Shaker Heights, OH, USA HEMANT K. MISHRA • Cellinfinitybio, San Francisco, CA, USA JOHN G. MORRIS JR. • Emerging Pathogens Institute, University of Florida, Gainesville, FL, USA SABARINATH NAIR • Department of Comparative, Diagnostic, and Population Medicine (CDPM), College of Veterinary Medicine, University of Florida, Gainesville, FL, USA; Laboratories of Comparative Immunology & Virology for Companion Animals, CDPM, College of Veterinary Medicine, University of Florida, Gainesville, FL, USA INNA G. OVSYANNIKOVA • Mayo Clinic Vaccine Research Group, Mayo Clinic, Rochester, MN, USA SOFIE PATTYN • ImmunXperts, a Q Solutions Company, Belgium SEBASTIAN PFEIL • JPT Peptide Technologies, Berlin, Germany

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MADELEINE M. RASCHE • Mayo Clinic Vaccine Research Group, Mayo Clinic, Rochester, MN, USA TAMAR RATISHVILI • Mayo Clinic Vaccine Research Group, Mayo Clinic, Rochester, MN, USA ELIZABETH REAP • Translational Science and Innovation Laboratory (TSAIL), Q Solutions, Durham, NC, USA ULF REIMER • JPT Peptide Technologies, Berlin, Germany DIANA R. ROEN • Department of Research & Development, Cellular Technology Limited, Shaker Heights, OH, USA BIKASH SAHAY • Laboratories of Comparative Immunology & Virology for Companion Animals, CDPM, College of Veterinary Medicine, University of Florida, Gainesville, FL, USA; Department of Infectious Diseases and Immunology, College of Veterinary Medicine, University of Florida, Gainesville, FL, USA KARSTEN SCHNATBAUM • JPT Peptide Technologies, Berlin, Germany R. HAL SCOFIELD • Arthritis and Clinical Immunology Program, Oklahoma Medical Research Foundation, Oklahoma, OK, USA; Department of Veterans Affairs Medical Center, Oklahoma City, Oklahoma, OK, USA; Department of Medicine, University of Oklahoma Health Sciences Center, Oklahoma, OK, USA CHRISTIAN SMEDMAN • Mabtech AB, Nacka Strand, Sweden ILYA M. SWANSON • Mayo Clinic Vaccine Research Group, Mayo Clinic, Rochester, MN, USA STEPHEN TODRYK • Faculty of Health & Life Sciences, Northumbria University, Newcastle upon Tyne, UK; Translational & Clinical Research Institute, Newcastle University, Newcastle upon Tyne, UK; Cellular Technology Limited (CTL) Europe GmbH, Rutesheim, Germany ALEXIS V. VALENTE • Research and Development Department, Cellular Technology Limited, Shaker Heights, OH, USA JUNBO WANG • Research and Development Department, Cellular Technology Limited, Shaker Heights, OH, USA HOLGER WENSCHUH • JPT Peptide Technologies, Berlin, Germany JANET K. YAMAMOTO • Department of Comparative, Diagnostic, and Population Medicine (CDPM), College of Veterinary Medicine, University of Florida, Gainesville, FL, USA; Laboratories of Comparative Immunology & Virology for Companion Animals, CDPM, College of Veterinary Medicine, University of Florida, Gainesville, FL, USA LINGLING YAO • Research & Development Department, Cellular Technology Limited, Shaker Heights, OH, USA

Chapter 1 Important Considerations for ELISpot Validation Sylvia Janetzki Abstract The ELISpot assay has a solid place in the immune monitoring field for over 40 years. It is an assay that can assess the function of single immune cells in a straightforward and easy-to-learn approach. Its use in basic research, translational, and clinical work has been documented in countless publications. Harmonization guidelines and invaluable tools for optimal assay performance and evaluation exist. However, the validation of an established ELISpot protocol has been left to diverse opinions about how to interpret and tackle typical validation parameters. This chapter addresses important considerations for ELISpot validation, including the interpretations of validation parameters for a meaningful description of assay performance. Key words ELISpot, ELISpot validation, Limit of Detection, Precision, Accuracy, Specificity, Robustness

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Introduction More than 40 years ago, ELISpot was introduced to the scientific community [1]. While it has an impressive sensitivity, it is robust and easy to perform. Every aspect of the assay has been described, dissected, and improved [2]. Harmonization guidelines for its performance and evaluation ensure easy adaptation and implementation [3, 4]. Statistical guidance and tools allow an appropriate data analysis [5, 6]. But, the final validation of ELISpot performance has been subject to a multitude of interpretations. This comes as a surprise since the purpose of validation is well-defined [7]. A validation is done to provide data that support the use of an assay for a specific purpose, to identify the sources of potential assay variability and address the quantification of these errors, and to describe in mathematical and quantifiable terms the performance characteristics of an assay to demonstrate its reliability. The validation of an ELISpot assay requires scientific judgment. It considers the assay itself, its typical known challenges, and the issues that can arise during real-time monitoring of clinical trial samples. There is no place for addressing validation parameters

Alexander E. Kalyuzhny (ed.), Handbook of ELISPOT: Methods and Protocols, Methods in Molecular Biology, vol. 2768, https://doi.org/10.1007/978-1-0716-3690-9_1, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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with complicated approaches without the specific assay in mind and the usefulness of the parameter values determined. Initial recommendations have already been given in the very first Handbook of ELISpot [8]. More recent publications from the gene therapy field have elegantly built on those with useful details for ELISpot validation [9–11]. ELISpot is not a new assay. Its general performance characteristics and challenges are well described; hence, readdressing those has little value. It is rather about designing validation experiments while keeping those performance characteristics in mind and demonstrating how an optimized standard operating procedure (SOP) performs in the given study setting in a specific laboratory. However, across the fields that utilize ELISpot, validation approaches have been described that, instead of focusing on the latter, readdress general assay characteristics, as evident in even more recent white papers [11]. Scientists performing ELISpot often come from different backgrounds, which do not always involve single-cell functional assays. For other assays, validation can be straightforward, and validation steps are well-defined [12]. But trying to apply those validation strategies to ELISpot often makes little sense. ELISpot is an assay that assesses the function of immune cells on a single-cell level. No gold standard exists that the results obtained with an ELISpot assay can be compared to and verified with. An inherited variability simply comes with an assay that is used to identify specific cells in very low frequencies. Let us take the simple but telling and oftenused example of a pool filled with red and blue marbles. The probability that a handful of marbles taken blindly out of a large pool of red and blue marbles represents the correct ratio of red and blue marbles in that pool (and with that, the correct frequency of one or the other colored marbles) is rather low. Figure 1 demonstrates that challenge on a much smaller scale. The probability of grabbing a sample of 1 red and 4 blue marbles out of a pool of 10 red and 40 blue marbles is just 4.3%. However, the likelihood of grabbing 5 blue marbles and no red at all is over 30% instead. It is easy to convert this example to the challenge of testing a small sample of cells for rare antigen-specific T or B cells. In addition, we need to account for the many technical steps and missteps that can influence the assay outcome [2]. Hence, it is imperative to know the assay in detail so that a set of experiments can be diligently designed to provide valuable information for a reviewer of study results about their trustworthiness. A typical assay validation encompasses the investigation of the following assay performance parameters: A. Accuracy. B. Specificity. C. Precision.

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Fig. 1 Probability example of capturing the correct frequency of two different objects by analyzing a limited-sized sample

D. Limit of Detection. E. Limit of Quantitation (Upper and Lower Limit of Quantitation; Limit of Blank). F. Linearity. G. Range. H. Robustness and Ruggedness. While guidance documents for validation from regulatory agencies exist [7], their actual guidance for assays like ELISpot with its particularities is rather limited. The qualification of an ELISpot assay requires scientific judgment in determining the most applicable and descriptive parameters that require assessment. Based on such judgment, a two-tier priority system can be established as follows: Essential: 1. Accuracy – Does my assay provide accurate results? 2. Precision – What is the variability associated with my assay; can results be repeated? 3. Specificity – Is my assay read-out specific? (Addressing this question is far more complex than it appears at first). 4. Limit of Detection (LOD) – What noise (background) does my assay produce? 5. Robustness – Does my assay withstand small changes in its execution, and to what extent? Less important: 6. Linearity – Over what range of cell number plated per well are results linear?

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7. Range – Over what linear range is the precision of my assay acceptable? 8. Upper and Lower Limit of Detection – What is the lowest and highest spot number produced with my assay that can be measured with acceptable precision? (This sounds very important, but further below, it will be explained why this information may not be that important at all). 9. Limit of Blank – How many spots are detected if no cells are added to the well? Before an assay protocol can be validated, it must be optimized. Optimization is about making the assay as perfect as possible, as sensitive as possible, and as robust as possible. This includes important assay parameters like the cell number plated per well, the incubation time of cells and antigen, the incubation time of the substrate, as well as the choice of important reagents and materials, and more. It is easy to realize that the assay protocol optimization addresses various validation parameters already, namely linearity (optimal seeding number of cells per well) and robustness (which defines ranges for deliberate changes like incubation time of cells, e.g., “at least 16 hours but no longer than 24 hours”; or incubation time with substrate, e.g., “3-4 minutes at room temperature”). Once the assay is optimized, validation runs should provide more insight into how the assay behaves or performs with the optimized SOP applied, what the limitations are, and what needs to be known about the assay to feel comfortable about conclusions made from the results obtained. The “Method” section gives a summary of important considerations for validation parameters. It does not provide a validation protocol.

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Materials and Reagents 1. Validation Plan (see Note 1). 2. Peripheral blood mononuclear cells (PBMC) samples (see Note 2). 3. All Materials and Reagents as described in the optimized SOP. 4. Control peptides (see Note 3). 5. All experimental antigen preparations (see Note 4).

3 3.1

Method Accuracy

Perhaps the most pertinent question related to ELISpot assay performance is the question about its accuracy (see Note 5). And

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while it is the most pertinent question, it is also the most difficult question to answer. Accuracy describes the closeness of agreement of the measured value and the true value. But accuracy for functional single-cell assays is difficult to determine since no gold standard exists to compare the results to (even if a known number of TCR-specific cells are spiked into a sample, it is not exactly known how many of these cells are functionally active and hence detectable). The only way to check on the accuracy of an established SOP is via participation in proficiency panels, during which all participants test the same samples against the same antigens but with their own SOP and reagents (see Note 6). The comparison of results provides a relative accuracy measure. 3.2

Precision

Precision is an extremely important parameter. It describes the closeness of agreement between a series of measurements for the same sample. Precision data are essential for the overall trustworthiness of data, even more so for data for the same study subject obtained from different study time points. Three different precision measurements should be assessed: 1. Intra-assay precision = variability between replicate wells. 2. Inter-assay precision = variability between assays performed on different days. 3. Reproducibility = variability between assays performed by different operators. For operator-related precision, the influence of sample handling by different operators should also be assessed, best in a crisscross testing approach during which each operator thaws and processes a separate vial of the same PBMC on the same day; plates those PBMC in his own assay and gives the remaining PBMC to the other operator who tests them in his own assay, next to the selfprocessed cells. The “magic” number for precision testing is 3: a minimum of three assessments of three samples with three different response levels (high, medium, and low responders) with three replicates, for the intra-assay variability, the inter-assay variability (same operator), and between operators (all operators that will be involved in the study). A precision below 50% coefficient of variability (CV) should be reached, and an average CV of 25% is desirable for cell-based assays like ELISpot. The mathematical issues below the Lower Limit of Quantitation (LLOQ) (see Subheading 3.9) require consideration. In that range, it is recommended to express the precision as standard deviation.

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Specificity

Forty years of ELISpot use [1] and more than 7000 publications have proven that ELISpot is an assay that measures specific responses. A recheck of that well-known specificity fact is of course possible by testing cells against no antigen and against different antigens, including a positive peptide pool with a high likelihood of eliciting a response (e.g., CEFx) and a negative control peptide (pool) (see Note 7). The results obtained may not add much to the 40 years of ELISpot wisdom about assay specificity. A much more important issue that should be addressed is related to the possibility of the experimental peptide pools eliciting responses in healthy and/or treatment-naı¨ve donors. This topic often appears neglected; hence, a closer look is provided here: Extensive pretesting of 20–30 healthy and/or treatment-naı¨ve donors has been recommended for quite a while [13]. Obtained data provide essential information about the level of preexisting responses among the study population. Data on response distribution is important for the data interpretation obtained from the study. It allows assumptions about how many study subjects may exhibit responses against the experimental antigens at baseline, before exposure to the antigens. This may have important implications for the study itself. It further assists in defining a suitable response definition to treatment before the start of the study, whether it is a newly developed response or an enhanced response in case of preexisting responses. Furthermore, pretesting provides insight into the general suitability of the experimental antigen preparations for clinical immune monitoring, especially if responses are elicited in multiple donors, which follow an unexpected pattern caused by specific antigen pools. One of the central issues in immunogenicity testing is the avoidance of false positive responses. This issue is essentially related to peptides, a major source for false positive responses. The underlying mechanisms can be as follows: A. Cross-contamination due to line-clearance issues. B. Residual solvents, residual hydrophobic protection groups, and hydrophobic peptides. C. Neoepitopes due to missing capping. While (A) can be avoided by using a reputable manufacturer, issues under (B) are more difficult to resolve, but in general also rare in occurrence. However, the lack of capping (C) constitutes a major source for potential false positive responses. Peptide manufacturers typically assure clients of necessary protection steps during synthesis, but it needs to be clarified if this assurance reaches beyond the fmoc protection step. Hence, a closer explanation related to capping is given in Note 8 and Fig. 2, which should allow scientists to ask the appropriate questions.

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Fig. 2 Peptide synthesis. Peptide strands are synthesized on a solid phase (resin). Side groups and the N-terminus of amino acids are fmoc protected (green triangle and orange rectangle) (1.). After selective deprotection of the N-terminus of coupled amino acids, new fmoc-protected amino acids are added (2.). Uncoupled and unprotected N-termini require capping to avoid coupling with a new amino acid out of sequence (3) 3.4 Limit of Detection (LOD)

The LOD describes the noise level of an assay, which is defined by the background reactivity level observed under the given SOP. The background reactivity is evaluated by testing cells plus medium only (see Note 9). The two-fold median background reactivity (see Note 10) is set as the LOD [7]. Responses cannot be detected below that cut point because any possible differences between medium and antigen spot counts are observed within the assay’s noise range. Should the SOP produce very low background reactivity levels (e.g., two spots per well or less), the threefold median can be used instead for setting the LOD [7]. Well-optimized ex vivo IFNγ ELISpot assays commonly produce an LOD within the range of 5 and 12 spots. The LOD range for assays working with in vitro expanded cells has not been well described in literature but can be expected to be likely higher. It must also be anticipated that background levels between healthy donor samples used for validation and obtained under optimal conditions and actual study samples, often obtained at multiple clinical sites, may differ (see Note 11). It is recommended that the LOD is rechecked and possibly adapted with the actual study data since it drives the cutoff for response acceptance for the study cohort.

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3.5

Robustness

The robustness of an ELISpot SOP refers to the allowed flexibility in protocol steps without affecting spot counts. Typically, those protocol steps and ranges are already defined during SOP optimization. The most important parameters that need to be addressed are the incubation time of cells and antigens, incubation time with substrate, length of cell resting periods if implemented, and concentrations for essential reagents and antigens. For instance, incubation with the substrate for 3–5 minutes may not result in any discernable differences; however, incubation for only 2 minutes may lead to noticeably lower spot counts, or incubation for 20 minutes may lead to very large spots and high level of noise signals that can interfere with spot counting. Robustness testing should be guided by available information from own experience, scientific publications, and manufacturers’ recommendations. It typically comprises sets of experiments with a limited number of samples, addressing logical ranges for protocol steps.

3.6

Linearity

Approaching linearity testing to answer the question if ELISpot is linear has little value. An ELISpot SOP that tests for responses using soluble antigens does not produce results that follow a linear distribution, meaning the number of spots measured is not equivalent to the number of cells added per well [2, 14, 15]. A different notion demonstrates a lack of knowledge about ELISpot principles. The only circumstances when ELISpot exhibits comprehensive linearity is when separate antigen-presenting cells are used in numbers that ensure the coverage of all effector cells added to the plate. Nowadays, experiments run under those conditions are seldom found. The correct question to be asked is the one addressing the range (range refers to the range of the number of cells plated per well) over which ELISpot testing behaves linearly. While this has been addressed repeatedly in scientific publications, an operator may find it comforting to show to a reviewer that a decrease in plated cells per well falls in the linear range of the assay, and hence extrapolations of spot numbers to a common denominator are acceptable (see Note 12). A limited set of experiments with donors exhibiting different response levels to positive control antigens, during which the number of plated cells is serially diluted, is appropriate for addressing linearity. The focus of that dilution is on the lower limit of acceptable plated cells, e.g., 200,000, 150,000, and 100,000 PBMC per well.

3.7

Range

The range defines the linear range of an assay that has acceptable precision. The experiment set for linearity testing suffices for addressing the range. The parameter of interest is if the lowest cell number in the linear range still has acceptable precision.

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3.8 Upper Limit of Quantitation (ULOQ)

The ULOQ defines the highest spot number created under a given SOP that allows the enumeration of all spots with confidence. That parameter is defined by the number and size of spots per well, and the evaluation method. A typical colorimetric ELISpot assay starts to show confluency of spots with image analysis in a range of 400–1000 spots per well. The parameter should be addressed with high responders to a positive control antigen. The CEFx pool often elicits very strong responses; hence, it may be an optimal choice for testing the ULOQ. The testing should separately be addressed with reactivity to mitogen, since spots created under mitogen stimulation often appear different, mostly smaller, than under antigen-specific stimulation conditions. Importantly, while the ULOQ parameter should be evaluated when classic image analysis is used for spot counting, it has limited meaning for next-generation spot counting with applied RAWSpot™ technology [16]. The advanced technology is not restricted by the limitations of image analysis, but rather processes the RAW signal captured by the camera sensor; hence, it is able to discriminate spots even if they appear confluent in an image (see Note 13).

3.9 Lower Limit of Quantitation (LLOQ)

An ELISpot LLOQ is the lowest spot number that can be quantified with acceptable precision. Replicate low spot counts often exhibit a high coefficient of variability (CV). That high variability reflects the mathematical issue related to the CV, which simply presents the ratio of the standard deviation to the mean. The lower the spot count is, the higher the CV will be (see Note 14). The LLOQ has a low value for ELISpot validation. It describes the spot number below which variability will simply increase above 25–30% [15]. There is no actual need to create a new data set that confirms the mathematical CV issue and hence the well-known “ELISpot LLOQ,” which lies around 20–30 spots per well.

3.10 Limit of Blank (LOB)

The limit of blank describes the spot number counted in wells that have no cells plated but are otherwise coated and developed as all other wells. The LOB should ALWAYS be 0. If the LOB in the hands of an operator is not zero, one of the following two scenarios is likely: A. Insufficient optimization of the SOP: The detection antibody was not filtered before addition to the plate. Antibody aggregates can create false positive spots, which appear like regular spots created by cells. If this is the case, the ELISpot SOP requires optimization. B. Insufficient plate evaluation: Technical artifacts (e.g., dust particles) can create signals that may be picked up by the reader and counted as spots. Such counting errors are a well-known phenomenon in ELISpot [4, 17] and were already described in early proficiency panels as a factor attributing to variability

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[3]. The issue has been addressed in ELISpot harmonization guidelines with the call for auditing ELISpot evaluation results and necessary parameter adjustments or application of masks to exclude artifacts from being counted [3, 4]. If technical artifacts are allowed to be counted, the ELISpot plate reading SOP requires optimization.

4

Notes 1. A validation plan is established that details the assay SOP, and the validation strategy for each validation parameter. 2. It is often difficult to obtain samples from a donor cohort with the disease background of the study. That is in most cases not a problem since the validation is about the assay performance per se. Hence, surrogate samples, namely samples from healthy “normal” donors are acceptable for use in ELISpot validation experiments. Such samples can be commercially obtained from a variety of reliable sources. While three to six different donor samples suffice for most optimization runs, proper validation requires 20–30 samples [13]. 3. Positive control peptide pools can be used as a surrogate for most validation runs. Such peptide pools should optimally be of the same format as the experimental antigens, e.g., in the case of a protein-spanning peptide pool of 15 mers with an amino acid overlap of 11, a Cytomegalovirus (CMV) peptide pool of the same format could be used [18]. Other excellent choices are peptide pools of the CEF pool family [19], which exist in various compositions. The CEFx peptide pool has an incredible reliability of eliciting responses in donors from a wide range of ethnic backgrounds. 4. All experimental antigen preparations should be tested to obtain information about their integrity (see Specificity) and preexisting responses in a treatment-naive cohort, if applicable. However, in the case of patient-specific antigens (e.g., tumorspecific neoantigens), the pretesting of such is clearly not feasible. 5. Even and especially when ELISpot work is outsourced to a contract research organization (CRO), insist on accuracy reports. Don’t buy the pig in a poke. 6. An open proficiency panel program is accessible once a year to everyone performing IFNγ ELISpot: https://www.immudex. com/services/proficiency-testing/t-cell-elispot-proficiencytesting/. This proficiency panel program was established by the CIC/CRI and the CIP/CIMT and was later outsourced.

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Performance reports are provided, as shown in this online example: https://www.immudex.com/media/1883/tf12 6301-t-cell-elispot-proficiency-panel-report-2021.pdf. 7. Negative peptide pools are commercially available, e.g., for MOG or actin. However, it is possible that occasional donor samples show reactivity against those peptide pools, as has been demonstrated for myelin oligodendrocyte glycoprotein (MOG) [20]. The exposure of the sample donor to foreign antigens that share sequence or structural similarities to these pools is also possible (molecular mimicry) and should hence be kept in mind. 8. Standard peptide synthesis is the addition of one peptide at a time after the next, forming a growing peptide strand on a solid phase, typically resin (for this example, the wanted amino acid sequence is A-B-C-D). Peptides are linked via amide bonds, and the peptide chain grows from the C-terminus to the N-terminus. Those bond formations are slow and are hence activated. During synthesis, the protection of the N-terminus and side chains of the amino acids is essential to avoid undesirable side reactions like self-coupling of the activated amino acid. This is done via the widely known fmoc protection (fmoc stands for fluorenylmethoxycarbonyl). Once the amino acid is coupled to the growing strand, a selective deprotection of the N-terminus is done, and the next amino acid is added with the side groups and N-terminus protected again, and so on. The challenge is that no chemical reaction is 100% certain. In addition to the wanted coupled ends of the growing peptide chain with protected amino groups, uncoupled ends with free amino groups can occur, to which no newly added amino acid had bound (see 3. in Fig. 2). Those uncoupled ends need to be capped; otherwise, the next added amino acid will likely bind, skipping the one in a sequence that was not coupled. Capping is typically done by acetylation, e.g., with phenoxyacetic acid anhydride. This adds a cap to the uncoupled end of a peptide chain and with that stops the reaction of that specific peptide strand only. Capping must be done after the addition of each amino acid before the synthesis can continue. With capping applied, truncated peptides may be produced, e.g., A only, A-B only, A-B-C only, or A-B-C-D. If capping is not performed, deletion peptides will also be produced, caused by skipped amino acids during synthesis (e.g., A-C-D or A-B-D). 9. Data can easily be deduced from the specificity testing. 10. Do not base the LOD calculation on the mean background reactivity because of possible distortion by outlier measurements.

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11. Conditions related to patients (e.g., immune suppression or infection), reagents (e.g., different freezing media), or applied SOPs (e.g., sample processing) can unexpectedly affect background reactivity levels. Enrichment of samples adds another uncertainty factor. 12. The typical reliable linear range of an ELISpot performed with soluble antigens is between 200,000 and 300,000 cells plated. Some reports extend that range to 100,000–400,000 cells per well. In either case, it is permissible to plate out cells at the lowest concentration that falls within the linearity range, if sample availability is limited, and later extrapolate the counted spot number to the cell number typically required to be plated as per SOP. 13. RAWSpot™ technology applies a highly advanced algorithm that can backtrack the movement of cytokine molecules that led to the formation of spots, and hence allows the exact determination of the spot center and spot volume [21, 22]. 14. Here is an example demonstrating the CV issue driven by low spot counts: Replicate spot counts of 1 and 2 have a very small SD (0.7), but a very high CV (47%), while, using common sense, the values are very close (as close as it gets in ELISpot while not being the same; just one spot difference). References 1. Czerkinsky CC, Nilsson LA, Nygren H et al (1983) A solid-phase enzyme-linked immunospot (Elispot) assay for enumeration of specific antibody-secreting cells. J Immunol Methods 65(1–2):109–121 2. Janetzki S (2016) Elispot for rookies (and experts too) in techniques in life science and biomedicine for the non-expert. Springer International Publishing. https://doi.org/10. 1007/978-3-319-45295-1 3. Janetzki S, Panageas KS, Ben-Porat L et al (2008) Results and harmonization guidelines from two large-scale international Elispot proficiency panels conducted by the cancer vaccine consortium (CVC/SVI). Cancer Immunol Immunother 57(3):303–315. https://doi. org/10.1007/s00262-007-0380-6 4. Janetzki S, Price L, Schroeder H et al (2015) Guidelines for the automated evaluation of Elispot assays. Nat Protoc 10(7):1098–1115. https://doi.org/10.1038/nprot.2015.068 5. Moodie Z, Price L, Gouttefangeas C et al (2010) Response definition criteria for Elispot assays revisited. Cancer Immunol Immunother 59(10):1489–1501. https://doi.org/10. 1007/s00262-010-0875-4

6. Moodie Z, Price L, Janetzki S et al (2012) Response determination criteria for Elispot: toward a standard that can be applied across laboratories. Methods Mol Biol 792:185–196. https://doi.org/10.1007/978-1-61779-3257_15 7. FDA (2021) Guideline for industry. Text on validation of analytical procedures. https:// www.fda.gov/regulatory-information/searchfda-guidance-documents/q2r1-validation-ana lytical-procedures-text-and-methodology-guid ance-industry. Accessed 09 June 2023 8. Janetzki S, Cox JH, Oden N et al (2005) Standardization and validation issues of the Elispot assay. Methods Mol Biol 302:51–86. https:// doi.org/10.1385/1-59259-903-6:051 9. Patton KS, Harrison MT, Long BR et al (2021) Monitoring cell-mediated immune responses in AAV gene therapy clinical trials using a validated ifn-gamma Elispot method. Mol Ther Methods Clin Dev 22:183–195. https://doi. org/10.1016/j.omtm.2021.05.012 10. Gorovits B, Azadeh M, Buchlis G et al (2023) Evaluation of cellular immune response to adeno-associated virus-based gene therapy.

ELISpot Validation AAPS J 25(3):47. https://doi.org/10.1208/ s12248-023-00814-5 11. Islam R, Vance J, Poirier M et al (2022) Recommendations on Elispot assay validation by the GCC. Bioanalysis 14(4):187–193. https://doi.org/10.4155/bio-2022-0010 12. Ederveen J (2010) A practical approach to biological assay validation. Hoofddorp: Prog 106(1) 13. CLSI (2013) Performance of single cell immune response assays; approved guideline second edition. CLSI document I/LA26-A2. Clinical and Laboratory Standards Institute, Wayne, PA 14. Smith JG, Liu X, Kaufhold RM et al (2001) Development and validation of a gamma interferon Elispot assay for quantitation of cellular immune responses to varicella-zoster virus. Clin Diagn Lab Immunol 8(5):871–879. h t t p s : // d o i . o r g / 1 0 . 1 1 2 8 / C D L I . 8 . 5 . 871-879.2001 15. Maecker HT, Hassler J, Payne JK et al (2008) Precision and linearity targets for validation of an ifngamma Elispot, cytokine flow cytometry, and tetramer assay using CMV peptides. BMC Immunol 9:9. https://doi.org/10.1186/ 1471-2172-9-9 16. Jahnmatz P, Sundling C, Yman V et al (2020) Memory b-cell responses against merozoite antigens after acute plasmodium falciparum malaria, assessed over one year using a novel multiplexed Fluorospot assay. Front Immunol 11:619398. https://doi.org/10.3389/ fimmu.2020.619398

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17. Janetzki S (2018) Mastering the computational challenges of Elispot plate evaluation. Methods Mol Biol 1808:9–30. https://doi.org/10. 1007/978-1-4939-8567-8_2 18. Maecker HT, Dunn HS, Suni MA et al (2001) Use of overlapping peptide mixtures as antigens for cytokine flow cytometry. J Immunol Methods 255(1–2):27–40. S0022175901004161 [pii] 19. Currier JR, Kuta EG, Turk E et al (2002) A panel of MHC class I restricted viral peptides for use as a quality control for vaccine trial Elispot assays. J Immunol Methods 260(1–2): 157–172. S002217590100535X [pii] 20. Koehler NK, Genain CP, Giesser B et al (2002) The human T cell response to myelin oligodendrocyte glycoprotein: a multiple sclerosis family-based study. J Immunol 168(11): 5920–5927. https://doi.org/10.4049/ jimmunol.168.11.5920 21. Jalde´n J, del Aguila Pla P (2018) Cell detection by functional inverse diffusion and non-negative group sparsity—part i: modeling and inverse problems. IEEE Trans Signal Process 60(20):5407–5421. https://doi.org/10. 1109/TSP.2018.2868258 22. Jalde´n J, del Aguila Pla P (2018) Cell detection by functional inverse diffusion and non-negative group sparsity—part ii: proximal optimization and performance evaluation. IEEE Trans Signal Process 66(20): 5422–5437. https://doi.org/10.1109/TSP. 2018.2868256

Chapter 2 Current Trends in Validating Antibody Specificities for ELISpot by Western Blotting Biji T. Kurien and R. Hal Scofield Abstract The enzyme-linked immunospot (ELISpot) assay is a highly useful and sensitive method to detect total immunoglobulin and antigen-specific antibody-secreting cells. In addition, this method can measure biological activity and immunological secretions from immune cells. In general, membrane-bound antigen allows binding of antibody secreted by B cells, or a membrane-bound analyte-specific antibody binds to the specific analyte (e.g., cytokines) elicited from cells added to the well containing the bound antibody. The response from added cells is then detected by using an anti-Ig antibody and a colorimetric substrate, while in the case of non-B cells, the elicited antigen is detected with appropriate antibodies and enzymeconjugated antibodies. Specificity of antibodies binding the protein of interest is necessary to achieve correct results. Western blotting can be used for this with/without siRNA knockdown of proteins of interest or with the use of peptide inhibitors to inhibit the binding of specific antibodies to the target protein. Despite its general simplicity, western blotting is a powerful technique for immunodetection of proteins (notably low abundance proteins) as it provides simultaneous resolution of multiple immunogenic antigens within a sample for detection by specific antibodies. Now, we have plethora of immunoblotting methods to validate antibodies for ELISpot. Key words Western blotting, SDS-PAGE, Nitrocellulose

1

Introduction The difficult task of probing electrophoresed gel proteins with specific antibodies led to the development of western blotting [1, 2], following the development of Southern blotting. The western blotting method includes both manual sample deposition (dot blotting) and transfer from polyacrylamide gels (PAGE). The proteins electrophoresed on sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) gels are transferred electrophoretically to microporous membranes like nitrocellulose or polyvinylidene dichloride (PVDF) membranes. The protein blotting method evolved as an offshoot of DNA (Southern) blotting [3] and RNA (northern) blotting [4]. The method was named “western

Alexander E. Kalyuzhny (ed.), Handbook of ELISPOT: Methods and Protocols, Methods in Molecular Biology, vol. 2768, https://doi.org/10.1007/978-1-0716-3690-9_2, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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Western Blotting and Detection

SDS-PAGE

E

A

Western blot

Develop color

B

D

Blot from gel

C Add primary antibody

Add enzyme conjugate

Fig. 1 Schematic representation of western blotting and detection procedure. (a) Unstained SDS-PAGE gel prior to western blot. The bands shown are hypothetical. (b) Exact replica of SDS-PAGE gel obtained as a blot following western transfer. (c) Primary antibody binding to a specific band on the blot. (d) Secondary antibody conjugated to an enzyme (alkaline phosphatase or horse radish peroxidase) binding to primary antibody. (e) Color development of specific band. (Reproduced from Ref [10] with permission from Elsevier)

blotting” [5] to retain the “geographic” naming tradition pioneered by Sir Southern [3]. The transferred proteins form a literal copy of the SDS PAGE gel [6] on the membrane, and this membrane forms the starting step for a number of experiments (Fig. 1). The ensuing use of antibody probes directed against the membrane-bound proteins (immunoblotting) has revolutionized the field of immunology. Immunoblotting [1, 2, 5] is a powerful tool to detect and characterize different proteins, especially low-abundance proteins. Protein blotting is advantageous because (i) moist membranes are pliable and are easy to handle compared to gels, (ii) proteins immobilized on the membrane are easily accessible to different antibodies/ligands, (iii) only small amount of reagents are required for transfer analysis, (iv) it provides several replicas of a gel, (v) it allows prolonged storage of transferred patterns before use, and (vi) the same protein transfer can be used for multiple successive analyses [7–9].

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While the original protein blotting method was described in 1979, several methods to transfer proteins have been described since then [10]. However, western blot sensitivity is dependent on blotting or transfer efficiency, retention of antigen during processing and the final detection/amplification system employed. Results are compromised if there are deficiencies in any of these steps [11]. Transfer of proteins from a gel to a solid membrane support is dependent on (a) the nature of the gel, (b) the molecular mass of the proteins being transferred, and (c) the type of membrane used. It is best to run the softest gel in terms of acrylamide and crosslinker, which produces the essential resolution. Transfer will be complete and faster when thinner gels are used. The use of ultrathin gels, however, may cause handling problems and a 0.4 mm thickness represents the lower practical limit [12]. High molecular proteins transfer poorly from SDS-PAGE gels, resulting in low levels of detection on immunoblots. The use of heat, special buffers, and partial proteolytic digestion of the proteins prior to transfer [11, 13–17], however, has enabled the efficient transfer of high molecular proteins.

2

Membrane Supports for Protein Transfer The generally used solid microporous phases for protein blotting are made up of microporous surfaces and membranes like cellulose, nitrocellulose (NC), polyvinylidene difluoride, cellulose acetate, polyethane sulfone, and nylon. The exceptional properties of microporous surfaces that make them suitable for “protein blotting” are (i) large volume-to-surface area ratio, (ii) high binding capacity, (iii) short- and long-term storage of immobilized molecules, (iv) ease of processing by allowing a solution phase to interact with the immobilized molecule, (v) lack of interference with the detection strategy, and (vi) reproducibility. These properties are useful for the high-throughput assays used also in the postgenomic era [2, 4, 14, 18, 19]. These microporous surfaces, typically, are used in the form of membranes or sheets with a thickness of 100 μm and possess an average pore size that ranges from 0.05 to 10 μm in diameter. The interaction of biomolecules with each of these membranes is not completely understood, except that in general the interaction is noncovalent [20, 21].

2.1 Nitrocellulose Membranes

Nitrocellulose (NC) membranes are used in many applications, including high-throughput array, immunodiagnostic as well as mass-spectrometry coupled proteomic applications, filtration/concentration, ion exchange, and amino acid sequencing in addition to traditional immunoblotting methods. Southern first showed in

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1975 the usefulness of NC to capture nucleic acids. Towbin in 1979 [4] and Burnette in 1981 [1] demonstrated that these membranes could also be used for proteins. NC continues to be useful in the postgenomic era technology [19], since high-throughput methodologies for proteomics and genomics depend greatly on traditional notions of molecular immobilization followed by hybridization binding/analysis. 2.1.1 Mechanism of Immobilization

The exact type of interaction of biomolecules with NC is unclear. However, studies suggest that the interaction is noncovalent and hydrophobic. One evidence favoring hydrophobic interaction is the fact that since most proteins at pH values above 7 are negatively charged, it is surprising that NC, which is also negatively charged, can bind proteins efficiently. An additional fact is that nonionic detergents, like Triton X-100 are effective in removing bound antigens from NC [8]. High salt concentrations and low methanol concentrations increase efficiency of immobilization [22]. Protein bound to NC membranes can be stained with dyes like amido black [4], Coomassie brilliant blue (CBB) [1], aniline blueblack, Ponceau S, fast green, or toluidine blue. Amido black staining can detect a 25-ng spot of bovine serum albumin easily with acceptable background staining. CBB gives a higher background staining, while Ponceau S gives a very clean pattern but with slightly less sensitivity than amido black.

2.1.2 Disadvantages of Nitrocellulose Membrane

NC membranes are fragile and, therefore, cannot be stripped and reprobed many times. It can become brittle when dry. In addition, small proteins move through NC membranes, and only a small fraction of the total amount actually binds. Using membranes with smaller pores can prevent this issue [12]. Gelatin-coated NC has been used for quantitative retention [10, 23]. The mechanical strength of the membrane has been improved by integrating a polyester support web in supported NC (e.g., Hybond-C Extra) to enable easier handling.

2.2 Polyvinylidene Difluoride

PVDF (polyvinylidene difluoride) is a linear polymer with repeating –(CF2-CH2)- units [2]. The membrane was renamed as Immobilion-P™ Transfer Membrane after being initially referred to as Immobilon™ PVDF transfer membrane to differentiate it from other PVDF and non-PVDF-based blotting membranes referred to collectively as Immobilon family and marketed by Millipore. Immobilon-PSQ membrane with a 0.2 μm pore size suitable for proteins with a molecular weight < 20 kD (to prevent blow through) and immobilon-FL membrane optimized for all fluorescence applications also form part of the Immobilon family of PVDF membranes, added recently. Sequelon [24], a PVDF-based sequencing membrane, sold by Milligen/BioSearch, a Millipore subsidiary, is useful due to its chemical stability, physical strength, and high protein binding scope.

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2.2.1 Mechanism of Binding to Membranes

Proteins transferred to the Immobilon-P membrane during western blotting are retained efficiently on its surface throughout the immunodetection process via a combination of dipole and hydrophobic interactions. The antigen-binding capacity of the membrane is 170 μg/cm2 (for bovine serum albumin), and this is proportionate with that of NC. Also, the immobilon-P membrane has very good mechanical strength and like Teflon™ (a related fluorocarbon polymer) is compatible with chemicals and organic solvents like acetonitrile, trifluoroacetic acid, hexane, ethylacetate, and trimethylamine [2, 25]. It is necessary to wet the PVDF membrane in either methanol or ethanol prior to using with aqueous buffers. Except for this, the blotting mechanics are not different from that seen with NC. This is because there is no added surfactant in PVDF and also due to its highly hydrophobic nature.

2.2.2 Advantages of PVDF

An important advantage of transferring proteins to PVDF membranes is that replicate lanes from a single gel can be used for several purposes such as N-terminal sequencing, proteolysis/peptide separation/internal sequencing along with western analysis. Proteins blotted into PVDF membranes can be stained with amido black, India ink, and silver nitrate [26]. These membranes are also amenable to staining with CBB, thus allowing excision of proteins for N-terminal protein sequencing [25, 27].

2.3

Proteins can bind to activated paper (diazo groups) covalently. However, this is disadvantageous as the coupling method is incompatible with many gel electrophoresis systems. Linkage is through primary amines, and therefore, systems that use gel buffers without free amino groups must be used with this paper. In addition, the paper is expensive and the reactive groups have a limited half-life once the paper is activated.

Activated Paper

2.4 Nylon Membranes

Nylon-based membranes are thin, with a smooth surface like NC, but they have much better durability. Two kinds of membranes are available commercially: Gene Screen and Zetabind (ZB). ZB is a nylon matrix (polyhexamethylene adipamine or Nylon 66) modified by the addition of many tertiary amino groups during the manufacturing process (extensive cationization). It has excellent mechanical strength and also offers the potential of very significant (yet reversible) electrostatic interactions between the membrane and polyanions. Nylon has a greater protein-binding capacity compared to NC (480 μg versus 80 μg bovine serum albumin (BSA) bound/cm2). Also, nylon gives the benefits of more consistent transfer results and a significantly increased sensitivity compared to other membranes [7, 18]. This effect is possible owing to the extra potential difference created by the positive charge of Zetabind.

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2.4.1 Shortcomings of Nylon

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Higher nonspecific binding is an issue owing to the high binding capacity of these membranes. Another problem with using nylon membranes is that they bind strongly to the commonly used anionic dyes like Coomassie blue, amido black 10B [18], aniline blue-black, Ponceau S, fast green, or toluidine blue. SDS, dodecyl trimethylammonium bromide or Triton X-100 at low concentrations (0.1% in water) remove the dyes from the membrane while simultaneously destaining the transferred proteins, with SDS being the best. Destaining of this membrane is thus not possible, unlike NC, and therefore, the background remains as high as the signal [8]. On account of these problems, NC membranes have remained the best compromise for most situations. However, an immunological stain and India ink have been used to detect proteins on ZB [28–30] and NC membranes. Nylon membranes have been found very useful in binding the negatively charged DNA, especially the positively charged Zetabind membranes. So, they have been used more for DNA blotting than for protein blotting.

Antibody Evaluation The assessment of antibodies by western blot is dependent on the antibody binding the protein of interest at about the correct molecular weight. One can use several ways to prove that the antibody is targeting the protein of interest. The first method is to use siRNA knockdown technology to reduce the concentration of the protein of interest and, study the protein by western blot, and compare it with controls that do not have the protein knocked down. The second way is to use peptide inhibitors to inhibit the binding of specific antibodies to the target protein. A third approach would be to use knockout technology to show specificity of antibody against the protein of interest.

4

Antibody Considerations Protein blots are often used in combination with enzyme-linked immunosorbent assays, which are important alternative antibodybased detection methods. As seen from the previous section, immunoblotting is a “must” to determine specificity of antibodies used for any immunological application, including ELISpot. A main characteristic of any successful western blot is the highly specific interaction between an antigen and an antibody. The actual point of interaction takes place between a small portion of the antigen (an epitope) and the recognition sites located on the Fab region of the antibody molecule (a paratope). Antibodies selected for

Western Blotting

21

immunodetection protocols should be tested by western blot analysis, when possible, and experimental conditions recommended by the antibody supplier must be followed carefully [31]. Electrophoretic separation of proteins is commonly carried out under denaturing conditions. Therefore, the western blots derived from such SDS-PAGE gels would contain its replica of denatured proteins. Western blot-positive antibodies normally recognize a short linear portion of amino acids found in the nonlinearized target protein, which become available for binding under denaturing and reducing conditions. However, antibodies recognizing conformational epitopes, regions forming a three-dimensional structural configuration of amino acids, lose their binding ability upon denaturation of the protein. However, western blotting protocols are flexible. Since one can choose gel electrophoresis and protein blotting conditions, it is possible to modify buffers to retain sufficiently higher-order protein structures for detection by some antibodies. The datasheet obtained with the antibody typically gives information regarding buffers best suited for specific antigen–antibody binding [31]. 4.1 Polyclonal Antibodies Versus Monoclonal Antibodies

5

Polyclonal antibodies are usually made in experimental animals like mice, rabbits, sheep, goats, and donkeys by immunization with a specific protein or peptide. These antibodies contain a pool of immunoglobulin molecules that bind to different epitopes found on a single protein. On the other hand, monoclonal antibodies bind only to a single epitope within an antigen. These antibodies contain homogenous cloned immunoglobulins made by fusing antibody-producing B cells from the spleen of the immunized animal (rat or mouse) with an immortalized cell line. Both polyclonal and monoclonal antibodies are used in protein blotting, and the choice should be made depending on the downstream application. Polyclonal antibodies can give a higher background and cross-reactivity, compared to monoclonal antibodies, due to the detection of multiple epitopes. However, polyclonal antibodies are more sensitive than monoclonals since the signal is amplified from the binding of several antibodies per protein target [31, 32].

Methods to Transfer Proteins from Gel to Membrane Investigators have used three basic methods to transfer protein from SDS-PAGE or native gels to nitrocellulose or PVDF membranes. These methods are (a) simple diffusion, (b) vacuum-assisted solvent flow, and (c) “Western” blotting or electrophoretic elution [4, 12, 33–39].

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Gel

Glass plate

Filter paper

Membrane

Plastic container

Moist paper towel

Clamp

Fig. 2 Bidirectional nonelectrophoretic transfer of proteins from SDS-PAGE gels to NC membranes to obtain up to 12 blots. The PAGE gel is sandwiched between two membranes, filter paper, and glass plates and incubated at 37 °C for varying periods of time to obtain up to 12 blots. (Reproduced from Ref [10] with permission from Elsevier) 5.1

Simple Diffusion

Simple diffusion or diffusion blotting was originally developed for transferring proteins separated by isoelectric focusing on thin gels to membranes. This method was later expanded to other gel systems [40–46]. In this technique, a membrane is placed on the gel surface with a stack of dry filter papers on top of the membrane. A glass plate and an object with a certain weight are placed on this assembly to allow the diffusion process. However, since quantitative transfer of protein was lacking, this protocol did not gain widespread acceptance. Interest began to pick up, when it was shown that it was possible to get up to 12 blots from a single gel by sandwiching it between two membranes in a sequential manner (Fig. 2) [33]. Membrane transfers from SDS-PAGE gels by simple diffusion provide a useful way for identifying proteins by mass spectrometry [47, 48]. The gel can be stained with Coomassie following diffusion blotting. The antigens on the blot are detected by immunostaining, and the immunoblotted target band can be compared with the Coomassie stained gel by superimposing the blot and the stained gel, permitting the identification of the band to be excised for tryptic digestion and subsequent matrix-assisted laser desorption time of flight mass spectrometric scrutiny. The main benefit of diffusion blotting compared to electroblotting is that several transfers or imprints can be obtained from the same gel, and different antisera can be tested on identical transfers. Quantitative data regarding protein transfer during diffusion blotting were obtained soon using 14C labeled proteins. A 3-minute diffusion blotting procedure was shown to allow a transfer of 10% compared to electroblotting. Diffusion blotting of the same gels carried out multiple times for prolonged periods at 37 °C causes the gel to shrink, a problem that was overcome using gels cast on plastic supports [44, 45].

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Activity gel electrophoresis or zymography has also been studied with regard to the utility of diffusion. This procedure involves the electrophoresis of enzymes (either nucleases or proteases) through discontinuous polyacrylamide gels containing enzyme substrate (either type III gelatin or β-casein). After electrophoresis, SDS is removed from the gel by washing in 2.5% Triton X-100. This permits the enzyme to renature and the substrate to be degraded. Staining of the proteins with CBB allows the bands of enzyme activity to be detected as clear bands of lysis against a blue background [49]. An additional immunoblotting analysis using another gel is often needed in this technique to examine a particular band that is involved. Diffusion blotting has been used to circumvent the use of a second gel for this purpose [45]. The activity gel was blotted onto PVDF for immunostaining, and the remaining gel after blotting was used for routine “activity staining.” Since the blot and the activity staining are derived from the same gel, the signal localization in the gel and the replica can be easily aligned for comparison. Diffusion blotting transfers 25–50% of the [45] proteins to the membrane compared to electroblotting. However, the benefit of getting several blots from the same gel could outweigh the loss in transfer and actually could be compensated for by using sensitive detection methods. The gel remains on its plastic support, which prevents stretching and compression; this warrants identical imprints and enables more reliable molecular mass determination. If only a few imprints are made, sufficient protein remains within the gel for general protein staining. These benefits make diffusion blotting the method of choice when quantitative protein transfer is not essential. 5.2

Vacuum Blotting

Peferoen et al. established vacuum blotting [50] as an alternative to diffusion blotting and electroblotting. The suction power of a pump connected to a slab gel dryer system drives the separated polypeptides from the gel to the nitrocellulose membrane. Both low and high-molecular-weight proteins get transferred using this process. Small molecular weight proteins (~14,000 Daltons) are not well adsorbed by the 0.45 μm nitrocellulose membranes. Therefore, when using low molecular weight proteins, one should use membranes with a small pore size (0.2 or 0.1 μm). One drawback of this procedure is that the gel can dry out if the vacuum blotting is carried out over 45 minutes. In such a scenario, enough buffers should be used. In some cases, low-concentration polyacrylamide gels become stuck to the membrane following transfer. In such a scenario, one should rehydrate the gel to detach the gel residues from the nitrocellulose membrane.

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Biji T. Kurien and R. Hal Scofield Support pads

Gel Filter paper

Transfer membrane Support pads

Positive electrode

Fig. 3 The western blot transfer assembly. (Reproduced from Ref [10] with permission from Elsevier) 5.3

5.3.1

Electroblotting

Wet Transfer

5.3.2 “Semi-Dry” Transfer

Research laboratories most commonly use the electroblotting process to transfer proteins from a gel to a membrane. The major advantages are speed and the completeness of transfer compared to diffusion or vacuum blotting. Electroblotting can be achieved either by (a) wet transfer, namely the complete immersion of a gel-membrane sandwich (Fig. 3) in a buffer, or by (b) semi-dry transfer, namely placing the gel-membrane sandwich between absorbent paper soaked in transfer buffer. The conditions for transfer are dependent on gel type, the immobilization membrane, and the transfer apparatus used as well as the protein itself. SDS gels, urea gels [4], lithium dodecyl sulfatecontaining gels, nondenaturing gels, two-dimensional gels, and agarose gels have been used for protein electrophoretic blotting [18]. The electric charge of the protein should be determined, and the membrane should be placed on the appropriate side of the gel. Proteins from SDS-PAGE gels are eluted as anions, and therefore, the membrane should be placed on the anode side of the gel. When using urea gels, the membrane should be placed on the cathode side of the gel [4]. The wet transfer procedure involves placing the sandwich in a buffer tank with platinum wire electrodes. A large number of different apparatus are available to efficiently transfer proteins (or other macromolecules) transversely from gel to membrane. Most of these, however, are based on the design of Towbin et al. [1], which is that they have vertical stainless steel/platinum electrodes in a large tank. The “Semi-dry” transfer protocol requires the gel-membrane sandwich to be placed between carbon plate electrodes. “Semi-dry” or “horizontal” blotting uses two plate electrodes (stainless steel or graphite/carbon) for uniform electrical field over a short distance, and sandwiches between these up to six gel/membrane/filter paper

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assemblies, all soaked well in transfer buffer. The assembly is clamped or otherwise secured on its side, and electrophoretic transfer is carried out in this position, using as transfer buffer only the liquid contained in the gel and filter papers or other pads in the assembly. The advantages of this procedure compared to the conventional upright protocol are (a) gels can be blotted concurrently, (b) electrodes can be cheap carbon blocks, and (c) less power is needed for transfer. Consequently, this method requires only a simpler power supply instrument.

6

Conclusion Western blotting or protein blotting has evolved significantly since its establishment in 1979. There are numerous ways and means of transferring and detecting proteins currently. The benefits of protein blotting come from its capability of providing concurrent resolution of multiple immunogenic antigens within a sample for detection by specific antibodies. This has made it a very valuable technique, particularly for testing the specificity of antibodies to be used in ELISpot studies [10, 51, 52].

References 1. Towbin H, Staehelin T, Gordon J (1979) Electrophoretic transfer of proteins from polyacrylamide gels to NC sheets: procedure and applications. Proc Natl Acad Sci U S A 76: 4350–4354 2. LeGendre N (1990) Immobilon-P transfer membrane: applications and utility in protein biochemical analysis. BioTechniques 9(6 Suppl):788–805. Review 3. Southern EM (1975) Detection of specific sequences among DNA fragments separated by gel electrophoresis. J Mol Biol 98:503–517 4. Alwine JC, Kemp DJ (1977) Stark, G.R. Method for detection of specific RNAs in agar gels by transfer to diazobenzyloxymethyl-paper and hybridization with DNA probes. Proc Natl Acad Sci U S A 74:5350–5354 5. Burnette WN (1981) “Western blotting”: electrophoretic transfer of proteins from sodium dodecyl sulfate—polyacrylamide gels to unmodified NC and radiographic detection with antibody and radioiodinated protein A. Anal Biochem 112:195–203 6. Laemmli UK (1970) Cleavage of structural proteins during assembly of the head of bacteriophage T4. Nature 227:680–685

7. Kost J, Liu L-S, Ferreira J et al (1994) Enhanced protein blotting from PhastGel media to membranes by irradiation of low-intensity. Anal Biochem 216:27–32 8. Gershoni JM, Palade GE (1982) Electrophoretic transfer of proteins from sodium dodecyl sulfate-polyacrylamide gels to a positively charged membrane filter. Anal Biochem 124: 396–405 9. Gershoni JM (1988) Protein blotting: a manual. Methods Biochem Anal 33:1–58. Review 10. Kurien BT, Scofield RH (2006) Western blotting. Methods 38:283–293 11. Karey KP, Sirbasku DA (1989) Glutaraldehyde fixation increases retention of low molecular weight proteins (growth factors) transferred to nylon membranes for Western blot analysis. Anal Biochem 178:255–259 12. Harlow E, D (1988) Lane Immunoblotting. In: Antibodies. A laboratory manual. Cold Spring Harbor Laboratory, p 485 13. Renart J, Reiser J, Stark GR (1979) Transfer of proteins from gels to diazobenzyloxymethyl paper and detection with anti-sera: a method for studying antibody specificity and antigen

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structure. Proc Natl Acad Sci U S A 76:3116– 3120 14. Elkon KB, Jankowski PW, Chu JL (1984) Blotting intact immunoglobulins and other highmolecular-weight proteins after composite agarose-polyacrylamide gel electrophoresis. Anal Biochem 140:208–213 15. Gibson W (1981) Protease-facilitated transfer of high-molecular-weight proteins during electrotransfer to NC. Anal Biochem 118:1–3 16. Bolt MW, Mahoney PA (1997) High efficiency blotting of proteins of diverse sizes following sodium dodecyl sulfate-polyacrylamide gel electrophoresis. Anal Biochem 247:185–192 17. Kurien BT, Scofield RH (2002) Heat mediated, ultra-rapid electrophoretic transfer of high and low molecular weight proteins to NC membranes. J Immunol Methods 266: 127–133 18. Gershoni JM, Palade GE (1983) Protein blotting: principles and applications. Anal Biochem 131:1–15 19. Thornton DJ, Carlstedt I, Sheehan JK (1996) Identification of glycoproteins on nitrocellulose membranes and gels. Mol Biotechnol 5: 171–176 20. Tonkinson JL, Stillman B (2002) NC: a tried and true polymer finds utility as a post-genomic substrate. Front Biosci 7:c1–c12. Review 21. Lauritzen E, Masson M, Rubin I et al (1993) Peptide dot immunoassay and immunoblotting: electroblotting from aluminum thinlayer chromatography plates and isoelectric focusing gels to activated NC. Electrophoresis 14:852–859 22. Masson M, Lauritzen E, Holm A (1993) Chemical activation of NC membranes for peptide antigen-antibody binding studies: direct substitution of the nitrate group with diaminoalkane. Electrophoresis 14:860–865 23. Too CK, Murphy PR, Croll RP (1994) Western blotting of formaldehyde-fixed neuropeptides as small as 400 daltons on gelatin-coated NC paper. Anal Biochem 219:341–348 24. Coull JM, Dixon JD, Laursen RA et al (1989) Development of membrane supports for the solid-phase sequence analysis of proteins and peptides. In: Witmann-Liebold B (ed) Methods in protein sequence analysis. Springer-Berlag, Berlin, pp 69–78 25. Matsudaira P (1987) Sequence from picomole quantities of proteins electroblotted onto polyvinylidene difluoride membranes. J Biol Chem 262:10035–10038 26. Pluskal MF, Przekop MB, Kavonian et al (1986) BioTechniques 4:272–282

27. Xu QY, Shively JE (1988) Microsequence analysis of peptides and proteins. VIII. Improved electroblotting of proteins onto membranes and derivatized glass-fiber sheets. Anal Biochem 170:19–30 28. Kittler JM, Meisler NT, Viceps-Madore D et al (1984) A general immunochemical method for detecting proteins on blots. Anal Biochem 137: 210–216 29. Hughes JH, Mack K, Hamparian VV (1988) India ink staining of proteins on nylon and hydrophobic membranes. Anal Biochem 173: 18–25 30. Tovey ER, Baldo BA (1989) Protein binding to NC, nylon and PVDF membranes in immunoassays and electroblotting. J Biochem Biophys Methods 19:169–183 31. Moore C (2009) Introduction to western blotting. AbD serotec 32. Signore M, Reeder KA (2012) Antibody validation by Western blotting. Methods Mol Biol 823:139–155 33. Kurien BT, Scofield RH (1997) Multiple immunoblots after non-electrophoretic bidirectional transfer of a single SDS-PAGE gel with multiple antigens. J Immunol Methods 205:91–94 34. Otter T, King SM, Witman GB (1987) A two-step procedure for efficient electro transfer of both high-molecular weight (greater than 400,000) and low-molecular weight (less than 20,000) proteins. Anal Biochem 162:370–377 35. Harper DR, Kit ML, Kangro HO (1990) Protein blotting: ten years on. J Virol Methods 30: 25–39. Review 36. Egger D, Bienz K (1994) Protein (western) blotting. Mol Biotechnol 1:289–305 37. Wisdom GB (1994) Protein blotting. Methods Mol Biol 32:207–213 38. Kurien BT, Scofield RH (2003) Protein blotting: a review. J Immunol Methods 274:1–15. Review 39. Kurien BT, Scofield RH (2015) Electrophoresis – blotting techniques. In: Reedijk J (ed) Elsevier reference module in chemistry, molecular sciences and chemical engineering. Elsevier, Waltham. https://doi. org/10.1016/B978-0-12-409547-2. 11157-6 40. Reinhart MP, Malamud D (1982) Protein transfer from isoelectric focusing gels: the native blot. Anal Biochem 123:229–235 41. Jagersten C, Edstrom A, Olsson B et al (1988) Blotting from PhastGel media after horizontal sodium dodecyl sulfate-polyacrylamide gel electrophoresis. Electrophoresis 9:662–665

Western Blotting 42. Kazemi M, Finkelstein RA (1990) Checkerboard immunoblotting (CBIB): an efficient, rapid, and sensitive method of assaying multiple antigen/antibody cross-reactivities. J Immunol Methods 128:143–146 43. Heukeshoven J, Dernick R (1995) Effective blotting of ultrathin polyacrylamide gels anchored to a solid matrix. Electrophoresis 16:748–756 44. Olsen I, Wiker HG (1998) Diffusion blotting for rapid production of multiple identical imprints from sodium dodecyl sulfate polyacrylamide gel electrophoresis on a solid support. J Immunol Methods 220:77–84 45. Chen H, Chang GD (2001) Simultaneous immunoblotting analysis with activity gel electrophoresis in a single polyacrylamide gel. Electrophoresis 22:1894–1899 46. Bowen B, Steinberg J, Laemmli UK et al (1980) The detection of DNA- binding proteins by protein blotting. Nucleic Acids Res 8: 1–20 47. Kurien BT, Scofield RH (2000) Association of neutropenia in systemic lupus erythematosus

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with anti-Ro and binding of an immunologically cross-reactive neutrophil membrane antigen. Clin Exp Immunol 120:209–217 48. Kurien BT, Matsumoto H, Scofield RH (2001) Purification of tryptic peptides for mass spectrometry using polyvinylidene fluoride membrane. Indian J Biochem Biophys 38:274–276 49. Bischoff KM, Shi L, Kennelly PJ (1998) The detection of enzyme activity following sodium dodecyl sulfate-polyacrylamide gel electrophoresis. Anal Biochem 260:1–17. Review 50. Peferoen M, Huybrechts R, De Loof A (1982) Vacuum-blotting: a new simple and efficient transfer of proteins from sodium dodecyl sulfate-polyacrylamide gels to NC. FEBS Lett 145:369–372 51. Kurien BT, Scofield RH (2009) A brief review of other notable protein blotting methods. Methods Mol Biol 536:367–384. Review 52. Kurien BT, Dorri Y, Dillon S et al (2011) An overview of Western blotting for determining antibody specificities for immunohistochemistry. Methods Mol Biol 717:55–67. Review

Chapter 3 An Overview of Peptides and Peptide Pools for Antigen-Specific Stimulation in T-Cell Assays Karsten Schnatbaum, Pavlo Holenya, Sebastian Pfeil, Michael Drosch, Maren Eckey, Ulf Reimer, Holger Wenschuh, and Florian Kern Abstract The analysis of antigen-specific T-cell responses has become routine in many laboratories. Functional T-cell assays like enzyme-linked-immuno-spot (ELISPOT), which depend on antigen-specific stimulation, increasingly use peptides to represent the antigen of interest. Besides single peptides, mixtures of peptides (peptide pools) are very frequently applied. Such peptide pools may, for example, represent entire proteins (with overlapping peptides covering a protein sequence) or include noncontiguous peptides such as a collection of T-cell-stimulating peptides. The optimum specification of single peptides or peptide pools for T-cell stimulation assays will depend on the purpose of the test, the target T-cell population, the availability of sample, requirements regarding reproducibility, and, last but not least, the available budget, to mention only the most important factors. Because of the way peptides are produced, they will always contain certain amounts of impurities such as peptides with deletions or truncated peptides, and there may be additional by-products of peptide synthesis. Optimized synthesis protocols as well as purification help reduce impurities that might otherwise cause false-positive assay results. However, specific requirements with respect to purity will vary depending on the purpose of an assay. Finally, storage conditions significantly affect the shelf life of peptides, which is relevant especially for longitudinal studies. The present book chapter addresses all of these aspects in detail. It should provide the researcher with all necessary background knowledge for making the right decisions when it comes to choosing, using, and storing peptides for ELISPOT and other T-cell stimulation assays. Key words Peptides, Solid phase peptide synthesis, Peptide libraries, Peptide pools, Protein-spanning peptide pools, Peptide purity, Deletion peptides, Capping, T-cell activation assays, ELISPOT, Peptide storage, Peptide solvents, Epitope mapping

1

Introduction Synthetic peptides have been used in immunological experimentation around the world for decades. The basis for using peptides in T-cell stimulation assays was the seminal discovery in the late 1980s that the T-cell receptor recognizes short peptides embedded in Major Histocompatibility Complex (MHC) molecules

Alexander E. Kalyuzhny (ed.), Handbook of ELISPOT: Methods and Protocols, Methods in Molecular Biology, vol. 2768, https://doi.org/10.1007/978-1-0716-3690-9_3, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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[1]. However, researchers often use peptides without knowledge of how these peptides are made and how certain peptide features may affect the outcome of their experiments. Designing optimum peptides for T-cell stimulation can be a complex task. Regarding peptide length, a number of different considerations must be made, including MHC presentation, target cell populations, target amino acid sequence coverage, core epitope length, sample availability, and budget. Also, peptide preparations may differ significantly in terms of the purity of the “target product(s)” and the presence of by-products that could have an effect on the assay. There are measures that can be taken to reduce the toxicity of peptide preparations and to avoid the presence of impurities that might lead to false-positive stimulations. Knowing about and understanding these options is helpful when planning experiments. Once all the important criteria for successful T-cell activation are met, there is still a lot of room for creativity when designing the optimum peptide pool(s) for T-cell stimulation. Overlapping peptides of various lengths may be pooled to cover longer stretches and even entire proteins. Peptide pools that cover entire proteins may additionally include known sequence variations. Alternatively, peptide pools may contain several or many noncontiguous T-cell-stimulating peptides. There are several reasons why peptides are preferable over proteins in T-cell stimulation assays: peptides can be manufactured reproducibly and with high purity, quality control is simple and can be automated, no protein expression systems are required, sequence variations of interest can be covered easily, and, finally, specific post-translational modifications can be introduced as required. This book chapter covers the topics that we believe are most relevant to scientists using peptides in their T-cell assays whether that involves research or clinical assay development. Our intention is to provide useful background knowledge while focusing on the application. Understanding the principles and mechanisms that lead to successful T-cell activation while avoiding pitfalls is paramount to designing suitable peptides and peptide libraries (i.e., the list of peptides that go into a pool). The central topic of this chapter is the use of peptides in ELISPOT assays; however, what we discuss also applies to other types of T-cell stimulation assays that use peptides, irrespective of the readout.

2

Peptide Synthesis, Purification, Analysis, and QC/QA

2.1 Peptide Synthesis

Peptides for T-cell activation assays are usually synthesized by Fluorenylmethoxycarbonyl (Fmoc)-based solid-phase peptide synthesis (SPPS) [2]. In this process, Fmoc amino acid couplings and Fmoc group de-protections are carried out in an iterative manner

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Fig. 1 Capping during solid-phase peptide synthesis (SPSS) avoids deletion peptides. (a) General principle of SPPS, (b) SPPS without capping, and (c) SPPS with capping. Black circles represent the solid support, lighter circles represent amino acids. Fmoc = 9-fluorenylmethyloxycarbonyl; TFA = trifluoroacetic acid

(one amino acid at a time from the C-terminal to the N-terminal end) on solid support until the desired amino acid sequence is synthesized. Subsequent treatment with trifluoroacetic acid (TFA) induces cleavage from the solid support and removal of temporary side chain-protecting groups, thus releasing the desired peptide (Fig. 1a). Solid supports in SPPS are usually polystyrene-based resins. However, in recent years, peptides synthesized on cellulose membranes by a high-throughput method called Spot synthesis [3] have also been successfully used in ELISPOT experiments [4, 5]. Spot peptides are especially advantageous for epitope mapping studies that would not be economically viable using classical resin-based synthesis approaches. Because amino acid coupling reactions do not always proceed with 100% efficacy, peptides synthesized by SPPS frequently contain impurities composed of shorter peptides. Such shorter peptides may have missing amino acids along the sequence (referred to as “deletion peptides”) or at the end (“truncation peptides”). Truncation peptides may form when the coupling of one amino acid is incomplete, and no further amino acid couples in subsequent synthesis steps (Fig. 1b). As a result, truncation peptides are simply shortened versions of the target peptides but without sequence deviations. Deletion peptides, by contrast, lack one or more amino acid(s) within the target peptide sequence and as a result show sequence deviations. They are formed when an incomplete coupling reaction occurs at one step but one or more subsequent

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coupling steps are successful (Fig. 1b). Whereas truncation peptides might stimulate the same T-cells as the full-length peptide (depending on their length), deletion peptides may give rise to de novo T-cell epitopes recognized by different T-cell populations. As a result deletion peptides may give rise to unwanted false-positive T-cell responses [6]. The potentially huge consequences of falsepositive results in immunoassays are obvious. In the worst case, such false-positive results could lead to the selection of a suboptimal candidate peptide in vaccine or immunotherapy development. For example, if the preparation of a peptide that is nonstimulating in itself contains a stimulating deletion peptide, the nonstimulating peptide might erroneously be taken forward as a candidate. In this context, it is important to note that even small amounts of deletion peptides may have large effects on immunological assays. This is because memory T-cells are often highly sensitive to even small amounts of recognized peptides, so contamination could represent only 1% or even less of the preparation but still provide efficient stimulation. Because of the potentially significant problems arising from deletion peptides, strategies for avoiding them were developed. One of them is the purification of peptides after synthesis. However, as deletion peptides are usually structurally and chemically similar to the target peptide, their separation is often difficult [7]. An alternative strategy is to reduce the extent of incomplete coupling events by performing multiple coupling steps for each amino acid (e.g., triple couplings instead of the more usual single or double couplings). However, this is more laborious and does not entirely eliminate the problem. A final strategy to avoid deletion peptides that we would like to discuss here is a process called “capping” (Fig. 1c). Peptide capping uses an additional synthesis step that chemically modifies free amino functions resulting from incomplete couplings in such a way that they cannot react further. Practically, this is done by acetylation with the highly reactive reagent acetic anhydride [8]. SPPS with capping mainly produces acetylated (Ac-modified) truncation peptides as side products, which are usually easy to remove during peptide purification (Fig. 1c). Most importantly, after capping, further extension of the peptide in subsequent coupling steps is not possible, which prevents the formation of deletion peptides. The concept of minimizing deletion peptides by capping has been applied in a number of studies [9–12]. 2.2 Peptide Analysis and Purification

Peptide analysis is routinely carried out by high-pressure liquid chromatography coupled with mass spectrometry (HPLC-MS). HPLC separates the target peptide from by-products. This is achieved by chromatography on a hydrophobic stationary phase using mixtures of water and organic solvents, mostly acetonitrile, as mobile phase. The mobile phase eluting from the column is

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normally analyzed by two methods: (a) UV absorption using a UV lamp (wavelength 220 nm) and a UV light detector and (b) mass spectrometry. Analysis by UV absorption is done to provide information on the relative amounts of the target peptide as well as peptidic side products. The ratio of target peptide/all peptide products is commonly referred to as peptide purity. The wavelength of 220 nm is the standard for determining peptide purity, because this is the wavelength of UV light absorbed by the amide bonds of the peptide backbone. In the next step, different mass spectrometric approaches may be applied to confirm the correct molecular mass (i.e., the identity) of the target peptide and identify the molecular masses of its accompanying impurities. MS approaches may include different types of ionization, for example, electrospray ionization (ESI), as well as a range of detection methods such as time of flight (TOF), ion trap, or quadrupole. Which of these technologies is best suited to identify a peptide depends on its chemical and physicochemical properties. In addition to HPLC-MS, a range of additional analytical methods can be applied for the assessment of peptide composition and purity. These include high-resolution exact molecular weight determination, peptide sequencing, peptide quantitation/peptide content determination by amino acid analysis (AAA), determination of enantiomeric integrity, residual solvent determination, residual water determination, and residual counterion determination. Peptide purification is usually done by reversed-phase HPLC (RP-HPLC) [13]. The main advantage of RP-HPLC for peptide purification is that it can be automated relatively easily, is reproducible, and works well for the majority of peptides. However, handling MHC ligand peptides that display a certain degree of hydrophobicity requires experience [14]. Disadvantages of RPHPLC are its relatively high cost and consumption of large amounts of organic solvents. “Greener,” i.e., more environmentally sustainable procedures aiming to reduce organic solvent consumption are thus in development [15]. A further disadvantage of RP-HPLC is that contaminating peptides whose physicochemical properties are very similar to those of the target peptide (e.g., deletion peptides) are difficult to remove. Several other methods for preparative peptide purification are in use but are less common. These include normal-phase HPLC [16], capillary electrophoresis [17], or chromatography-free purification systems [18, 19]. The level to which peptides for T-cell assays should be purified strongly depends on the application. While peptides for T-cell immune monitoring in clinical studies are usually purified to >70% or higher, other applications do not require such high purities. For example crude peptides are often used for T-cell epitope mapping because of the relatively high number of required peptides in combination with the requirement for only small amounts.

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In a peptide pool of defined purity, every single peptide should meet the desired specification (i.e., in a pool with >90% purity, every peptides has to be >90% pure). The use of “average” or “mean” purity to characterize peptide pools is not useful since in this case some peptides in the pool might have very high purity while others, which could be the most important ones, may have very low purity. When crude peptides are used in T-cell assays like ELISPOT it should be ascertained that each and every individual target peptide is present in the pool, as otherwise critical immune responses might be missed. This is especially relevant for hydrophobic peptides, which may be difficult to synthesize but are more often found to give rise to T-cell epitopes than hydrophilic ones [20]. In any case, the optimum peptide specification for a given purpose should be determined carefully and, ideally, with expert advice. 2.3 Peptide Impurities, quality assurance (QA), and quality control (QC)

Depending on the target specification, synthetic peptides will contain varying amounts of impurities [21]. Table 1 provides an overview of common impurities in peptide preparations and strategies to minimize their presence. While it is common to use HPLC to

Table 1 Impurities in peptides and strategies for their minimization

Source

Nature of impurity

Synthesis

Deletion peptides Truncation peptides Various other peptides, e.g., cyclized, oxidized, deamidated, beta-eliminated, and racemized peptides

Countermeasure/strategy for reduction • Capping • HPLC purification • Optimized coupling and Fmoc deprotection conditions, e.g., using advanced coupling reagents [23] • HPLC purification • Optimized synthesis conditions, e.g., using special building blocks • Optimized TFA cleavage conditions • HPLC purification

Synthesis and/or Residual solvents purification Water

• Lyophilization using optimized conditions • Lyophilization using optimized conditions

Storage/ Various peptides, e.g., cyclized, oxidized, decomposition deamidated, and beta-eliminated

• Low storage temperatures • Minimization of freeze–thaw cycles • See also Chap. 5

Other peptides handled in parallel (e.g., Crosscontamination synthesized/purified on same from other instruments) syntheses

• Line clearance • Stringent QC/QA measures • Pretesting in T-cell assay

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remove impurities, optimized synthesis conditions may avoid them in the first place. While the amount of deletion peptides is most efficiently minimized by capping, truncation peptides should be minimized by efficient coupling and Fmoc de-protection conditions. Optimized synthesis conditions are key to the reduction of many potential additional impurities, including protecting group artifacts, partially oxidized, hydrolyzed, rearranged or decomposed products. Unless present in excessive concentrations, residual solvents may not be detrimental in T-cell assays. However, their amount should be minimized and can be quantified if desired. Stringent line clearance and QA/QC measures are crucial to reducing impurities from cross-contaminations (see the following chapter). Rarely, false-positive responses in T-cell assays resulting from cross-contamination [22] have been reported. Measures to minimize impurities could have avoided such complications. An important lesson from the cited study is that even small amounts of peptides with aberrant sequences may create significant problems in T-cell assays. Theoretically, peptide impurities with sequence deviations are the greatest problem with respect to false-positive results but can be very effectively reduced by capping and avoidance of cross-contamination. Even though counterions of peptides are not considered impurities, they are typically present in considerable amounts. This is because every basic group in a peptide (free N-terminus, lysine, and arginine side chain) is typically protonated and, therefore, requires a counterion by default. Trifluoroacetate (TFA) is the most common counterion in peptides because TFA is used during peptide cleavage and purification by HPLC. Counterions like TFA are usually present in sub-toxic concentrations and buffered by the reconstitution medium before applied in biological experiments. However, as for minimization of residual solvents, measures like thorough lyophilization, which is preferred over other vacuum drying methods such as the use of SpeedVacs, should be taken to eliminate an unnecessary excess of TFA. If there is any doubt, it is recommended to measure counterion content or to perform cell toxicity testing. Low-bioburden manufacturing processes are recommended for all peptides used in T-cell assays. This is already ensured to a large extent by using organic solvents in the manufacturing process. Additional measures can be applied to minimize the risk for biological contaminations (hygiene, pest control, etc.). In highly regulated environments, the presence of biological contaminations and/or their impact should be measured. Typical assays for this purpose are bacterial endotoxin testing, peptide sterility testing, and bioburden determination. Line clearance is another important element of QA/QC. Cross-contamination of synthetic peptides and peptide pools with unrelated peptides is not uncommon and represents a

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serious risk for T-cell assay performance [22] and regulatory compliance. This is because peptide manufacturers often process many orders in parallel for economic reasons. Depending on the QA/QC measures and standard operating procedures (SOPs) in use, multiple peptides might be produced in the same working space using the same peptide synthesizer, the same HPLC column for purification, the same freeze-drying instrument, etc. In such a setting the risk of cross-contamination must be mitigated as effectively as possible. Depending on the specific requirements of a given synthesis such measures may include the exclusive use of thoroughly cleaned and/or new components where these are replaceable (e.g., columns), dedicated instruments, specialized cleaning procedures, organizationally or spatially separated laboratory environments, and more. Proof of presence of each peptide in a peptide pool is very important for quality and regulatory compliance. But even stateof-the-art analytical methods are not always able to identify all peptides in a pool, especially in a large pool. Therefore, alternative workflows, for example, based on sub-pool generation and analysis, are recommended to provide evidence for the presence of all peptides. Finally, it should be mentioned that peptide stability and solubility testing are additional QA/QC measures that are increasingly applied to peptides for (clinical) immune monitoring. Degradation of peptides during storage may be attributed to a variety of chemical reactions such as oxidation, hydrolysis, structural rearrangements, and more. Despite this knowledge, the stability of a given peptide is hard to predict. Stability testing is, therefore, advisable for certain applications. More specifically, it is recommended to perform regular stability testing during long-term storage. The so obtained information is useful to define expiry dates and to inform optimum storage conditions. A stress stability test at elevated temperatures (e.g., 40 °C) can be performed in order to estimate longterm stability of peptides in a short time frame. Like peptide stability, peptide solubility is influenced by many factors and is also hard to predict from the amino acid sequence alone. In addition, the applied reconstitution medium or solvent together with the concomitant procedure has a great impact on the final solubility. The nature of accompanying peptides in a pool may also affect the solubility of peptides in a peptide pool. It is, therefore, recommended to perform solubility testing for applications where peptide solubility is critical.

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Peptide Length and Applications The binding of peptides to the MHC (or human leukocyte antigen (HLA) in humans) is a prerequisite for the recognition by the TCR

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(“TCR-MHC/peptide recognition”) [24, 25]. The TCR recognizes a portion of the peptide itself but also a portion of the MHC molecule, both of which make a contribution to the actual T-cell epitope. Without an MHC molecule, there is no T-cell epitope. We usually speak about “T-cell stimulating peptides” when talking about the peptide component of an epitope in isolation. A range of peptide properties determines whether or not a peptide can bind to an MHC molecule. Such binding is not covalent but driven by a mix of physical interactions (mostly hydrogen bonds and hydrophobic interactions) between the amino acids of the peptide and the amino acids in the binding groove. The MHC binding groove contains pockets that can accommodate certain amino acid side chains but not others, thus providing the basis for its specificity toward certain peptides. To begin, maybe the most basic determinant for the binding of peptides to class-I MHC molecules is peptide length [26]. The class-I MHC binding groove is closed at both ends, which limits the length of the peptides that can be accommodated to typically 8–10 amino acids. So, even if a peptide otherwise perfectly matches the binding groove, it cannot be too long and has to be trimmed to the right length before it can bind. Such trimming is thought to occur by extracellular peptide processing [27–29]. Exceptionally, longer peptides can form a loop that sticks out of the binding groove [30, 31]. The class-II MHC binding groove, by contrast, is open at both ends, so that longer peptides can be accommodated. Nevertheless, the length of the peptide “core,” i.e., the portion of the peptide that is interacting with the binding groove, is the same as the length of class-I MHC binding peptides, i.e., 8–10 amino acids. It is very likely, though, that flanking amino acids have a role in TCR-MHC II/peptidecomplex recognition. Whereas length is a very simple parameter that influences peptide-MHC binding, the sequence of amino acids in the peptide is a complex parameter. With 20 natural amino acids, the total number of possible peptide sequences in a 9-aa peptide is 20Exp9 or 512,000,000,000. MHC binding grooves are designed in such a way that only peptides with certain side-chain properties in certain positions along the binding groove will actually bind. These include so-called anchor residues, which provide a particularly strong binding of peptides to MHC molecules [32]. For each MHC allele there are specific binding requirements, which can be the same or similar for very closely related alleles but tend to be quite different for alleles that are not closely related. T-cell stimulating peptides are generally used for identifying or monitoring T-cell responses to selected proteins or specific epitopes, or to identify T-cell epitopes. The exact purpose of the research, the amount of biological samples, and the available budget tend to be key determinants of the selected experimental design. For example, if the research focuses on CD8 T-cells, there is a sufficient amount of biological samples, and a sufficiently large

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budget, short peptides optimized for CD8 T-cell stimulation are a good choice. In that case, a whole protein could be represented by consecutive 9-amino acid peptides with an overlap of eight amino acids. Covering a stretch of 100 amino acids would thus require 92 peptides. However, if the amount of sample is not sufficient for that many tests, or the required budget is not available, a different design might be chosen. An alternative would be a peptide length of 15 amino acids and an overlap of 11 amino acids. In that case, representing the same stretch of 100 amino acids would require only 23 peptides, with the last peptide being shorter. In fact, there is ample experimental evidence that 15 amino acid peptides are still short enough to allow reliable identification of CD8 T-cell responses in ELISPOT experiments, and, on the other hand, such peptides also work very well for CD4 T-cell stimulation [33]. This particular design appears to be the most common one to date for T-cell response detection/monitoring with protein-spanning peptide pools. Even longer peptides may still work well for CD4 T-cells, but appear to be less efficient for stimulating CD8 T-cells. Actually, it was previously shown that when using 15-aa peptides for stimulating CD8 T-cell responses, slightly higher concentrations of these peptides (in micrograms/milliliter) were needed to achieve the same level of stimulation (% of CD8 T-cells responding) compared with 9-aa peptides [33]. Any design that deviates from the optimum length for CD4 or CD8 peptides is obviously a compromise, but generally, some compromise has to be made. The typical concentration of each 15-aa peptide in a shortterm stimulation assay is in the range of 1 ug/mL, but the optimum concentration might depend on the peptide sequence and purpose. When designing a peptide pool covering a whole protein, it is important that the overlap between peptides is sufficient to ascertain that potential epitopes are not missed. The overlap of 11 aa between two 15-aa peptides ensures that any possible stretch of 12 aa length is found in at least one of the overlapping 15-aa peptides, whereas any peptide of 11 aa length is contained in at least two 15-aa peptides. There is some evidence that certain amino acids may block the above-mentioned trimming (“clipping”) of peptides, so having the identical 11-aa sequence contained at different positions in two adjacent 15-aa peptides is of advantage [33]. Taken together, protein-spanning peptide pools with 15-aa peptides and an overlap of 11 amino acids represent a good compromise to achieve both MHC I and MHC II stimulation. They are widely used for a broad range of applications.

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Peptide Libraries and Pools Peptide pools are established tools for stimulating antigen-specific T-cell responses in T-cell assays for a wide variety of applications. These applications range from antigen and epitope discovery to

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Fig. 2 The best peptide library design depends on the application. Commonly used types of peptide libraries for peptide pool generation are shown. Left: overlapping peptides representing a whole protein antigen (protein or antigen-spanning peptide pool). Middle: peptides covering sequence variants. Right: selection of T-cellstimulating peptides from various proteins of one virus. Beads represent single amino acids

immune monitoring in research and clinical trials to the development of diagnostic tests. Depending on the application, different types of ‘peptide libraries’ are used to generate peptide pools (Fig. 2). As explained above, for the purpose of this chapter, a list of peptides compiled for generating a peptide pool is referred to as ‘peptide library’. The collection of actual physical peptides, by contrast, is referred to as peptide pool. 4.1 Pool Size and Limitations to the Number of Peptides PerPool

There is no general rule when it comes to the optimum/maximum number of individual peptides in peptide pools. Too many peptides in a pool might impair solubility and lead to precipitation of peptides, thus potentially resulting in lower spot numbers and potentially less reproducible results in ELISPOT assays. Competition for MHC binding sites [34] might also be an issue for very large pools, although at the same time, there is evidence that outcompeting a good binder requires excessive concentrations of competing peptides that are not present in peptide pools containing the same amount of each. Based on the above considerations and our longterm experience, we recommend using a maximum number of 200 peptides per peptide pool as a reasonable approach. Examples where this limit was applied are the CEFX pool (176 peptides) [35] and the SARS-CoV-2 spike glycoprotein pool (315 peptides divided in two sub-pools of 158 and 157 peptides) [36]. It is, however, possible to exceed this ‘reasonable’ limit under certain circumstances. This will depend on the solubility of the peptides, the amount of available sample and the specific application, to name the most important determinants [37].

4.2 ProteinSpanning Peptide Pools

Protein-spanning peptide pools are generated by dividing the complete sequence of a protein into overlapping peptides of a specific length and with a specific overlap or ‘offset’. The optimum peptide

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Fig. 3 Peptide pools can effectively represent entire proteins. Design of a protein-spanning peptide library. The example shows peptides of 15 amino acids (aa) length overlapping by 11 aa along the original protein sequence (top). Peptides can be synthesized and combined in one single pool (whole protein-specific stimulation) or divided into several pools (e.g., for epitope mapping). This is the most common type of peptide library used to provide stimulation at the whole protein level

length for epitope mapping may vary and could range from 8 to 20 amino acids with an offset of 1–4 amino acids. However, it was found that 15-aa peptides with an overlap of 11 aa covering an entire protein represent a good compromise for stimulating and detecting both CD8 and CD4 T-cells [33]. The schematic diagram in Fig. 3 shows an example of a peptide pool with a peptide length of 15 aa and an overlap of 11 aa. 4.3 Peptide Pools Covering Sequence Variants

Sequence diversity is a hallmark of many pathogenic viruses and also tumors. Somatic mutations in tumors cause considerable sequence heterogeneity between and even within individuals, which can significantly alter tumor immune recognition. For example, 29,881 somatic mutations were previously identified for TP53, one of the most important tumor driver genes [38]. Likewise, many pathogenic viruses exhibit a high sequence diversity because of the accumulation of mutations and recombination events. This

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sequence diversity should be taken into account when designing peptide libraries, irrespective of whether these pools are intended for immune monitoring or vaccination. Therefore, complex peptide libraries are required to reflect the full spectrum of sequence variation among virus strains or across a patient population. Bioinformatics algorithms have been developed for this purpose, creating all possible peptides and scoring these according to their frequency of occurrence across all samples. In this way, the algorithms identify the optimal layout for providing homogeneous overall sequence coverage. If the number of input sequences is too large or too diverse to be covered in a single library, the library size can be limited by selecting peptides according to their contribution to the overall coverage. For example, a conventional peptide library for the HIV-Nef protein generated from the model sequence HXB2 alone (49 peptides) attains a coverage of 10-10) [26], markedly different spot morphologies can be expected to arise. Depending on the extent of affinity maturation that the memory B-cell (Bmem) repertoire has undergone with respect to an antigen, which increases with repeated and long-lasting exposure [27–29], this affinity distribution (and hence spot morphology in the direct assay) will show fundamental variations from antigen to antigen, between different individuals, and possibly even between sequential bleeds of the same individual. On the one hand, this diversity is a challenge to automated image analysis using legacy counting parameters. However, on the other hand, if the individual secretory footprints are accurately assessed for high-content information, such diversity can provide invaluable insights into the affinity distribution of the antigen-specific Bmem repertoire in any given individual, at any given time point (for more on this issue, we refer to the chapter by Becza et al. in this volume) [8]. High-content analysis (HCA) of spot morphologies in antigenspecific direct ImmunoSpot® assays can, therefore, provide thus far underexploited information about individual ASC [25]. For practical use, HCA should allow the identification and quantification of subpopulations of B cells within the antigen-specific ASC repertoire, assessing affinity distributions and productivities as shaped by vaccinations, infections, allergens, and autoantigens. 1.1 More Diverse Spot Morphologies Are Observed in Direct ImmunoSpot® Assays Than in Pan-Ig Assays

Due to the diverse affinity of the individual B cells, in assays in which the antigen itself serves as the capture reagent for binding the ASC-derived Ig, antigen-specific ASC produce a wider range of spots with different morphologies than when these ASC-derived Ig are captured using anti-Ig-specific capture antibodies in pan-Ig assays (of which images of representative wells are shown in Fig. 2 and see Note 5). Such visually observable heterogeneity of spot sizes and intensities can be quantitatively evaluated using dot plots of spot intensity versus spot size (Fig. 3). In the pan-Ig assay, spots are bright and compact, and the intensity versus size dot plots accordingly show a single population of spots with a narrow distribution for both parameters. This also applies to T-cell and inverted B-cell assays (see Note 5). In the antigen-specific direct assay, in contrast, the size/density distributions are much broader and distinct spot (ASC) subpopulations can be seen.

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Fig. 2 Representative images of (a) pan-Ig and (b) antigen-specific direct B cell ImmunoSpot® assay wells detecting secretory footprints of IgG+ ASC. For schematic representation of the two assay types, please refer to Fig. 1, the protocols are detailed in Materials and Methods. In (a), anti-human Igκ/ Igλ-specific antibodies were used to capture the Ig produced, in (b) the membrane was affinity-coated with SARS-CoV-2 Spike protein. Polyclonally preactivated peripheral blood nuclear cells (PBMC) from a COVID-19 mRNAvaccinated donor were tested in serial dilution. In both cases, spots were visualized with anti-human IgG-specific detection antibodies. Representative wells containing secretory footprints in the Goldilocks range are shown: (a) 4 x 103 PBMC/well and (b) 2 x 104 PBMC/well. In panels (c) and (d), images from panels (a) and (b) are represented in 3D format for the best visualization of morphological differences between spots in pan-Ig and antigen-specific direct assays. Note that maximal intensity peak values do not exceed camera dynamic rage (no peak “trimming” observed) permitting accurate HCA analysis of total “spot mass”

1.2 Size Distribution of Antigen-Specific ASC Does Not Follow Log Normal Distribution and Cannot Be Analyzed Using Statistics-Based Size Gating

In T-cell ELISPOT, the cytokine of interest (e.g., interferongamma) can also be produced by non-T bystander cells such as monocytes, basophils, dendritic cells, and other cell types [30, 31]. However, we have shown that the spot size distribution for antigen-stimulated memory T cells consistently follows a log-normal distribution pattern that permits discrimination between T-cell- and bystander cell-derived spots in order to count the former while neglecting the latter. This approach is based on an automated gating strategy that subjects spot size distributions to statistical analysis [14, 32]. In contrast to T-cell cytokines, only B cells can secrete immunoglobulin/antibody and thus all detectable spots (see Note 6) must be counted. Establishing the upper limit of the number of spots per well originating from individual ASC in antigen-specific direct assays is critical: this limit depends

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Fig. 3 Representative dot-plots of mean spot intensities versus spot sizes generated by ImmunoSpot® Studio Software for (a) pan-IgG, and (b) SARS-CoV-2 Spike-specific IgG+ ASC detected in the respective B cell ImmunoSpot® assay. For the raw data, the legend to Fig. 2 applies. Secretory footprints at the respective Goldilocks numbers were subjected to high content image analysis using the IntelliCount® module of the ImmunoSpot® Software by merging data from multiple replicate wells. The individual secretory footprints (“events” in flow cytometry terminology) are represented in flow cytometry standard dot-plot format. Mean spot intensities (Y axis, calculated as a sum of all pixels intensities in an individual spot divided by the number of pixels) are plotted versus the respective spot’s size (X axis). Such high content spot morphology data are automatically generated as graphs in the ImmunoSpot® software. The raw data containing 12 numeric parameters to detail each SFU’s morphology are automatically captured as FCS files and can be subjected to more detailed analysis using any FACS software suite

upon the morphology of ASC-derived secretory footprints which, in turn, reflects the affinity of secreted antibodies (see Note 7). In this same context, ELISA effects (see Note 8) and merging of secretory footprints can interfere with accurate quantification. We, therefore, tested whether ASC-derived spots in antigen-specific direct ImmunoSpot® assays follow a distribution that can be leveraged for automated gating. As shown in Fig. 4, spot size distributions for antigen-specific ASC in direct assay do not follow a log-normal function. This deviation from normality is related to the polyclonal nature of an antigen-specific B-cell repertoire: spot size distributions of individual ASC not only reflect differences in their productivity, as for T cell cytokines, but also in their affinity and fine epitope specificity. 1.3 Artificial Intelligence-Based IntelliCount™ Provides Accurate User- and Assay-Independent Counts of AntigenSpecific ASC-Derived Spots

Because of the relatively simple spot morphologies involved, traditional thresholding and fixed parameter-based image analysis algorithms are suited for providing accurate and scientifically validated spot-forming unit (SFU) counts for T-cell ELISPOT [14], and multicolor T-cell FluoroSpot assays [5, 13]. Such conventional parametric counting is also well suited for pan-Ig and inverted B-cell assays [7, 11, 18, 20, 21], but it frequently disappoints when applied to antigen-specific, direct B-cell assays. This is the

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Fig. 4 Spot size distributions in antigen-specific B cell ImmunoSpot® assay do not follow a log-normal function. Left panel (a) shows representative spot size (mm2) histogram calculated for ~800 IgG+ spots from SARS-CoV-2 Spike-specific B cell ImmunoSpot® assays. For the underlying raw data, legends of Figs. 2 and 3 apply. Green line corresponds to the best fit of these data with the log-normal distribution function. Right panel (b) represents QQ plot for experimental versus theoretical log-normal distributions. Systematic deviation of QQ plot (round dots) from the straight line proves that the experimental spot size distribution does not follow a log-normal function. Shapiro–Wilk (0.96) and Lilliefors (0.09) statistical tests also rejected the log-normal distribution hypothesis with a 5% significance level

case when frequencies of antigen-specific ASC in test samples (e.g., PBMC) show considerable variability (which is generally observed, see Note 7). In particular, this is the case when the intent of the assay is to extract high-content information on the wide spectrum of secretory footprints, beyond obtaining mere SFU counts (e.g., to assess the affinity distribution of an antigen-specific B-cell repertoire). Even when counting parameters are fine-tuned manually well-by-well (see Note 9), it remains challenging to establish parameters to simultaneously detect fuzzy, spread-out, low-intensity spots along with bright compact spots in the same well, and to accurately establish the boundaries for each SFU. The latter, however, is essential for the precise quantification of the ASC-derived Ig retained within each SFU, i.e., the “spot mass” (see Note 10). An artificial-intelligence (AI)-based spot recognition algorithm, however, inherently lends itself better to such a task. IntelliCount™ is built around CTL’s proprietary deep neural network that was trained using tens of thousands of ELISPOT and FluoroSpot images. Thus, it does not require special knowledge for setting counting parameters, and instead, data analysis becomes fully automated and objective. High-content analysis of secretory footprints may also require high dynamic range (HDR) imaging to accurately quantify fluorescence intensity for spots whose luminosity spans a sufficiently wide range and cannot be fully captured using a single fixed exposure image. IntelliCount™ fully supports HDR-based HCA integration to quantitatively assess the characteristics of individual spots.

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ImmunoSpot® Studio software generates comprehensive HCA data outputs in the form of Flow Cytometry Standard (FCS) files. Such data can be processed and viewed within the ImmunoSpot® software as dot plots and histograms, or can be readily exported into advanced flow cytometry suites for more detailed statistical analysis using multidimensional gating and other tools. 1.3.1 IntelliCount™ Automatically Establishes Accurate Spot Boundaries for High-Content Analysis

One of the goals of HCA is to quantify the net analyte captured within each secretory footprint, i.e., the “total spot intensity” or “spot mass” (see Note 10). To obtain this information from antigen-specific B-cell assays, spot boundaries have to be defined accurately to include the entire spot area. Images of a representative well that was counted using threshold-based parametric counting versus IntelliCount™ are presented in Fig. 5. In this sample, well

Fig. 5 Precision of thresholding-based assessment of secretory footprint outlines (a) versus the IntelliCount™-based automated detection of such in the same well (b). The same well as shown in Fig. 1B from a direct SARS-CoV-2 Spike-specific (IgG+) assay was analyzed using (a) conventional parametric or (b) IntelliCount™ counting algorithms in ImmunoSpot® Studio Software. Zoomed-in sections of the same well analyzed with parametric and IntelliCount™ modes are shown in panels (c) and (d), respectively. The spot counts are shown in yellow

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containing SFU at the “Goldilocks number” was analyzed (see Note 11), both approaches yield similar overall SFU counts, but the spot areas are largely underestimated by intensity thresholdbased parametric counting. Importantly, in parametric spot recognition, spot boundaries are defined by an intensity threshold set at a discrete background level above which “total spot intensity” is calculated (much as the sea level defines the shoreline of an island). Any fluctuations of the background within and between wells (see Note 12) will affect spot boundaries and do so dramatically for faint spots. The challenge of properly defining SFU sizes, or even recognizing individual SFU, increases when their frequencies exceed the Goldilocks number since this can result in both local and global ELISA effects (see Note 8). In IntelliCount™ mode, in contrast, the morphology of each spot is analyzed in its entirety, and spot boundaries are calculated with precision for each spot’s individual modeled shape, irrespective of background fluctuations. These boundaries are used to assess the amount of analyte retained within each secretory footprint via the fluorescent intensity of the captured label. 1.3.2 IntelliCount™ Counting Mode Provides Extended Linear Ranges for Accurate Calculation of ASC Frequencies

Antigen-specific, direct B-cell ImmunoSpot® assays are primarily performed to determine the frequency of antigen-specific, memory B-cell-derived ASC producing different classes and subclasses of Ig in PBMC (or other primary cell material) [22]. By doing so, one can predict the magnitude and quality of effector functions mediated by Bmem upon antigen reencounter, when they engage in secondary-type antibody responses (see Notes 13 and 14). A technical challenge in doing so is that frequencies of antigen-specific Bmem-derived ASC-producing antibodies of a given class/subclass occur at markedly different interindividual frequencies, even when assessed in individuals at the same time point after infection/ vaccination [18] (see Note 7). Moreover, the frequencies of Bmem producing different classes and subclasses of Ig are also typically orders of magnitudes apart in individuals [18]. Furthermore, within an individual, the frequency of Bmem recognizing different antigens is also highly variable, dependent on the individual’s memory status relative to each antigen [12]. True frequencies of antigen-specific Bmem can only be established in ImmunoSpot® assays under conditions when secretory footprints of individual ASC are clearly discernable; in such cases, the number of SFU counts per well reveals the number of antigenspecific ASC among all PBMC plated into that well, i.e., their frequency. When SFU numbers per well increase, the expected direct linear relationship between numbers of cells plated and spots counted breaks down because of merging of secretory footprints and ELISA effects (see Note 8). At low spot counts, however, frequency estimates become imprecise, at least when a limited

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number of replicate wells are tested, due to the onset of Poisson noise (see Notes 15 and 16). Therefore, there is a certain range of SFU numbers per well from which reliable data can be extracted for precise ASC frequency calculations (and even more stringently, for high-content spot morphology analysis). Therefore, frequencies of ASC are best estimated when PBMC are seeded in serial dilution into ImmunoSpot® wells and are calculated by extrapolation from the linear portion of the graph in which SFU counts per well/ PBMC seeded per well are plotted [12]. The ImmunoSpot® Studio software implements a Linear Range Finder function for automatic frequency calculations: using statistical analysis, it finds the initial linear part in the cell titration results and calculates frequencies by linear regression from these accurate SFU counts. The chapter by Yao et al. in this volume [22] introduces protocols on how to readily measure frequencies of antigen-specific Bmem-derived ASC producing distinct Ig classes (or IgG subclasses) with only 4 × 105 PBMC per antigen, leveraging four-color ImmunoSpot® analysis; it also established that the frequency of ASC can be established by a single-well serial dilution approach as precisely as when done involving four replicates in order to maximize utility of precious cell material. With the ImmunoSpot® Software, the generation of cell titration graphs and the frequency extrapolations are fully automated, as shown in Fig. 6. Figure 6 shows the results of a serial dilution experiment utilizing PBMC in which the frequencies of pan-IgG+ memory B-cellderived ASC were determined by the Linear Range Finder regression analysis using legacy parametric counting versus the IntelliCount™ approach. Note the higher number of data points falling into the linear range with IntelliCount™ (seven with IntelliCount™ versus four by legacy counting in the example shown), which reduced the regression error and increased the precision of the extrapolated frequency (see Note 16). These data also illustrate the robustness of IntelliCount™ to discern individual secretory footprints even as background levels rise with increasing ASC numbers per well. 1.4 Further Advantages of AIBased SFU Analysis

Unlike any other spot-counting algorithm presently available on the market, IntelliCount™ does not require special knowledge for setting counting parameters; thus, data analysis becomes easy, objective, user-independent, and fully automated (see Note 17). Moreover, AI-based counting, being more forgiving with fluctuations of image intensity, background staining, and spot contrast over the background, will reduce interassay variability of test results when aliquots of the same PBMC are retested in the same or different laboratories (see Note 18). IntelliCount™, due to the way the deep neural networks are trained, is also rather insensitive to variations in well-image properties resulting from image pixel

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Fig. 6 Representative PBMC serial dilution for pan-IgG detecting B cell ImmunoSpot® assay evaluated via (a) parametric counting algorithm versus (b) IntelliCount™. For the raw data analyzed, the legend to Fig. 2 applies. SFU counts were established by ImmunoSpot® Studio software automatically using the specified counting module. Four replicate wells were used, +/-SD for each dilution is shown (except for the highest cell number per well, they were the same size or smaller than symbols size). The linear regression line (in blue) in both cases was automatically calculated by the Linear Range Finder Function integrated into the ImmunoSpot® Studio software. The number of data points fitting the linear range were in this case 4 for parametric counting versus 7 for IntelliCount™, respectively. The corresponding frequencies of ASC extrapolated were 8.5 versus 8.0% of PBMC, respectively. While both numbers are similar (in this case four data points were in the linear range for the parametric count), the precision of frequency calculations is higher for IntelliCount™ with more points in the linear portion of the titration graph. The standard error of regression, calculated as a square root of sum of quadratic errors divided by the number of points in the linear range, were 11.7 and 8.2 for the parametric count and IntelliCount, respectively

resolution or varying image acquisition parameters with the same or different reader(s). IntelliCount™ practically eliminates the necessity for harmonization when several instruments are operating in the same laboratory, or in multicenter studies. SFU counts reported by different independent groups should become more comparable using IntelliCount™, representing a major step toward count harmonization [33]. A further advantage of IntelliCount™ is its faster speed of data processing compared to legacy counting, due to the optimized utilization of modern GPU and TPU accelerator cards. Typically, the counting time for a 96-well plate for a single fluorescence channel does not exceed 1 min with IntelliCount™ versus 3–4 min with legacy counting. Such time-saving particularly benefits multicolor/channel analysis. With this technology, ImmunoSpot® assays, in particular their multiplexed HCA-inclusive B cell analysis variants, will become truly high-throughput suitable methods that can serve the advanced needs of both immunemonitoring efforts in regulated environments and academic research laboratories.

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Materials

2.1 Single-Color ELISPOT Assay

1. Commercially available, single-color Human Ig class (IgA, IgE, IgG, or IgM) or subclass (IgA1, IgA2, IgG1, IgG2, IgG3 or IgG4) ELISPOT kit. 2. 190 proof (95%) EtOH. 3. Cell culture-grade water. 4. Ninety-six-well, round-bottom dilution plate. 5. 0.05% Tween-PBS wash solution. 6. 0.1 μm low-protein binding syringe filter. 7. Plate washer. 8. ImmunoSpot® S6 Ultimate 4 LED Analyzer, or suitable instrument equipped with the appropriate detection channels, running CTL’s ImmunoSpot® Studio Software Suite.

2.2 Single-, Three-, or Four-Color FluoroSpot Assays

1. Commercially available single-color Human Ig class (IgA, IgE, IgG, or IgM) or subclass (IgA1, IgA2, IgG1, IgG2, IgG3, or IgG4) FluoroSpot kit. 2. Commercially available, three-color Human Ig class (IgA, IgG, and IgM) FluoroSpot kit. 3. Commercially available, four-color Human Ig class (IgA, IgE, IgG, and IgM) FluoroSpot kit. 4. Vacuum manifold.

2.3 Four-Color Antigen-Specific Direct FluoroSpot Assay (Affinity Capture Coating)

1. Commercially available, four-color Human Ig class (IgA, IgE, IgG, and IgM) affinity capture (His) FluoroSpot kit.

2.4 Single-Color, Antigen-Specific Inverted ImmunoSpot® Assay

1. Commercially available, single-color inverted (His) human B cell ImmunoSpot® kit.

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2. Commercially available, four-color Human IgG subclass affinity capture (His) FluoroSpot kit. 3. His-tagged recombinant protein.

Methods

3.1 Pan-Ig ImmunoSpot® Assay (total ASC, Irrespective of Specificity)

1. One day before plating cells (Day 1), prepare 70% EtOH and pan anti-Ig capture antibody solutions. 2. Remove underdrain and pipet 15 μL of 70% EtOH solution into the center of each well (or designated wells) of the assay plate. Immediately after the addition of the 70% EtOH

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solution to the entire plate (or designated wells), add 180 μL/ well of PBS (see Note 19). Decant and wash wells again with 180 μL/well of PBS. 3. Decant the assay plate, replace underdrain, and immediately add 80 μL/well of the pan anti-Ig capture antibody solution into each well (or designated wells) of the low autofluorescence PVDF-membrane plate provided with the kit. 4. Incubate the plate overnight at 4 °C in a humidified chamber. 5. On the day of the assay (Day 0), decant the assay plate and wash wells with 180 μL/well of PBS. Next, decant the plate and add 150 μL/well of prewarmed BCM to block the plate (≥1 h at RT). 6. If using PBMC following polyclonal activation in vitro, collect the cell suspension(s) and transfer into labeled conical tube(s). Keep the cells warm during processing. Wash culture vessel’s interior with sterile warm PBS to collect residual PBMC and transfer into the corresponding conical tube(s). Increase volume to fill the tube with additional warm PBS and then centrifuge balanced tubes at 330 × g for 10 min nonrefrigerated, centrifuge with brake on (see Notes 20 and 21). 7. Decant supernatant and resuspend the cell pellet(s) using prewarmed BCM to achieve a cell density of ~2–5 × 106 cells/mL (the cell number recovered at this point can be estimated to be 50% of the number of cells frozen). 8. Pipet 15 μL of live/dead cell counting dye onto a piece of parafilm to form a droplet. 9. Remove 15 μL of cell suspension and combine with droplet of live/dead cell counting dye. Pipet up and down 3–5 times to mix the sample while avoiding the formation of bubbles. 10. Transfer 15 μL of the cell and dye suspension into each chamber of a hemocytometer. 11. Determine live cell count and viability using CTL’s Live/Dead Cell Counting™ suite. 12. Increase volume of cell suspension(s) with additional sterile warm PBS and centrifuge balanced tubes at 330 × g for 10 min with centrifuge brake on, unrefrigerated (see Notes 22 and 23). 13. Decant supernatant and resuspend the cell pellet(s) using prewarmed BCM at 2 × 105 PBMC/mL. 14. Decant the BCM used for blocking the ImmunoSpot® assay plate and replace with 100 μL/well of prewarmed BCM. 15. Prepare PBMC serial dilution series in a round-bottom 96-well polystyrene plate. For this, we recommend the following

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procedure: into the round-bottom 96-well dilution plate add 120 μL of prewarmed BCM into all wells, except for Row A. Into Row A, add 240 μL of diluted single-cell suspension at 2 × 105 PBMC/mL in one or more replicates (see Note 24). Using a multichannel pipettor, perform a two-fold dilution series of the PBMC test sample(s) by transferring 120 μL from each row to the next, diluting the cells by gently aspirating and ejecting twice at each dilution step. Once the cell dilution in the round-bottom dilution plate is completed, using a multichannel pipettor and fresh tips, transfer 100 μL of the serially diluted cells from the dilution plate into the actual ImmunoSpot® test plate. 16. Incubate cells in the ImmunoSpot assay plate for 16–18 h at 37 °C, 5% CO2. 17. After completion of the assay incubation period, decant (or reutilize) cells and wash plate two times with warm PBS (200 μL/well), followed by two additional washing steps with 0.05% Tween-PBS wash solution (see Note 25). 18. Prepare anti-Ig class/subclass-specific detection antibody solution(s) according to kit protocol and pass through 0.1 μm low-protein binding syringe filter to remove any protein aggregates. 19. Decant 0.05% Tween-PBS wash solution, add 80 μL/well of the anti-Ig class/subclass-specific detection antibody solution into designated wells, and incubate for 2 h at RT (protected from light). 20. Wash plate(s) two times with 0.05% Tween-PBS wash solution. 21. Prepare tertiary solution by following kit protocol and pass through 0.1 μm low-protein binding syringe filter to remove any aggregates. 22. Decant 0.05% Tween-PBS wash solution, add 80 μL/well of tertiary solution into designated wells, and incubate for 1 h at RT (protected from light). 23. Wash plates(s) twice with distilled water. 24. Remove protective underdrain and place plate face down on vacuum manifold. Completely fill the backside of the plate with distilled water and apply vacuum to draw water through the membrane (“back to front”) (see Note 26). 25. Allow plate to dry completely, protected from light (see Note 27). 26. Scan and count plate(s) with suitable analyzer equipped with the appropriate detection channels.

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1. One day before plating cells (Day 1), prepare 70% EtOH and antigen coating solutions (see Notes 28 and 29). 2. Remove underdrain and pipet 15 μL of 70% EtOH solution into the center of each well (or designated wells) of the assay plate. Immediately after the addition of the 70% EtOH solution to the entire plate (or designated wells), add 180 μL/well of PBS (see Note 19). Decant and wash wells again with 180 μL/well of PBS. 3. Decant the assay plate, replace underdrain, and immediately add 80 μL/well of the antigen coating solution into each well (or designated wells) of the low autofluorescence PVDFmembrane plate provided with the kit. 4. Incubate the plate overnight at 4 °C in a humidified chamber. 5. On the day of the assay (Day 0), decant the assay plate and wash wells with 180 μL/well of warm PBS. Next, decant the plate and add 150 μL/well of pre-warmed BCM to block the plate (≥1 h at RT). 6. If using PBMC following polyclonal activation in vitro, collect the cell suspension(s) and transfer into labeled conical tube(s). Keep the cells warm during processing. Wash culture vessel’s interior with sterile warm PBS to collect residual PBMC and transfer into the corresponding conical tube(s). Increase volume to fill the tube with additional warm PBS and then centrifuge balanced tubes at 330 × g for 10 min non-refrigerated, centrifuge with brake on (see Notes 21 and 22). 7. Decant supernatant and resuspend the cell pellet(s) using pre-warmed BCM to achieve a cell density of ~2–5 × 106 cells/mL (the cell number recovered at this point can be estimated to be 50% of the number of cells frozen). 8. Pipet 15 μL of live/dead cell counting dye onto a piece of parafilm to form a droplet. 9. Remove 15 μL of cell suspension and combine with droplet of live/dead cell counting dye. Pipet up and down 3–5 times to mix the sample while avoiding the formation of bubbles. 10. Transfer 15 μL of the cell and dye suspension into each chamber of a hemocytometer. 11. Determine live cell count and viability using CTL’s Live/Dead Cell Counting™ suite. 12. Increase volume of cell suspension(s) with additional sterile warm PBS and centrifuge balanced tubes at 330 × g for 10 min with centrifuge brake on, unrefrigerated. 13. Decant supernatant and resuspend the cell pellet(s) using pre-warmed BCM at 2–5 × 106 PBMC/mL (see Note 30). 14. Decant the BCM used for blocking the ImmunoSpot® assay plate and replace with 100 μL/well of pre-warmed BCM.

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15. Prepare PBMC serial dilution series in a round-bottom 96-well polystyrene plate. For this, we recommend the following procedure. Into the round-bottom 96-well dilution plate add 120 μL of pre-warmed BCM into all wells, except for Row A. Into Row A, add 240 μL of diluted single-cell suspension at 2–5 × 106 PBMC/mL in one or more replicates (see Note 24). Using a multichannel pipettor, perform a two-fold dilution series of the PBMC test sample(s) by transferring 120 μL from each row to the next, diluting the cells by gently aspirating and ejecting twice at each dilution step. Once the cell dilution in the round-bottom dilution plate is completed, using a multichannel pipettor and fresh tips, transfer 100 μL of the serially diluted cells from the dilution plate into the actual ImmunoSpot® test plate. 16. Incubate cells in the ImmunoSpot® assay plate for 16–18 h at 37 °C, 5% CO2. 17. After completion of the assay incubation period, decant (or reutilize) cells and wash plate two times with warm PBS (200 μL/well), followed by two additional washing steps with 0.05% Tween-PBS wash solution (see Note 25). 18. Prepare anti-Ig class/subclass-specific detection antibody solution(s) according to kit protocol and pass through 0.1 μm low-protein binding syringe filter to remove any protein aggregates. 19. Decant 0.05% Tween-PBS wash solution, add 80 μL/well of the anti-Ig class/subclass-specific detection antibody solution into designated wells, and incubate for 2 h at RT (protected from light). 20. Wash plate(s) two times with 0.05% Tween-PBS wash solution. 21. Prepare tertiary solution by following kit protocol and pass through 0.1 μm low-protein binding syringe filter to remove any aggregates. 22. Decant 0.05% Tween-PBS wash solution, add 80 μL/well of tertiary solution into designated wells, and incubate for 1 h at RT (protected from light). 23. Wash plates(s) twice with distilled water. 24. Remove protective underdrain and place plate face down on vacuum manifold. Completely fill the backside of the plate with distilled water and apply vacuum to draw water through the membrane (“back to front”) (see Note 26). 25. Allow plate to dry completely, protected from light (see Note 27). 26. Scan and count plate(s) with suitable analyzer equipped with the appropriate detection channels.

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1. One day before plating cells (Day 1), prepare 70% EtOH and anti-human IgG capture antibody solutions. 2. Remove underdrain and pipet 15 μL of 70% EtOH solution into the center of each well (or designated wells) of the assay plate. Immediately after the addition of the 70% EtOH solution to the entire plate (or designated wells), add 180 μL/well of PBS (see Note 19). Decant and wash wells again with 180 μL/well of PBS. 3. Decant the assay plate, replace underdrain, and immediately add 80 μL/well of the anti-human IgG capture antibody solution into each well (or designated wells) of the low autofluorescence PVDF-membrane plate provided with the kit. 4. Incubate the plate overnight at 4 °C in a humidified chamber. 5. On the day of the assay (Day 0), decant the assay plate and wash wells with 180 μL/well of warm PBS. Next, decant the plate and add 150 μL/well of prewarmed BCM to block the plate (≥1 h at RT). 6. If using PBMC following polyclonal activation in vitro, collect the cell suspension(s) and transfer into labeled conical tube(s). Keep the cells warm during processing. Wash culture vessel’s interior with sterile warm PBS to collect residual PBMC and transfer into the corresponding conical tube(s). Increase volume to fill the tube with additional warm PBS and then centrifuge balanced tubes at 330 × g for 10 min nonrefrigerated, centrifuge with brake on (see Notes 21 and 22). 7. Decant supernatant and resuspend the cell pellet(s) using prewarmed BCM to achieve a cell density of ~2–5 × 106 cells/mL (the cell number recovered at this point can be estimated to be 50% of the number of cells frozen). 8. Pipet 15 μL of live/dead cell counting dye onto a piece of parafilm to form a droplet. 9. Remove 15 μL of cell suspension and combine with droplet of live/dead cell counting dye. Pipet up and down 3–5 times to mix the sample while avoiding the formation of bubbles. 10. Transfer 15 μL of the cell and dye suspension into each chamber of a hemocytometer. 11. Determine live cell count and viability using CTL’s Live/Dead Cell Counting™ suite. 12. Increase volume of cell suspension(s) with additional sterile warm PBS and centrifuge-balanced tubes at 330 × g for 10 min with centrifuge brake on, unrefrigerated. 13. Decant supernatant and resuspend the cell pellet(s) using prewarmed BCM at 1 × 106 PBMC/mL (see Notes 31, 34, and 35).

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14. Decant the BCM used for blocking the ImmunoSpot® assay plate and replace with 100 μL/well of prewarmed BCM. 15. Prepare PBMC serial dilution series in a round-bottom 96-well polystyrene plate. For this, we recommend the following procedure. Into the round-bottom 96-well dilution plate add 120 μL of prewarmed BCM into all wells, except for Row A. Into Row A, add 240 μL of diluted single-cell suspension at 1 × 106 PBMC/mL in one or more replicates (see Note 24). Using a multichannel pipettor, perform a two-fold dilution series of the PBMC test sample(s) by transferring 120 μL from each row to the next, diluting the cells by gently aspirating and ejecting twice at each dilution step. Once the cell dilution in the round-bottom dilution plate is completed, using a multichannel pipettor and fresh tips, transfer 100 μL of the serially diluted cells from the dilution plate into the actual ImmunoSpot® test plate. 16. Incubate cells in the assay plate for 16–18 h at 37 °C, 5% CO2 (see Note 36). 17. After completion of the assay incubation period, decant (or reutilize) cells and wash plate two times with warm PBS (200 μL/well), followed by two additional washing steps with 0.05% Tween-PBS wash solution. 18. Prepare His-tagged antigen probe solution (see Notes 34 and 37) according to kit protocol and pass through 0.1 μm low-protein binding syringe filter to remove any protein aggregates. 19. Decant 0.05% Tween-PBS wash solution, add 80 μL/well of His-tagged antigen probe solution and incubate for 2 h at RT (protected from light). 20. Wash plate(s) two times with 0.05% Tween-PBS wash solution. 21. Prepare Anti-His detection antibody solution according to kit protocol and pass through 0.1 μm low-protein binding syringe filter to remove any aggregates. 22. Decant 0.05% Tween-PBS wash solution, add 80 μL/well of Anti-His detection antibody solution into designated wells, and incubate for 1 h at RT (protected from light). 23. Wash plate(s) two times with 0.05% Tween-PBS wash solution. 24. Prepare tertiary solution according to kit protocol and pass through 0.1 μm low-protein binding syringe filter to remove any aggregates. 25. Wash plates(s) twice with distilled water. 26. Remove protective underdrain and place plate face down on vacuum manifold. Completely fill the backside of the plate with distilled water and apply vacuum to draw water through the membrane (“back to front”) (see Note 26).

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27. Allow plate to dry completely, protected from light (see Note 27). 28. Scan and count plate(s) with suitable analyzer equipped with the appropriate detection channel. 3.4 Automatic Scanning and Counting of ImmunoSpot® Plates

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1. ImmunoSpot® plates were scanned on CTL Series 6 Ultimate Analyzer equipped with the appropriate fluorescent detection channels. 2. SFUs were counted using ImmunoSpot® Studio Software with integrated IntelliCount™ mode and Linear Range Finder for accurate frequency calculations.

Notes 1. In B-cell ImmunoSpot® assays, there is no inherent lower limit of detection. The PBMC numbers plated per well into a 96-well plate should not exceed 1 × 106 cells per well, because with higher numbers, the cells no longer form a monolayer on the membrane [34] and the resulting cell layering can interfere with the capture of ASC-derived antibodies. If, e.g., ten million PBMC are plated at 1 × 106 PBMC across 10 replicate wells, one cell in ten million is the detection limit, etc. Owing to increased Poisson noise occurring at such low frequencies, however, the number of replicate wells needs to be increased accordingly to obtain accurate measurements [35]. As shown in Fig. 3 of the chapter by Lehmann et al. in this volume [12], antigen-specific memory B cells (Bmem) quite frequently occur in low frequencies. 2. In ELISPOT assays, the enzymatic amplification of the signal leads to loss of direct proportionality between the amount of labeled detection antibody bound and the eventual substrate precipitate color density. Once the density of the substrate precipitate deposition on the membrane reaches a certain point, the spot’s optical density/color intensity does not increase even if more substrate is converted and more precipitate is deposited (much like applying many layers of non-trasparent paint). With fluorescent detection, however, the number of fluorescent tags bound is proportional to the number of detection antibodies retained on the membrane. 3. As enzyme-linked ImmunoSpot (ELISPOT) and FluoroSpot assays differ only in the modality of detecting secretory footprints of cells on membranes, we collectively refer to both as ImmunoSpot® assays. In the former, the detection antibody is tagged to enable the engagement of an enzymatic reaction that results in the local precipitation of a converted substrate that is

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visible under white light. In the latter, the plate-bound detection antibodies are visualized via fluorescent tags using appropriate excitation and emission wavelengths. Data provided in a chapter in this volume by Yao et al. [22] establish that ELISPOT and FluoroSpot assays have equal sensitivity for detecting numbers of antibody-secreting cell (ASC)-derived secretory footprints. However, they are not equally suited for high-content analysis (HCA) of spot morphologies (see Note 2). 4. Our introduction of the PVDF membrane to ELISPOT assays [9, 10], with its by far superior adsorption capacity for capture antibodies [36], was key for improving our ability to detect secretory footprints to the point needed for transforming ImmunoSpot® into the robust immune monitoring platform it has become for detecting rare -- even extremely rare -antigen-specific lymphocytes ex vivo, in freshly isolated PBMC or other lymphoid cell material. We refer to Fig. 1 in [37] to appreciate the difference in assay performance using the PVDF membrane versus the previously used mixed cellulose ester membrane. 5. The 3D shape of secretory footprints (spot morphologies) produced by T cells follows defined rules since the capture antibody’s (i.e., an anti-cytokine-specific mAb) affinity for the analyte to be detected is high and fixed. Consequently, only the quantity of analyte (cytokine) produced by the T cell will define the morphology of the resulting secretory footprint [25]. Predictable (log-normal [32]) spot sizes permit objective automated size gating [13, 14]. 6. Although only B cells can secrete antibodies, even in B-cell ImmunoSpot® assays, there can be small background spots resulting primarily from aggregated detection reagents. Such artifacts can be reduced/eliminated by filtering or centrifuging at high speed the reagents to eliminate aggregates. To identify such spots, it is important also to include negative control wells that are subject to the entire test procedure, but do not contain cells. Background spots should and can be readily gated out during ImmunoSpot® analysis. 7. We refer to Fig. 3 of the chapter by Lehmann et al. in this volume [12] to convey the high degree of variability in frequency of antigen-specific Bmem in PBMC. 8. ASC secrete Ig in an undirected fashion into 3D space above the membrane. In ImmunoSpot® assays, the antibody released/diffusing toward the membrane will be captured as a secretory footprint while the remainder of the secreted antibody will diffuse away from the surface and will be diluted in the bulk of the culture supernatant. As the concentration of

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such free bulk diluted antibodies increases in the culture medium, they are captured on the membrane distantly from the source ASC, increasing the background signal in the assay and undermining the resolution of individual secretory footprints. Such an elevated background in an ImmunoSpot® assay is termed an ELISA effect. If ASC—by chance—settle in clusters on the membrane, local ELISA effects can occur surrounding these cells resulting in regions with increased local background. The ImmunoSpot® software implements powerful local background correction, and, therefore, such ELISA effects do not interfere with the detection of SFU; however, they affect threshold-based detection of spot outlines needed for HCA. 9. Fine-tuning of parameters manually not only requires expert knowledge but also takes considerable time, and thus it can rarely be done for analyzing an entire assay. Due to global and local ELISA effects in wells, the background level is variable in most assays preventing the accurate detection of outlines of secretory footprints. When using parametric counting for the initial machine reading of the plate, under such conditions, well-by-well recounting in quality control mode may be required for finalizing the results. IntelliCount™ greatly streamlines this process. 10. In FluoroSpot assays, the overall fluorescence intensity of a spot (“spot mass”) is proportional to the quantity of analyte captured within the secretory footprint, i.e., “total spot intensity” of a given ASC (where “total spot intensity” is equal to “mean spot intensity” multiplied by spot size). In antigenspecific direct assays, spot morphologies can include all possible variations of sizes and intensities (see Fig. 2). A multitude of morphological parameters is readily captured for each SFU and stored in FCS format, to perform in-depth HCA. 11. The so-called “Goldilocks” number is defined as the maximal number of cells that can be plated in a B-cell ImmunoSpot® assay well while still being able to discern clearly secretory footprint boundaries derived from individual antigen-specific ASC. For HCA, i.e., for the accurate definition of secretory footprint boundaries, the Goldilocks number is lower than the breaking point for linearity in mere SFU counts. As it is assaydependent, it needs to be experimentally established by serial dilution of PBMC in the respective assay, but ~50 SFU/well is a safe estimate. 12. Frequently, the background membrane staining of individual wells is not perfect even in ImmunoSpot® assays and that can interfere with accurate SFU detection, in particular when relying on fixed counting parameters. Lowering nonspecific

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background staining, and reduction of “hot spots” in the center of the assay wells can be achieved through performing the “back to front” water filtration technique. Regarding regional and global ELISA effects, see Note 8. 13. Antibodies occur in four classes (IgM, IgG, IgA, and IgE), and in four subclasses (IgG1, IgG2, IgG3, and IgG4). ASC producing different classes or subclasses can be detected simultaneously in multiplexed ImmunoSpot® assays using only 4 × 105 PBMC/antigen (see chapter by Yao et al. in this volume, [22]). The different Ig classes and subclasses are endowed with distinct effector functions and each contributes nonredundant roles toward maintaining host defense (reviewed in [38]). Stimulating optimal Ig class usage during an infection or following vaccination is vital to successful host defense and the avoidance of collateral immune-mediated pathology (reviewed in [39]). 14. During the primary immune response, B cells can transition from IgM-expressing naive B cells into effector cells (antibodysecreting plasma cells) and resting Bmem that have undergone class switch recombination (CSR). CSR is an irreversible process that involves the excision of DNA encompassing the exons of the Igμ heavy chain required for expression of IgM and the juxtaposition of upstream variable region genes with downstream exons encoding alternative Ig classes or IgG subclasses [40]. Class switching of the BCR to downstream Ig classes or IgG subclasses is an instructed process and can be influenced by the cytokine milieu and co-stimulation provided by CD4+ T helper cells. Thus far, we have not seen evidence for in vivo class-switched Bmem to undergo further during short-term polyclonal stimulation in vitro using R848 plus IL-2, as needed for their detection in ImmunoSpot® assays. Thus, it has to be assumed that the Ig calls subclass utilization of ASC observed in ImmunoSpot® assays ex vivo reflects on the corresponding Bmem commitment for Ig class/subclass utilization upon antigen reencounter in vivo. 15. Once activated by polyclonal stimulation, ASC are autonomous. Thus, the well-to-well variation in numbers of ASC in B cell ImmunoSpot® assays is dependent on their concentration in the test sample following the rules of a Poisson distribution: the rarer the cells, the higher the well-to-well variation when an equal set volume is sampled/plated. This knowledge permits to precisely calculate the number of replicate wells needed to establish frequencies with required precision when ASC frequencies are low [35]. 16. For low-frequency antigen-specific B-cell assay results, the conventional parametric approach can establish SFU counts (but to a lesser extent HCA-pertaining parameters) with a

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similar accuracy as IntelliCount™; however, it requires expertise to set up parameters, whereas IntelliCount™ does it automatically. 17. Subjective counting is a considerable challenge for count harmonization among individuals and laboratories [33]. 18. Even slight changes in assay conditions (e.g., incubation times and temperature) as well in reagent properties over time (e.g., storage-dependent aggregation or decay) can have an effect on the SFU staining intensity seen in repeat ImmunoSpot® assays. By being less sensitive to such qualitative differences, IntelliCount™ helps the assay’s robustness in the evaluation phase. 19. Activation of the PVDF membrane with 70% EtOH is instantaneous and can be seen visually as a graying of the membrane. It is important to be sure that the EtOH solution has spread across the entire membrane before adding the first wash of PBS. If needed, tapping the plate can promote contact of the EtOH solution with the PVDF membrane. We recommend only prewetting one plate at a time with 70% EtOH to ensure that the contact time is ≤1 min; longer contact times may promote leaking of the membrane and result in suboptimal assay performance. 20. We refer to the chapter of Yao et al. in this volume [22] for detailed procedures covering the isolation of peripheral blood mononuclear cells (PBMCs), their cryopreservation and thawing, as well as the polyclonal in vitro stimulation culture needed to trigger antibody production by resting memory B cells. 21. PBMC, or other primary cell material, collected acutely following known antigen encounter, which may contain spontaneous (in vivo differentiated) ASC, can also be evaluated in such assays. 22. If the cells are not washed thoroughly, antibodies in the cell suspension(s) can compete with the binding of ASC-derived Ig in the assay, resulting in elevated membrane staining that can interfere with the accurate detection of individual ASC’s secretory footprints. 23. Using a serial dilution approach, an ideal starting cell input of 2 × 104 is appropriate for typical pan (total) IgA/IgG/IgM measurements following in vitro differentiation of PBMC. However, higher cell inputs may be more appropriate for measurements of spontaneous (in vivo differentiated) ASC. 24. Serial dilutions involving single wells for each cell dilution, progressing in a 1 + 1 (two-fold) dilution series, is a valid option for establishing accurate SFU frequencies and greatly reduces the cell numbers and reagents required (see the chapter

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by Yao et al. in this volume) [22]. In Fig. 5a of that chapter, the recommended plate layout for such a serial dilution assay is shown. 25. For automated washing, the pin height and flow rate should be customized to avoid damaging the assay membranes, the CTL 405LSR plate washer supports such adjustments. Plate washes may also be performed manually. See also Note 22. 26. Optimal removal of background staining, fibers, and other debris, along with reduction of “hot spots” in the center of the assay wells, is achieved through performing the “back to front” water filtration technique. 27. To completely dry plates, blot assay plate(s) on paper towels to remove residual water before either placing them in a running laminar flow hood at a 45° angle for >20 min or placing face down on paper towels for >2 h in a dark drawer/cabinet. Do not dry assay plates at temperatures exceeding 37 °C as this may cause the membrane to warp or crack. Fluorescent spots may not be readily visible while the membrane is still wet and the background fluorescence may be elevated. Scan and count plates only after membranes have dried completely. 28. Direct application of an antigen to the PVDF membrane can result in variable and often low-efficiency coating owing to weak, nonspecific binding (primarily via hydrophobic interaction). Alternatively, our recent introduction of affinity capture coating [11] enables specific and high-affinity binding of antigen to the assay membrane. 29. Optimizing the concentration of His-tagged protein(s) used for affinity capture coating is recommended. A concentration of 10 μg/mL His-tagged protein has yielded well-formed secretory footprints for most antigens, but increased concentrations of the anti-His affinity capture antibody and/or His-tagged protein may be required to achieve optimal assay performance. 30. Using a serial dilution approach, a starting cell input of 2–5 × 105 is appropriate for typical antigen-specific ImmunoSpot® tests following in vitro differentiation of PBMC. However, higher cell inputs may be more appropriate for measurements of spontaneous (in vivo differentiated) ASC. 31. Owing to polyclonal stimulation of Bmem to trigger their terminal differentiation, a large majority of IgG+ ASC will not be antigen-specific yet will compete for “real estate” on the lawn of anti-IgG capture reagent used for coating. Consequently, inverted assays aimed at studying lower frequency ASC specificities are directly limited by the maximal number of total IgG+

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ASC that can be input into a single well while still maintaining the ability to resolve individual antigen-specific secretory footprints. 32. Prior to performing an inverted ImmunoSpot® assay using limiting quantities of antigen detection probe, it is recommended to first determine the Goldilocks cell input to achieve ~50 SFU/well using an aliquot of cryopreserved cell material. 33. In instances when the frequency of antigen-specific ASC is low among all ASC, we recommend increasing the number of replicate wells and seeding at lower cell inputs. Moreover, to conserve on cell material required, increasing the fold dilution of the antigen probe and/or testing only at predetermined concentrations are both valid options. 34. If the Goldilocks cell number input is already known, and the intent of the assay is to assess the affinity spectrum of the antigen-specific ASC compartment, the relevant assay procedures are described in detail in the chapter by Becza et al. [8]. 35. Using a serial dilution approach, a starting cell input of 1 × 105 is appropriate for antigen-specific, inverted ImmunoSpot® tests following in vitro differentiation of PBMC. However, higher cell inputs may be more appropriate for measurements of spontaneous (in vivo differentiated) ASC. 36. Shorter B-cell ImmunoSpot® assay incubation times are suggested if using an enzymatic-based detection approach to avoid merging of spots and/or elevated membrane background staining. 37. The optimal concentration of affinity (His)-tagged antigen probe used for detection of all antigen-specific secretory footprints (i.e., SFU), low- or high-affinity alike, should be determined empirically.

Acknowledgments We wish to thank the R&D and the Software Development teams at CTL for their continued support and technological innovation that made our B-cell ImmunoSpot® endeavor possible. We also thank Dr. Graham Pawelec and Diana Roen for carefully proofreading the manuscript and providing constructive feedback. Lastly, we thank Gregory Kovacs for his support in generation of graphic illustrations. All efforts were funded from CTL’s research budget. Conflicts of Interest P.V.L. is Founder, President, and CEO of CTL, a company that specializes in immune monitoring by ImmunoSpot®. A.Y.K, M.K., Z.M, and G.A.K are employees of CTL.

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References 1. Lehmann PV, Zhang W (2012) Unique strengths of ELISPOT for T cell diagnostics. Methods Mol Biol 792:3–23 2. Bucheli OTM, Sigvaldadottir I, Eyer K (2021) Measuring single-cell protein secretion in immunology: Technologies, advances, and applications. Eur J Immunol 51(6): 1334–1347 3. Lehmann PV, Suwansaard M, Zhang T et al (2019) Comprehensive evaluation of the expressed CD8+ T cell epitope space using high-throughput epitope mapping. Front Immunol 10(655):1–13 4. Caspell R, Lehmann PV (2018) Detecting all Immunoglobulin classes and subclasses in a multiplex 7 color ImmunoSpot® assay. Methods Mol Biol 1808:85–94 5. Karulin AY, Megyesi Z, Caspell R et al (2018) Multiplexing T- and B-Cell FLUOROSPOT assays: experimental validation of the multicolor ImmunoSpot® software based on center of mass distance algorithm. Methods Mol Biol 1808:95–113 6. Roen DR, Hanson J, Lehmann PV (2018) Multiplex ImmunoSpot® assays for the study of functional B cell subpopulations. Methods Mol Biol 1808:73–83 7. Hadjilaou A, Green AM, Coloma J et al (2015) Single-cell analysis of B cell/antibody crossreactivity using a novel multicolor FluoroSpot assay. J Immunol 195(7):3490–3496 8. Becza N, Liu Z, Chepke J et al (2023) Assessing the affinity spectrum of the antigenspecific B cell repertoire in freshly isolated cell material via ImmunoSpot®. J Methods Mol Biol. ibid 9. Forsthuber T, Yip HC, Lehmann PV (1996) Induction of TH1 and TH2 immunity in neonatal mice. Science 271(5256):1728–1730 10. Lehmann PV, Karulin AY, Trezza RP et al (2002) Methods for measuring T cell cytokines. United States Patent US 6,410,252 B1 11. Koppert S, Wolf C, Becza N et al (2021) Affinity tag coating enables reliable detection of antigen-specific B cells in ImmunoSpot assays. Cell 10(8):1–22 12. Lehmann PV, Becza N, Liu Z et al (2023) Monitoring memory B cells by next generation ImmunoSpot® provides insights into humoral immunity that measurements of circulating antibodies do not reveal. Methods Mol Biol. ibid 13. Megyesi Z, Lehmann PV, Karulin AY (2018, 1808) Multi-color FLUOROSPOT counting

using ImmunoSpot® Fluoro-X Suite. Methods Mol Biol:115–131 14. Zhang W, Lehmann PV (2018) Objective, user-independent ELISPOT data analysis based on scientifically validated principles. Methods Mol Biol 792:155–171 15. Yip HC, Karulin AY, Tary-Lehmann M et al (1999) Adjuvant-guided type-1 and type-2immunity: infectious/noninfectious dichotomy defines the class of response. J Immunol 162(7):3942–3949 16. Czerkinsky CC, Nilsson LA, Nygren H et al (1983) A solid-phase enzyme-linked immunospot (ELISPOT) assay for enumeration of specific antibody-secreting cells. J Immunol Methods 65(1–2):109–121 17. Sedgwick JD, Holt PG (1983) A solid-phase immunoenzymatic technique for the enumeration of specific antibody-secreting cells. J Immunol Methods 57(1–3):301–309 18. Wolf C, Koppert S, Becza N et al (2022) Antibody levels poorly reflect on the frequency of memory B cells generated following SARSCoV-2, seasonal influenza, or EBV Infection. Cell 11(22):1–21 19. Murata T, Sugimoto A, Inagaki T et al (2021) Molecular basis of epstein-barr virus latency establishment and lytic reactivation. Viruses 13(12):1–19 20. Terlutter F, Caspell R, Nowacki TM et al (2018) Direct detection of T- and B-memory lymphocytes by ImmunoSpot® assays reveals HCMV exposure that serum antibodies fail to identify. Cell 7(5):1–18 21. Kuerten S, Pommerschein G, Barth SK et al (2014) Identification of a B cell-dependent subpopulation of multiple sclerosis by measurements of brain-reactive B cells in the blood. Clin Immunol 152(1–2):20–24 22. Yao L, Becza N, Maul-Pavicic A et al (2023) Four color ImmunoSpot® assays performed in serial dilution permit precise frequency measurements of antigen-specific B cells secreting immunoglobulins of all classes and subclasses. Methods Mol Biol. ibid 23. Holt PG, Cameron KJ, Stewart GA et al (1984) Enumeration of human immunoglobulin-secreting cells by the ELISA-plaque method: IgE and IgG isotypes. Clin Immunol Immunopathol 30(1):159–164 24. Franke F, Kirchenbaum GA, Kuerten S et al (2020) IL-21 in conjunction with anti-CD40 and IL-4 constitutes a potent polyclonal B cell stimulator for monitoring antigen-specific memory B cells. Cell 9(2):1–15

AI-based ImmunoSpot analysis 25. Karulin AY, Lehmann PV (2012) How ELISPOT morphology reflects on the productivity and kinetics of cells’ secretory activity. Methods Mol Biol 792:125–143 26. Moulana A, Dupic T, Phillips AM et al (2023) The landscape of antibody binding affinity in SARS-CoV-2 Omicron BA.1 evolution. elife 12:1–18 27. Di Niro R, Lee SJ, Vander Heiden JA et al (2015) Salmonella infection drives promiscuous B cell activation followed by extrafollicular affinity maturation. Immunity 43(1):120–131 28. Liao HX, Lynch R, Zhou T et al (2013) Co-evolution of a broadly neutralizing HIV-1 antibody and founder virus. Nature 496(7446):469–476 29. Wysocki L, Manser T, Gefter ML (1986) Somatic evolution of variable region structures during an immune response. Proc Natl Acad Sci U S A 83(6):1847–1851 30. Karulin AY, Hesse MD, Yip HC et al (2002) Indirect IL-4 pathway in type 1 immunity. J Immunol 168(2):545–553 31. Guerkov RE, Targoni OS, Kreher CR et al (2003) Detection of low-frequency antigenspecific IL-10-producing CD4(+) T cells via ELISPOT in PBMC: cognate vs. nonspecific production of the cytokine. J Immunol Methods 279(1–2):111–121 32. Karulin AY, Karacsony K, Zhang W et al (2015) ELISPOTs produced by CD8 and CD4 cells follow log normal size distribution permitting objective counting. Cell 4(1):56–70

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33. Sundararaman S, Karulin AY, Ansari T et al (2015) High reproducibility of ELISPOT counts from nine different laboratories. Cell 4(1):21–39 34. Hanson J, Sundararaman S, Caspell R et al (2015) ELISPOT assays in 384-well format: up to 30 data points with one million cells. Cell 4(1):71–83 35. Karulin AY, Caspell R, Dittrich M et al (2015) Normal distribution of CD8+ T-Cell-derived ELISPOT counts within replicates justifies the reliance on parametric statistics for identifying positive responses. Cell 4(1):96–111 36. Weiss AJ (2012) Overview of membranes and membrane plates used in research and diagnostic ELISPOT assays. Methods Mol Biol 792: 243–256 37. Lehmann PV, Lehmann AA (2020) Aleatory epitope recognition prevails in human T Cell responses? Crit Rev Immunol 40(3):225–235 38. Lu LL, Suscovich TJ, Fortune SM et al (2018) Beyond binding: antibody effector functions in infectious diseases. Nat Rev Immunol 18(1): 46–61 39. Webb NE, Bernshtein B, Alter G (2021) Tissues: the unexplored frontier of antibody mediated immunity. Curr Opin Virol 47:52–67 40. Stavnezer J, Guikema JE, Schrader CE (2008) Mechanism and regulation of class switch recombination. Annu Rev Immunol 26:261– 292

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 6 Comparing Flow Cytometry and ELISpot for Detection of IL-10, IL-6, and TNF Alpha on Human PBMCs Kristina Boss, Jodi Hagen, Megan Constans, Christine Goetz, and Alexander E. Kalyuzhny Abstract ELISpot and flow cytometry are two methods often utilized side-by-side for detecting secreted and intracellular cytokines, respectively. Each application has its own advantages and challenges. ELISpot is more sensitive compared to ELISA and appears to be more consistent in detecting IL-10 production than flow cytometry. ELISpot can be used for detecting the secretion of multiple cytokines but not from the same cells simultaneously, whereas flow cytometry allows for the concurrent detection of multiple intracellular cytokines by the same cells. Flow cytometry is a convenient technique allowing for the detection of many cytokines at the same time in a population of cells. The restimulation cocktails used for cytokine detection in flow cytometry are hard on cells and lead to decreased cell viability. Using a live dead dye allows for the exclusion of dead cells when analyzing data. We illustrated the differences between ELISpot and flow cytometry by stimulating cells with two toll-like receptor (TLR) agonists, LPS or Pam3CSK4. Both activators increase production of various cytokines, including IL-10, IL-6, and TNF-alpha. The TLR2 antagonist, MMG-11, was used to inhibit this increased cytokine production. We observed some inhibition of IL-6 and IL-10 from Pam3CSK4 stimulation in the presence of MMG-11 by flow cytometry. TNF-α remains largely unchanged as its basal expression is high, but there is some reduction in the presence of MMG-11 for both methods. However, IL-10 was difficult to detect by ELISpot given the low seeding density. Overall, both ELISpot and flow cytometry are good methods for detecting secreted and intracellular cytokines, respectively, and should be used as complimentary assays. Key words Flow cytometry, ELISpot, Peripheral blood mononuclear cells (PBMCs), Monocytes, IL10, IL-6, TNF-alpha

1

Introduction The detection and measurement of cytokine production in situ plays a critical role in the study of immune responses. Current methods for detecting and analyzing cytokines range from singletarget immunoassays such as ELISAs to antibody arrays, which measure cytokines on a large scale. Ultimately, which approach is used to measure cytokine production will be based on specific

Alexander E. Kalyuzhny (ed.), Handbook of ELISPOT: Methods and Protocols, Methods in Molecular Biology, vol. 2768, https://doi.org/10.1007/978-1-0716-3690-9_6, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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requirements for each study. Within the large field of cytokine detection methods, ELISpot and flow cytometry have emerged as powerful tools for visualizing cytokine-secreting cells and measuring intracellular cytokine responses [1–3]. Here we will focus on the advantages and inherent challenges of each method as well as important factors that should be considered when selecting a suitable assay to detect and analyze cytokine production. Flow cytometry can be used to simultaneously detect multiple secreted cytokines at a single-cell level using fluorophoreconjugated antibodies [3, 4]. Secreted cytokines are detected using intracellular staining methods that perforate the cell membrane so that antibodies can bind to their respective targets within the cell. A Golgi stop or Golgi plug, such as brefeldin A or monensin, must be used to prevent the secretion of your cytokines of interest out of the cell. Intracellular staining can be done in conjunction with other flow cytometry surface staining protocols for further immunophenotyping of specific cell subsets as well as functional studies of these cells. Multicolor flow cytometry permits the detection of multiple cytokines within the same experiment for a more comprehensive cell profile. These features allow for better distinguishing and characterization of cells making flow cytometry a versatile method for immune profiling. While flow cytometry requires a greater cell density for staining procedures than ELISpot, it allows for a greater detection of complex cytokine phenotypes. Furthermore, the addition of a viability dye prior to staining ensures that end users are analyzing cytokine production in live cells. Dead cells are not completely eliminated through doublet exclusion gating, and antibody often sticks to dead cells giving a false-positive signal. When analyzing flow cytometric data, it is important to understand both the percentage of positively labeled cells and mean fluorescence intensity (MFI). Knowledge of both statistics will allow you to fully understand the expression pattern of your protein of interest. MFI is a measure of the fluorescence intensity of a population of cells. It allows you to ascertain the relative amount of antibody, on a per-cell basis, that is bound to the marker of interest. Higher MFI values are indicative of larger amounts of antibodies bound to each cell. Alternatively, percent or shift allows for the separation of cells that have antibodies bound to them versus those that do not have antibodies bound. Combining these statistics allows you to understand both how many cells have antibodies bound to the marker of interest and how much antibody is bound on a per-cell basis in that positive population. In order to detect the secreted cytokines of interest, cells must first be stimulated in the presence of an activator such as LPS or Pam3CSK4 prior to staining. Both of these compounds work by activating different toll-like receptors (TLRs). TLRs are initiators of inflammatory responses by first detecting various microbial

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pathogens [5]. LPS is a potent TLR2/TLR4 agonist of monocytes and macrophages responsible for stimulating immune responses through the surface receptor CD14 [6]. This interaction is responsible for the secretion of various cytokines such as IL-6, IL-10, and TNF-alpha. Similar to LPS, monocytes and macrophages can be stimulated using the synthetic lipopeptide Pam3CSK4, an important TLR2/TLR1 agonist responsible for the differentiation of monocytes into M2-like macrophages [7]. Unlike LPS and Pam3CSK4, MMG-11 is a TLR2 antagonist with a preferential blockage of TLR2/TLR1 [8]. Thus, prestimulation of cells with MMG-11 should inhibit the increased expression of IL-6 and TNF-alpha from Pam3CSK4 stimulation. While LPS and Pam3CSK4 are powerful inducers of cytokine production, it is also beneficial to add either brefeldin A or monensin to your culture in order to allow cytokines to accumulate within the cell. When using agents that induce cytokine secretion, it is important to note that cell death may occur during stimulation depending on the length of stimulation and the concentration used in culture. We highly recommend including a dead cell exclusion marker to ensure you are detecting cytokine production in live cells. In our previous studies, we did a side-by-side comparison of IFNγ secretion and production in human PBMCs using ELISpot, ELISA, and flow cytometry and reported that there can be a lack of correlation between different techniques [9]. In our current study, we hypothesized that LPS and Pam3CSK4 will result in increased production of IL-6, IL-10, and TNF-alpha in human PBMCs via TLR2/TLR4 and TLR2/TLR1 activity. Additionally, prestimulation of cells with MMG-11 should inhibit the increased production of IL-6 and TNF-alpha to a greater degree than IL-10 due to its preferential inhibition of TLR2/ TLR1. To help illustrate the differences between ELISpot and flow cytometry, we compared the production of IL-6, IL-10, and TNF-alpha using both detection methods.

2

Materials

2.1 Isolation of Human Peripheral Blood Mononuclear Cells (PBMCs)

1. Sterile Ficoll-Pacque Premium (GE Biosciences, Uppsala, Sweden). 2. Sterile 1× phosphate-buffered saline (PBS) (145 mM NaCl, 2.7 mM KCl, 8.1 mM Na2HPO4, and 1.5 mM KH2PO4), pH 7.3. 3. Sterile flow cytometry human lyse buffer (FC002, Bio-techne, Minneapolis, MN). 4. Sterile HEPES 1× wash buffer.

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5. Sterile 1× column wash buffer (from MagCellect Human CD4+ T cell isolation kit (MAGH102)). 6. Centrifuge capable of spinning of 50 mL culture tubes at 1100–2000 rpm. 7. Hematocytometer (Thermo Fisher Scientific, Rochester, NY) to count lymphocytes under the microscope or flow cytometer such as Quanteon or Novocyte (Agilent Technologies, Santa Clara, CA) with the ability to count cells via the absolute count function. 8. Trypan Blue dye (Gibco BRL, Grand Island, NY) or Sytox™ Blue Dead Cell Stain (Thermo Fisher Scientific, Rochester, NY). 9. Upright microscope equipped with bright-field illumination and phase contrast condenser. 2.2 Stimulation and Restimulation of PBMCs for Flow Cytometry

1. RPMI 1640 medium with 2 mM L-glutamine (1 Liter) supplemented with 1× Glutamax-1 (Gibco, Billings, MT), 1 mM sodium pyruvate (Gibco, Billings, MT), 1× penicillin–streptomycin (Gibco, Billings, MT), 50 μM 2-mercaptoethanol (Sigma-Aldrich, St. Louis, MO), 1× MEM nonessential amino acids (Gibco, Billings, MT), and 5% pooled human AB serum plasma derived heat inactivated (Innovative Research Inc., Novi, MI). 2. 5% CO2 incubator set at 37 °C. 3. Lipopolysaccharide (LPS) (Sigma-Aldrich, St. Louis, MO). 4. Pam3CSK4 (4633) (Tocris Bioscience, Bristol, UK). 5. MMG-11 (6858) (Tocris Bioscience, Bristol, UK). 6. Brefeldin A (1231) (Tocris Bioscience, Bristol, UK). 7. Monensin (5223) (Tocris Bioscience, Bristol, UK). 8. FACs tubes.

2.3 Flow Cytometry Assays

1. FACs tubes. 2. Ninety-six-well v-bottom plate. 3. Human (1-001-A) and mouse (Bio-techne, Minneapolis, MN). 4. Live-or-Dye™ Fremont, CA).

405/545

(2-001-A)

(Biotium

Aqua)

FC

block

(Biotium,

5. Antibodies to hIL-10 A488 (IC9210G), hTNF-alpha FITC (IC210F), hIL-6 A647 (IC206R), and hCD14 A750 (FAB3832S). Isotypes to rabbit IgG A488 (IC1051G), mouse IgG1 FITC (IC002F), and mouse IgG2B A647 (IC0041R) (Bio-techne, Minneapolis, MN).

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6. Fixation and permeabilization of stimulated PBMCs: 1% flow cytometry fixation buffer (FC004; dilute to 1% in 1× PBS), 1× flow cytometry permeabilization/wash buffer I (FC005) (Bio-techne, Minneapolis, MN). 7. 1× PBS. 8. Hanks-buffered saline solution (HBSS) (Gibco, Billings, MT). 9. Agilent Quanteon™ flow cytometer. 10. Flow cytometry staining buffer (RDFII) (FC001) (Bio-techne, Minneapolis, MN). 2.4 Stimulation of PBMCs for ELISpot

1. RPMI 1640 medium with 2 mM L-glutamine (1 Liter) supplemented with 1× Glutamax-1 (Gibco, Billings, MT), 1 mM Sodium Pyruvate (Gibco, Billings, MT), 1× penicillin–streptomycin (Gibco, Billings, MT), 50 μM 2-mercaptoethanol (Sigma-Aldrich, St. Louis, MO), 1× MEM nonessential amino acids (Gibco, Billings, MT), and 5% pooled human AB serum plasma derived heat inactivated (Innovative Research Inc., Novi, MI). 2. Lipopolysaccharide (LPS) (Sigma-Aldrich, St. Louis, MO). 3. Pam3CSK4 (4633) (Tocris Bioscience, Bristol, UK). 4. MMG-11 (6858) (Tocris Bioscience, Bristol, UK). 5. 5% CO2 incubator set at 37 °C.

2.5

ELISpot Assays

1. Commercially available, ELISpot development modules to measure secretion of human IL-6 (SEL206), IL-10 (SEL217B), and TNF-alpha (SEL210) (R&D Systems, Minneapolis, MN). 2. MultiScreen HTS IP Filter Plates, 0.45 μM, white, nonsterile (Millipore Sigma, St. Louis, MO). 3. ELISpot Blue Color Module (SEL002) (R&D Systems, Minneapolis, MN). 4. PBS – 137 mM NaCl, 2.7 mM KCl, 8.1 mM Na2HPO4, 1.5 mM KH2PO4, pH 7.2–7.4, 0.2 μm filtered. 5. Wash buffer – 0.05% Tween®20 in PBS. 6. Blocking buffer – 1% BSA, 5% sucrose in PBS. 7. Reagent diluent – 1% BSA in PBS, pH 7.2–7.4. 8. Hand-held Nunc-Immuno™ 12-plate washer. 9. ELISpot plate reader QuantiHub (http://www.mvspacific. com).

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Methods

3.1 Isolation of PBMCs

1. Isolate PBMCs from human whole blood trima cone (Innovative Blood Resources, St. Paul, MN) using density centrifugation with Ficoll-Pacque Premium (GE Biosciences, Uppsala, Sweden) at room temperature (RT) (21–23 °C; see Note 1). All steps must be carried out in a laminar flow hood using an aseptic technique. Allow the trima cone to empty into a 50 mL centrifuge tube. Rinse the trima cone twice with 10 mL of 1× PBS. Bring the centrifuge tube volume up to 50 mL with 1× PBS. 2. Centrifuge at 1800–2000 rpm (centrifuge must be set at 21–23 °C for this and all subsequent steps; see Note 2) for 10 min at RT. Pipet out the plasma top layer and add 1× PBS to the blood pellet up to the 30 mL mark for each tube and mix. 3. Slowly overlay the diluted blood on top of 20 mL of Ficoll. Centrifuge at 1800–2000 rpm for 40 min at RT with the brake off. 4. Pipet out the top layer and discard. Then transfer the middle buffy layer into a sterile 50 mL centrifuge tube. Avoid pipetting deep into the bottom layer or you will contaminate your cells. Bring up volume to 50 mL with 1× PBS and spin at 1100 rpm for 10 min at RT. 5. Remove supernatant, break the pellet by gently vortexing with cap on and bring volume to 50 mL with 1× PBS. Spin at 1100 rpm for 10 min at RT. These washes remove contaminating neutrophils. 6. Remove supernatant, break the pellet by gently vortexing with cap on, add 10 mL of 1× human lyse (FC002), and incubate for 10 min at RT. After incubation with lyse, fill the tube up to 50 mL with 1× HEPES. 7. Centrifuge at 1500 rpm for 5 min at RT. Remove supernatant, break the pellet, and resuspend cells in 50 mL 1× PBS. 8. Centrifuge at 1500 rpm for 5 min at RT. Remove supernatant, break the pellet, and resuspend cells 5 mL 1× column wash buffer (see Note 3). 9. Make a 1:20 dilution with 1× PBS and Sytox Blue™ as follows; 90 μL 1× PBS, 10 μL cell suspension, and 100 μL 2× Sytox Blue™ and count on the Agilent Quanteon flow cytometer using the absolute count function. The relative purity of the leukocyte populations (lymphocytes and monocytes vs. granulocytes) can be determined at the same time using the forward and side scatter parameters. Avoid using PBMCs with high contamination of granulocytes or platelets. 10. The PBMCs are now ready for downstream applications.

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Table 1 Experiment layout for flow cytometry MMG-11 Antagonist (μM) stim time (antagonist) (h)

LPS (e-coli) (ng/mL) (agonist)

Pam3CSK4 (ng/mL) (agonist)

Agonist stim time (h)

Monensin and brefeldin A (Y/N)

Condition

–

–

–

–

–

Y

1

–

–

100

–

20–24

Y

2

–

–

–

50

20–24

Y

3

100

1

–

–

20–24

Y

4

100

1

100

–

20–24

Y

5

100

1

–

50

20–24

Y

6

3.2 Stimulation and Restimulation of Naive PBMCs for Flow Cytometry

1. Perform stimulations in sterile FACs tubes. Use 12 × 106 PBMCs per condition. Set up your stimulations as laid out in Table 1. Use the same donor for all conditions. 2. Add media and naive PBMCs to FACs tubes so each condition has a final volume of 2 mL. 3. Add 100 μM MMG-11 to the appropriate tubes and incubate with the cells at 37 °C for 1 h. 4. Add 100 ng/mL LPS or 50 ng/mL Pam3CSK4 to appropriate tubes and incubate overnight at 37 °C. 5. In the morning, add 3 μM monensin and 5 μg/mL brefeldin A to each condition for 6 h at 37 °C to facilitate restimulation (Note 4). Total stimulation time (from the addition of agonist to the end of restimulation) should be 20–24 h.

3.3 Flow Cytometric Staining Protocol

1. After restimulation, wash cells with 2 mL 1× PBS twice (see Note 5). 2. All conditions are stained with 1× Biotium Aqua, a fixable dead cell exclusion marker, for 30 min (see Notes 6 and 7). Cells are Fc blocked with human IgG and mouse IgG for the first 10 min of the live/dead incubation (4 μL of human IgG/1 × 106 cells and 2 μL of mouse IgG/1 × 106 cells). After 10 min, all conditions are stained with hCD14 A750 (FAB3832S) for an additional 20 min. These incubations take place at RT in the dark (see Note 8). 3. Wash cells one time with 2 mL 1× RDFII followed by one wash with 2 mL 1x HBSS. All conditions are then fixed with 1% flow cytometry fixation buffer (FC004) for 15 min at RT in the dark (see Note 9).

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Table 2 Flow cytometric staining panels with ideal working concentrations of each marker Isotype (μg/mL)

Antibody (μg/mL)

Panel 1 RbIgG A488 @20

IL-10 A488 @20

CD14 A750 @100

CD14 A750 @100

Biotium Aqua

Biotium Aqua

Panel 2 MsIgG1 FITC @25

TNFα FITC @25

MsIgG2B A647 @20

IL-6 A647 @20

CD14 A750 @100

CD14 A750 @100

Biotium Aqua

Biotium Aqua

4. The cells are washed twice with 2 mL 1× HBSS to remove excess flow cytometry fixation buffer. To permeabilize the cell membrane, the cells are washed with 1× flow cytometry permeabilization/wash buffer I (FC005). Resuspend each condition in 400 μL 1× flow cytometry permeabilization/wash buffer I. Because there are two staining panels, split cells from each condition into four separate wells in a 96-well v-bottom plate (see Note 10). Use one well for each of the following staining conditions (see Table 2): 5. The desired antibodies are added simultaneously (hIL-10 A488 (IC9210G), hTNFalpha FITC (IC210F), hIL-6 A647 (IC206R)), and incubated for 20 min at RT in the dark (see Note 7). 6. The cells are washed once with 2 mL 1× flow cytometry permeabilization/wash buffer I and resuspended in cell staining buffer for flow cytometric analysis. 7. Compensation tubes should also be set up for each of the fluorochromes listed in Table 2 using compensation beads (see Note 11). 3.4 Flow Cytometric Analysis of IL-6, TNFAlpha, and IL-10 on the Agilent Quanteon

1. Set up the gates as follows: FSC/SSC plot to gate on monocytes, Biotium Aqua/SSC to gate on live cells (see Note 12), then FSC-H/FSC-A plot to exclude doublets. Gating on singlets and live monocytes, set up density plots with CD14 on the x-axis to examine IL-6, TNF-alpha, and IL-10 expression on CD14-positive monocytes (Fig. 1) (see Note 13).

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Fig. 1 Gating strategy for flow cytometry analysis of IL-10, IL-6, and TNF-alpha. PBMCs have been stimulated for 20–24 h with 100 ng/mL LPS at 37 °C. They were restimulated for the last 6 hours with 3 μM monensin and 5 μg/mL brefeldin A. Above data are representative of a single donor

2. Run antibody-labeled samples to determine optimal voltages for each parameter if running samples on the BD Fortessa (see Note 14). Then run the compensation beads to optimally compensate each fluorochrome. 3. Analyze data in FlowJo (see Note 15). 3.5

ELISpot Assay

1. Make working solutions of capture antibodies by diluting concentrated lyophilized capture antibody 1:60 with PBS. 2. Add 100 μL of capture antibody working solution into each well and incubate ELISpot plates overnight (18 h) at 4 °C. 3. After finishing the incubation, aspirate capture antibodies from the plates and block the ELISpot plates with blocking buffer (200 μL/well) for 2 h at RT. 4. After blocking, aspirate the blocking buffer and add cell culture media (200 μL/well) to ELISpot plates for a minimum of 30 min to prepare the plates for cells.

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5. Aspirated cell culture media and add unstimulated PBMCs (100 μL/well) into the appropriate wells on the ELISpot plates at cell concentrations of 5 × 104 and 5 × 105 cells/mL (see Note 16). 6. Stimulate PBMCs (cell concentrations of 5 × 104 and 5 × 105) with 100 μM MMG-11 (100 μL/well) added directly to cells in appropriate wells of ELISpot plate and incubate in a CO2 incubator at 37 °C for 1 h. 7. Add culture media stimulated with 100 μM MMG-11 and either 50 ng/mL Pam3CSK4 or 100 ng/mL LPS (100 μL/ well) to wells containing cells previously stimulated with MMG-11. 8. Stimulate other PBMCs (cell concentrations of 5 × 104 and 5 × 105) with either 50 ng/mL Pam3CSK4 or 100 ng/mL LPS (100 μL/well) and add to appropriate wells on the ELISpot plate. Incubate in a CO2 incubator at 37 °C for 20–24 h (see Note 17). 9. After finishing the incubations, aspirate cells from the plates and wash the plate by rinsing wells four times with wash buffer (see Notes 18–20). 10. Make working solutions of detection antibodies by diluting concentrated detection antibody 1:60 in reagent diluent. 11. Add 100 μL of detection antibody working solution into each well and incubate ELISpot plates overnight at 4 °C (see Note 21). 12. Wash plates three times with the wash buffer. 13. Prepare working solution of streptavidin-alkaline phosphatase by mixing the concentrated stock solution 1:60 with reagent diluent. 14. Add 100 μL of streptavidin-alkaline phosphatase working solution into each well and incubate for 2 h at RT. 15. Wash plates three times with wash buffer. 16. Add 100 μL of ready-to-use BCIP/NBT substrate into each well and incubate for 30–60 min at RT in a place protected from direct light. 17. Wash plates three times with distilled water and let them dry completely (see Note 22). 18. Quantify spots using an automated ELISpot reader. 3.6 Flow Cytometry Results

High levels of donor variability came into play with these experiments. The experiment was carried out on multiple donors (n = 3). Expression of all three cytokines varied widely both in their basal expression and how much they responded to the stimulations. Both LPS and Pam3CSK4 stimulations lead to increased IL-6 percent

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Fig. 2 Average fold change of percent positive monocytes for IL-10, IL-6, and TNF-alpha. PBMCs were stimulated with either media alone, 100 ng/mL LPS, Pam3CSK4 (50 ng/mL), MMG-11 (100 μM), LPS (100 ng/ mL) with a prestimulation of MMG-11 (μM), or Pam3CSK4 (50 ng/mL) with a prestimulation of MMG-11 (μM). Figure shows averaged data from three donors. Data is normalized to the media only condition

positive monocytes and MFI (Figs. 2 and 3). Prestimulation with MMG-11 inhibited some of the increased IL-6 expression when looking at the number of positive cells (Fig. 2) and decreased the MFI (Fig. 3). IL-10 expression increased with LPS stimulation and that increased expression was blocked with MMG-11 prestimulation (Fig. 2). This increased expression was limited to an increase in the number of IL-10-positive monocytes (percent positive or shift) (Fig. 2). There does not seem to be an increase in the amount of IL-10 produced per cell (MFI) (Fig. 3). Due to donor variability and the small differences in IL-10 expression with LPS stimulation, other applications that are more sensitive (such as ELISpot or Simple Plex) are better for IL-10 detection. TNF-alpha’s steadystate level is high. Both LPS and Pam3CSK4 stimulations did not alter the number of TNF-alpha-expressing monocytes (Fig. 2). There is a slight increase in TNF-alpha MFI with LPS stimulation, and that slight increase in expression is inhibited with MMG-11 (Fig. 3). Other stimulations that activate or inhibit the P2X7 receptor may be better for inducing changes in TNF-alpha expression in monocytes [10]. 3.7

ELISpot Results

ELISpot assays were carried out with multiple donors (n = 3). Prestimulation with MMG-11 inhibited all the cytokine’s assayed compared to LPS or Pam3CKS4 stimulation alone. IL-6 and TNF-alpha secretion was seen with all three donors even in unstimulated cells but was mostly increased with LPS or Pam3CKS4

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Fig. 3 Average fold change of MFI on monocytes for IL-10, IL-6, and TNF-alpha. PBMCs were stimulated with either media alone, 100 ng/mL LPS, Pam3CSK4 (50 ng/mL), MMG-11 (100 μM), LPS (100 ng/mL) with a prestimulation of MMG-11 (μM), or Pam3CSK4 (50 ng/mL) with a prestimulation of MMG-11 (μM). Figure shows averaged data from three donors. Data is normalized to the media only condition

stimulation except for TNF-alpha in Donor 1. IL-10 secretion in PBMCs with the stimulations used in this experiment was not ideal given the cell concentration used. Other stimulations or a higher cell concentration may be more adequate for inducing IL-10 secretion (Table 3). 3.8 Benefits of Flow Cytometry and ELISpot for Visualization of Secreted Cytokines 3.8.1 Benefits of Flow Cytometry

1. Flow cytometry allows for the visualization of cytokines produced in live cells by utilizing viability dyes and doublet exclusion gates (see Note 6). 2. Lineage markers and FSC/SSC gates can be used to include or exclude certain populations when working with heterogenous populations of cells such as PBMCs. For example, you can gate on monocytes in the FSC/SSC plot to exclude most lymphocytes. CD14 and CD16 can then be used to evaluate differences in protein expression on classical (CD14+ and CD16-), intermediate (CD14+ and CD16+), and nonclassical monocytes (CD14- and CD16+). This method of gating may inadvertently exclude small monocytes or include large lymphocytes. Another option is to gate on both monocytes and lymphocytes in the FSC/SSC plot. Next, gate on HLADR+ cells (to include all monocytes and B cells) and CD20- and CD19- cells to exclude all B cells. At this point, monocytes have been isolated and can be further separated into different classifications for analysis of differential protein expression.

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Table 3 Average spot counts for cells secreting IL-6, IL-10, and TNF-a IL-6 Stimulation Donor

MMG-11 1 0 2 0 3 18

MMG-11 + LPS 111 4 152

MMG-11 + Pam3CSK4 44 8 110

LPS 598 393 886

Pam3CSK4 353 274 877

Unstimulated 7 71 432

MMG-11 + LPS 0 0 0

MMG-11 + Pam3CSK4 0 0 0

LPS 9 0 5

Pam3CSK4 0 0 6

Unstimulated 0 0 5

MMG-11 + LPS 261 45 491

MMG-11 + Pam3CSK4 97 17 404

LPS 789 553 1059

Pam3CSK4 563 451 1006

Unstimulated 767 94 440

IL-10 Stimulation Donor

MMG-11 1 0 2 0 3 0

TNF-a Stimulation Donor

MMG-11 1 0 2 0 3 88

Cells were plated at 50,000 per well for IL-6 and TNF-alpha assays. For IL-10 assays, cells were plated at 500,000 per well. Cells were stimulated with either 100 μM MMG-11 and incubated overnight, 100 μM MMG-11 and incubated for 1 hour then restimulated with 100 μM MMG-11 and 100 ng/mL LPS and incubated overnight, 100 μM MMG-11 and incubated for 1 hour then restimulated with 100 μM MMG-11 and 50 ng/mL Pam3CSK4 and incubated overnight, 100 ng/mL LPS and incubated overnight, 50 ng/mL Pam3CSK4 and incubated overnight, or not stimulated (unstimulated)

3. MFI allows for an approximation of how much cytokine is produced by a population of cells. Larger MFI values (relative to negative controls for that same fluorochrome) translate to larger amounts of the cytokine of interest produced by that population of cells. 4. Flow cytometry allows for the quantification of cytokine production in specific cell populations. This can be achieved by combining the use of gating and lineage markers to isolate specific populations with MFI analysis to quantify cytokine levels present in each of the cell populations of interest. 5. Multiple cytokines can be evaluated on the same cells at the same time if the antibodies to the cytokines of interest are conjugated to different fluorochromes for a single panel. 3.8.2

Benefits of ELISpot

1. Because ELISpot visualizes cytokine that is secreted, restimulation with a protein transport blocker (i.e., monensin or brefeldin A) is unnecessary. This will result in better viability of the cells during the stimulation process.

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2. ELISpot does not require many cells per well to visualize the number of cells that are secreting a cytokine of interest. Thus, experiments can still be carried out when you do not have many cells. 3. ELISpot is more sensitive than other applications because secreted protein is directly captured onto the well of the plate before it is diluted into the culture or captured by receptors on adjacent cells.

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Notes 1. Ficoll-Pacque Premium can be either stored at RT or brought to RT prior to the isolation of PBMCs. Failure to do so will result in a poor PBMCs isolation as the density separation is temperature sensitive. 2. Failure to keep the centrifuge at RT during isolation of PBMCs will result in a poor PBMC isolation because the density separation is temperature sensitive. 3. Ensure that 1× column wash buffer is kept cold (4 °C) throughout the purification procedure. Failure to do so will result in compromised cell purity. 4. When staining for secreted cytokines via flow cytometry, it is vital to use monensin, a Golgi stop, or brefeldin A, a Golgi plug, to induce accumulation of the cytokine in the cells. Golgi stops and Golgi plugs work by inhibiting protein transport thus allowing for the accumulation of secreted cytokines within the cell. If a Golgi plug or stop is not used prior to fixation and permeabilization of the cells, the cytokine of interest will be secreted into the media and will not be properly detected via flow cytometry. Determining the proper cytokine secretion blocker to use will depend on your cytokine of interest. BioLegend’s “Stimulation Guide for Intracellular Staining of Cytokines/Chemokines” is an available resource for determining the proper protein transport blocker to use for your experiment [11]. Both brefeldin A and monensin are toxic to cells and will decrease cell viability, so it is important to include a live dead stain in your panel [3]. 5. Ensure you wash your cells with 1× PBS or 1× HBSS prior to your live/dead stain. Many fixable live/dead stains, including Biotium Aqua, do not work in buffers containing BSA or other proteins/serums. These stains should be done in PBS and washed out with either a serum-containing buffer to stop the stain or washed multiple times with 1× PBS to dilute any remaining dead cell exclusion marker [3].

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6. Use a dead cell exclusion marker, such as Biotium Aqua, to gate out dead cells. Doublet exclusion helps (FSC-A vs. FSC-H) but does not eliminate all the dead cells. This is important as antibodies will stick to dead cells and give false-positive signals. Some stimulations, such as LPS, can negatively impact cell viability. Additionally, excluding dead cells is important when using Alexa 488 or FITC conjugated antibodies because dead cells auto-fluoresce in the same channel as these fluorochromes. 7. Do not leave the Biotium Aqua viability dye on the cells for longer than the indicated time. Extending the time past 30 min will give a false representation of cell viability as the cells will start dying in the viability dye. When using a fixable live/dead, such as Biotium Aqua, it is important to stain your cells with it prior to fixation and permeabilization. If it is added after these steps all cells will appear dead. 8. Staining cells with fluorescently conjugated antibodies should be done in the dark. Examples of how this could be achieved are the following: a drawer if the staining is done at RT, a refrigerator for 4 °C (given the light turns off when the door is closed), or a cell culture incubator for 37 °C. Some antibodies are prone to photobleaching and detection of certain protein markers can be affected if this occurs. 9. Always fix before permeabilization of cells. If you reverse this order, you will destroy your cells (i.e., perm buffers “poke” holes into the cells to allow for optimal staining of intracellular markers). You need to preserve the cell membrane first with fixation, then permeabilize for optimal intracellular staining. 10. It is important to use v-bottom 96-well plates for flow staining if you are targeting a small population of cells or expect to have low cell viability. Multiple washes will be done, and this will minimize cell loss due to the formation of a tight cell pellet. A flat-bottom plate will not allow for the formation of a tight cell pellet. A round-bottom plate or FACS tube is acceptable, but a v-bottom plate is better for the reasons listed above. 11. Compensation beads are the best way to optimally compensate the fluorochromes on your stimulated PBMCs cells. This is critical as some of the fluorochromes have high spectral overlap. False-positive signals may be obtained for certain markers (e.g., CD14 A750 and IL-6 A647) if compensation is not carried out [3]. 12. Viability dyes such as Biotium Aqua stain dead cells; therefore, the positive population will be the dead cells (this is also the case with SYTOX™ blue). Gate on the Biotium Aqua negative population for live cell analysis. This is best achieved by creating a dot plot with the Biotium Aqua on the x-axis and SSC-H on the y-axis. Using a histogram for gating can be difficult since

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it can be hard to distinguish between the break in the live versus dead cells. In addition, you can further clean up your cell population with a doublet exclusion gate. This is achieved by creating a dot plot with FSC-A on the x-axis and FSC-H on the y-axis. The cells that are in the diagonal are singlets; draw a polygon gate around these. Cells that fall outside the diagonal are doublets; do not include these in your analysis. Doublets take up fluorescent antibodies and can give a false-positive signal. Gating should be done as follows: FSC-H vs. SSC-H gate (cell population) ! FSC-A vs. FSC-H gate (doublet exclusion) ! Biotium Aqua vs. SSC-H gate (live cells). Apply this final gate to all your dot plots for surface marker analysis (Ref. [3] provides more in-depth explanation of live cell gating using dead cell exclusion dyes and doublet exclusion). 13. It is important to run all samples and donors on the same flow cytometer. The median fluorescence may differ between instruments due to differences in lasers. 14. Determine the optimal voltages for each fluorochrome in your multicolor sample before proceeding to compensation beads. This way you ensure that none of the fluorochromes on your cells will be off scale or set at too low a voltage. This is especially important if you are running your samples on a less automated cytometer, such as the BD Fortessa. 15. When analyzing MFI in FlowJo, there are three statistical options available. Median should be used instead of geometric mean or mean. This will give you the most accurate MFI value for your sample because it is less affected by outliers. 16. Diluting cells at different concentrations for ELISpot ensures that when you are counting the completed assay, you will have wells that do not have too little or too many cells [12]. 17. Adding aluminum foil to the bottom of the plate allows for even heat distribution and can help prevent the “edge effect” of the plate in which the cells migrate to the side wells of the plate where the plastic is warmer [13]. 18. Multiple washes ensure that cells are removed from the plate. This eliminates wells from being developed with a “sand” effect on the membrane from cells sticking to it. It is also acceptable to have a brief incubation (5 min) with a cell dissociation media to ensure complete removal of cells. 19. It is important to completely tap out the excess liquid in the wells onto a paper towel after washing is completed to prevent diluting the subsequent reagents that will be added to the ELISpot plate. 20. To ensure accurate results, reagent addition should be continuous and completed within 15 min.

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21. Incubation of detection antibodies for 2 h on a plate rocker at RT is also acceptable, but background noise may be higher. 22. Wells that have not dried completely will give inaccurate spot counts and spots will appear to be fuzzy or not well defined. References 1. Czerkinsky CC, Nillson LA, Nygren H et al (1983) A solid-phase enzyme-linked immunospot (ELISPOT) assay for enumeration of specific antibody-secreting cells. J Immunol Methods 65:109–121 2. Kalyuzhny AE (2005) Chemistry and biology of the ELISPOT assay. Methods Mol Biol 302: 15–31 3. Goetz C, Hammerbeck C, Bonnevier J (2018) Flow cytometry basics for the non-expert. In: Kalyuzhny A (ed) Techniques in life science and biomedicine for the non-expert. Springer, Minneapolis, pp 1–222 4. Bienvenu J, Monneret G, Fabien N et al (2000) The clinical usefulness of the measurement of cytokines. Clin Chem Lab Med 38:267–285 5. O’Neill LAJ (2006) How Toll-like receptors signal: what we know and what we don’t know. Curr Opin Immunol 18:3–9. https:// doi.org/10.1016/j.coi.2005.11.012 6. Park B, Lee JO (2013) Recognition of lipopolysaccharide pattern by TLR4 complexes. Exp Mol Med 45:e66 7. Horuluogli B, Bayik D, Kayraklioglu N et al (2019) PAM3 supports the generation of M2-like macrophages from lupus patient monocytes and improves disease outcome in murine lupus. J Autoimmun 99:24–32

8. Grabowski M, Murgueitio MS, Bermudez M et al (2020) The novel small-molecule antagonist MMG-11 preferentially inhibits TLR2/1 signaling. Biochem Pharmacol 171:113687. ht t p s : //d o i . o r g / 1 0. 10 1 6 / j . b c p . 20 1 9 . 113687. Epub 2019 Nov 1 9. Hagen J, Zimmerman R, Goetz C et al (2015) Comparative Multi-Donor Study of IFNγ Secretion and Expression by Human PBMCs Using ELISPOT Side-by-Side with ELISA and Flow Cytometry Assays. Cell 4:84–95 10. Maria B-C, Go´mez AI, Alberto B-M et al (2017) P2X7 receptor induces tumor necrosis factor-α converting enzyme activation and release to boost TNF-α production. Front Immunol 8:1–11. https://doi.org/10.3389/ fimmu.2017.00862 11. Biolegend Stimulation Guide: https://www. biolegend.com/Files/Images/media_assets/ support_protocol/BioLegend_ StimulationGuide_101711.pdf 12. Sedgewick JD, Holt PG (1983) A solid-phase immunoenzymatic technique for the innumeration of specific antibody-secreting cells. J Immunol Methods 57:301–309 13. Kalyuzhny A, Stark S (2001) A simple method to reduce the background and improve well-towell reproducibility of staining in ELISPOT assays. J Immunol Methods 257:93–97

Chapter 7 Reagent Tracker™ Platform Verifies and Provides Audit Trails for the Error-Free Implementation of T-Cell ImmunoSpot® Assays Alexander A. Lehmann, Diana R. Roen, Zolta´n Megyesi, and Paul V. Lehmann Abstract ELISPOT and FluoroSpot assays, collectively called ImmunoSpot assays, permit to reliable detection of rare antigen-specific T cells in freshly isolated cell material, such as peripheral blood mononuclear cells (PBMC). Establishing their frequency within all PBMC permits to assess the magnitude of antigen-specific T-cell immunity; the simultaneous measurement of their cytokine signatures reveals these T-cells’ lineage and effector functions, that is, the quality of T-cell-mediated immunity. Because of their unparalleled sensitivity, ease of implementation, robustness, and frugality in PBMC utilization, T-cell ImmunoSpot assays are increasingly becoming part of the standard immune monitoring repertoire. For regulated workflows, stringent audit trails of the data generated are a requirement. While this has been fully accomplished for the analysis of T-cell ImmunoSpot assay results, such are missing for the wet laboratory implementation of the actual test performed. Here we introduce a solution for enhancing and verifying the error-free implementation of T-cell ImmunoSpot assays. Key words ELISPOT, FluoroSpot, ImmunoSpot®, T cells, Immune monitoring, Immune memory, High throughput

1

Introduction ELISPOT and FluoroSpot assays both visualize secretory footprints of individual cells [1]. The two test systems differ only at the level of the visualization of the plate-bound analyte: in the former, the detection antibody is coupled to an enzyme that catalyzes the generation of a precipitating substrate detectable under white light; in the case of the latter, the plate-bound analyte is visualized using fluorescence-tagged detection antibodies. ELISPOT is suited for single- and dual-color analysis [2], while the unambiguous measurement of more than 2 analytes requires FluoroSpot tests using fluorochromes with nonoverlapping spectra [3]. As otherwise

Alexander E. Kalyuzhny (ed.), Handbook of ELISPOT: Methods and Protocols, Methods in Molecular Biology, vol. 2768, https://doi.org/10.1007/978-1-0716-3690-9_7, © The Author(s) 2024

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the two assays are identical, we refer to both collectively as ImmunoSpot®. ImmunoSpot® assays have found wide use in the field of T-cell immune monitoring. This is due to the unprecedentedly high sensitivity of the test system to detect antigen-specific T cells (which, with rare exceptions, always occur in very low frequency) in PBMC and other freshly isolated cell material [4]. The assay’s success is further based on its economy in cell utilization (see Note 1), its performance is unaffected when cryopreserved PBMC samples are tested [5], and last but not least, the simplicity and robustness of the ImmunoSpot® assay [6, 7] resulting in its suitability for regulatory validation in clinical trials [8]. The economy of PBMC utilization, along with ImmunoSpot®‘s high-throughput capability, enables batch testing of dozens, and even hundreds, of subjects’ PBMC for T-cell reactivity to hundreds of individual antigens/peptides per donor [9]. Such is essential, for example, for high-resolution CD8+ T-cell immune monitoring. As epitope/peptide recognition by CD8+ T cells is highly variable and unpredictable (“aleatory”) among outbred individuals [10, 20, 21], only the testing of extensive peptide libraries is suited for the comprehensive assessment of the magnitude and finespecificity of the antigen-specific CD8+ T-cell repertoire in individuals. There is a substantial danger of human error, however, when complex tests are performed involving the pipetting of the PBMC of numerous donors and many antigens. To overcome this problem, we have introduced neutral “regent tracker” dyes [11] that can assist in the visual verification of whether (a) the right antigen has been plated into the right well, and (b) whether the right amount/ volume of antigen has been plated (c) and whether the cells have been added (d) in the right numbers/volume. A typical 96-well plate with Reagent-Tracker®-marked antigens is shown in Fig. 1 (see Note 2). In this chapter, we introduce a software tool, the Reagent Tracker (RT) Software™, which permits to verify the accuracy of ImmunoSpot® assay implementation via automated image analysis while also providing tamper-proof audit trails (see Note 3). Figure 2a shows that Reagent Tracker® dyes can be readily distinguished by image analysis in the 3D color space: the color of each dye can be represented as a tight cluster of RGB coordinates. This permits the RT Software™ to unambiguously verify that the color actually present in a well indeed matches the color (antigen/ peptide) that was intended to be present in that well. Deviations from these color coordinates indicate impurities, e.g., resulting from spilling of antigen, and are automatically identified by the software as such. Optical densities of these defined colors reflect on the volume plated per well, and as such permit to identify quantitative antigen pipetting errors. The RT Software™ also automatically assesses whether such deviations from the planned volumes have occurred, permitting to flag deviant wells.

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Fig. 1 Color-code verification of pipetting accuracy. (a) Standard 96-well T-cell ImmunoSpot® plate is shown, without Reagent Tracker™ dyes, after plating 100 μL of antigen and the PBMC in 100 μL per well, both in medium containing phenol red. As all wells look alike, it is not possible to verify the accuracy of plating. (b) ImmunoSpot® plate after plating color-coded antigens using medium that has not been supplemented with phenol red, in 100 μL. In the example shown, eight Reagent Tracker™ dyes were used to color-code the antigens; each antigen was plated in duplicate wells underneath each other. (c) The plate shown in B after the addition of the PBMC in 100 μL medium containing phenol red

Fig. 2 Definition of Reagent Tracker™ dyes in RGB space. (a) The plate shown in Fig. 1b was scanned on an ImmunoSpot® Reader, and the wells containing the eight RT dyes were each defined in the red–green–blue color space. The axes are shown in the respective colors. (b) The plate shown in Fig. 1c was scanned and the color space was defined as above. Note the marked shift of the RGB coordinates for each color after adding the cells in phenol red-containing medium

As can be seen in Figs. 1b vs. 1c, and 2a vs. 2b, the addition of the PBMC in phenol red-containing medium causes a marked color shift of the RT dyes. Such shifts are assessed by the RT Software, permitting to automatically verify that the cells had indeed been added to each well, and that the planned numbers (volume) were added. The Reagent Tracker® Software guides the actual implementation of the Reagent Tracker™ strategy for T-cell ImmunoSpot® testing. As the first step, using this software’s planning module, the plate layout is defined, assigning to each well the antigens/peptide to be tested in it, as well as the PBMC donors’ identity whose cells are to be interrogated, including the number of replicate wells for

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each test condition, if applicable (see Notes 5 and 6). Now the color coding follows: color combinations are to be selected that provide visually clearly discernible patterns on the plates, like that shown in Fig. 1b using 8 RT Dyes™ (see Note 7). The color-coded reagents are prepared and plated (see Note 8) according to the plan. Before adding the PBMC, the plates are scanned and analyzed via the RT Software™ to establish and document the accuracy of the antigen transfer (see Note 9). Now the PBMCs are added, followed by a renewed scan, testing for the color shift resulting from the addition of the cells in phenol red media (see Note 9). As before, reports and audit trails are generated to document in a tamper-proof manner the accuracy of the plating process. Once the actual ImmunoSpot® assay has been concluded, the spot forming units (SFUs) per well are automatically counted by the ImmunoSpot® Software [12, 13], means of replicate wells are calculated, positive responses are automatically identified via statistical calculations, the results are exported into a database for further analysis, and, if desired, for instant graphic representation of the results. Barcoding is used to assure plate identity, and orientation, during the entire course of the test. The following provides technical details of performing RT-enhanced high-throughput T-cell-ImmunoSpot® assays. The protocol for loss-free cryopreservation and thawing of PBMC, and for the implementation of the basic T-cell assay, are provided elsewhere [14]. Dissolving antigens in RT Dyes™ and creating master plates for en bloc antigen transfers are also described in [14]. The automated evaluation of T-cell ImmunoSpot® assay results is described in [15, 16] (see Note 10).

2

Materials 1. ePBMC® samples or test samples (freshly isolated or cryopreserved human PBMC). 2. Precoated human IFN-γ 96-well ImmunoSpot® kit, enzymatic, or ELISPOT kit detecting any other analyte. 3. 50 mL conical tubes (polypropylene). 4. CTL Anti-Aggregate™ medium. 5. CTL-Test™ medium with phenol red. 6. CTL Live/Dead™ Cell Counting Dye and disposable hemocytometers. 7. Parafilm. 8. 70% ethanol 9. CTL Live/Dead Cell Counting™ suite. 10. Antigens/peptides of interest. 11. CTL-Reagent Tracker (RT)™ 1.

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12. CTL-Reagent Tracker (RT)™ 2. 13. CTL-Reagent Tracker (RT)™ 3. 14. CTL-Reagent Tracker (RT)™ 4. 15. Uncoated ELISPOT plate, 96 wells (for Reagent Tracker™ software calibration). 16. PBS. 17. 0.05% Tween-PBS. 18. Distilled water. 19. CTL Reagent Tracker™ software, version 1.1.3.1 or higher. 20. CTL ImmunoSpot® software. 21. CTL ImmunoSpot® analyzer.

3

Methods

3.1 Plan Experiment and Assign Reagent Tracker™ Dyes to Color-Code Each Assay Condition for Verification of Pipetting Accuracy

1. Open the CTL Reagent Tracker™ software. 2. Select “New Experiment”. 3. Click on “Add New Plate”. 4. Enter the plate name. 5. Select “Undefined Colors” from the Experiment template drop-down menu. 6. Begin by assigning Reagent Tracker™ 1 to the first assay condition (negative control). 7. The first peptide/antigen stimulation (condition 2) will be prepared with Reagent Tracker™ 2. 8. The second peptide/antigen stimulation (condition 3) with Reagent Tracker™3, and so forth until all assay conditions have been assigned a color code. If more than four conditions are used, the fifth condition will be assigned Reagent Tracker™ 1 and the cycle will begin again, until all conditions and controls have been assigned a Reagent Tracker™ dye. The software will display the completed plate diagram, which serves as a guide for generation of the calibration plate.

3.2 Preparation of Reagent Tracker™ Calibration Plate for Calibration of the Software

1. Following the plate diagram generated by the software, load 100 μL/well of the appropriate Reagent Tracker™ dye to each well of the calibration plate. For instance, if the negative control will be plated into wells A1 and A2 of the ELISPOT assay, add 100 μL of Reagent Tracker™ 1 dye to wells A1 and A2 of the calibration plate. 2. Continue loading the appropriate dyes into the calibration plate, 100 μL/well, until all wells have been filled. 3. Place the lid on the plate to preserve sterility during scanning.

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3.3 Perform Reagent Tracker™ Calibration Scan to Generate Reference Data

1. Click the eject button to eject the stage. 2. Load the calibration plate onto the stage, and click the load button to load the plate into the analyzer. 3. Click the scan button to perform a scan of the calibration plate. 4. Click the options button to the right of the plate name. Select “Update Dye Map.” The software will now save the reference scan data and will display 96 green wells under the “Pipetting Result” heading, indicating the successful generation of the reference data.

3.4 Thawing of Cryopreserved PBMC Samples (Sterile Conditions)

Warm all media to 37 °C prior to use. Perform all steps in a biological safety cabinet observing all applicable safety precautions. 1. Check the ID of the ePBMC® sample vials and place them into a bead (or water) bath at 37 °C for 8–10 min to thaw. 2. Wipe the cryovials with 70% ethanol and transfer them to a rack in the biological safety cabinet. 3. Invert the vials twice gently to resuspend the cells. Transfer the contents of a single donor’s sample into a labeled 50 mL conical tube. Up to 5 cryovials from a single donor can be pooled into one 50 mL conical tube. 4. Rinse the cryovial with 1 mL prewarmed Anti-Aggregate™ thawing medium, and transfer slowly, dropwise into the 50 mL conical tube while gently swirling the tube to ensure adequate mixing. 5. Add 2 mL Anti-Aggregate™ thawing medium to the conical tube dropwise while swirling gently to mix. 6. Add the remaining 6 mL Anti-Aggregate™ medium to the conical tube to bring the volume up to 10 mL total (for each cryovial thawed, e.g., if thawing 2 vials per donor, the total volume should be 2 × 10 mL = 20 mL), cap, and invert gently to resuspend the PBMC. 7. Repeat steps 3–6 for each donor sample to be tested, until all samples are resuspended in a warm thawing medium. 8. Centrifuge samples at 300 × g for 10 min with max brake. 9. Decant supernatant and flick the bottom of the tube gently to resuspend the pellet (do not vortex as this is detrimental to cell health). 10. Resuspend the pellet in 10 mL Anti-Aggregate™ medium. 11. Place a 20 μL aliquot of CTL Live/Dead™ dye onto a piece of parafilm for each sample to be counted.

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12. Remove 20 μL of PBMC suspension and add it to the droplet of CTL Live/Dead™ dye on the parafilm. Pipet up and down several times to mix. 13. Transfer 20 μL of the mixture into each chamber of the disposable hemocytometer. Only 10 μL will be drawn into the chamber, and the remainder will pool in the reservoir on the side, to prevent evaporation during counting. 14. Load the hemocytometers into the adapter tray and count the cells using CTL’s Live/Dead Cell Counting™ suite. 15. Centrifuge samples at 300 × g for 10 min with max brake. 16. Decant supernatant and flick the bottom of the tube gently to resuspend the cell pellet. 17. Resuspend each sample to the desired cell concentration (e.g., 4 × 106 PBMC/mL) in prewarmed CTL-Test™ medium containing phenol red. 18. Place the samples into a 37 °C incubator with 5–9% CO2 with loosened caps to allow for gas exchange until ready to plate them into the assay. 3.5 Preparation of Antigens and First Verification of Pipetting Accuracy Using Reagent Tracker™ Dyes and Software (Sterile Conditions)

1. Prepare antigens/peptides at 2× final concentration in Reagent Tracker dye. Utilize Reagent Tracker dyes such that no two consecutive peptides share the same dye color, following the plate diagram generated by the software. 2. Plate 2× antigen solutions, 100 μL/well, into the precoated ELISPOT plate (Note: after the addition of the cell suspension, the final concentration will become 1×). 3. Place the lid on the plate to preserve sterility and perform a scan of the precoated ELISPOT plate containing antigen solutions using the Reagent Tracker™ software and ImmunoSpot® analyzer to confirm correct concentration, volume, and dye color for each well. If all wells are within acceptable limits, the plate overview will show 96 green wells to signify accurate pipetting. 4. Once the accuracy of antigen loading has been verified, proceed to the addition of PBMC samples.

3.6 Plating of PBMC into ELISPOT Assay and Second Verification of Pipetting Accuracy Using Reagent Tracker™ Dyes and Software (Sterile Conditions)

1. Once the pipetting of the antigens has been confirmed to be correct, plate the cells in CTL-Test containing phenol red, 100 μL/well. This will cause all Reagent Tracker™ dye color profiles to shift slightly toward the red end of the spectrum, which can then be detected by the software. 2. Place the lid on the plate and perform a Reagent Tracker™ scan to determine that the correct volume of cells has been added to each well. If all concentrations, volumes, and dye colors are correct, the plate overview will display 96 green wells to signify accurate pipetting.

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3. Tap the plate gently on all sides to ensure even distribution of the cells across the membrane, and incubate at 37 °C with 5–9% CO2 for 18–24 h following the kit protocol. Do not move plates during the incubation to avoid disruption of spot formation. 3.7 Development of the ELISPOT Assay (Nonsterile Conditions)

1. Prepare detection solution according to the kit protocol, 10 mL per plate. 2. Remove plate(s) from the incubator and decant. 3. Wash plates 2× with PBS and 2× with 0.05% Tween-PBS, 200 μL per well. 4. Add detection solution, 80 μL/well. Cover and protect from light. Incubate at room temperature (RT) for 2 h. 5. Prepare tertiary solution according to the kit protocol, 10 mL per plate. 6. Decant plates and wash 3× with 0.05% Tween-PBS, 200 μL/ well. 7. Add tertiary solution, 80 μL/well. Cover plates and protect from light. Incubate for 30 min at RT. 8. Prepare substrate solution according to the kit protocol, 10 mL per plate. 9. Decant plates and wash 2× with 0.05% Tween-PBS and 2× with distilled water, 200 μL/well. 10. Add substrate solution, 80 μL/well. Cover plates and protect from light. Incubate for 10–15 min at RT. 11. Decant substrate solution and remove underdrain. Rinse plates, front and back, with tap water. 12. Dry plates completely prior to image acquisition and analysis. 13. Scan dried plates on an ImmunoSpot® analyzer using appropriate excitation and emission channels to visualize the assay prior to data analysis.

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Notes 1. PBMCs are always limited with clinical samples. In the typical 96-well plate format, 100,000–one million PBMC are tested per well, assuring a cell density on the membrane that results in a linear relationship between PBMC numbers plated and antigen-induced SFUs. In 384-well format, the number of PBMCs needed can be reduced to one-third, permitting to test 30 wells with only one million PBMC [17] obtainable from 1 mL of blood.

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2. The likelihood of human error is even larger when manually pipetting 384-well plates. 3. We have tested the biologic neutrality of Reagent Tracker dyes only in T-cell ELISPOT assays so far, but assume that this approach is also suited for other test systems involving complex pipetting patterns, which, however, would need to be verified for each additional test system. Reagent Tracker dyes are autofluorescent, however, which interferes with their use in FluoroSpot assays. 4. Systematic studies, in which 50–150 μL of color-coded antigen was pipetted per well in 10 μL increments showed that volume deviations of ±20% can be verified via color intensity analysis. 5. The number of replicates to be selected, as well as the choice of testing in 96- or 384-well plate format should be guided by the expected frequency of the antigen-specific T cells within the PBMC. As the ImmunoSpot® assay detects the cytokine footprint of individual antigen-triggered T cells, the Poisson distribution applies; it permits to select the PBMC numbers to be plated per well, the number of replicates, and the plate format [15]. 6. We recommend color coding with RT Dyes™ when many antigens/peptides are tested on PBMC from a high number of donors. 7. While for machine reading, the selection of colorful plate design can be random as the colors are clearly discernable in RGB space (see Fig. 2), visually discernable, repetitive color patterns provide guidance during the actual plating of the antigens. 8. When a high number of antigens are to be tested, we recommend preparing “antigen master plates” that follow the antigen layout of the planned test(s), and from which multichannel pipettors can transfer the antigens in the predefined sets [14]. 9. Using backlight, the plates can be scanned with the lid on to maintain sterility. 10. The rules for ELISPOT and FluoroSpot analysis of T-cell assay results are similar, but the latter needs to also reliably define co-expression patterns by individual T cells for which simple overlays of centers of masses do not suffice, but that requires accounting for Stokes shifts and movement of the cells [18] in particular when the production of the analytes is asynchronous, being the case for most cytokines produced by T cells [19].

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Acknowledgments We wish to thank R&D and the Software Development teams at CTL for their continued support and technological innovation that made our Reagent Tracker Software development possible. All efforts were funded from CTL’s research budget. Conflicts of Interest P.V.L. is Founder, President, and CEO of CTL, a company that specializes in immune monitoring by ImmunoSpot®. A.A.L. is CTL’s COO. D.R.R. and Z.M. are employees of CTL.

References 1. Hesse MD, Karulin AY, Boehm BO et al (2001) A T cell clone’s avidity is a function of Its activation state. J Immunol 167:1353– 1361 2. Karulin AY, Hesse MD, Tary-Lehmann M et al (2000) Single-cytokine-producing CD4 memory cells predominate in type 1 and type 2 immunity. J Immunol 164:1862–1872 3. Megyesi Z, Lehmann PV, Karulin AY (2018) Multi-color FLUOROSPOT counting using ImmunoSpot® Fluoro-X Suite. Methods Molec Biol 1808:115–131 4. Helms T, Boehm BO, Asaad RJ et al (2000) Direct visualization of cytokine-producing recall antigen-specific CD4 memory T cells in healthy individuals and HIV patients. J Immunol 164:3723–3732 5. Kreher CR, Dittrich MT, Guerkov R et al (2003) CD4+ and CD8+ cells in cryopreserved human PBMC maintain full functionality in cytokine ELISPOT assays. J Immunol Methods 278:79–93 6. Sundararaman S, Karulin AY, Ansari T et al (2015) High reproducibility of ELISPOT counts from nine different laboratories. Cell 4:21–39 7. Zhang W, Caspell R, Karulin AY, et al., (2009) ELISPOT assays provide reproducible results among different laboratories for T-cell immune monitoring – even in hands of ELISPOTinexperienced investigators. J Immunotoxicol 6:227–234 8. Tary-Lehmann M, Hamm CD, Lehmann PV (2008) In: Kelley eUPaM (ed) Validating reference samples for comparison in a regulated ELISPOT assay. Validation of Cell-Based Assays in the GLP Setting, pp 127–146. h t t p s : // d o i . o r g / 1 0 . 1 0 0 2 / 9780470987810.ch9

9. Lehmann PV, Suwansaard M, Zhang T et al (2019) Comprehensive evaluation of the expressed CD8+ T cell epitope space using high-throughput epitope mapping. Front Immunol 10:655–668 10. Moldovan I, Targoni O, Zhang W et al (2016) How frequently are predicted peptides actually recognized by CD8 cells? Cancer Immunol Immunother 65:847–855 11. Lehmann A, Megyesi Z, Przybyla A et al (2018) Reagent tracker dyes permit quality control for verifying plating accuracy in ELISPOT tests. Cell 7:3–13 12. Karulin AY, Caspell R, Dittrich M et al (2015) Normal distribution of CD8+ T-cell-derived ELISPOT counts within replicates justifies the reliance on parametric statistics for identifying positive responses. Cell 4:96–111 13. Zhang W, Lehmann PV (2012) Objective, user-independent ELISPOT data analysis based on scientifically validated principles. Methods Molec Biology 792:155–171 14. Lehmann PV, Roen DR, Lehmann AA (2023) Unbiased, high-throughput identification of T cell epitopes by ELISPOT. Methods Molec Biology 2673:69–88 15. Dittrich M, Lehmann PV (2012) Statistical analysis of ELISPOT assays. Methods Molec Biol 792:173–183 16. Karulin AY, Karacsony K, Zhang W et al (2015) ELISPOTs produced by CD8 and CD4 cells follow log normal size distribution permitting objective counting. Cell 4:56–70 17. Hanson J, Sundararaman S, Caspell R et al (2015) ELISPOT assays in 384-well format: up to 30 data points with one million cells. Cell 4:71–83

Reagent Tracker Software 18. Karulin AY, Megyesi Z, Caspell R et al (2018) Multiplexing T- and B-cell FLUOROSPOT assays: experimental validation of the multicolor ImmunoSpot® Software based on center of mass distance algorithm. Methods Molec Biol 1808:95–113 19. Duechting A, Przybyla A, Kuerten S et al (2017) Delayed activation kinetics of Th2and Th17 cells compared to Th1 cells. Cell 6: 29–34

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20. Lehmann AA, Zhang T, Reche PA et al (2021) Discordance between the predicted vs. the actually recognized CD8+ T cell epitopes of HCMV pp65 antigen and aleatory epitope dominance. Front Immunol 11:Art. 618428 21. Lehmann AA, Lehmann PV (2020) Aleatory epitope recognition prevails in human T cell responses? Crit Rev Immunol 40:225–235

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 8 Detection of SARS-CoV-2-Specific Cells Utilizing Whole Proteins and/or Peptides in Human PBMCs Using IFN-ƴ ELISPOT Assay Madeleine M. Rasche, Ella C. Kaufmann, Tamar Ratishvili, Ilya M. Swanson, Inna G. Ovsyannikova, and Richard B. Kennedy Abstract SARS-CoV-2 continues to threaten global public health, making COVID-19 immunity studies of utmost importance. Waning of antibody responses postinfection and/or vaccination and the emergence of immune escape variants have been ongoing challenges in mitigating SARS-CoV-2 morbidity and mortality. While a tremendous amount of work has been done to characterize humoral immune responses to SARS-CoV2 virus and vaccines, cellular immunity, mediated by T cells, is critical for efficient viral control and protection and demonstrates high durability and cross-reactivity to coronavirus variants. Thus, ELISPOT, a standard assay for antigen-specific cellular immune response assessment, allows us to evaluate SARS-CoV2-specific T-cell response by quantifying the frequency of SARS-CoV-2-specific cytokine-secreting cells in vitro. We have outlined a detailed procedure to study T-cell recall responses to SARS-CoV-2 in human peripheral blood mononuclear cells (PBMCs) following infection and/or vaccination using an optimized IFN-γ ELISPOT assay. Our methodologies can be adapted to assess other cytokines and are a useful tool for studying other viral pathogen and/or peptide-specific T-cell responses. Key words IFN-ƴ ELISPOT, SARS-CoV-2, COVID-19, COVID-19 vaccination, PBMCs, Cellular immunity

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Introduction COVID-19, caused by the novel coronavirus SARS-CoV-2, has killed over 1.1 million people in the United States and over 15 million people worldwide as of March 2023 [1–3]. SARS-CoV-2 has had an unprecedented impact on individuals, societies, and economies, and persists as a global public health threat. The availability of safe and effective COVID-19 vaccines has played a crucial role in pandemic mitigation efforts. However, the waning of disease- and vaccine-induced immunity (especially waning antibody titers) [4– 6] and continuous viral evolution resulting in the emergence of

Alexander E. Kalyuzhny (ed.), Handbook of ELISPOT: Methods and Protocols, Methods in Molecular Biology, vol. 2768, https://doi.org/10.1007/978-1-0716-3690-9_8, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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viral immune escape variants (i.e., variants of concern) [7, 8] pose ongoing challenges toward these mitigation efforts. Assessment of disease- or vaccine-induced, as well as hybrid, immunity to SARS-CoV-2 is critical to the evaluation of immune status at individual and population levels. This knowledge also guides the determination of correlates of protection from infection or reinfection and of durable and cross-protective immunity – which do not yet exist for SARS-CoV-2. Humoral immune responses, specifically neutralizing antibodies (nAb), have been the primary focus of SARS-CoV-2 immune response and vaccine immunogenicity/efficacy studies. However, T-cell-mediated cellular immunity has gained increasing interest, as it has been found to be crucial for viral control, disease resolution, and generation of fine-tuned coordinated immune response against SARS-CoV-2 [9– 13]. Compared to humoral responses, the T-cell response to SARS-CoV-2 demonstrates higher durability and extensive crossreactivity to emerging viral variants. T cells make important contributions to protection against severe disease [9, 14–17]. As IFN-γ secretion from antigen-stimulated T (CD4+/CD8+) cells is a sensitive marker for immune recognition by Th1-biased cell-mediated immunity (CMI), IFN-γ ELISPOT allows for enumeration of antigen-specific cytokine-secreting cells at a single-cell level. Though no gold-standard assay has yet been defined for rapid high-throughput quantification of a SARS-CoV-2-specific adaptive cellular immune response, IFN-γ ELISPOT is extensively used for assessment of CMI to various pathogens and vaccines and is now being applied to studying SARS-CoV-2 adaptive immunity after both COVID-19 disease and vaccination [18–23]. To this end, we have optimized an IFN-γ ELISPOT assay with multiple SARSCoV-2 antigens (whole proteins and peptide pools) to detect SARS-CoV-2-specific cellular immune responses in cryopreserved peripheral blood mononuclear cells (PBMCs). We have successfully used this method for SARS-CoV-2-specific cell quantification in healthy COVID-19 convalescent individuals, vaccine recipients with no history of disease, and those with hybrid SARS-CoV2 immunity. As the IFN-γ ELISPOT assay measures cellular immunity, it is ideal for evaluating immune responses in individuals with defects in the humoral arm of the immune system [24–26]. We have demonstrated this potential in patients with chronic lymphocytic leukemia/lymphoma (CLL) [27], whose humoral immune responses to COVID-19 disease and vaccination are suboptimal due to the underlying disease and use of B cell targeting treatments [26, 28]. Our optimized IFN-γ ELISPOT method works well with fresh PBMCs, but here we report the procedure utilizing cryopreserved human PBMCs without the need to isolate T cells (although this can be done to focus on CD4, CD8, or other T cell subset-specific responses). Nor is there a need to prestimulate T cells as is necessary

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with some other cytokines (e.g., IL-17). These assay characteristics reduce the technical barriers associated with the use of fresh blood samples, and the increased cost/complexity of cell purification procedures and lengthy incubation protocols that may introduce in vitro artifacts. Thus, this assay can be utilized as a high-throughput screening assay for quantifying SARS-CoV-2-specific CMI in large diverse study cohorts in different laboratory settings. Here we present the detailed optimized protocol for cell thawing, cell culture, assay development, and spot quantification to obtain accurate and reproducible SARS-CoV-2-specific IFN-γ ELISPOT results.

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Materials

2.1 Preparation of Antigen Stocks

1. RPMI 1640 culture media with 5% FCS: 930 mL of RPMI 1640 with glutamine, 50 mL of FCS, 10 mL of penicillin– streptomycin, 10 mL of sodium pyruvate. 2. PepTivator® SARS-CoV-2 Prot_S Complete (Miltenyi Biotec). 3. SARS-CoV-2 (2019-nCoV) Spike S1+S2 ECD-His Recombinant Protein (Sino Biological). 4. SARS-CoV-2 (2019-nCoV) Nucleocapsid-His Recombinant Protein (Sino Biological). 5. Phytohemagglutinin (PHA). 6. 1× PBS. 7. Sterile water. 8. Dimethyl sulfoxide (DMSO). 9. 1.5 mL tubes.

2.2 Collection and Isolation of PBMCs

1. Ficoll-Paque Plus. 2. Accuspin tube or Leucosep tube. 3. Human whole blood. 4. 1× PBS. 5. Pasteur pipettes. 6. 0.4% Trypan blue. 7. Hemocytometer. 8. 50 mL conical tubes. 9. ACK lysis buffer. 10. Cell strainer. 11. FACS tubes. 12. RPMI 1640 freezing media: 700 mL of RPMI 1640 with glutamine, 200 mL of FCS, 100 mL of DMSO.

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13. 1.8 mL cryogenic freezing tubes. 14. Freezing container (Mr. Frosty). 15. 99% Isopropanol. 2.3 Coating of ELISPOT Plates

Coating of ELISPOT plates should be done a day prior to assay setup. 1. Capture antibody: Concentration: 1.0 mg/mL. BD NA/LETM purified anti-human IFN-γ (sterile) (included in the BD Human IFN-γ ELISPOT Set kit. BD Biosciences). 2. Coating buffer: phosphate buffered saline (PBS). 3. ELISPOT plate (included in the BD Human IFN-γ ELISPOT Set kit. BD Biosciences). 4. Dilute capture antibody in coating buffer (PBS) 1:200. 5. Add 100 μL of diluted antibody solution to each well of an ELISPOT plate. 6. Store plates at 4 °C overnight with the plate lids on. Wrapping in aluminum foil is optional.

2.4 Thawing of Cryopreserved PBMCs

1. Cryopreserved PBMCs. 2. RPMI 1640 thaw media: 879 mL of RPMI 1640 with glutamine, 100 mL of FCS, 10 mL of penicillin–streptomycin, 10 mL of sodium pyruvate, 1 mL of DNAse. 3. 15 mL conical tubes. 4. FACS tubes. 5. 1× PBS. 6. 0.4% Trypan blue. 7. Hemocytometer. 8. RPMI 1640 culture media: 930 mL of RPMI 1640 with glutamine, 50 mL of FCS, 10 mL of penicillin–streptomycin, 10 mL of sodium pyruvate. 9. 50 mL conical tubes.

2.5 Preparation of Antigen Dilutions (Working Solutions)

1. RPMI 1640 culture media: 930 mL of RPMI 1640 with glutamine, 50 mL of FCS, 10 mL of penicillin–streptomycin, 10 mL of sodium pyruvate. 2. Stock antigens. Specific antigens and controls for cell stimulation vary by experiment and are defined by the investigator. Stock antigens should be diluted in RPMI 1640 culture media at appropriate dilutions. For this assay, we used the aliquots of stock antigens/controls listed in 2.1 at concentrations recommended by the manufacturers (see Note 1).

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3. PepTivator® SARS-CoV-2 Prot_S Complete: 50 μg/peptide/ mL. 4. SARS-CoV-2 (2019-nCoV) Spike S1+S2 ECD-His recombinant protein: 200 μg/mL. 5. SARS-CoV-2 (2019-nCoV) Nucleocapsid-His recombinant protein: 200 μg/mL. 6. Phytohemagglutinin PHA: 200 μg/mL. 7. 15 mL conical tubes. 2.6 Human IFN-γ ELISPOT Assay Setup: Plating and Culture

1. RPMI 1640 blocking media: 879 mL of RPMI 1640 with glutamine, 100 mL of FCS, 10 mL of penicillin–streptomycin. 2. Human PBMCs (concentration of 2 × 106 cells/mL). 3. Antigen dilutions. 4. Phytohemagglutinin PHA (positive control). 5. RPMI 1640 culture media (negative control).

2.7 Assay Development

1. Detection antibody: Concentration: 0.5 mg/mL. Biotinylated anti-human IFN-γ (included in the BD Human IFN-γ ELISPOT Set kit. BD Biosciences). 2. Streptavidin-HRP: Concentration: 100×. Enzyme conjugate (included in the BD Human IFN-γ ELISPOT Set kit. BD Biosciences). 3. Wash buffer I: 1× PBS containing 0.05% Tween-20. 4. Wash buffer II: 1× PBS. 5. Dilution buffer: 1× PBS containing 10% FCS. 6. Substrate solution: MossBio TMB-H peroxidase substrate. 7. Deionized (DI) water.

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Methods The following method describes the process to detect and quantify SARS-CoV-2-specific cells that secrete IFN-γ in human PBMCs using the BD Biosciences Human IFN-γ ELISPOT Set. Note that similar reagents or kits are available from many other manufacturers. The general procedure outlined here can be easily modified to adapt to those kits.

3.1 Preparation of Antigen Stocks

Prepare antigen stocks with manufacturer-recommended diluents. Store at recommended concentrations or concentrations optimized in the lab at recommended temperatures. For this assay: 1. Add 500 μL of sterile water or PBS to SARS-CoV-2 (2019nCoV) Spike S1+S2 ECD-His recombinant protein for a stock concentration of 200 μg/mL. Aliquot as needed and store at

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-20 °C to -80 °C. Thaw and allow to equilibrate to room temperature shortly before use. Do not refreeze leftover material. 2. Add 500 μL of sterile water or PBS to SARS-CoV-2 (2019nCoV) Nucleocapsid-His recombinant protein for a stock concentration of 200 μg/mL. Aliquot as needed and store at -20 °C to -80 °C. 3. Add 200 μL of sterile water to a 6 nmol/peptide vial (2 mL to 60 nmol/peptide vials) of PepTivator® SARS-CoV-2 Prot_S Complete to have a stock at 50 μg/mL. Aliquot as needed and store at -20 °C. Thaw and allow to equilibrate to room temperature shortly before use. Do not refreeze leftover material. 4. Add 25 mL of sterile water or RPMI 1640 culture media to 5 mg lyophilized PHA for a stock concentration of 200 μg/mL (assay positive control). Aliquot 1 mL in 1.5 mL tubes and store at -20 °C. Thaw and allow to equilibrate to room temperature shortly before use. Do not refreeze leftover material. 3.2 Collection and Isolation of Human PBMCs

1. Bring Ficoll-Paque Plus to room temperature, ensuring that it is protected from light. 2. Pipet 15.5 mL of Ficoll-Paque Plus into the upper chamber of each Accuspin tube or Leucosep tube. You need 4 Accuspin tubes/subject assuming 90–100 mL whole blood is collected. Adjust the procedure as needed. 3. Centrifuge Accuspin tubes at 800× g (1800 rpm) for 30 s at room temperature. After centrifugation, Ficoll-Paque Plus will be in the chamber below the frit. If there are bubbles below the frit add 1 mL of Ficoll-Paque Plus and spin again. If there is some extra Ficoll-Paque Plus on the frit, remove it via aspiration. 4. Label 4 Accuspin tubes with the subject ID. 5. If blood was collected in 10 mL heparin tubes, carefully remove the tube caps, and gently pipet whole blood volume into Accuspin tubes. Pipet the volume of 2.5 heparin tubes into each Accuspin tube. If blood was collected in a bag with heparin, mix the blood by inverting the bag. Then place the bag inside a beaker so it is secure. (The side with the metal crimp should be on the top.) Take scissors and cut off one of the upper corners of the blood bag. Using a 25 mL serological pipet, gently transfer 25 mL of blood into the upper chambers of each Accuspin tube. 6. After each Accuspin tube has been filled with approximately 25 mL blood, bring the volume up to 50 mL with sterile 1× PBS. Mix the blood with the PBS by gently pipetting the mixture with a 25 mL pipet, being careful not to force the blood below the frit.

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7. Centrifuge at 2070 rpm for 15 min at 25 °C, with the centrifuge brake OFF. 8. While centrifuging the Accuspin tubes, label two 50 mL sterile centrifuge tubes with the subject ID for each subject. 9. After centrifugation, aspirate and discard the topmost layer of supernatant (diluted plasma) above the buffy coat (white layer above the frit). Be careful not to disturb the buffy coat layer. 10. Pour off the buffy layer and the remaining supernatant from the 4 Accuspin tubes into 2 prelabeled 50 mL centrifuge tubes. 11. Wash the cells by bringing the volume to 50 mL with sterile 1× PBS. 12. Gently resuspend the lymphocytes by inverting the tube several times. 13. Centrifuge at 1500 rpm for 10 min at 25 °C. Centrifuge brake ON. 14. Remove the supernatant by aspiration using a sterile Pasteur pipette. Do not disturb the cell pellet. Use a new pipette for each subject sample. 15. Add 5 mL of ACK lysis buffer to the cell pellet and resuspend the cells by vortexing or pipetting up and down. Let the cells sit for 5 min at room temperature. 16. Add sterile 1× PBS to bring the volume up to 50 mL. 17. Centrifuge at 1500 rpm for 10 min at 25 °C. Centrifuge brake ON. 18. Remove the supernatant by aspiration using a sterile Pasteur pipette being careful not to disturb the cell pellet. Use a new pipette for each subject. 19. Resuspend the cells in 10 mL of 1× PBS per tube (20 mL total). 20. Label one 50 mL centrifuge tube for each subject with the subject ID. 21. Place a cell strainer on top of each 50 mL centrifuge tube and pipet the cells from each 50 mL tube for one subject into the strainer to remove the platelet aggregates (you should pool the cells from one subject at this point). Mix well by inverting the centrifuge tube. If the total volume is not 20 mL, bring it to 20 mL with 1× PBS. 22. Cell counting: Place 200 μL of 1× PBS, 37.5 μL of trypan blue, and 12.5 μL of your cell suspension into a falcon tube. Mix the sample well. Fill a hemacytometer with 10 μL of the sample and count unstained (live) cells in the four outer quadrants.

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23. Calculate cell concentration and record cell recovery on the checklist for each subject. Cell concentration formula: Total cell count = (# of live cells/4) × 10,000 × 20 (dilution factor [250/12.5]) × total volume of cells (2 mL). 24. Centrifuge the cell suspension at 1500 rpm for 10 min at 25 °C. Centrifuge brake ON. 25. Remove the supernatant by aspiration using a sterile pipette. Use a new pipette for each subject. 26. Adjust the cells to a concentration of 1 × 107 cells/mL with cold (4 °C) RPMI freezing media. 27. Pipette the cell suspension into prelabeled cryogenic freezing tubes. Inserts need to be placed in the caps. 34 sets of labels will be provided for each subject and each cryogenic freezing tube needs 2 labels (barcode label on the cap insert and the detailed label on the side). Aliquot 1.0 mL of the cell suspension into each cryogenic freezing tube (see Note 2). 28. Place the cryogenic freezing tubes into a controlled rate freezing container. Place freezing container in -80 °C freezer overnight. 29. Transfer the cryogenic freezing tubes to a liquid nitrogen storage tank for long-term storage. 3.3 Coating of ELISPOT Plates

Coating of ELISPOT plates should be done a day prior to assay setup.

3.4 Thawing of Cryopreserved PBMCs

In this method, we use cryopreserved PBMCs, which allows us to perform IFN-y ELISPOT assays later from the collection date of these cells. This allows for the use of large batches and minimizes assay drift/variability and batch effects. Previous studies have shown that cell viability is not compromised due to cryopreserving PBMCs [19]. 1. Warm RPMI 1640 thawing media in 37 °C water bath for 15 min (see Note 3). 2. Remove PBMC vials from liquid nitrogen tanks. Store in -80 °C freezer until ready to start. 3. Label one 15 mL conical tube with subject/sample ID for each subject/sample. 4. Thaw PBMCs in 37 °C water bath for 2–3 min. 5. In a sterile tissue culture hood, add 500 μL RPMI 1640 thawing media to each 15 mL conical tube. 6. Wipe PBMC vials with 70% ethanol and place them in a sterile tissue culture hood. 7. Transfer PBMCs to designated 15 mL conical tubes.

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8. Add RPMI 1640 thawing media slowly, doubling the volume (starting with 500 μL of RPMI 1640 thaw media and 1 mL from PBMC vial, add 1.5 mL of media, then 3 mL of media, and then 6 mL of media) until a final volume of 12 mL is reached. This should be done over a 5-minute period (see Note 4). 9. Cap each tube and invert 5 times to mix, do not vortex. 10. Centrifuge at 1200 rpm at room temperature for 7 min. 11. Remove supernatant by vacuum aspiration, leaving approximately 300 μL in the tube. Do not aspirate the cell pellet. 12. Resuspend the cells in 10 mL thawing media. 13. Cap each tube and invert 5 times to mix, do not vortex. 14. Place 15 mL conical tubes in 37 °C water bath for 20 min. Invert tubes to mix at 10 minutes. 15. Cool cells on ice for 7 min. 16. Centrifuge at 1200 rpm at 4 °C for 7 min. 17. Carefully remove supernatant by vacuum aspiration without disturbing the cell pellet. 18. Resuspend cells in each tube in 2 mL RPMI 1640 culture media (or customize based on expected cell recovery). 19. Prepare falcon tubes for counting. To count the number of live and dead cells, place 200 μL of 1× PBS, 37.5 μL of trypan blue, and 12.5 μL of cell suspension into a falcon tube. Mix well and pipette 10 μL onto a hemocytometer. Count and record the number of live cells in the four outer quadrants of the hemocytometer. 20. Total cell count = (# of live cells/4) × 10,000 × 20 (dilution factor [250/12.5]) × total volume of cells (2 mL). 21. Add or remove the amount of RPMI 1640 culture media to the appropriate conical tube to adjust the cell concentration to 2 × 106 cells/mL (see Note 5). 22. The recommended cell viability for this assay is >80%. 3.5 Preparation of Antigen Dilutions (Working Solutions)

Antigen concentrations are defined by the manufacturer or in the lab during assay optimization. 1. Label 15 mL conical tubes with the names of each stimulant/ control. Add needed volume of RPMI 1640 culture media to tubes. 2. Bring the stock preparations to room temperature and prepare dilutions as soon as possible (see Note 6). 3. Prepare Spike S1+S2 ECD-His recombinant protein dilutions by diluting the stock (200 μg/mL) 1:80 in RPMI 1640 culture media for a 2.5 μg/mL working concentration ! 1.25 μg/mL final concentration.

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4. Prepare Nucleocapsid-His recombinant protein dilutions by diluting the stock (200 μg/mL) 1:80 in RPMI 1640 culture media for a 2.5 μg/mL working concentration ! 1.25 μg/mL final concentration. 5. Prepare PepTivator® SARS-CoV-2 Prot_S Complete dilutions by diluting the stock (50 μg/peptide/mL) 1:25 in RPMI 1640 culture media for a 2 μg/mL working concentration ! 1 μg/ mL final concentration after plating with cells. 6. Prepare PHA dilutions by diluting the stock (200 μg/mL) 1: 20 in RPMI 1640 culture media for a 10 μg/mL working concentration ! 5 μg/mL final concentration. 3.6 Human IFN-γ ELISPOT Assay Setup: Plating and Culture

1. Label the lid of the ELISPOT plate with the following information (before plating). (a) Subject ID (b) Unstimulated (to appropriate wells) (c) SARS-CoV-2 antigens (Spike protein, Nucleocapsid protein, Peptivator) (d) PHA (e) Your initials, date, and plate label on BOTH the lid and on the ELISPOT plate. 2. From this point on, all steps in this section should be done in a tissue culture hood to maintain sterility. 3. Remove blocking media by dumping contents of the plate into a container. Pat the plate dry. 4. Use a multichannel pipette to add cells and antigens. We used quadruplicates for each condition (see Notes 7 and 8). 5. Add 100 μL of cells in each indicated well (per predefined plate template). 6. Add 100 μL of each antigen in each indicated well (per predefined plate template). 7. Add 100 μL RPMI 1640 culture media to unstimulated wells (assay negative control). 8. Add 100 μL PHA to the indicated wells (assay positive control). 9. Incubate at 37 °C in 5% CO2 incubator for 18–24 h with the plate lid on. We recommend wrapping the plates in aluminum foil for incubation to ensure even distribution of temperature in wells (see Notes 9 and 10).

3.7 Assay Development

1. Prepare dilution buffer and wash buffer I. 2. Remove the plates from the incubator. 3. Sterile conditions are not necessary from now on.

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4. Cell culture supernatant can be collected at this stage if desired. Any analyte, excluding the one captured by antibodies used in the ELISPOT assay (here: IFN-γ) can be measured. Remove 150–175 μL supernatant using 200–300 μL pipette tips and a multichannel pipette and pipette into round- or v-bottom plates. Supernatants can be used freshly or can be frozen at 80 °C (cover the plate with a plate sealer) and thawed at another time. 5. If supernatant collection is not intended, remove cell suspension from the plates by deliberately flicking into a sink or waste receptacle. 6. Wash wells 2× with deionized (DI) water. Allow wells to soak for 3–5 min at each wash step. 7. Wash wells 3× with 200 μL/well wash buffer I. Discard wash buffer. Pat dry to remove as much liquid as possible. 8. Dilute detection antibody in dilution buffer (1:250) and add 100 μL/well. 9. Incubate 2 h at room temperature with the plate lid on. 10. Discard detection antibody solution. Wash wells 3× with 200 μL/well wash buffer I. Allow wells to soak for 1–2 min at each wash step. Pat dry to remove as much liquid as possible. 11. Dilute enzyme conjugate (streptavidin-HRP) in dilution buffer (1:100) and add 100 μL/well. Make just before using. 12. Incubate 1 h in the dark at room temperature with the plate lid on. 13. Discard enzyme conjugate solution. Wash cells 4× with 200 μL/well wash buffer I. Allow cells to soak for 1–2 min at each wash step. 14. Wash cells 2× with 200 μL/well wash buffer II. Pat dry to remove as much liquid as possible. 15. Add 100 μL of substrate solution to each well and protect plates from light. Monitor spot development for 7 min (unless indicated otherwise). Do not allow spots to overdevelop, as this will lead to high background (see Note 11). 16. Stop substrate reaction by rinsing the plate 3× with DI water. Remove the plastic drain and rinse the bottom of the plate. 17. Pat the plate dry on paper towels. Wipe the bottom of the plate dry with paper towels without damaging the well membranes. 18. Air-dry plate at room temperature for 2 h or overnight until it is completely dry. Removal of plastic tray under plate will facilitate drying.

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3.8 Determination of IFN-γ Secreting Cell Frequencies (in SFUs per 2 × 105 cells) (See Notes 12–14)

1. Scan the IFN-γ ELISPOT plate after drying and make sure that the plate is in focus, is dry, looks uniform, membranes are intact, and that contrast or exposure parameters are optimal for counting these plates. Save the results. Use the same parameters for all plates. 2. Count a batch of IFN-γ ELISPOT plates using appropriate counting parameters based on spot detection sensitivity, minimum and maximum spot sizes, and spot separation. Keep the same counting parameters for all batches in the same project.

4

Notes 1. Avoid freeze–thaw cycles by using aliquots of antigens and controls. 2. When aliquoting isolated PBMCs for freezing in cryotubes, pipette the cell suspension up and down several times after distributing every 5 aliquots to mix and ensure uniform cell concentrations. 3. Do not omit the thaw media warming step for maximal recovery of live PBMCs. 4. Where slow addition of the thawing media to cells is recommended, add media in a slow, drop-wise manner to avoid osmotic and thermal shock to PBMCs. 5. Plate cells as soon as possible after counting to maintain cell viability and function. Prepare antigen dilutions during the last centrifugation (before counting), vortex, and keep them at room temperature until cells are counted and resuspended. If using numerous antigens for the setup, one person can prepare and plate the antigens while another person is preparing cells. 6. Prepare enough antigen for at least 10% more wells than needed. This will help avoid potential delays during plating. 7. Due to inherent variability of the ELISPOT assay readouts, the use of technical replicates is strongly recommended. Testing in triplicates or quadruplicates is a recommended minimum. 8. The membranes on the bottom of the ELISPOT plate microwells are fragile. Take care not to touch the well membranes with pipette tips during cell and antigen plating and washes. Most plates have an underdrain protective cover – do not remove it until after the plates have developed and you are washing off the substrate solution. 9. For overnight incubation, wrap the plates in aluminum foil to permit uniform temperature distribution in all wells and thus helps minimize evaporation and edge effects. Check the wrapping for folds and bumps on the bottom to avoid tilting the plate.

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10. Do not stack the plates when placing them in the 5% CO2 incubator. The plates should be spaced and incubated separately. This allows even temperature distribution and further reduces edge effects. Do not disturb the plates during incubation to avoid development of ambiguous spots or streaks. 11. Several common problems can be encountered after the substrate solution is washed off and the plate is developed. (a) Dark background complicates accurate spot quantification in all wells. Shortly after stopping the substrate-enzyme reaction and final DI water wash, the entire membrane will be dark blue and the color contrast between spots and the background will not be sufficient for adequate visualization of spots. (i) This could mean the plates are still wet. Plates cannot be scanned and accurately read until plates are dried thoroughly. Allow at least 1–2 h (preferably overnight) for plates to dry at room temperature after patting them dry on a pile of paper towels following final DI washes. Plates should be protected from light during this time. (ii) If the contrast between the background and spots is still low after the plates are fully dry and high background color is present across the plate (or batch) and not limited to one or several samples, it is likely a procedural issue rather than a cell stimulation issue. Stringently follow the blocking and plate washing protocol to prevent high background. Make sure not to skip the PBS washing step before adding the substrate solution to the plates after PBS-Tween-20 washes, because Tween-20 can impair substrate reaction. Also, leaving the substrate solution on too long can result in overdeveloping the plates. Avoiding lengthy spot development time will minimize background. (b) Background in the unstimulated wells is high, that is, the spot counts are high despite no stimulation. In contrast to the situation in the previous paragraph, high spot counts in unstimulated wells indicate a problem in the cell stimulation conditions. The cells could have been inadvertently stimulated both specifically and nonspecifically due to several reasons. Make sure to follow stringent sterile protocol where appropriate (see above), including during media preparation to avoid contamination (e.g., fungus and LPS). Change pipettes and/or pipette tips between antigens and samples. It is not

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uncommon for the cells to inadvertently get exposed to antigens or mitogens included in the experimental setup if some antigen dilution is accidentally put into unstimulated condition wells. Make sure your pipetting is careful and consistent, and if such errors happen, indicate on the plate cover. (c) Spot counts are adequate/within the expected range, but spot intensity and contrast to background are low, causing errors in counting. This could be caused by underdevelopment of the plates/spots due to suboptimal incubation time with substrate solution, or use of cold enzyme-streptavidin conjugate or substrate solution. Make sure to bring the reagents to room temperature as indicated in the kit. Closely monitor spot development after adding the substrate solution to the plate to avoid under- and overdevelopment of spots. (d) Spot counts are lower than expected in all stimulated wells, including those stimulated with a positive control, such as PHA. You should ensure all antigens/stimuli and assay reagents were used at appropriate concentrations and remain biologically active. Some kits contain an IFN-γ positive control, which could help highlight the primary problem. If the IFN-γ control works robustly, then the problem is more likely to be the cell stimulation conditions. In contrast, low spot counts in all wells, including IFN-γ positive control, points to the assay setup and/or plate development steps as the problem. Another reason behind lower-than-expected spot counts across all stimulation conditions is the use of cells at suboptimal concentrations. We conventionally plate 2 × 105 PBMCs per well based on our previous optimization experiments. Higher (up to 5 × 105) or lower cell concentrations may be used after testing different cell dilutions and defining the optimum for your studies. 12. Once optimized for an assay, the scanning and counting parameters should not be changed for the complete batch. Most ELISPOT readers allow the user to save these parameters for use with subsequent plates. 13. If modifications to scanning or counting parameters are necessary (plate is drastically different such as darker wells, high background, etc.) from other plates in the batch, always record these modifications. If satisfactory modifications are not feasible, consider repeating the plate.

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14. After counting the spots before analyzing the data, define a cutoff for mean spot numbers in positive and negative controls (e.g., mean spot counts in PHA-stimulated wells >100–150) based on your optimization experiment(s) to determine if the cells were viable and capable of producing cytokines or that the background is sufficiently low to allow accurate spot counting in the experimental wells. Apply these rules to the whole dataset for consistency.

Acknowledgments We thank the Mayo Clinic Vaccine Research Group team for technical assistance and participation in the development, optimization, and execution of this assay. This work was supported by funding from ICW Ventures, and NIH-funded grants R01AI132348, R01 AI127365, and R01 AI121054. Conflict of Interest Dr. Ovsyannikova holds patents related to vaccinia and measles peptide vaccines. Drs. Kennedy and Ovsyannikova hold a patent related to vaccinia peptide vaccines, and a patent related to the impact of single nucleotide polymorphisms on measles vaccine immunity. Drs. Kennedy and Ovsyannikova have received grant funding and royalties from ICW Ventures for preclinical studies on a peptide-based COVID-19 vaccine. Dr. Kennedy has received funding from Merck Research Laboratories to study waning immunity to mumps vaccine. Dr. Kennedy also offers consultative advice on vaccine development to Merck & Co. and Sanofi Pasteur. These activities have been reviewed by the Mayo Clinic Conflict of Interest Review Board and are conducted in compliance with the Mayo Clinic Conflict of Interest policies. This research has been reviewed by the Mayo Clinic Conflict of Interest Review Board and was conducted in compliance with the Mayo Clinic Conflict of Interest policy.

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Chapter 9 Interferon-γ/IL-2 ELISpot and mRNA Responses to the SARS-CoV2, Feline Coronavirus Serotypes 1 (FCoV1), and FCoV2 Receptor Binding Domains by the T Cells from COVID-19-Vaccinated Humans and FCoV1-Infected Cats Sabarinath Nair, Bikash Sahay, Ananta P. Arukha, Lekshmi K. Edison, Chiquitha D. Crews, John G. Morris Jr, Subhashinie Kariyawasam, and Janet K. Yamamoto Abstract The receptor binding domain (RBD) of SARS-CoV-2 (SCoV2) has been used recently to identify the RBD sequences of feline coronavirus serotypes 1 (FCoV1) and 2 (FCoV2). Cats naturally infected with FCoV1 have been shown to possess serum reactivities with FCoV1 and SCoV2 RBDs but not with FCoV2 RBD. In the current study, COVID-19-vaccinated humans and FCoV1-infected laboratory cats were evaluated for interferon-gamma (IFNγ) and interleukin-2 (IL-2 ELISpot responses by their peripheral blood mononuclear cells (PBMC) to SCoV2, FCoV1, and FCoV2 RBDs. Remarkably, the PBMC from COVID-19vaccinated subjects developed IFNγ responses to SCoV2, FCoV1, and FCoV2 RBDs. The most vaccinated subject (five vaccinations over 2 years) appeared to produce hyperreactive IFNγ responses to all three RBDs, including the PBS media control. This subject lost IFNγ responses to all RBDs at 9 months (9 mo) post-last vaccination. However, her IL-2 responses to FCoV1 and FCoV2 RBDs were low but detectable at 10 mo post-last vaccination. This observation suggests that initially robust IFNγ responses to SCoV2 RBD may be an outcome of robust inflammatory IFNγ responses to SCoV2 RBD. Hence, the T-cell responses of vaccine immunity should be monitored by vaccine immunogen-specific IL-2 production. The PBMC from chronically FCoV1-infected cats developed robust IFNγ responses to SCoV2 and FCoV2 RBDs but had the lowest IFNγ responses to FCoV1 RBD. The constant exposure to FCoV1 reinfection may cause the IFNγ responses to be downregulated to the infecting virus FCoV1 but not to the cross-reacting epitopes on the SCoV2 and FCoV2 RBDs. Key words SARS-CoV2, Feline coronaviruses, Receptor binding domain, COVID-19 vaccines

Sabarinath Nair and Bikash Sahay contributed equally with all other contributors. Alexander E. Kalyuzhny (ed.), Handbook of ELISPOT: Methods and Protocols, Methods in Molecular Biology, vol. 2768, https://doi.org/10.1007/978-1-0716-3690-9_9, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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Introduction In a recent publication, the sera and plasma from cats naturally infected with feline coronavirus serotype 1 (FCoV1) cross-reacted with SARS-CoV2 receptor binding domain (SCoV2 RBD) [1]. The SCoV2 RBD was used in this chapter to identify the RBD sequences of FCoV1 and FCoV serotype 2 (FCoV2). The FCoV1 and FCoV2 RBDs were produced and evaluated as described [1]. The plasma from FCoV1-KU2-infected laboratory cats reacted with FCoV1 RBD but did not cross-react with FCoV2 and SCoV2 RBDs. However, the plasma from FCoV2-79-1146infected laboratory cats reacted with FCoV2 RBD and crossreacted with SCoV2 RBDs but not with FCoV1 RBD. The serum samples from clinic cases reacted with FCoV1 RBD but not with FCoV2 RBD. These samples also cross-reacted with SCoV2 RBD when the cats and the owners were negative for SCoV2 infection [1]. The serum from group-housed laboratory cats accidentally infected with FCoV1 was also tested and found to react with both FCoV1 and SCoV2 RBDs but not with FCoV2 RBD. The fact that FCoV1 infection in cats is more common than FCoV2 infection in the United States supports their findings that naturally or accidentally infected cats were most likely infected with FCoV1 as supported by their reactivity with FCoV1 RBD and not with FCoV2 RBD [1, 2]. Their results further suggest that not all FCoV1 strains will infect and produce cross-reactive antibodies to SCoV2 RBD, even when FCoV2 inoculated cats developed cross-reactive antibodies to SCoV2 RBD [1]. Their serological studies demonstrate that there are most likely common epitopes on SCoV2 RBD and FCoV1 and FCoV2 RBDs. The goals of the current studies are to determine if crossreactive T-cell responses in the PBMC of COVID-19-vaccinated humans induce reactivity specifically to SCoV2 RBD rather than those to FCoV1 and/or FCoV2 RBD(s). Since there is no effective commercial FCoV1 vaccine, FCoV1-infected cats were evaluated for the T-cell reactivity of their peripheral blood mononuclear cells (PBMC) to FCoV1, FCoV2, and SCoV2 RBDs. The PBMC from COVID-19-vaccinated subjects and FCoV1-infected laboratory cats were stimulated with SCoV2, FCoV1, and FCoV2 RBDs in sterile ELISpot plate or in sterile round-bottom 96-well plate for mRNA analyses and analyzed as described. The value of using ELISpot over intracellular staining (ICS) for IL-2 and IFNγ is that the antibodies to human IL-2 and IFNγ are available for ICS but not for feline counterparts. Both human and feline IL-2 and IFNγ ELISpot modules are commercially available (Materials). Furthermore, cDNA primers for both human and feline IL-2 and IFNγ mRNAs are readily available (Materials), but this method will

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not demonstrate or support the production of IL-2 and IFNγ proteins in responses to these RBDs. Hence, current studies combined two approaches to address the overarching goals of these studies.

2 2.1

Materials Human Subjects

1. COVID-19-vaccinated human subjects were recruited and bled according to approved University of Florida IRB202002902. 2. The vaccination and background profiles of the five consented and coded subjects (Y2, Y3, Y8, Y9, and Y10) are shown in Table 1 with age at the most recent bleeding in 2023.

2.2

Animal Subjects

1. Specific pathogen-free (SPF) cats were bred at the University of Florida (UF) by the Laboratories of Comparative Immunology and Virology for Companion Animals (LCIV-CA) program using IACUC protocols 202001838 and 202300000347. 2. Accidental FCoV1 infection of SPF cats (Y2E and 4GA) occurred in 2022. These cats were individually housed for over 4 months to clear FCoV1 infection. 3. The blood from chronically FCoV1-infected laboratory cats (G1, G2, and G7) were donated using approved IACUC 202103990. 4. The vaccination and background profiles of the five laboratory cats (G1, G2, G7, 4GA, and Y2E) are shown in Table 2, with the estimated age at most recent bleeding in 2023.

Table 1 Profiles of the five vaccinated subjects Subject Code

Y2

Y3

Y8

Y9

Y10

Age in years at 2023

69

69

48

48

21

Gender

Female

Female

Male

Male

Male

4

5

3

3

4

Vaccination Frequency a

a

b

Vaccine type

M/M/M/Mb

P/P/P/P/Mb

M/P/Pb

P/P /P

P/P/P/P

SCoV2 exposure

0

0

0

1b

0

FCoV cat exposure

+

–

–

–

+

Pfizer monovalent (P) and bivalent (Pb) vaccines and Moderna monovalent (M) and bivalent (Mb) vaccines were used. Subject Y9 was infected with SCoV2 post-second vaccination.

b

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Table 2 Chronically FCoV1-Infected and FCoV1-Cleared Laboratory Cats Lab Cat Code

G1

G2

G7

4GA

Y2E

Age in years at 2023

>3.5

>3.5

>3.5

2.25

3.25

Gender

Neutered Female

Neutered Male

Male

FCoV1 infection statusa + Infected

+ + Infected Infected

– – Cleared Cleared

Feline vaccinationsb

+

+

+

–

–

SCoV2 exposure

None

None

None

None

None

FCoV1 exposure

Chronic

Chronic Chronic

Cleared Cleared

a

Chronically FCoV1-infected cats (G1,G2, and G7) and FCoV1-infection-cleared cats based on presence or absence of anti-FCoV antibodies. b These laboratory cats were vaccinated with RABVAC3 vaccine against rabies and with Fel-O-Vax PCT + Calicivax vaccine against feline panleukopenia, calici, rhinotracheitis viruses, and hemorrhagic feline calicivirus strain.

2.3 Peripheral Blood Mononuclear Cell Preparation

1. Lymphocyte separation medium (LSM or Ficoll-Hypaque; Corning/Mediatech, Inc., Manassas, VA).

2.4

1. SCoV2, FCoV2, and FCoV1 receptor binding domain (RBD) were synthesized by our laboratory using the method described [1].

Stimulants

2. Hanks’ balanced salt solution 10× without sodium bicarbonate, calcium, and magnesium (Corning, Cat. No. 20-021-CV, Manassas, VA). Sodium bicarbonate at 7.5% w/v solution (Mediatech Inc., Cat. No. 25-035-CI, Manassas, VA).

2. T-cell mitogens are phytohemagglutinin P (PHA-P, SigmaAldrich, Inc., Cat. No. L1668-5MG, St. Louis, MO) for humans and concanavalin A (ConA, Sigma Cat. No. C52755MG) for cats. 2.5 IFNγ and IL2 ELISpot for Humans and Cats

1. Ninety-six-well polyvinylidene fluoride (PVDF)-membrane white plates (Millipore Multiscreen plates, Merck Millipore, Cork, Ireland). 2. Culture medium for ELISpot: AIM-V medium (Gibco/Life Technologies, Cat. No. 12055-083, Grand Island, NY) supplemented with gentamicin (25 μg/mL) and with 5% heatinactivated (56 °C, 30 min) fetal bovine serum (FBS, R&D Systems/Bio-Techne, Cat. No. S11550, Atlanta GA). 3. Phosphate-buffered saline (PBS, Corning, Cat. No. 21040CM), 11.9 mM phosphate,137 mM sodium chloride, 2.7 mM potassium chloride, pH 7.4. endotoxin low, DNasefree, RNase-free, and protease-free and sterile filtered (0.2 μm) to decrease the background.

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4. Reagent diluent: PBS supplemented with 1% (w/v) fraction V bovine serum albumin (Fisher Scientific, Cat. No. bp16000100) for reconstituting lyophilized antibodies. 5. Blocking buffer: 1% BSA, 5% (w/v) sucrose in PBS. 6. Human IFNγ ELISpot development module (R&D Systems Inc., Cat. No. SEL285, Minneapolis, MN), human IL-2 ELISpot development module (R&D Systems Inc., Cat. No. SEL202) included a matching pair of antibodies used for capture and detection of individual feline cytokine and ELISpot blue color module (R&D Systems Inc.). (a) Respective feline cytokine (IFNγ, IL-2). Capture antibody concentrate reconstituted in 1 mL PBS and diluted 1 in 50 parts PBS. (b) Respective feline cytokine (IFNγ, IL-2). Detection antibody concentrate was reconstituted in 1 mL reagent diluent further 50-fold with the reagent diluents. 7. Feline IFNγ ELISpot development module (R&D Systems Inc., Cat. No. SEL764) and feline IL-2 ELISpot development module (R&D Systems Inc., Cat. No. SEL1890) included a matching pair of antibodies used for capture and detection of individual feline cytokine and ELISpot Blue color module (R&D systems Inc.). 8. ELISpot Blue color module (R&D Systems Inc., Cat. No. SEL002) (a) Streptavidin-AP concentrate (streptavidin conjugated to alkaline phosphatase) diluted 1 in 50 parts with reagent diluent. (b) 5-Bromo-4-chloro-3′ indolyl phosphate p-toluidine salt (BCIP)/nitro blue tetrazolium (NBT) chromogen consists of BCIP and NBT chloride in an organic solvent. 2.6 PBMC Culture Medium

1. RPMI-1640 (CORNING, Mediatech Inc. 10-040-CM, Manassas, VA) 2. Supplements: gentamicin sulfate (Gemini Byproducts, Cat. No. 400-108, Sacramento, CA), fetal bovine serum (FBS, R&D Systems/Bio-Techne, Cat. No. S11550, Atlanta GA).

2.7 Primers and Probes for Cytokine mRNAs

1. The primers and probes for human IFNγ, IL2, and GAPDH were produced by Integrated DNA Technologies (IDT, Coralville, IO). 2. The forward and reverse primers and probes used for each amplification of IFNγ, IL2, and GAPDH sequences are shown in Table 3.

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Table 3 Primers and Probe for IFNγ, IL2, and GAPDH Human IFNγ Amplification Forward Reverse Probe

5′GCATCGTTTTGGGTTCTCTTG 3′ 5′AGTTCCATTATCCGCTACATCTG 3′ 5′TCTGCTTCTTTTACATATGGGTCCTGGC 3′

Human IL2 Amplification Forward Reverse Probe

5′AAAGAAAACACAGCTACAACTGG 3′ 5′GAAGATGTTTCAGTTCTGTGGC 3′ 5′TGTGAGCATCCTGGTGAGTTTGGG 3′

GAPDH Amplification Forward Reverse Probe

2.8

mRNA Extraction

5′ACATCGCTCAGACACCATG 3′ 5′TGTAGTTGAGGTCAATGAAGG 3′ 5′AAGGTCGGAGTCAACGGATTTGGT 3′

Content of Zymo Direct-zol RNA micro prep kit: 1. Zymo Direct-zol RNA micro prep (ZYMO, Cat. No. R2050, Irvine, CA) 2. Tri reagent (Cat. No. R2050-1-200) 3. Prewash buffer (Cat. No. R2050-2-40) 4. Wash buffer (Cat. No. R1003-3-6).

2.9 Reverse Transcription and PCR of cDNA

3

1. Luna script reverse transcriptase (Luna Script RT Super mix Kit, New England Biolabs cat no. NEB E3010, Ipswich, MA). 2. Sso advanced universal probes super mix (BIORAD Cat. No. 1725284, Hercules, CA).

Methods Two approaches were undertaken to compare the preliminary IFNγ and IL-2 responses to SCoV2, FCoV1, and FCoV2 RBDs by the PBMC from COVID-19-vaccinated subject Y3 for both ELISpot and mRNA analyses. All remaining COVID-19-vaccinated subjects and FCoV1-infected laboratory cats were evaluated by IFNγ ELISpot analyses. Both IFNγ and IL-2 are mainly produced by T cells present in the PBMC [3–5]. Current results will determine if the IFNγ and IL-2 responses to SCoV2, FCoV1, and FCoV2 RBDs can be measured by ELISpot analyses for the T-cell immunity generated in the PBMC from the COVID-19-vaccinated human subjects. Since there is no effective commercial FCoV1 vaccine, the PBMC from chronically FCoV1-infected laboratory cats will be compared to the human results and to the FCoV1-infection-cleared laboratory cats for RBD-specific IFNγ responses by ELISpot analyses.

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Fig. 1 Anti-CoV RBD IFNγ and IL-2 ELISpot responses of the PBMC from a COVID-19-vaccinated subject Y3. Subject Y3 has been vaccinated five times with COVID-19 vaccine since 2020–2022. The last vaccination consisted of Moderna bivalent COVID-19 vaccine but those prior to this vaccine were all monovalent Pfizer COVID-19 vaccine. The IFNγ and IL-2 responses to human (SCoV2) and feline (FCoV1, FCoV2) receptor binding domains (RBDs) are shown with the months post-last vaccination, when the blood was collected for PBMC isolation. The red dotted line represents the threshold of the average PBS control response. Those responses to RBD(s), substantially above this threshold, would indicate a substantial or significant PBMC response. The statistical significant difference between the PBS control and the respective responses to the RBDs is shown p < 0.025 (*) and p < 0.05 (**), based on the two-tailed paired t-test. This subject has never been infected with SCoV2. The description of this subject’s profile is shown in Table 1 (Subheading 2.1). 3.1 IFNγ and IL2 Analyses of Subjects Y3

1. Subject Y3 received five COVID-19 vaccinations starting from fall 2020 to fall 2022. This individual was initially 67 years old when SCoV2 first spread in the United States in 2020 and was recently evaluated for IFNγ responses to SCoV2, FCoV1, and FCoV2 RBDs. At 4 mo post-5th vaccination (p5v), her PBMC had significantly higher IFNγ responses to all RBDs than her average PBS control value (Fig. 1, IFNγ 4 mo). More importantly, IFNγ responses to all RBDs were close to the average IFNγ levels of the PBMC stimulated by the T-cell mitogen, phytohemagglutinin P (PHA-P). Her average PBS control level was also highly elevated, showing spontaneous IFNγ production. Thus, the IFNγ responses to RBDs were still hyperreactive at 4 mos p5v but completely disappeared by 9 mo p5v (Fig. 1, 4 mo vs. 9 mo). One month later, at 10 mo p5v, her PBMC had low but substantial IL-2 ELISpot responses to FCoV1 and FCoV2 RBDs (Fig. 1, IL-2 10 mo). The cytokine IL-2 is known as a strong T-cell immune mediator and is not known as an inflammatory cytokine [5]. However, IFNγ is well known to be an inflammatory cytokine and, at a moderate dose, an important T-cell immune mediator [3, 4]. Current results suggest that the COVID-19 vaccine-induced IFNγ for this subject is responding as an inflammatory cytokine that crashes in IFNγ production with no recall memory by 9 mo p5v. The fact that IL-2 responses still existed at 10 mo p5v indicates that vaccine immunity should be monitored by vaccine immunogen-specific IL-2 responses.

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2. In our study, female subject Y3 was vaccinated five times over the 2 years (2020–2022), and her PBMC collected from recent bleedings in 2023 were tested. The vaccination profile of subject Y3 is shown in Table 1 (Subheading 2.1). The PBMC from this subject was stimulated with these RBDs for 18 hours at 1 × 106 cells/well, and the extracted mRNAs were tested for IFNγ and IL-2 mRNA levels. The IFNγ mRNA levels of the PBMC from Y3 were hyperreactive and produced high IFNγ and IL2 mRNA levels to SCoV2 and FCoV1 RBDs but not to FCoV2 RBD at 3 mo p5v. These levels were at the same level as PBS media control (see Note 1). The PHA-P mitogen was not available for this study. The same PBMC preparation at 10 mo p5v as Fig. 1 induced high IFNγ and IL-2 mRNA levels to SCoV2 and FCoV1 RBDs but not to FCoV2 RBD. The high IFNγ and IL2 mRNA levels were similar to the PHA-P and PBS media control levels. Thus, all samples had similar high IFNγ mRNA levels, demonstrating hyperstimulation for spontaneous IFNγ production, most likely due to overvaccination (see Note 1). The high IL-2 mRNA levels at both time points may suggest that mRNAs are produced at high levels but potentially not translated to proteins at high levels. The most conflicting results are that both IFNγ and IL-2 mRNA levels in response to FCoV2 RBD were not detected when ELISpot results consistently showed responses to FCoV2 in PBMC from Y3 in Fig. 1. The culture conditions between ELISpot and mRNA analyses were considerably different. The mRNA analysis had a shorter duration of stimulation (18 h), RPMI medium with less FBS (2%), and the number of PBMC twofold less (0.5 × 106 PBMC/well). Furthermore, it is much easier to degrade mRNA during purification, which is most likely the cause of the lack of IFNγ and IL-2 mRNA levels to FCoV2 RBD. Current studies indicate that ELISpot analysis is far better than cytokine mRNA analysis (see Note 2). 3.2 IFNγ Analysis of Other Vaccinated Subjects

The female subject Y2 is the same age as subject Y3 but received less vaccination than Y3. Both male subjects, Y8 and Y9, are of the same age with the same frequency of vaccinations. However, subject Y9 is the only one infected with SCoV2 (Table 1). Subject Y10 is the youngest at age 21 years at the time of blood collection. The PBMC were isolated from the blood collected from subject Y2 at 5 months (5 mo) post-last 4th vaccination (5 mo p4v), Y10 at 9 mo post-last 4th vaccination (9 mo p4v), Y9 at 12 mo post-last 3rd vaccination (12 mo p3v), and Y8 at 9 mo post-last 3rd vaccination (9 mo p3v), as shown next to their identification code in the figure. The PBMC from subjects Y2 and Y10 have substantially to significantly more IFNγ responses to SCoV2, FCoV1, and FCoV2 RBDs than their average PBS control value. Both Y2 and Y10 have multiple FCoV1infected cats, which may have altered their COVID-19 vaccination

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results to be higher by constant shedding of FCoV1 by their cats in their households. However, both subjects, Y8 and Y9, have no cats in their household. The PBMC from subject Y9 has significantly more IFNγ responses to FCoV1 and FCoV2 RBDs than his average PBS control value. Unexpectedly, his PBMC had no IFNγ responses to the SCoV2 RBD substantially above the average PBS control value. The PBMC from Y9 at 12 mo p3v is approximately 1.5 years postSCoV2 infection, suggesting that the SCoV2 infection at 12 mo p2v and 6 mo before the p3v has augmented the cross-reactive CoV immunity. Unexpectedly, the PBMC at 9 mo p3v from subject Y8 had no IFNγ responses to any of the RBDs, which were above his average PBS control value. Subject Y8, who is the same age as Y9 (48 years), has no responses to any RBDs even though he was last vaccinated 9mo ago with Pfizer bivalent. Such a result is similar to the IFNγ responses observed with PBMC of subject Y3 at 9 mo p5v. Hence, his PBMC should be tested for IL-2 responses since his immunity at age 48 years should be more robust than Y3 at age 69 years. In contrast, the PBMC of subject Y10 at 9 mo p4v had significant IFNγ responses to SCoV2 and FCoV1 RBDs and substantial IFNγ responses to FCoV2 RBD when compared to his average PBS control value. Subject Y10, at the time of bleeding, was 21 years old, has retained the CoV RBD responses at 9 mo p4v far better than Y8 at 9 mo p3v, Y9 12 mo p3v, and Y2 at 5 mo p4v, and even higher than IL-2 response of subject Y3 at 9 mo p5v (Fig 1). Unexpectedly, the IFNγ responses to SCoV2 RBD for Y2, Y9, and Y10 were slightly to substantially lower in levels than those levels to FCoV1 RBD, and even with IL-2 responses for Y3 at 10 mo p5v. Such results suggest immunogen-specific downregulation of immune responses. Overall, Figs. 1, 2, and 3 demonstrate that COVID-19 vaccines induce FCoV1 and FCoV2 RBD responses more strongly than those to SCoV2 RBD by ELISpot analyses. These responses to FCoV1 and FCoV2 RBDs by the PBMC from COVID-19-vaccinated subjects suggest that such responses are more likely due to the cross-reactive epitope(s) on SCoV2 RBD residing also on FCoV1 and FCoV2 RBDs. Although unlikely, there is a remote possibility that all six vaccinated subjects were all recently infected with human common cold α-coronaviruses with RBD epitope (s) cross-reactive with both SCoV2 RBD and FCoV RBDs. Further studies should be performed with a larger number of subjects and also evaluate their anti-CoV serology. 3.3 IFNγ Analysis of FCoV1-Infected Cats

Chronically FCoV1-infected cats (G1, G2, and G7) have antibodies to FCoV (Fig. 4) whole-virus (WV) and live together in a multiplecat housing, which causes the constant FCoV1 reinfection [1]. While infected, they should have IFNγ responses to at least FCoV1 RBD. Remarkably, two of the three cats had more IFNγ

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Fig. 2 The IFNγ and IL-2 mRNA levels in response to CoV RBDs of the PBMC of subjects Y3. The PBMC from subject Y3 for cytokine mRNA analysis were performed at two timepoints, 3mo p5v (top) and 10mo p5v (bottom). The results are shown in relative quantification (RQ) to the housekeeping gene, GAPDH. The mRNA analysis for IFNγ (left) and IL-2 (right) shows responses to only SCoV2 (blue bars) and FCoV1 (red bars) RBDs and high levels to PBS media control (MC, white bars) and mitogen phytohemagglutinin-P (PHA, bottom gray bars). The PHA bar is missing at 3 mo p5v

responses to SCoV2 and FCoV2 RBDs than to FCoV1 RBD. However, all three cats had slight-to-substantial IFNγ responses to all RBDs above their average PBS control value. FCoV infection cycles among the infected/reinfected cats based on our serology study described in our previous studies [1]. Hence, the variation in the IFNγ responses to FCoV1 RBD by these cats was expected. However, the fact that IFNγ responses to SCoV2 and FCoV2 RBDs were higher than those to FCoV1 RBD in two of the three infected cats suggests downregulation of IFNγ responses occurs specifically to the infecting virus serotype, FCoV1. Such downregulation did not occur to the cross-reactive epitopes on SCoV2 and FCoV2 RBDs. More cat studies will be needed to confirm these results. As expected, the two FCoV1-infection-cleared cats (4GA, 2.25-year-old male, and Y2E, 3.25-year-old male) had no IFNγ responses to any of the RBDs. In fact, although slight, all IFNγ responses to RBDs were lower than their average PBS control

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Fig. 3 Anti-CoV RBD IFNγ ELISpot responses of the PBMC from COVID-19-vaccinated subjects. Three COVID19-vaccinated subjects (Y2, Y8, and Y10) were reported to be free of SCoV2 exposure, whereas one subject Y9 was infected with SCoV2 one year after the second vaccination, which is also 3 mo before the third vaccination. The red dotted line for each subject represents the threshold of the average PBS control response. Those responses to RBD(s), substantially above this threshold, would indicate a substantial or significant PBMC response. The statistical significant difference between the PBS control and the respective responses to the RBD(s) is shown as p < 0.05 (*), based on the two-tailed paired t-test. The description of these subjects’ profiles is shown in Table 1 (Subheading 2.1).

Fig. 4 Anti-CoV RBD IFNγ ELISpot responses of the PBMC from chronically FCoV1-infected cats and FCoV1 infection-cleared cats. Chronically FCoV1-infected cats are laboratory cats G1, G2, and G7. FCoV1-infectioncleared cats are laboratory cats 4GA and Y2E. The red dotted line for each cat represents the threshold of the average PBS control response. These results suggest that downregulation of IFNγ responses to FCoV1 RBD in the infecting coronavirus FCoV1 occurs with cats G1, G2, and 4GA. Those responses to RBD(s) substantially above this threshold would indicate a substantial or significant PBMC response. The statistical significant difference between the PBS control and the respective responses to the RBD(s) is shown as p < 0.05 (*) and p ≤ 0.025 (**), based on the two-tailed paired t-test. The description of these cats’ profile is described in Table 2 (Subheading 2.2)

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value, demonstrating the clearing of FCoV1 infection. The IFNγ responses to FCoV1 and FCoV2 RBDs in cat 4GA were statistically lower than the average PBS control value. These two cats were considered cleared of FCoV1 infection based on the total loss of their antibodies to FCoV-WV by living individually in a single housing unit for at least 4 months. The single housing prevents reinfection from occurring from another FCoV1-infected cat and causes no cycling of reinfection. Their lack of IFNγ responses to all RBDs clearly demonstrates clearance of FCoV1 infection. The ConA mitogen stimulation is lower in the PBMC from laboratory cats 4GA and Y2E than those from cats G1, G2, and G7. Besides the difference in age range, cats 4GA and Y2E were not vaccinated with feline vaccines as they were bred in SPF condition at UF (Table 2). Hence, they are closer to SPF status by being unvaccinated and not being constantly exposed to FCoV1. 3.4 Isolation of Human and Feline PBMC

1. Prepare the 1× HBSS by adding 100 mL of 10× HBSS to 900 mL of sterile deionized water and 2.99 mL of 7.5% sodium bicarbonate and mix. 2. For every 1 mL of blood, add at least 1 mL of 1× HBSS (i.e., 10 mL of whole blood should be diluted to a total volume of 20 mL by adding 10 mL of 1× HBSS). 3. Mix the whole blood and HBSS by inversion (at least 10×) of the 50 mL conical tube or by pipetting. Mix until a homogenous suspension is formed. 4. Load 3 mL lymphocyte separation media (LSM) into each sterile 15 mL sterile conical tube. 5. Carefully load 10 mL of whole blood/HBSS mixture gently on the slightly tilted side of the tube onto the 3 mL LSM in the 15 mL conical tube. Use caution not to disturb the LSM. 6. Centrifuge the samples for 20 min at 500 g (1570 rpm) 7. Harvest the interphase/buffy coat (3–5 mL) with a sterile transfer pipette into 10 mL of HBSS and mix by vortex. 8. Spin for 5 min at 1570 rpm to pellet the PBMC. 9. Decant the supernatant and resuspend the PBMC pellet using 10 mL HBSS. 10. Centrifuge the PBMC suspension for 5 min, at 1570 rpm. 11. Decant the supernatant and repeat steps 9–11 for an additional two more washes to remove platelets. 12. Upon decanting the fluid, resuspend the PBMC pellet in the appropriate culture media.

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1. Charging/activating the PVDF plate (a) Ninety-six-well PVDF membrane plates are primed by adding 30 μL of 35% ethanol (molecular biology grade) in each well and incubating at room temperature for 60 s. 2. Addition of capture antibody (a) The wells are washed 3× with PBS (250 μL per well) using a multichannel pipette. Remove any remaining PBS by decanting the plate and blotting it against a clean paper towel (see Note 3). (b) Once the plates are primed immediately (see Note 3), add 100 μL of capture antibody solution at 1:60 dilution to each well, cover, and incubate for at least 10 h at 2–8 °C. 3. Addition of mitogen/stimulant (a) Remove the capture antibody solution from each well by aspirating or by decanting the plate, blot, and then wash 3× with PBS (250 μL per well). (b) Block the membranes by adding 250 μL of blocking buffer (see Note 4) to each well and incubate the plates for 2 h at 37 °C. (c) The blocking buffer is removed by aspiration or by decanting the plates. Subsequently, the wells are washed with 1× with AIM-V medium. (d) AIM-V medium (50 μL; see Note 5) is added to each well and kept at room temperature until the addition of stimulants and peripheral blood mononuclear cells (PBMC). (e) Add the total volume of 50 μL per each well with mitogen, 5 μg/50 μL/well FCoV1 RBD, 5 μg/50 μL/well FCoV2 RBD, and 5 μg/50 μL/well SCoV2. The mitogen consists of 2 μg/50 μL/well phytohemagglutinin P (PHA-P; see Note 6) for Human ELISpot and 0.8 μg/50 μL/well concanavalin A (ConA) for Feline ELISpot (see Note 7). Each stimulant or diluent control should have at least duplicate wells. Thus, the final volume per well at this point will be 100 μL. (f) PBMC (100 μL) is added to the IL-2 or IFNγ ELISpot plate at a concentration of 2.0 × 106 PBMC per well in the AIMS-V culture medium. The final volume per well with PBMC/stimulant is 200 μL. (g) Incubate the covered plates for 48 h at 37 °C in 5% CO2 (see Note 8). 4. Addition of detection antibody (a) The wells are washed 5× with PBS (250 μL per well). After the final wash, remove any remaining liquid by decanting the plate and blotting.

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(b) Detection antibody solution (100 μL) is added at 1:50 dilution to each well and incubated for 12 h or overnight at 2–8 °C. 5. Addition of streptavidin-AP conjugate (a) Remove the detection antibody solution by decanting the plates. The wells are washed 5× with PBS (250 μL per well). After the final wash, remove any remaining liquid by inverting the plate and blotting. (b) Streptavidin-AP solution (100 μL; 1:50 dilution in reagent solution that is 1% BSA in PBS) is added to each well and incubated for 2 h at room temperature. 6. Addition of the chromogenic substrate, development, and counting (a) Wash the wells 5× with PBS (250 μL per well). After the final wash, remove any remaining liquid by inverting the plate and blotting. (b) BCIP/NBT chromogen solution (100 μL) is added to each well, covered, and incubated for 30–45 min at room temperature in the dark by wrapping the plates in foil. (c) Rinse the plates with deionized water in a plastic container. Decant and blot the plates to remove excess water. Allow the plates to dry at room temperature in the dark. (d) Once the plates are fully developed and dried, the spots are quantitated by an automatic ELISpot reader as spot forming unit (SFU) per number of PBMC, and photographs are taken by the reader. (e) Whenever there is a dark background, the machine count is taken with the help of the AID ELISPOT highresolution reader system, or otherwise, it was manually counted with the help of an ELISpot reader (Quantihub color 4.1 ELISpot reader). 3.6 Human Cytokine mRNA Analyses

1. PBMC culturing with stimulants (a) PBMC from the collected blood is prepared by using the LSM separation method (Subheading 3.4). (b) The cells should be counted with a hemocytometer and resuspend the PBMC at a concentration of 1 × 107 PBMC/mL in RPMI medium consisting of 2% heat (56 °C, 30 min)-inactivated FBS and 25 μg/mL gentamycin.

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(c) The sterile 96-well round-bottom plate is labeled next to the duplicate wells as follows: row 1 (none), row 2 (FCoV1), row 3 (FCoV2), row 4 (SCoV2), row 5 (none), row 6 (mitogen), row 7 (none), and row 8 (PBS media control). (d) Add 0.007 mL of PBS to the duplicate wells with 0.093 mL RPMI medium as PBS media control. (e) Add each RBD at 5 μg/0.007 mL/well to duplicate wells with 0.093 mL RPMI medium, resulting in a total volume of 0.1 mL (Note 7). (f) Next add 2 μg/0.007 mL/well T-cell mitogen PHA-P to the duplicate wells with 0.093 mL RPMI medium, resulting in a total volume of 0.1 mL per well (see Note 7). (g) Add to each well 0.1 mL of 1 × 107 PBMC/mL in RPMI medium, resulting in 1 × 106 PBMC per well in a total volume of 0.2 mL. (h) The plate is incubated for 18 h at 37 °C in 5% CO2 incubator. (i) Spin the plate for 1 min at peak speed of 2500 rpm, carefully remove the supernatant with multichannel pipet, add 0.1 mL of sterile RNAse-free PBS to each well, mix the duplicates together in a sterile 1.5 mL microfuge tube, micro-centrifuge the tubes at 45 s at 8000 rpm, remove the fluid by pipetting, and freeze the pellet at 80 °C. 2. RNA isolation from PBMC (a) Add 150 μL of Zymo Tri reagent to the frozen PMBC pellet and mix by vortex for 25 s. (b) Add 300 μL ethanol to the sample and vortex for 10 s. (c) Zymo RNA micro prep column tubes are labeled, and each mixture is transferred to a column tube. (d) Column tubes are centrifuged at 13,000 rpm for 1 min, and effluent is discarded. (e) The columns are washed twice with 400 μL RNA prewash buffer at 13,000 rpm for 1 min and decant. (f) Subsequently, the columns are washed once with RNA wash buffer 700 μL at 13,000 rpm for 1 min and decant. (g) Finally, the columns are dry-run without any buffer for 1 min. at 13,000 rpm to remove any residual ethanol fluid. (h) The columns are transferred to new Eppendorf tubes, RNA is eluted with 15 μL of RNase-/DNase-free water into the tube, and the columns are removed. The 15 μL of RNA in each tube is stored at -80 °C ultralow freezer.

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3. Reverse transcription using Luna script Reverse transcriptase (a) 3.5 μL RNA is transferred to 0.5 mL PCR tube and 3.5 μL of DNAse/RNAse-free water is added. (b) 0.5 μL random primer is added to the above mixture. (c) 1.0 μL Luna RT enzyme is added to the above mixture. (d) Amplify with a real-time PCR machine using the following cycling to obtain the amplified DNA. (e) The cycling condition of the reverse transcription is performed on MJ Research PCR machine. Primer annealing

25 °C for 2 min

cDNA synthesis

55 °C for 10 min

Heat inactivation

95 °C for 1 min

4. Real-time polymerase chain reaction for cDNA (a) 1.0 μL of amplified DNA is subjected to real-time polymerase chain reaction using the Sso advanced universal probes super mix. (b) The cycling condition of real-time PCR is performed on a QuantaBio PCR machine. Initial denaturation

95 °C for 3 min

Cycling (50 cycles) of

95 °C for 5 s 60 °C for 15 s 72 °C for 30 s

(c) The data depicted in the graph show the reciprocal of cT (threshold cycle), which is graphed using GraphPad Prism Software (V10; Boston, MA).

4

Notes 1. In order to quantitate mRNA levels among the stimulated samples, the mRNA amounts must be the same among all stimulated samples, even though the relative quantitation is based on the housekeeping gene GAPDH in each sample. 2. All tubes and cotton-plugged pipet tips must be sterile DNAsefree and RNAse-free to prevent contamination resulting in loss of mRNA. 3. Do not dry the plates before the capture antibody solution is fully ready. 4. Blocking buffer should be filter sterilized with a 0.2 μM filter before use. 5. AIMS-V medium should be stored at 2–8 °C, with caution not to freeze.

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6. PHA is a strong T-cell mitogen for human T cells but will not stimulate feline T cells. 7. The final concentration per mL for each stimulant is PHA (10 μg/mL), ConA (4 μg/mL), SCoV2 RBD (5 μg/mL), FCoV1 RBD (5 μg/mL), and FCoV2 (5 μg/mL). 8. Care should be taken not to move, bump, or expose the plates to vibrations that may cause the cells to move, as the assay is dependent on cells remaining in place to create a spot.

Acknowledgment This work was supported by the JKY Miscellaneous Donors Fund. J.K.Y., S.K., and B.S. are the inventors of record on a patent (US Provisional Patent: 63/373,474) held by the University of Florida and may be entitled to royalties from companies developing the commercial products related to the research described in this chapter. References 1. Yamamoto JK, Edison LK, Rowe-Haas DK et al (2023) Both feline coronavirus serotypes 1 and 2 infected domestic cats develop cross-reactive antibodies to SARS-CoV-2 receptor binding domain: its implication to pan-CoV vaccine development. Viruses 15(4):914. https://doi. org/10.3390/v15040914 2. Addie DD, Schaap IAT, Nicolson L et al (2003) Persistence and transmission of natural type I feline coronavirus infection. J Gen Virol 84 (Pt 10):2735–2744. https://doi.org/10. 1099/vir.0.19129-0 3. Roff SR, Noon-Song EN, Yamamoto JK (2014) Significance of interferon-γ in HIV-1

pathogenesis, therapy, and prophylaxis. Front Immunol 4:498. https://doi.org/10.3389/ fimmu.2013.00498 4. Henrie R, Cherniawsky H, Marcon K et al (2022) Inflammatory diseases in hematology: a review. Am J Physiol Cell Physiol 323(4): C1121–C1136. https://doi.org/10.1152/ ajpcell.00356.2021 5. Spolski R, Li P, Leonard WJ (2018) Biology and regulation of IL-2: from molecular mechanisms to human therapy. Nat Rev Immunol 18(10): 648–659. https://doi.org/10.1038/s41577018-0046-y

Chapter 10 Frequencies of SARS-CoV-2 Spike Protein-Specific Memory B Cells in Human PBMCs, Quantified by ELISPOT Assay Ilya M. Swanson, Iana H. Haralambieva, Madeleine M. Rasche, Inna G. Ovsyannikova, and Richard B. Kennedy Abstract Vaccination against SARS-CoV-2 with coronavirus vaccines that elicit protective immune responses is critical to the prevention of severe disease and mortality associated with SARS-CoV-2 infection. Understanding the adaptive immune responses to SARS-CoV-2 infection and/or vaccination will continue to aid in the development of next-generation vaccines. Studies have shown the important role of SARS-CoV-2specific antibodies for both disease resolution and prevention of COVID-19 serious sequelae following vaccination. However, antibody responses are short-lived, highlighting the importance of studying antigenspecific B-cell responses to better understand durable immunity and immunologic memory. Since the spike protein is the main target of antibody-producing B cells, we developed a SARS-CoV-2 memory B cell ELISPOT assay to measure the frequencies of spike-specific B cells after COVID-19 infection and/or vaccination. Here, we describe in detail the methodology for using this ELISPOT assay to quantify SARSCoV-2 spike-specific memory B cells produced by infection and/or vaccination in human PBMC samples. Application of this assay may help better understand and predict SARS-CoV-2 recall immune responses and to develop potential B cell correlates of protection at the methodological level. Key words B cell ELISPOT, SARS-CoV-2, PBMCs, Humoral immunity, COVID-19 vaccines

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Introduction Since its emergence in 2019, severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) has been argued to have caused one of the most impactful public health crises in recent history. Responsible for almost seven million global deaths at the time of this publication, SARS-CoV-2 has demanded the attention of research efforts from institutions worldwide [1]. Proactive disease prevention through vaccination against SARS-CoV-2 has been adopted worldwide, as the World Health Organization reports over five billion persons are fully vaccinated against SARS-CoV-2 [1]. Both innate and adaptive immunity are a critical part of the antiviral immune response against SARS-CoV-2 [2]. Our current

Alexander E. Kalyuzhny (ed.), Handbook of ELISPOT: Methods and Protocols, Methods in Molecular Biology, vol. 2768, https://doi.org/10.1007/978-1-0716-3690-9_10, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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understanding of antiviral immune defense implicates the B-cell response and the production of antigen-specific IgM, IgA, and IgG, in particular, spike-specific neutralizing IgG/IgA antibodies (Ab) as crucial for the resolution of disease and prevention of severe illness upon subsequent exposure to SARS-CoV-2, as well as for the diagnosis of viral infection (absent vaccination). Humoral immune responses to SARS-CoV-2 are mediated by antibodies that are dependent on CD4+ T cell help and directed to structural proteins, primarily the surface glycoprotein spike (S), the nucleocapsid (N), and other SARS-CoV-2 proteins [3]. Functional SARS-CoV-2specific neutralizing antibodies following infection and/or vaccination (anti-S and anti-RBD) are considered important for viral neutralization and virus clearance and are quantified using in vitro neutralization assays. For these reasons, neutralizing antibody titer is a good biomarker for protective antibody efficacy upon SARS-CoV-2 exposure and a marker for successful mounting of the humoral immune responses after vaccination [4]. In addition to the production of neutralizing antibodies, an effective immune response involves the generation of long-lived memory B cells (MBCs) that are an important component of the recall response to SARS-CoV-2 [5]. SARS-CoV-2 has multiple structural proteins with the spike glycoprotein being one of the immunodominant targets of the antibody response. The spike protein has two functional subunits: the S1 subunit containing the N-terminal domain (NTD) and the receptor-binding domain (RBD) mediating the attachment to the angiotensin-converting enzyme 2 (ACE2) receptor, and the S2 subunit, mediating the fusion of viral and cellular membranes [4]. As the SARS-CoV-2 spike antigen is the main target of neutralizing/protective antibody-producing B cells, we developed a functional memory B-cell ELISPOT assay to detect frequencies of spike-specific B cells after COVID-19 infection and/or vaccination. This assay has been successfully applied to study and longitudinally characterize memory immune responses after SARS-CoV-2 messenger RNA (mRNA) vaccination and to assess the cross-reactivity of vaccine-induced MBCs to the emerging SARS-CoV-2 variants of concerns following primary vaccination series [5]. Figure 1 illustrates the dynamics of spikespecific MBC response (S1-, S2-, RBD-, and NTD-specific MBC frequencies) overtime in a cohort of 17 healthy adults after primary vaccination series with BNT162b2 (Pfizer/BioNTech) or mRNA1273 (Moderna) vaccine [5]. Developing novel assays to reliably measure components of the recall immune response to SARS-CoV2 other than antibodies may provide additional insights into the markers associated with protection and risk of reinfection, clinical disease severity, and/or vaccine efficacy. We have previously published some of the materials and methods for this or similar ELISPOT procedures [5–12].

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S1 Wuhan-Hu-1 RBD Wuhan-Hu-1 S2 Wuhan-Hu-1 NTD Wuhan-Hu-1

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Fig. 1 SARS-CoV-2 Spike-specific memory B-cell response pre- and COVID-19 mRNA vaccination Dynamics of memory B-cell (MBC) response after coronavirus disease 2019 (COVID-19) messenger RNA (mRNA) vaccination for severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2). The frequencies of immunoglobulin G (IgG)–positive MBCs were measured using the Mabtech ELISPOTPLUS kit (enzyme-linked immunosorbent spot assay [ELISPOT]) for human IgG, using the indicated antigens (dark/light gray and white boxes), after in vitro peripheral blood mononuclear cell 3-day stimulation with human recombinant interleukin2 and R848. Detected responses are presented in spot-forming units (SFUs) per 2 × 105 cells, as subjects’ antigen-specific medians (from 3 replicates with subtracted subject-specific no-antigen background measure). The values are plotted on the log2 scale, but the scales of the axis reflect the untransformed values for easier interpretation. Each box was plotted using the interquartile range (IQR) and the median was represented by the bold line in the box. The “whiskers” extend up to 1.5x the IQR above or below the 75th or 25th percentiles, respectively. Black circles represent naive subjects (at baseline), while white circles represent COVID-19–recovered subjects (at baseline). The MBC responses and changes over time of S1/receptorbinding domain (RBD)–specific MBC response and S2/N-terminal domain (NTD) MBC response are compared using the Wilcoxon signed-rank test and reported in the Results. (Printed with permission from Journal of Infectious Diseases [5])

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Materials

2.1 SARS-CoV2 Spike-Specific Recombinant Proteins

The following SARS-CoV-2 Wuhan-Hu-1 purified recombinant proteins (Sino Biologicals, Beijing, China) were used for this assay: 1. SARS-CoV-2 Spike S1-His recombinant protein (cat. number 40591-V08H). 2. SARS-CoV-2 Spike S2 ECD-His recombinant protein (cat. number 40590-V08H1). 3. SARS-CoV-2 Spike RBD-His recombinant protein (cat. number 40592-V08H). 4. SARS-CoV-2 Spike S1 NTD-His and AVI recombinant protein (cat. number 40591-V49H).

2.2 Collection and Isolation of Peripheral Blood Mononuclear Cells (PBMCs)

1. HISTOPAQUE-1077. 2. Accuspin™ tube. 3. 1x sterile PBS. 4. ACK lysis buffer. 5. Cell strainers. 6. 0.4% Trypan blue. 7. Hemacytometer. 8. RPMI freezing medium: RPMI 1640 with L-glutamine supplemented with 20% FCS and 10% DMSO. 9. 1.8 mL cryogenic freezing tubes. 10. Sterile Pasteur pipettes.

2.3 Thawing of Cryopreserved PBMCs

1. 15 mL sterile conical centrifuge tubes. 2. RPMI culture medium supplemented with DNAse: RPMI 1640 with L-glutamine supplemented with 10% FCS, 100 U/mL penicillin-100 μg/mL streptomycin, 1 mM sodium pyruvate, and 10 μg/mL DNase. 3. RPMI culture medium: RPMI 1640 with L-glutamine supplemented with 5% FCS, 100 U/mL penicillin-100 μg/mL streptomycin, and 1 mM sodium pyruvate. 4. Cell strainers. 5. 50 mL sterile conical centrifuge tubes. 6. 0.4 % Trypan blue. 7. 1x sterile PBS.

2.4 Resting PBMCs in the Presence of IL2 and R848

1. 24-well sterile tissue culture plates. 2. Recombinant human IL-2 (rhIL-2) and R848 are provided as reagents for B-cell prestimulation in the Mabtech ELISPOTPLUS kit for human IgG (Mabtech Inc., Cincinnati, OH).

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3. RPMI culture medium: RPMI 1640 with L-glutamine supplemented with 5% FCS, 100 U/mL penicillin-100 μg/mL streptomycin, and 1 mM sodium pyruvate. 4. 0.4% Trypan blue. 5. 1x sterile PBS. 2.5 Coating of ELISPOT Plates

1. Millipore Immobilon-P-membrane multiscreen filter plate. 2. Purified recombinant SARS-CoV-2 Wuhan-Hu-1 proteins (Sino Biologicals, Beijing, China; see Subheading 2.1). 3. Anti-human total IgG capture antibody: mAb MT91/145 provided in the Mabtech ELISPOTPLUS kit for human IgG. 4. PBS, pH 7.4.

2.6 Human B-Cell ELISPOT Assay Setup: Plating and Culture

1. Mabtech ELISPOTPLUS kit for human IgG (Mabtech Inc., Cincinnati, OH, product Code: 3850-2H). 2. RPMI culture medium: RPMI 1640 with L-glutamine supplemented with 5% FCS, 100 U/mL penicillin-100 μg/mL streptomycin, and 1 mM sodium pyruvate. 3. 0.25% Trypsin-EDTA. 4. 0.4% Trypan blue. 5. 1x sterile PBS. 6. Aluminum foil.

2.7 Assay Development

1. Mabtech ELISPOTPLUS kit for human IgG (Mabtech Inc., Cincinnati, OH, product Code: 3850-2H). 2. Tetramethylbenzidine (TMB) substrate solution. 3. Automated ELISPOT reader or stereomicroscope.

2.8 Determination of Memory B-Cell Frequencies/MBCs (in SFUs per 2 × 105 Cells)

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1. An automated scanning/counting on an ELISPOT reader: ImmunoSpot® S6Macro696 Analyzer (Cellular Technology Ltd., Cleveland, OH, USA) with the ImmunoSpot® version 5.1 software or manually using a stereomicroscope.

Methods The following methods describe the steps required to detect and quantify SARS-CoV-2 spike-specific MBCs and total IgG-expressing B cells in human PBMCs. Some of the methods and procedures for this or similar ELISPOT assays have been previously published [5–12].

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3.1 Collection and Isolation of Peripheral Blood Mononuclear Cells (PBMCs)

The protocol is based on the manufacturer’s procedure for separating PBMCs using Accuspin™ tubes and has been previously published [8]. Blood is collected in tubes treated with anticoagulant (heparin or EDTA) to prevent coagulation. 1. Warm HISTOPAQUE-1077 to room temperature using a 37 °C water bath. Keep HISTOPAQUE-1077 out of direct light. 2. Pipet 15 mL of HISTOPAQUE-1077 into the upper chamber of each Accuspin™ tube. 3. Centrifuge tubes at 800× g for 30 s to move HISTOPAQUE1077 into the lower chamber of the tube. 4. Gently pipet whole blood into the upper chamber of the tube (15–20 mL/tube, see Note 1). 5. Add sterile 1x PBS into the tube up to 45 mL. 6. Gently mix blood and PBS, being careful not to drive blood below the frit. 7. Centrifuge tubes at 1000× g for 15 min at 25 °C with the centrifuge brake OFF. 8. After centrifugation, carefully remove approximately half of the plasma layer using a sterile Pasteur pipette. Do not disturb the buffy coat (white layer) of PBMCs located above the frit. 9. Using a sterile Pasteur pipette, carefully remove the layer of PBMCs (the white-hued layer directly above the frit) and transfer it to a 15 mL sterile conical centrifuge tube. 10. To wash cells, add 1x sterile PBS to PBMCs, bringing the volume of liquid in the 15 mL conical centrifuge tube up to the 10 mL mark. 11. Resuspend PBMCs by inverting the tube several times. 12. Centrifuge at 500× g for 10 min at 25 °C with brake ON. 13. Remove supernatant without disturbing the cell pellet. 14. Add 5 mL of ACK lysis buffer to the cell pellet. Resuspend cells by pipetting cell suspension up and down. 15. Allow cells to incubate at room temperature for 5 min in the ACK lysis buffer. 16. Add 1x sterile PBS to the cells + ACK lysis buffer, to bring the volume of liquid in the 15 mL conical centrifuge tube up to the 10 mL mark. 17. Centrifuge at 500× g for 10 min at 25 °C with brake ON.

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18. Remove supernatant without disturbing the cell pellet, then resuspend pellet in 5 mL of 1x sterile PBS. 19. Place a cell strainer on top of a 50 mL conical centrifuge tube. Transfer the cell suspension from the 15 mL conical centrifuge tube to the 50 mL conical centrifuge through the cell strainer. 20. To count the number of live and dead cells, place 200 μL of 1x PBS, 37.5 μL of Trypan blue, and 12.5 μL of cell suspension into a 5 mL falcon tube; mix well, then fill a hemacytometer with 10 μL of sample. Count and record the number of unstained (live) cells in the outer four quadrants of the hemocytometer. 21. Total number of cells = number of live cells/4 × 10,000 × 20 (dilution factor [250/12.5]) × total volume of cells (5 mL or pooled total). 22. Centrifuge cell suspension at 500× g for 10 min at 25 °C with brake ON. 23. Adjust cell concentration to 1 × 107 cells/mL with cold (4 °C) RPMI freezing medium (see Note 2). 24. Aliquot 1 mL of cell suspension into prelabeled cryogenic freezing tubes. 25. Place cryogenic freezing tubes into a -80 °C freezer in a controlled-rate freezing container overnight. 26. Transfer cells into a liquid nitrogen storage tank for long-term storage (see Note 3). 3.2 Thawing of Cryopreserved PBMCs [8]

Our protocol for the detection of SARS-CoV-2 antigen-specific IgG MBCs by ELISPOT is based on using cryopreserved PBMCs. This method allows testing of previously stored samples in larger batches (i.e., the method is more convenient for larger studies), thus minimizing assay drift/variability and batch effects. Alternatively, freshly isolated PBMCs can be also used in the ELISPOT assay. 1. Warm RPMI culture medium supplemented with DNAse in a 37 °C water bath for a minimum of 15 min. 2. Add 100 μL of RPMI culture medium supplemented with DNAse into a 15 mL conical centrifuge for each sample being thawed. 3. Remove one vial of PBMCs (cell concentration 1 × 107) for each sample from liquid nitrogen storage tank. 4. Rapidly thaw PBMCs stored in cryogenic freezing tubes using a 37 °C water bath by swirling the vial in the water bath until a small amount of ice remains. 5. Quickly wipe the vial with 70% ethanol and place it in a sterile tissue culture hood.

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6. Pipette each sample from the cryogenic freezing tube into a 15 mL conical centrifuge tube containing 100 μL of RPMI culture medium supplemented with DNAse. 7. Mix the cells and medium by gently shaking the 15 mL conical centrifuge tube. 8. Slowly add 500 μL of RPMI culture medium supplemented with DNAse while swirling the tube gently to mix the cells and medium together. 9. In 1 min, add double the amount (1 mL) of RPMI culture medium supplemented with DNAse to the cell suspension in the 15 mL conical centrifuge tube. 10. Continue adding double the amount of RPMI culture medium supplemented with DNAse every minute until the cell suspension reaches a final volume of 10 mL. 11. Cap each conical tube and invert it 5x to mix the cells; do not vortex cell suspension. 12. Centrifuge at 300× g for 7 min at 25 °C with brake ON. 13. Remove supernatant, then resuspend cells in 10 mL of RPMI culture medium supplemented with DNAse. 14. Cap each conical tube and invert it 5x to mix the cells; do not vortex cell suspension. 15. Incubate cells at 37 °C for 20 min by placing the 15 mL conical centrifuge tubes in a 37 °C water bath. Invert tubes once for 10 min into the incubation period. 16. After 20-minute incubation, place cells on ice for 7 min. 17. Centrifuge cells at 300× g for 7 min at 4 °C with brake ON. 18. Carefully remove all supernatant and resuspend cells in 1 mL of RPMI culture medium supplemented with 5% FCS. 19. Place a cell strainer on top of a 50 mL conical centrifuge tube. Transfer the cell suspension from the 15 mL conical centrifuge tube to the 50 mL conical centrifuge through the cell strainer (see Note 4). 20. To count the number of live and dead cells, place 200 μL of 1x PBS, 37.5 μL of trypan blue, and 12.5 μL of cell suspension into a falcon tube. Mix well and fill a hemacytometer with 10 μL of sample. Count and record the number of unstained (live) cells in the outer four quadrants of the hemocytometer. 21. Total number of cells = number of live cells/4 × 10,000 × 20 (dilution factor [250/12.5]) × total volume of cells (1 mL or pooled total). 22. Adjust the cell concentration to 2 × 106 cells/mL by adding RPMI culture medium supplemented with 5% FCS.

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3.3 Resting PBMCs in the Presence of IL2 and R848 (Nonspecific Stimulation of B Cells) [5–7]

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Antigen-specific MBCs have relatively low frequencies if measurement is carried out years after antigenic stimulation (vaccination/ infection). Protocols for MBC quantification using ELISPOT assay depend on in vitro nonspecific prestimulation of B cells to boost cell proliferation/activation of MBC. One of the most efficient ways for MBC prestimulation reported was a combination including the Toll-like receptor (TLR) agonist R848 and recombinant human interleukin (IL)-2 (rhIL-2) [13]. This prestimulation is recommended as per the manufacturer’s specifications for the Mabtech ELISPOTPLUS kit for human IgG, and rhIL-2 and R848 are provided as reagents in the kit. Investigators interested in quantitating the plasmablast response should omit this step. 1. Add 2 mL of PBMC suspension in RPMI with 5% FCS (from a concentration of 2 × 106 cells/mL) into each well of a 24-well sterile tissue culture plate (final 4 × 106 cells/well) (see Note 5). 2. Add 20 μL of rhIL-2 to each well to obtain 10 ng/mL final concentration (see Note 6). 3. Add R848 to obtain 1 μg/mL final concentration (i.e., add 2 μL of R848 to each well of 2 mL). 4. Incubate plate with cells at 37 °C in a 5% CO2 humidified incubator for 72 h.

3.4 Coating of ELISPOT Plates

The antigen coating of plates should be performed the day before assay setup. 1. Dilute SARS-CoV-2 recombinant proteins to a concentration of 2 μg/mL in phosphate-buffered saline (PBS, pH 7.4). 2. Dilute the anti-human total IgG capture mAb MT91/145 at 15 μg/mL (optimal coating is achieved with 1.5 μg per well; for antibody concentration/dilution, refer also to the insert of the Mabtech ELISPOTPLUS kit for human IgG). Include four antihuman total IgG wells for each subject. 3. Coat the wells of a Millipore Immobilon-P-membrane multiscreen filter (PVDF) 96-well plate with the diluted SARS-CoV2 recombinant proteins at 100 μL/well (0.2 μg SARS-CoV2 antigen/well). Design the plate map to include at least a quadruplicate for each antigen-specific assessment. Include control wells (e.g., four wells) coated with PBS, pH 7.4 only, to represent a subject-specific background measure (negative control). 4. Seal the plate with the appropriate plate sealer and incubate overnight at 4 °C.

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3.5 Human B-cell ELISPOT Assay Setup: Plating and Culture

1. After the prestimulation step, remove medium from wells and pool all media/cells from one subject into a single 15 mL conical centrifuge tube (or one 50 mL tube if more cells). 2. Add 0.5 mL of prewarmed (see Note 7) 0.25% trypsin-EDTA to each well. 3. Place plate in a 37 °C in a 5% CO2 humidified incubator for approximately 10 min until cells detach. Confirm detachment using a microscope. 4. Remove cells/trypsin suspension from each well and add to the corresponding 15 mL conical centrifuge tube that contains medium/cells harvested from the same wells. 5. Repeat the procedure. Add another 0.5 mL of prewarmed 0.25% trypsin-EDTA to each well, incubate the plate for 10 min in 37 °C in a 5% CO2 humidified incubator, and harvest the cells/trypsin suspension. 6. Add 0.5 mL of RPMI culture medium supplemented with 5% FCS to each well and mix by pipetting up and down. Pool suspension from each well and add to the corresponding 15 mL conical centrifuge tube that contains cells harvested from the same well/wells (subject). 7. Bring the volume of each 15 mL conical centrifuge tube up to 10 mL by adding RPMI culture medium supplemented with 5% FCS. 8. Centrifuge cells at 300× g for 7 min at 4 °C with the centrifuge brake ON. 9. Remove supernatant without disturbing the cell pellet, then resuspend cell pellet in 0.5 mL of RPMI culture medium supplemented with 5% FCS. Keep cells on ice. 10. Count the number of live and dead cells by placing 200 μL of 1x PBS, 37.5 μL of trypan blue, and 12.5 μL of cell suspension into a falcon tube. Mix well and fill a hemacytometer with 10 μL of sample. Count and record the number of unstained (live) cells in the outer four quadrants of the hemocytometer. 11. Total number of cells = number of live cells/4 × 10,000 × 20 (dilution factor [250/12.5]) × total volume of cells (0.5 mL). 12. Adjust the cells to the desired concentration (2 × 106 cells/ mL) by adding RPMI culture medium supplemented with 5% FCS (see Note 8). 13. Keep cells on ice until they are ready to be plated (see Note 9). 14. One hour before plating the cells, remove the precoated B-cell ELISPOT plate from 4 °C (cold room) in order to perform the blocking step. 15. For the blocking step, first discard the coating reagents. Wash 5x with sterile PBS (250 μL/well) to remove excess antibody/

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coating reagents. Add 200 μL/well-blocking solution (RPMI with 10% FCS) and incubate for 1 h at room temperature (in a biosafety hood). 16. Remove the blocking solution and pat the plate dry on paper towels until there is no media remaining in the ELISPOT plate (in a biosafety hood). 17. Add aliquots of cells (200,000 cells/well) from 2 × 106 cells/ mL cell suspension into SARS-CoV-2 antigen-coated wells (e.g., four wells per subject) and into the empty (no antigen) cell controls (four wells) (see Notes 8 and 10). 18. Perform 1:10 additional dilution of the cell suspension by adding 900 μL of media to 100 μL of cells. Add 50 μL cells (10,000 cells/well) into the total IgG-coated wells (e.g., four wells per subject). Add 50 μL of media (RPMI with 5% FCS) in the total IgG-coated wells to bring volume to 100 μL per well (see Notes 8 and 10). 19. Cover the plate with aluminum foil and place in 5% CO2 incubator at 37 °C for 24 h. Do not disturb the cells during the incubation period (see Note 11). 3.6 Assay Development (See Note 12)

1. For assay development, refer also to the manufacturer’s protocol (Mabtech). 2. After the 24-hour incubation, remove the media from the ELISPOT plate. 3. Using a 50–300 μL multichannel pipette, wash the plate 5x with sterile PBS (250 μL/well, allow the wells to soak for 1–2 min at each wash step). 4. Remove the wash buffer by “flicking” the plate. Pat the plate dry on paper towels between washes. 5. Dilute the detection biotinylated mAb MT78/145 to 1 μg/ mL in PBS-0.5% FBS (20 μL in 10 mL). Add 100μL/well and incubate the plate for 2 h at room temperature. 6. Remove the detection antibody by “flicking” the plate in the sink. Pat the plate dry on paper towels. 7. Wash wells 5x with sterile PBS (250 μL/well), as described above. 8. Before use, dilute the streptavidin-HRP enzyme conjugate (dilution 1:1,000) by adding 10 μL of streptavidin-HRP concentrate into 10 mL of dilution buffer (PBS-0.5% FBS). Add 100 μL of diluted streptavidin-HRP into each well and incubate for 1 h at room temperature (see Note 13). 9. Wash wells 5x with sterile PBS (250 μL/well), as described above.

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10. Add 100 μL of substrate solution (TMB substrate) to each well. Incubate the plate in the dark for 5–20 min at room temperature (see Note 14). 11. Remove the chromogen by “flicking” the plate into sink. Invert the plate and blot dry on paper towels. 12. Rinse the ELISPOT plate 3x with deionized water. Remove the plastic drain and rinse the bottom of the plate. Invert the plate and blot dry on paper towels. Wipe the bottom of the plate dry with paper towels. 13. Invert the plate and allow it to dry overnight in the dark. Once the plate is dry, cover it with the original lid. 3.7 Determination of MBC Frequencies (in SFUs per 2 × 105 Cells)

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Once the microplate is completely dry, the spots per well can be analyzed/counted using preoptimized counting parameters on an automated ELISPOT reader, or manually using a stereomicroscope (see Note 15). The results are presented in spot-forming units (SFUs) per 2 × 105 cells as subjects’ medians (i.e., median of SARS-CoV-2 antigen-specific response, measured in quadruplicate) or means. Alternatively (or in addition), the results for antigen-specific IgG MBCs can be presented as % of total IgG-secreting B cells (see Note 16).

Notes 1. Do not add more than 20 mL of whole blood into the Accuspin™ tube in order to achieve good cell separation. 2. Required volume (mL) freezing medium = 1 × 107/total number of cells. 3. Keep the cells on dry ice during handling to prevent thawing. 4. At this point, if you have multiple tubes for one subject/ sample, then they should be pooled into one 50 mL conical centrifuge tube before cell counting. 5. If any cells remain (with a volume less than 2 mL) after the 2 mL/well addition, add remaining cells to a new well and bring the volume (per well) up to 2 mL with RPMI culture medium supplemented with 5% FCS. 6. The rhIL-2 provided in the Mabtech ELISPOTPLUS kit for human IgG should be reconstituted in advance with 1 mL PBS to obtain 1 μg/mL concentration. The stock solution should be used immediately or stored in aliquots at -20 °C. 7. Warm 0.25% Trypsin-EDTA in a 37 °C water bath for a minimum of 15 min prior to usage. 8. The desired cell concentration for this assay is 2 × 106 cells/mL for plating the SARS-CoV-2 protein-coated wells of the

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ELISPOT plate, and 2 × 105 cells/mL for plating the total IgG-coated wells of the ELISPOT plate. Alternatively, higher or lower cell concentrations can be used for the detection of the antigen-specific MBCs, depending on the expected frequency of these cells. 9. Cells should be plated within 30 min after counting in order to maintain optimal viability. 10. Mix the cells well while pipetting; ensure the cells are in a homogenous suspension. 11. Wrapping microplate in aluminum foil during incubation reduces well-to-well variability. 12. The assay setup requires work in a biosafety hood; however, assay-development steps do not require aseptic conditions and can be performed outside of the hood. 13. During the incubation, warm the chromogen to room temperature. 14. Five to 20 min is usually sufficient for plate development, but longer incubation periods can be used if needed. 15. In larger studies, quality control (QC) needs to be performed by a single operator (visual checking for abnormalities/errors and QC of the plates on the ELISPOT reader) to ensure uniform QC and consistency of results. The scanning and counting of multiple plates (in larger studies) need to be performed with set parameters/gating of the ELISPOT reader for all plates. 16. In larger studies, statistical analysis should be used to assess assay drift, inter-operator differences, and study design variables (e.g., shipment of samples, time of year assay was performed), which could cause bias, distribution of raw data, etc.

Acknowledgments We would like to thank the Mayo Clinic Vaccine Research Group personnel for their technical assistance during the development and execution of these assays. Conflict of Interest Statement Dr. Ovsyannikova holds patents related to vaccinia and measles peptide vaccines. Drs. Kennedy and Ovsyannikova hold a patent related to vaccinia peptide vaccines. Drs. Kennedy and Ovsyannikova have received grant funding and royalties from ICW Ventures for preclinical studies on a peptidebased COVID-19 vaccine. Drs. Kennedy, Ovsyannikova, and Haralambieva hold a patent related to the impact of single nucleotide polymorphisms on measles vaccine immunity. Dr. Kennedy has

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received funding from Merck Research Laboratories to study waning immunity to mumps vaccine. Dr. Kennedy also offers consultative advice on vaccine development to Merck & Co. and Sanofi Pasteur. These activities have been reviewed by the Mayo Clinic Conflict of Interest Review Board and are conducted in compliance with the Mayo Clinic Conflict of Interest policies. This research has been reviewed by the Mayo Clinic Conflict of Interest Review Board and was conducted in compliance with the Mayo Clinic Conflict of Interest policy.

References 1. World Health Organizaion (WHO) Coronavirus (COVID-19) Dashboard May 2023 [cited 2023 May 3] 2. Poland GA, Ovsyannikova IG, Kennedy RB (2020) SARS-CoV-2 immunity: review and applications to phase 3 vaccine candidates. Lancet 396(10262):1595–1606 3. Castro Dopico X, Ols S, Lore´ K et al (2022) Immunity to SARS-CoV-2 induced by infection or vaccination. J Intern Med 291(1): 32–50 4. Qi H, Liu B, Wang X et al (2022) The humoral response and antibodies against SARS-CoV2 infection. Nat Immunol 23(7):1008–1020 5. Haralambieva IH, Monroe JM, Ovsyannikova IG et al (2022) Distinct homologous and variant-specific memory B-cell and antibody response over time after SARS-CoV-2 mRNA vaccination. J Infect Dis 226(1):23–31 6. Haralambieva IH, Ovsyannikova IG, Kennedy RB et al (2018) Detection and quantification of influenza A/H1N1 virus-specific memory B cells in human PBMCs using ELISpot assay. Methods Mol Biol 1808:221–236 7. Monroe JM, Haralambieva IH, Warner ND et al (2022) Longitudinal antibody titer, avidity, and neutralizing responses after SARSCoV-2 infection. Heliyon 8(11):e11676

8. Umlauf BJ, Pinsky NA, Ovsyannikova IG et al (2012) Detection of vaccinia virus-specific IFN-g and IL-10 secretion from human PBMC and CD8+ T cells by ELISPOT. In: Kalyuzhny AE (ed) Handbook of ELISPOT. Springer, pp 199–218 9. Ryan JE, Ovsyannikova IG, Poland GA (2005) Detection of measles virus-specific interferongamma-secreting T-cells by ELISPOT. Methods Mol Biol 302:207–218 10. Salk HM, Haralambieva IH, Ovsyannikova IG et al (2013) Granzyme B ELISPOT assay to measure influenza-specific cellular immunity. J Immunol Methods 398-399:44–50 11. Painter SD, Haralambieva IH, Ovsyannikova IG et al (2014) Detection of influenza A/ H1N1-specific human IgG-secreting B cells in older adults by ELISPOT assay. Viral Immunol 27(2):32–38 12. Haralambieva IH, Painter SD, Kennedy RB et al (2015) The impact of immunosenescence on humoral immune response variation after influenza A/H1N1 vaccination in older subjects. PLos One 10(3):e0122282 13. Jahnmatz M, Kesa G, Netterlid E et al (2013) Optimization of a human IgG B-cell ELISpot assay for the analysis of vaccine-induced B-cell responses. J Immunol Methods 391(1-2): 50–59

Chapter 11 Monitoring Memory B Cells by Next-Generation ImmunoSpot® Provides Insights into Humoral Immunity that Measurements of Circulating Antibodies Do Not Reveal Paul V. Lehmann, Zhigang Liu, Noe´mi Becza, Alexis V. Valente, Junbo Wang, and Greg A. Kirchenbaum Abstract Memory B cells (Bmem) provide the second wall of adaptive humoral host defense upon specific antigen rechallenge when the first wall, consisting of preformed antibodies originating from a preceding antibody response, fails. This is the case, as recently experienced with SARS-CoV-2 infections and previously with seasonal influenza, when levels of neutralizing antibodies decline or when variant viruses arise that evade such. While in these instances, reinfection can occur, in both scenarios, the rapid engagement of preexisting Bmem into the recall response can still confer immune protection. Bmem are known to play a critical role in host defense, yet their assessment has not become part of the standard immune monitoring repertoire. Here we describe a new generation of B cell ELISPOT/FluoroSpot (collectively ImmunoSpot®) approaches suited to dissect, at single-cell resolution, the Bmem repertoire ex vivo, revealing its immunoglobulin class/ subclass utilization, and its affinity distribution for the original, and for variant viruses/antigens. Because such comprehensive B cell ImmunoSpot® tests can be performed with minimal cell material, are scalable, and robust, they promise to be well-suited for routine immune monitoring. Key words ELISPOT, FluoroSpot, B cells, Immune monitoring, Antibodies, Antibody titers, Immune memory, Affinity

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Introduction Clinical immune diagnostics are currently confined to the detection of specific antibodies in serum and other bodily fluids. Diagnostic decisions on whether a person has been infected by, and subsequently developed immunity, e.g., to a particular virus or vaccination, and the magnitude of the ensuing immune response are presently deduced from measurements of serum antibody titers. In this chapter, we will describe why it is necessary to include memory B cell (Bmem) measurements into such considerations, and how this can be reliably accomplished with minimal cell

Alexander E. Kalyuzhny (ed.), Handbook of ELISPOT: Methods and Protocols, Methods in Molecular Biology, vol. 2768, https://doi.org/10.1007/978-1-0716-3690-9_11, © The Author(s) 2024

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material and labor by performing B cell ELISPOT/FluoroSpot, collectively “ImmunoSpot®” assays (see Note 1). There are conceptual and practical reasons why, thus far, mainstream efforts toward monitoring antigen-specific Bmem have not been sufficiently undertaken. Namely, it has largely been accepted that antibody titer measurements directly reflect Bmem frequencies. This notion has been based on the antiquated assumption that during the B cell response to antigen, long-lived Bmem and longlived plasma cells (PC) arise in a fixed ratio to each other. Thus, the feeling had been that cumbersome assays to detect Bmem utilizing fragile live cell material are unnecessary because simple measurements of stable proteins in serum reliably provide the sought-after information. However, recent advances in the understanding of B cell lineage differentiation pathways imply the requirement for a different approach. This topic has been the subject of several excellent reviews recently [1–3] and is only outlined briefly in the following section. 1.1 Differential Fate Decisions for B Cell Differentiation into the Memory vs. Plasma Cell Lineages

During the antigen-driven B cell response, within a germinal center (GC) present in the draining lymph node(s), Bmem and PC arise along differential, affinity-driven maturation pathways (Fig. 1). Briefly, when germinal center B cells (GCB) undergo proliferation and acquire somatic hypermutations (SHM) (see Note 2), daughter cells arise with mutated antigen-binding sites of their B cell antigen receptors (BCR) (see Note 3). Because SHM occurs somewhat randomly, a subset of these mutated BCR can acquire an increased affinity for the antigen, although the affinity of most BCR is either unaltered or attenuated as a result of these mutations. In subsequent steps, the progeny of GCB that gained an increased affinity for the antigen undergoes additional cycles of proliferation, SHM, and affinity-based positive selection. Multiple repetitions of this process eventually lead to the differentiation of PC, which constitutes the cellular basis of the affinity-matured antibody response. Progeny of GCB that do not meet the increasingly stringent antigen-driven affinity selection criterion (see Note 4) exit the GC and become long-lived Bmem. Bmem and PC, therefore, emerge along alternative routes driven by different selection criteria, and, subsequently, their frequencies are not necessarily linked. Consequently, one cannot expect antigen-specific serum antibody levels and Bmem frequencies to be proportional to each other. From the above understanding of antigen-driven B cell differentiation, it also follows that, while both PC and Bmem are antigen-specific (i.e., their BCR exceeds a minimal binding constant for antigen), PC (and hence secreted antibodies) constitute primarily the high affinity end of this affinity spectrum, while Bmem also encompass a lower affinity fraction. Recent interpretation of these findings even implies that PC and Bmem play fundamentally different roles in humoral immune defense [1].

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Fig. 1 Affinity-based differential differentiation of plasma- and memory B cells. In a secondary lymphoid tissue, e.g., a lymph node, a naive B cell (Bnaive) encounters the homotypic antigen presented by a follicular dendritic cell (FDC) for which its B cell receptor (BCR) has sufficient affinity to trigger activation. The B cell then presents BCR-acquired antigen-derived peptides to cognate T follicular helper (TFH) cells, resulting in the activation/polarization and proliferation of both cell types. The B cell then enters a germinal center within the lymph node, where it undergoes additional rounds of proliferation and somatic hypermutation (SHM) of its BCR. Because the latter is random, the daughter cells display BCR with a broad spectrum of affinities for the homotypic antigen. Cells endowed with high-affinity BCR for the homotypic antigen are positively selected and undergo additional cycles of proliferation, acquisition of further somatic hypermutations, and affinity-based positive selection, eventually differentiating into antibody-secreting plasma cells (PC). Daughter cells that express lower affinity BCR variants for the homotypic antigen exit the lymph node as memory B cells (Bmem). By chance cross-reactivity, however, some of these Bmem will have high affinity for heterotypic antigen variants

1.2 Serum Antibodies Reflect the First Wall of Adaptive Humoral Defense, Bmem the Second

While PC elicited during the primary immune response can secrete large amounts of antibodies, their lifespans are heterogenous and, contrary to the previous assumption that PC are long-lived, likely fall on a continuum with possibly only a fraction of them surviving long-term [4, 5]. The antibody molecules they secrete are also relatively short-lived in the body and possess half-lives of several weeks, at best (see Note 5). Thus, serum antibodies that are

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detected at any one time have been recently produced and, therefore, the maintenance of serum antibody levels depends on constant active replenishment by PC. Such antibodies in bodily fluids and their cell-bound variants (see Notes 5–7) constitute the first wall of the adaptive humoral immune defense since they are already present prior to antigen reencounter. Such antibodies (IgA and IgM) are transported across the mucosa and can confer instant host protection by preventing entry of the pathogen/antigen into the body. If the offending pathogen/antigen succeeds in crossing this interface and gains access into the body, preformed antibodies (IgG and IgM) already present in serum can still confer protection through direct neutralization. Serum antibodies also possess precipitating, opsonizing, and complement activating activity to further combat the dissemination of the pathogen/antigen. Preformed antibodies can additionally initiate inflammatory reactions through their interactions with Ig-binding receptors expressed on various immune cell populations (IgE, IgG, or IgM). Many times, however, the first wall of adaptive humoral immune defense fails to prevent reinfection. Such is the case when antibody levels either decline to sub-protective levels, or when antigenic viral variants are encountered that evade neutralization, as evidenced, e.g., in the recent COVID pandemic. Boosting the levels of specific antibodies that were already established through infection or vaccination against the previously circulating virus strain, the “homotype,” will not confer sterilizing immunity against a newly emerging antigenic variant strain, the “heterotype.” In this case, Bmem provide the second wall of adaptive humoral immune defense. Within the Bmem pool established during a preceding immune response, BCR specificities possessing a reduced affinity for the homotypic virus/antigen will be present that, by chance, have an increased affinity for the variant, heterotypic virus/antigen. As such, heterotypic antigen-reactive Bmem occur at higher frequencies than would occur within a B cell repertoire that is naive to the homotypic virus/antigen; moreover, as such Bmem have already undergone immunoglobulin (Ig) class switch recombination (CSR), they can readily engage into faster and more efficient recall responses, including potential reentry into a GC for acquisition of further affinity-enhancing SHM. While this Bmem-based second wall of adaptive humoral immunity is not able to prevent infection with a heterotypic virus, it still mediates immune protection against it by enabling the host to rapidly mount a secondarytype immune response at the first encounter with a heterotypic virus. Studying antigen-specific serum antibodies vs. Bmem, therefore, provides fundamentally different information on adaptive humoral immunity. The former reflects only on the past remnants of previously established, still deployable, but passive, and fading

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immunological memory. In contrast, studies of Bmem provide insights into the ability of an individual to actively mount secondary immune responses against homo- and/or heterotypic antigens. Thus, studying Bmem permits one to take a glance into the future. 1.3 Emerging Evidence for Serum Antibody Levels and Memory Cell Frequencies Providing Divergent Information on the Magnitude of B Cell-Mediated Immunity

Recently, we concluded a systematic study (to the best of our knowledge the first on this subject) in which we measured circulating antibody levels to SARS-CoV-2, EBV, and different seasonal influenza virus strains and compared them with the frequencies of Bmem reactive with the same antigens [6]. These ImmunoSpot®based Bmem detections were enabled by our newly acquired ability to achieve high-density antigen coating, thus reliable detection of Bmem specific for essentially any antigen ([11] and see Note 8). Representative results for the SARS-CoV-2 Spike protein are shown in Fig. 2. Notably, elevated frequencies of Spike-specific, Bmem-derived antibody-secreting cells (ASC) were detected in all of these convalescent COVID-19 donors despite variable levels of IgG antibody reactivity in the plasma at the time of sample collection. In

Fig. 2 Discordance between circulating antibody levels and Bmem frequencies in PBMC against SARS-CoV-2 Spike antigen. Correlation between SARS-CoV2 Spike (FL = full length)-specific IgG+ antibody-secreting cell (ASC) frequencies (x-axis) and plasma IgG levels (y-axis) in convalescent COVID-19 donors (n = 8). Pearson correlation analysis was performed using GraphPad Prism software on log-transformed antigen-specific ASC data and the corresponding antigen-specific IgG titers (expressed as μg/mL of IgG equivalents). Regression analysis was also performed using GraphPad Prism and the 95% confidence bands are plotted. These data are representative of similar findings reported previously for this and other viral antigens [6]

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this same study [6], discordance between antibody levels and Bmemderived ASC frequencies was also noted for the SARS-CoV2 NCAP antigen and multiple antigenically unrelated seasonal influenza strains, all of which represent respiratory viruses that cause acute, short-term infections. Furthermore, discordance between antibody levels and Bmem-derived ASC frequencies was also evidenced against the EBNA1 antigen of Epstein–Barr virus (EBV), a latent herpesvirus that occasionally reactivates [7]. 1.4 Emerging Evidence for Bmem Detection Being More Reliable for Revealing Past Antigen Exposure than Serum Antibody Measurements

It is not uncommon to find individuals who exhibit little if any seropositivity for an antigen, yet possess significant, often high, frequencies of antigen-specific, Bmem-derived ASC (Fig. 2, [6], and Kirchenbaum et al., manuscript in preparation). Among the antigens tested, SARS-CoV-2 proteins are the most informative in this regard because SARS-CoV-2 Spike and NCAP antigen-specific Bmem-derived ASC are not detected in samples from individuals cryopreserved prior to the onset of the COVID-19 pandemic (which, therefore, are verifiably immunologically naive to this virus owing to the time period in which these samples were collected). In contrast, each of the seronegative, yet Bmem-positive, donors in our study cohort had undergone PCR-verified SARSCoV-2 infections (see Note 9). In these instances, serum antibody measurements provided clearly false-negative results on the infection history of these individuals, while the presence of Bmemderived ASC reliably revealed it. We have also previously noted clearly false-negative serological assessments for human cytomegalovirus (HCMV) exposure [8]. Obtaining PBMC (peripheral blood mononuclear cells) from different FDA-approved blood banks, all of which screen their donors for HCMV seropositivity, we found that PBMC from many such HCMV-seronegative donors possessed HCMV antigen-reactive, Bmem-derived ASC in ImmunoSpot® assays following in vitro polyclonal stimulation. Such subjects also exhibited CD4+ and CD8+ T cell memory to HCMV. Thus, the presence of Bmem-derived ASC reactivity (and T cell memory) more reliably revealed the HCMV-exposed status of these donors than did serum antibody levels. In yet another independent line of investigation, studying patients with multiple sclerosis (MS), an autoimmune disease of the central nervous system (CNS), we detected neuroantigenspecific Bmem, in the absence of detectable serum antibody levels in most patients, whereas such Bmem were absent in healthy controls [9]. Collectively these findings suggest that detecting Bmem-derived ASC might be a more reliable way to diagnose past infections and possibly also autoimmune diseases than afforded presently by serum antibody measurements.

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1.5 Assessing Bmem Might Be Important, but how, and why?

Presently there are two major techniques that permit the detection of antigen-specific Bmem in blood, lymphoid tissues, and other primary cell material. One approach is based on labeling B cells with fluorescently tagged antigen(s) followed by their detection using flow cytometry; the other is by ImmunoSpot®. While the strength of the former approach is that it allows for the segregation of antigen probe-binding B cells into phenotypically distinct subsets based on their surface marker expression, it has several disadvantages compared to ImmunoSpot®. One crucial difference is the lower limit of detection of the former. While ImmunoSpot® enables measurement of a single ASC within a bulk population of cells with essentially no intrinsic lower detection limit (see Note 10), flow cytometry falls short of detecting antigen-specific B cells when they are present as low-frequency events. Bmem, however, frequently occurs in very low frequencies in PBMC (Fig. 3). Additionally, the number of PBMC required for flow cytometric detection of antigen-specific B cells is considerably higher compared to ImmunoSpot® (see Note 11). Notably, a surface staining-only approach for the identification of antigen-specific B cells does not reliably reveal the Ig class/subclass usage of ASC, while four-color ImmunoSpot® assays provide this key information with ease as part of routine Bmem frequency measurements (see Note 12). Lastly, unlike for flow cytometry, the actual wet lab implementation of B cell ImmunoSpot® assays is scalable for high-throughput analysis (see Note 13) and multi-color ImmunoSpot® analysis can be fully automated (see Chap. 5 by Karulin et al. in this volume [10]).

1.6 Affinity Coating: Enabling B Cell ImmunoSpot® Assays “to See”

With so many potential advantages in favor of ImmunoSpot® assays for the detection of Bmem, the question arises as to why this technique has not become more widely used for immune monitoring purposes. The answer is simple and practical: the original protocol (that involves direct coating of the antigen to the membrane) only works well for a rather limited set of antigens (see Note 14). Consequently, many investigators likely gave up on this assay after not being successful in their initial attempts to establish such assays for the antigen(s) of interest. Our recent introduction of affinity capture coating [11] represents a breakthrough to this end, as it provides a universal strategy for successful assay development for essentially any antigen: the membrane is first coated with an anti(His- or other) affinity tag-specific antibody followed by the addition of recombinant (His- or other) affinity-tagged recombinant antigen. In this way, low-affinity absorption of the antigen to the membrane via weak, non-specific binding forces (primarily hydrophobicity) is replaced by specific, high-affinity binding. The assay principle is depicted in Fig. 4b, and the protocol is described in detail below. This version of the B cell ImmunoSpot® assay is

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Fig. 3 Antigen-specific Bmem occur over a wide frequency range in blood. PBMC from healthy human subjects – each represented by a dot – were tested via ImmunoSpot® assays to establish the frequency of IgG+ Bmem-derived ASC specific for the panel of antigens shown: Tetanus toxoid heavy chain (TTHc), Epstein–Barr nuclear antigen (EBNA1) of EBV, gH pentamer complex (consisting of gH, gL, UL128, UL130, and UL131A proteins) of HCMV, recombinant hemagglutinin (rHA) proteins representative of A/California/2009 (H1N1), A/Texas/2013 (H3N2), and B/Phuket/2013 (Yamagata lineage) seasonal influenza vaccine strains, as well as SARS-CoV-2 Spike (full-length) and Nucleocapsid (NCAP) proteins. Serving as specificity controls for SARS-CoV2 antigens, donors whose PBMC were cryopreserved prior to widespread COVID infection and vaccination (before June 2021) are depicted as red dots, and subjects whose PBMC were frozen after June 2021 are denoted as black dots. Note, the data shown are results of testing PBMC without serial dilution at a fixed input of 3 × 105 PBMC per well, and because quantification in wells with >100 antigen-specific spot-forming units (SFU) is no longer precise, we defined the upper limit for accurate counts at 330 SFU per 106 PBMC. Values exceeding this threshold, denoted by the gray shading in the figure, require lower cell inputs for accurate quantification of antigen-specific ASC frequencies. Likewise, the lower gray-shaded region denotes low-frequency responses in which Poisson noise necessitates seeding higher PBMC numbers per well and larger numbers of replicate wells to be evaluated for accurate determination of antigen-specific ASC frequencies (see Note 10)

particularly well-suited for detecting and characterizing rare antigen-specific, Bmem-derived ASC in a test sample, whereas an alternative variant of the B cell ImmunoSpot® assay approach, which enables assessment of ASC functional affinity and is described in another chapter of this volume [12], is better suited for samples in which antigen-specific ASC are present at an elevated frequency among all ASC (see Note 15).

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Fig. 4 Principle of antigen-specific, (a) inverted vs. (b) direct B cell ImmunoSpot® assays (the latter, using the affinity capture coating approach). In (a), the PVDF membrane on the bottom of a 96-well plate is densely coated with a pan anti-Ig(G) class-specific (in this example IgG) capture antibody that will bind the ASC-secreted Ig(G) with high affinity irrespective of the ASC’s antigen specificity. In (b), the membrane is coated first with an anti-affinity tag-specific antibody (in this example anti-His) that captures the (His)-tagged antigen with high affinity. In this way, dense coating of the membrane with the antigen is accomplished. As the next step in both assay variants, the PBMC containing the ASC are plated. In (a), ASC-produced IgG is captured around each ASC that is secreting IgG and results in the formation of individual secretory footprints. In (b), only the antibody produced by antigen-specific ASC is captured on the lawn of antigen. After removal of the cells, (a) the affinity-tagged (in this example His) antigen is added at a sufficient concentration to be retained by antigen-specific secretory footprints generated by ASC producing low- or high-affinity antibody. Alternatively, after removal of cells, (b) antigen-bound antibody is visualized by biotinylated human Ig class/subclass(in this example IgG) specific detection antibodies. In (a), antigen-specific secretory footprints are then visualized using a biotinylated anti-affinity tag detection reagent, which is revealed by the addition of a fluorescently conjugated streptavidin (FluoroSpot, as shown) or via an enzymatic reaction (ELISPOT, not shown). In (b), antigen-specific secretory footprints are revealed by the addition of a fluorescently conjugated streptavidin (FluoroSpot, as shown) or via an enzymatic reaction (ELISPOT, not shown). In both B cell ImmunoSpot® assays, counting the spot-forming units (SFU) per well reveals the number of antigen-specific ASC within the PBMC plated. Furthermore, spot morphologies in such antigen-specific ImmunoSpot® assays also provide insights into the functional affinities of the antibody secreted by the individual ASC for the antigen, a topic covered in detail in the chapter by Becza et al., in this issue [12] 1.7 High-Content Information Provided by Assessing Individual ASC via ImmunoSpot® vs. Serum Antibody Measurements

Owing to the single-cell resolution afforded by ImmunoSpot® assays, they are ideally suited to study individual ASC that comprise an antigen-specific B cell response. While traditional serum antibody measurement techniques readily perceive large increases in antigen-specific antibody titers, they fail to appreciate more subtle changes in antibody levels, especially when the abundance of antibodies is very small, e.g., in the context of allergen-specific IgE [13], or when an elevated level of preexisting antibody reactivity is already present, e.g., in the context of studying seasonal influenza vaccine responses [14, 15]. In contrast, B cell ImmunoSpot® assays circumvent ambiguity in both cases through quantifying the precise number of cells that are actively secreting antigen-specific antibody and thus offer increased sensitivity and resolution. When optimal antigen-coating conditions exist in the direct B cell ImmunoSpot® assay, each secretory footprint will be composed of antibody originating from a single ASC and thus permits assessment of antibody affinity similar to that achieved through studying an individual monoclonal antibody (mAb). In this setup,

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the morphology of an ASC-derived secretory footprint is primarily defined by the affinity of the secreted antibody for the membranebound antigen. In agreement with prior computational modeling [16], we routinely observe variable spot morphologies in such antigen-specific B cell ImmunoSpot® assays [6, 11]. Importantly, beyond enumeration of such antigen-specific secretory footprints, a multitude of morphological features including metrics of intensity and size are also captured for each spot-forming unit (SFU) and are readily exported as flow cytometry standard (FCS) files by the ImmunoSpot® software. Such FCS files can be leveraged to visualize variable spot morphology, as was done using scatter plots in our recent publications [6, 11]. Here, we direct the reader to Chap. 5 by Karulin et al. in this volume that is dedicated to high-content analysis of spot morphologies [10]. Additionally, in the context of an inverted B cell ImmunoSpot® assay (see Note 16) in which ASC-derived secretory footprints are efficiently captured irrespective of antigen specificity, secretory footprints originating from individual antigen-specific ASC are revealed by their ability to retain an antigen probe. While also briefly introduced below, Chap. 13 in this volume by Becza et al. [12] describes in detail how the inverted ImmunoSpot® assay variant and high-content data analysis enable assessment of the functional affinity distribution present in a polyclonal population of ASC, such as the ASC response elicited following COVID-19 vaccination. 1.8 Frequency of Antigen-Specific, Bmem-Derived ASC Reflect Memory Potential

Because ImmunoSpot® assays detect the secretory footprints of individual ASC, frequency information is revealed by counting the numbers of antigen-specific SFU, either per cells plated per well, or, better, as the frequency of antigen-specific ASC secreting a given Ig class/subclass among all ASC producing that Ig class/subclass (see Note 16). In either case, however, the crowding of secretory footprints along with the ELISA effect (see Note 17) can lead to undercounting of secretory footprints [6]. Systematically studying this phenomenon previously, we found that there exists a range in which cell numbers plated per well are directly proportional to the number of SFU detected. Depending on the nature of the assay and morphology of the resulting SFU, however, the corresponding SFU counts measured at higher cell inputs may break down at approximately 100–200 SFU per well (see Note 18). This is a major concern for establishing accurate frequencies of antigenspecific, Bmem-derived ASC, especially in light of our previous data demonstrating that frequencies may span orders of magnitude for the same antigen in different individuals (Fig. 2 and [6]). Also, within any single individual, the frequency of Bmem-derived ASC specific for different antigens can span a similarly wide distribution [6]. Furthermore, even the number of ASC producing a given Ig class/subclass, irrespective of antigen specificity, exhibits considerable interindividual variation. ELISA effects in B cell

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ImmunoSpot® assays can also jeopardize high-content analysis of spot morphologies. A simple and efficient solution to this problem is to serially dilute the cell input to establish the linear range of SFU counts from which the accurate frequency of ASC can be extrapolated by linear regression ([6] and see Note 19). Karulin et al., Chap. 5 in this volume [10], introduce software for automatically establishing frequencies from such serial dilution experiments making this approach suitable for high-throughput workflows. 1.9 Defining the Ig Class/Subclass Usage of Antigen-Specific Bmem

The different Ig classes and subclasses are endowed with distinct effector functions and each contributes nonredundant roles toward maintaining host defense (see above and reviewed in greater detail in [17]). During the primary immune response, B cells can transition from IgM-expressing naive B cells into effector cells (PC) and Bmem that have undergone class switch recombination (CSR) [1]. CSR is an irreversible process involving the excision of DNA encompassing the exons of the Igμ heavy chain required for expression of IgM and juxtaposition of the upstream variable region genes (VDJ; which jointly define the antigen specificity of the BCR or secreted antibody) with downstream exons encoding alternative Ig classes or IgG subclasses [18]. Class switching of the BCR to downstream Ig classes or IgG subclasses is an instructed process and can be influenced by the cytokine milieu and co-stimulation provided by CD4+ T helper cells. For the latter, the differentiation of naive CD4+ T cells into different T helper cell classes (Th1, Th2, Th17, etc.) is defined by the circumstances of antigen encounter, in particular by Toll-like receptor (TLR)-derived signals [19, 20]. Triggering the “appropriate” type of T helper cells capable of stimulating the optimal Ig class usage during an infection or following vaccination is vital to successful host defense and the avoidance of collateral immune-mediated pathology (reviewed in [21]). The same applies to reinfection with the same pathogen. Upon antigen reencounter and subsequent reactivation, Bmem rapidly differentiates into PC that secrete the same Ig class/subclass expressed by the parental Bmem. Therefore, detecting the Ig class/ subclass that Bmem-derived ASC produced in B cell ImmunoSpot® assays permits the prediction of the specific types of antibodies that will be produced following the next antigen encounter (see Notes 20–22). Learning about the full spectrum of Ig classes/subclasses that the antigen-specific Bmem repertoire will produce when the antigen is reencountered is thus essential for predicting the protective efficacy of future antibody responses to an antigen. Moreover, such B cell ImmunoSpot® assays may also predict the likelihood of antibody-mediated complications occurring upon antigen re-exposure (as illustrated by the example of antibody-dependent enhancement leading to exacerbation of clinical disease following secondary dengue infection [22]). Importantly, the frequency of antigen-specific Bmem capable of secreting different Ig classes or

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IgG subclasses can be orders of magnitude apart [6] (and see also Yao et al., Chap. 15 in this volume [23]). Thus, it is not only recommended to assess all antigen-specific Ig classes/subclasses, but also to do so over a wide frequency range for each. The latter can be readily accomplished by combining the serial dilution strategy described above with four-color ImmunoSpot® analysis (for details, see the Chap. 15 of Yao et al. in this volume, [23]). 1.10 Assessing the Affinity Distribution of the Antigen-Specific Bmem Repertoire

As described above in this chapter, and briefly recapitulated here, SHM leads to the diversification of Bmem cells that previously participated in a GC reaction [24]. Upon reencounter with the same (homo-) or a modified (heterotypic) antigen, previously generated Bmem endowed with high-affinity BCR for the rechallenging antigen will readily differentiate into PC that are capable of rapidly increasing antibody titers [3]. Moreover, Bmem can also be re-recruited into a GC following antigen reencounter, where they will undergo further rounds of proliferation and acquire additional SHM necessary to refine and improve their BCR affinity for the offending antigen. Again, the GCB cell progeny endowed with the highest affinity BCR for the eliciting antigen will be selected for PC differentiation and ideally will enter the potentially long-lived fraction of the PC compartment. Thus, a Bmem repertoire that entails a higher frequency of high-affinity B cells specific for the (homo- or heterotypic) antigen can generate a faster and more robust anamnestic antibody response upon antigen (re)encounter. Thus far, the study of BCR/antibody affinity for a particular antigen has largely been confined to the generation of B cell hybridomas and/or paired IgH/IgL sequencing followed by expression and purification of individual mAb in order to establish their specificities and affinity using surface plasmon resonance or biolayer interferometry. While these approaches are the gold standard for studying individual mAbs (i.e., single B cells), it would require generation and subsequent characterization of hundreds of such mAbs to appreciate the underlying affinity distribution of antigenspecific Bmem repertoire in just one donor, at a single timepoint, and against only a single antigen of interest. While such an exercise is, in theory at least, conceptually possible for a very limited number of individuals, it would be inconceivable both in magnitude and cost to attempt such an objective for a larger donor cohort, e.g., as part of a clinical trial. Consequently, this traditional method for characterizing the affinity distribution of the antigen-specific B cell repertoire is not realizable for high-throughput immune monitoring efforts. The Chap. 13 in this volume by Becza et al. [12] demonstrates that the affinity distribution information of the antigen-specific B cell repertoire can be established with ease, and in a readily scalable manner, using two B cell ImmunoSpot® assay variants. For both, to characterize the affinity distribution of, e.g., vaccine-

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elicited IgG+ ASC, a first experiment would need to be performed that establishes the frequency of such antigen-specific IgG+ ASC using the serial dilution approach (see Note 23). This information is essential for being able to seed in a follow-up test the PBMC at a “Goldilocks” cell density in which the individual secretory footprints can be studied without interfering with each other. In this second experiment, using an independent aliquot of cryopreserved cell material, the donor PBMC are seeded at the so-called “Goldilocks number,” which is assay dependent and between 50 and 100 SFU per well. As each SFU represents the secretory footprint of an individual ASC, at 50 SFU per well, the secretory footprints of 50 distinct ASC would be assessed. Seeding additional replicate wells at this “Goldilocks number” would increase the number of antigen-specific IgG+ ASC being characterized. We recommend to study a fixed number of ≥300 ASC for each antigen concentration as this number gives a considerable (and representative) sample size for assessment of the antigen-specific Bmem repertoire elicited in a particular person, against an individual antigen, at the time of sample collection. Leveraging the inverted ImmunoSpot® approach for B cell affinity distribution measurements, the soluble antigen probe is added in decreasing concentrations to the same number of replicate wells for each antigen concentration, whereby each well was previously seeded with a “Goldilocks number” of cells. At the highest antigen probe concentration, the secretory footprints of all antigen-specific ASC, high- and low-affinity alike, will be detected as SFU when the membrane-retained antigen probe is visualized. In wells containing decreasing concentrations of the antigen probe, however, only the secretory footprints originating from ASC that produced an antibody with high affinity for the antigen probe will retain an adequate amount for their eventual detection as SFU; B cells with an affinity lower than dictated by the actual antigen concentration go undetected, and the SFU counts will decrease by their number. Therefore, establishing the SFU counts across a range of antigen probe concentrations provides insights into the affinity distribution of the antigen-specific B cell repertoire. The second B cell ImmunoSpot® assay variant suited for assessment of the affinity distribution present in a polyclonal antigenspecific ASC compartment relies on a direct assay in which the antigen-coating density is graded. Such assays (Fig. 4) inherently provide information on the affinity of the “monoclonal antibody” that each antigen-specific ASC secretes. Following the basic rules of antibody–antigen binding, ASC that produce high-affinity antibodies generate dense, bright, and tight (sharp and small) secretory footprints (“spots”), while ASC that produce antibodies with lower affinity yield secretory footprints that are fainter and more diffuse [16]. Coating the membrane with decreasing densities of antigen and monitoring the resulting changes in SFU number and

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morphology, therefore, provides information regarding the affinity distribution of an antigen-specific ASC repertoire (see Note 24). The Chap. 13 by Becza et al. in this volume [12] lays out in detail both assay variants for B cell affinity measurements. 1.11 Measuring Cross-Reactivities of Individual BmemDerived ASC

As described above, Bmem also provides a “second wall of adaptive humoral immunity” against mutated antigens/viruses (heterotypic antigens) that evolved to evade neutralizing antibodies induced by the original (homotypic) virus/antigen. While the primary response leads to fine tailoring of the B cell repertoire directed against the homotypic antigen, including affinity maturation, among the variants created by random SHM, Bmem endowed with BCR possessing affinity for the heterotypic antigen will be present. Even if such Bmem would be rare, and even if their initial affinity would be modest, they still would occur in much higher frequencies compared to the naive B cell repertoire. Moreover, most of these Bmem have already undergone Ig class switching. Therefore, when such clonally expanded, semi-affinity matured, and classswitched, cross-reactive Bmem engage in a primary immune response against the heterotypic antigen, they can generate a more rapid and efficient antibody response compared to individuals who have not been previously immunized/infected (see Note 25). Measuring existing serum antibodies does not detect such crossreactive Bmem, and thus does not provide predictive information about cross-reactive protection, whereas ImmunoSpot®-based assessments at the level of individual Bmem-derived ASC do. There are two ways to measure Bmem-derived ASC crossreactivity by ImmunoSpot®. For both, the appropriate cell input per well first needs to be defined, which will provide the “Goldilocks” SFU count for the homotypic antigen (between 50 and 100 SFU per well, see above). In the simple version of B cell cross-reactivity studies, a second experiment is performed, an inverted B cell assay, in which this Goldilocks input of PBMC for the particular donor is seeded into all wells. Setting up replicate wells permits to assess ≥300 individual ASC-derived secretory footprints in independent single-color ImmunoSpot® assays, comparing the numbers of SFU detected in all replicates for the homotypic antigen, vs. the cumulative SFU number in the same number of replicate wells detected using the heterotypic antigen probe. The ratio of these two numbers reveals the percentage of cross-reactive ASC at equimolar concentrations of the respective antigen probes (see Note 26 and Fig. 5). Raising/lowering the concentration of heterotypic antigen probe enables assessment of the affinity spectrum of Bmem-derived ASC that cross-react with the heterotypic antigen. In a second, somewhat more complex but perhaps more elegant, approach, Bmem-derived ASC cross-reactivity is evaluated using alternatively labeled homotypic and heterotypic antigens. As

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Fig. 5 Inverted ImmunoSpot® enables assessment of Bmem cross-reactivity against the receptor binding domain (RBD) of the homotypic prototype strain of SARS-CoV-2, and heterotypic antigenic variants of concern that emerged later in the COVID-19 pandemic. PBMC from nine convalescent COVID-19 donors with PCR-verified infection during the initial wave of SARS-CoV-2 infections caused by the prototype Wuhan-Hu1 strain (collected prior to November 1, 2021, i.e., before the onset of the Delta and Omicron waves), were cryopreserved. After thawing, the PBMC were stimulated for 5 days in vitro to transition resting Bmem into antibody-secreting cells (ASC). These in vitro stimulated PBMC were then seeded into an inverted ImmunoSpot® assay, leveraging IgG capture, to detect IgG+ secretory footprints that captured RBD protein representative of the homotypic Wuhan-Hu-1, or heterotypic B.1.617.2 (Delta) or B.1.1.529 (Omicron) strains of SARSCoV-2. Each of these RBD-variant specific secretory footprints was detected in single-color assays, first by the addition of the respective soluble His-tagged RBD antigen probe, followed by the addition of biotinylated antiHis mAb, and finally by the addition of SA-CTL-Red™. The individual secretory footprints, or spot-forming units (SFU), with reactivity against the His-tagged RBD probes were subsequently enumerated using ImmunoSpot® software. Representative well images for donor A are shown in panels a–c for the three RBD probes, as specified. (d) The percentage of IgG+ ASC, following in vitro differentiation, exhibiting reactivity with the homotypic Wuhan-Hu-1, or heterotypic Delta or Omicron RBD probes. (e) The percentage of IgG+ ASC reactive with the heterotypic Delta or Omicron RBD probes relative to the homotypic Wuhan-Hu-1 strain (denoted as 100%). Data are plotted as mean ± SD. Statistical significance was determined using an analysis of variance (ANOVA) and Bonferroni’s multiple comparisons post-hoc test. ***p < 0.001

before, the PBMC are plated at the Goldilocks input number into replicate wells of an inverted B cell ImmunoSpot® assay. The “tag 1” labeled homotypic antigen probe (e.g., His tag) and the “tag 2” labeled heterotypic antigen probe (e.g., FLAG tag) are added simultaneously, at equimolar concentrations, followed by dualcolor detection of SFU that bound an antigen probe. The ratio of “tag 1” single-color positive SFU and of “tag 1 + tag 2” doublecolor positive SFU is established, revealing the frequency of cross-

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reactive ASC under equimolar conditions. Raising and lowering the concentration of the heterotypic antigen probe provides insights into the affinity of the individual homotype-primed Bmem for the heterotypic antigen. 1.12 Concluding Remarks

The aim of this chapter was to draw attention to the tremendous, thus far largely unrealized, potential of B cell ImmunoSpot® assays, and their potential to revolutionize immune diagnostics. Although the ELISPOT assay was originally introduced 40 years ago for detecting antigen-specific ASC [25, 26], it was T cell ELISPOT assays (introduced 5 years later [27]) that took the limelight. For the latter to occur, however, we needed to introduce fundamental modifications to the originally described protocol so that the assay lived up to its potential and was capable of reliably revealing secretory footprints of individual T cells responding to antigen ([28] and see Note 27). Our introduction of automated, objective, and scientifically validated software-assisted machine reading of T cellderived secretory footprints [29] also largely contributed to the success of T cell ImmunoSpot® assays (see Note 28). Despite being introduced earlier into the literature, B cell ImmunoSpot® assays have, thus far, not become a mainstay in B cell immune monitoring efforts. One likely reason is that for most antigens the classic protocol of directly absorbing the antigen to the assay membrane simply does not work (see above); only our recent introduction of the affinity coating approach [11] enables one to rapidly develop ImmunoSpot® assays for detecting ASC with specificity for essentially any affinity-tagged antigen of interest. But perhaps the primary reason why B cell ImmunoSpot® assays have not been sufficiently pursued until now is the widely held, and flawed, assumption that serum antibody measurements yield sufficient insight on underlying B cell-mediated adaptive humoral immunity. As outlined above, from the basic science perspective, this standpoint is no longer tenable. Importantly, a major component of the critical information pertaining to Bmem-mediated immunity is now easily attainable through the implementation of B cell ImmunoSpot® assays, as outlined above, and is neglected by studying antigen-specific serum reactivity/titers alone. Ease of implementation, low cost, and moderate labor investment are major requirements for the success of any assay. The efficiency of B cell ImmunoSpot® in PBMC utilization has been repeatedly highlighted in this chapter (see also see Note 23) and is critical in clinical trial settings. Cryopreserved PBMC can be used without impairing B cell function [30], permitting batch testing of dozens of samples by a single investigator in a single experiment (and hundreds of samples with a well-trained team). Fully automated image analysis of the assay results is now available, along with retention of audit trails [10]. The reproducibility of B cell ImmunoSpot® is quite remarkable for a cellular assay [6], and the serial

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dilution strategy detailed above permits to extend the upper and lower detection limits of the assay over orders of magnitude [6]. Moreover, B cell ImmunoSpot® assays have been shown to be suitable for regulated testing [31]. For all of the above reasons, we believe B cell ImmunoSpot® testing will soon become an indispensable component of the immune monitoring repertoire. Detailed methodology for the affinity capture-based B cell ImmunoSpot® assay is described in another Chap. 15 contributed by Yao et al. in this volume [23]. This is the method of choice for establishing frequencies of antigen-specific ASC via the serial dilution strategy; including for all Ig classes, or IgG subclasses, simultaneously. Detailed methodology for the single-color, antigenspecific inverted B cell ImmunoSpot® assay described in Fig. 5 is detailed below. Both the direct and inverted ImmunoSpot® assay formats are also well-suited for assessment of ASC cross-reactivity. However, while direct ImmunoSpot® assays provide insights into the affinity distributions through studying spot morphologies, the inverted assay described in detail by Becza et al. [12] is the method of choice for such high-content analysis of antigen-specific ASC repertoires. Software solutions for B cell ImmunoSpot® analysis are detailed in another Chap. 5 of this volume by Karulin et al. [10].

2

Materials

2.1 Thawing of Cryopreserved PBMC

1. Class II biosafety cabinet (BSC). 2. Cryopreserved PBMC sample(s) (see Note 23). 3. 70% (v/v) ethanol (EtOH) 4. Conical tubes. 5. Sterile transfer pipette. 6. DNase-containing washing medium (prewarmed to 37 °C) (see Note 29). 7. Centrifuge capable of spinning 50 mL conical tubes at 330 × g (temperature set to 25 °C). 8. Complete B cell medium (BCM) (prewarmed to 37 °C) (see Note 30). 9. Ca2+, Mg2+-free phosphate-buffered saline (PBS), pH 7.2 (room temperature). 10. Parafilm. 11. CTL-LDC™ live/dead cell counting kit. 12. ImmunoSpot® S6 Ultimate 4 LED Analyzer, or suitable instrument equipped with the appropriate detection channels, running CTL’s live/dead cell counting suite software.

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2.2 In Vitro Polyclonal Stimulation of B Cells in PBMC

1. B-Poly-S. 2. Volume-dependent, 48- or 24-well plate, or sterile culture flask (see Note 31). 3. Humidified incubator set at 37 °C, 5% CO2.

2.3 Four-Color, Antigen-Specific FluoroSpot Assay (Affinity Capture Coating)

1. Commercially available, four-color Human Ig class (IgA, IgE, IgG, and IgM) affinity capture (His) FluoroSpot kit (see Note 32). 2. Commercially available, four-color Human IgG subclass affinity capture (His) FluoroSpot kit (see Note 32). 3. His-tagged recombinant protein (see Notes 33 and 34). 4. 190 proof (95% v/v) EtOH 5. Cell culture-grade water. 6. 96-well, round bottom dilution plate. 7. 0.05% Tween-PBS wash solution 8. 0.1 μm low-protein binding syringe filter 9. Plate washer. 10. Vacuum manifold. 11. ImmunoSpot® S6 Ultimate 4 LED Analyzer, or suitable instrument equipped with the appropriate detection channels, running CTL’s ImmunoSpot® UV.

2.4 Single-Color, Antigen-Specific Inverted ImmunoSpot® Assay

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1. Commercially available, single-color inverted (His) human B cell ImmunoSpot® kit (see Note 35). 2. His-tagged recombinant protein (see Note 36).

Methods

3.1 Thawing of Cryopreserved PBMC (Sterile Conditions)

1. Place cryovial(s) into a 37 °C bead bath (or water bath, we prefer the former for sterility reasons) for 8 min to thaw. 2. Remove cryovial(s) and wipe with 70% EtOH inside the biosafety cabinet (BSC) before unscrewing the cap(s). 3. Using a sterile pipette, transfer contents of cryovial(s) into a labeled conical tube. 4. Rinse each of the cryovials with 1 mL of prewarmed antiaggregate solution. Transfer the rinse solution(s) to the conical tube(s) dropwise while swirling the tube to ensure adequate mixing of the cells and thawing medium. 5. Double the volume of the cell suspension by dropwise addition of warm anti-aggregate solution while swirling the tube to ensure adequate mixing of the cells and thawing medium.

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6. Continue doubling the volume of the cell suspension by dropwise addition of warm anti-aggregate solution while swirling the tube until the cryopreserved cell material has been diluted tenfold. If multiple cryovials are pooled, calculate using 1 mL of cryopreserved cell suspension + 9 mL of anti-aggregate solution to determine the necessary final resuspension volume. 7. Centrifuge balanced tubes at 330 × g for 10 min with the centrifuge brake on, at room temperature (RT). 8. Decant supernatant and resuspend the cell pellet(s) using prewarmed B cell medium (BCM) to achieve a cell density of approximately 2–5 × 106 cells/mL. The number of PBMC recovered can be estimated at this point being 80% of the number frozen. 9. Pipet 15 μL of live/dead cell counting dye onto a piece of parafilm to form a droplet. 10. Remove 15 μL of cell suspension and combine with droplet of live/dead cell counting dye. Pipet up and down 3–5 times to mix the sample while avoiding the formation of bubbles. 11. Transfer 15 μL of the cell and dye suspension into each chamber of a hemacytometer. 12. Determine live cell count and viability using CTL’s live/dead cell counting suite. 13. Increase volume of cell suspension(s) with additional sterile PBS and centrifuge balanced tubes at 330× g for 10 min with centrifuge brake on, unrefrigerated. 14. Decant supernatant and gently resuspend the cell pellet (s) using prewarmed BCM at a cell density to 2 × 106 cells/mL. 3.2 In Vitro Polyclonal Stimulation of B Cells in PBMC (Sterile Conditions)

1. Dilute CTL’s B-Poly-S polyclonal stimulation reagent 1:500 into prewarmed BCM to achieve a final concentration of 2X. Into labeled sterile culture vessel, add the same volume of PBMC at 2–4 × 106 cells/mL (see Note 31). 2. Transfer culture vessels (flasks or plates) into humidified incubator set at 37 °C, 5% CO2 for 4–6 days (96–144 h).

3.3 Four-Color, Antigen-Specific FluoroSpot Assay (Affinity Capture Coating)

1. Two days before plating cells (Day 2), prepare 70% EtOH and anti-His affinity capture antibody solutions. 2. Remove underdrain and pipet 15 μL of 70% EtOH solution into the center of each well (or designated wells) of the assay plate. Immediately after the addition of the 70% EtOH solution to the entire plate (or designated wells), add 180 μL/well of PBS. Decant and wash wells again with 180 μL/well of PBS. 3. Decant the assay plate, replace underdrain, and immediately add 80 μL/well of the anti-His affinity capture antibody

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solution into each well (or designated wells, see Note 37) of the low autofluorescence PVDF-membrane plate provided with the kit. 4. Incubate the plate overnight at 4 °C in a humified chamber. 5. The following day (Day 1), dilute the His-tagged protein (s) into Diluent A (provided with the kit) to the previously determined optimal concentration. 6. Decant the assay plate and wash wells with 180 μL/well of warm PBS. Immediately, add 80 μL/well of the corresponding His-tagged protein coating solution(s) into the designated wells. 7. Incubate the plate overnight at 4 °C in a humidified chamber. 8. On the day of the assay (Day 0), decant the assay plate and wash wells with 180 μL/well of warm PBS. Next, decant the plate and add 150 μL/well of prewarmed BCM to block the plate (≥1 h at RT). 9. If using PBMC following in vitro polyclonal stimulation, collect the cell suspension(s) and transfer into labeled conical tube (s). Keep the cells warm during processing. Wash culture vessel’s interior with sterile warm PBS to collect residual PBMC and transfer into the corresponding conical tube(s). Increase volume to fill the tube with additional warm PBS and then centrifuge balanced tubes at 330 × g for 10 min nonrefrigerated, centrifuge with brake on. Alternatively, follow the procedures detailed above to obtain freshly isolated PBMC, or to thaw PBMC that were previously cryopreserved, if prior in vitro stimulation is not required to elicit antigen-specific ASC activity in the sample(s). 10. Decant supernatant and resuspend the cell pellet(s) using prewarmed BCM to achieve a cell density of ~2–5 × 106 cells/mL (the cell number recovered at this point can be estimated to be 50% of the number of cells frozen). 11. Pipet 15 μL of live/dead cell counting dye onto a piece of parafilm to form a droplet. 12. Remove 15 μL of cell suspension and combine with droplet of live/dead cell counting dye. Pipet up and down 3–5 times to mix the sample while avoiding the formation of bubbles. 13. Transfer 15 μL of the cell and dye suspension into each chamber of a hemacytometer. 14. Determine live cell count and viability using CTL’s live/dead cell counting suite. 15. Increase volume of cell suspension(s) with additional sterile warm PBS and centrifuge balanced tubes at 330× g for 10 min with centrifuge brake on, unrefrigerated.

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16. Decant supernatant and resuspend the cell pellet(s) using prewarmed BCM at 2 × 106 PBMC/mL. 17. Decant the BCM used for blocking the assay plate and replace with 100 μL/well of prewarmed BCM. 18. If applicable, e.g., for Goldilocks number determinations or accurate frequency measurements, prepare PBMC serial dilution series in a round-bottom 96-well polystyrene plate to match the plate layout shown in Fig. 5a of the chapter by Yao et al. in this volume [23] before transferring the PBMC into the ImmunoSpot® assay. 19. Incubate cells in the assay plate for 16–18 h at 37 °C, 5% CO2. 20. After completion of the assay incubation period, decant (or reutilize) cells and wash plate two times with warm PBS (200 μL/well), followed by two additional washing steps with 0.05% Tween-PBS wash solution (see Note 38). 21. Prepare anti-Ig class/subclass-specific detection antibody solution(s) according to kit protocol and pass through 0.1 μm low-protein binding syringe filter to remove any protein aggregates. 22. Decant 0.05% Tween-PBS wash solution, add 80 μL/well of the anti-Ig class/subclass-specific detection antibody solution into designated wells, and incubate for 2 h at RT (protected from light). 23. Wash plate(s) two times with 0.05% Tween-PBS wash solution. 24. Prepare tertiary solution by following kit protocol and pass through 0.1 μm low-protein binding syringe filter to remove any aggregates. 25. Decant 0.05% Tween-PBS wash solution, add 80 μL/well of tertiary solution into designated wells, and incubate for 1 h at RT (protected from light). 26. Wash plates(s) twice with distilled water. 27. Remove protective underdrain and place plate face down on vacuum manifold. Completely fill the backside of the plate with distilled water and apply vacuum to draw water through the membrane (“back to front”) (see Note 39). 28. Allow plate to dry completely, protected from light (see Note 40). 29. Scan and count plate(s) with suitable analyzer equipped with the appropriate detection channels (see Note 18). 3.4 Single-Color, Antigen-Specific Human IgG-Inverted ImmunoSpot® Assay

1. One day before plating cells (Day 1), prepare 70% EtOH and anti-human IgG capture antibody solutions. 2. Remove underdrain and pipet 15 μL of 70% EtOH solution into the center of each well (or designated wells) of the assay

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plate. Immediately after addition of the 70% EtOH solution to the entire plate (or designated wells), add 180 μL/well of PBS. Decant and wash wells again with 180 μL/well of PBS. 3. Decant the assay plate, replace underdrain, and immediately add 80 μL/well of the anti-human IgG capture antibody solution into each well (or designated wells) of the low autofluorescence PVDF-membrane plate provided with the kit. 4. Incubate the plate overnight at 4 °C in a humified chamber. 5. On the day of the assay (Day 0), decant the assay plate and wash wells with 180 μL/well of PBS. Next, decant the plate and add 150 μL/well of prewarmed BCM to block the plate (≥1 h at RT). 6. If using PBMC following polyclonal stimulation, collect the cell suspension(s) and transfer into labeled conical tube(s). Rinse interior of the culture vessel to recover all cells and transfer into the corresponding conical tube(s). Increase volume to fill up tube with additional PBS and then centrifuge balanced tubes at 330× g for 10 min with centrifuge brake on, at RT. Alternatively, thaw previously cryopreserved PBMC as detailed above if prior in vitro stimulation is not required to elicit antigen-specific ASC activity in the sample(s). 7. Decant supernatant and resuspend the cell pellet(s) using prewarmed BCM to achieve a cell density of approximately 2–5 × 106 cells/mL (this number can be estimated to be 50% of that prior to the stimulation culture). 8. Pipet 15 μL of live/dead cell counting dye onto a piece of parafilm to form a droplet. 9. Remove 15 μL of cell suspension and combine with droplet of live/dead cell counting dye. Pipet up and down 3–5 times to mix the sample while avoiding the formation of bubbles. 10. Transfer 15 μL of the cell and dye suspension into each chamber of a hemacytometer. 11. Determine live cell count and viability using CTL’s live/dead cell counting suite. 12. Increase volume of cell suspension(s) with additional sterile warm PBS and centrifuge balanced tubes at 330× g for 10 min with centrifuge brake on, at RT. 13. Decant supernatant and resuspend the cell pellet(s) using prewarmed BCM at approximately 1 × 106 PBMC/mL. 14. Decant the BCM used for blocking the assay plate and replace with 100 μL/well of prewarmed BCM to block the plate. 15. If the Goldilocks number has already been established for the PBMC(s), plate cells at that number into the ImmunoSpot®

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assay. If that number is unknown, prepare serial dilution of PBMC (or other single-cell suspension) accordingly, and then transfer into the ImmunoSpot® assay plate. 16. Incubate cells in the assay plate for 16–18 h at 37 °C, 5% CO2. 17. After completion of the assay incubation period, remove plate and decant cells. Wash plate two times with PBS (200 μL/ well), followed by two additional washing steps with 0.05% Tween-PBS wash solution (see Note 38). 18. Prepare His-tagged antigen probe solution at optimized concentration and pass through 0.1 μm low-protein binding syringe filter to remove any protein aggregates. 19. Decant 0.05% Tween-PBS wash solution, add 80 μL/well of His-tagged antigen probe solution into designated wells, and incubate for 2 h at RT (protected from light). 20. Wash plate(s) two times with 0.05% Tween-PBS wash solution. 21. Prepare anti-His detection antibody solution according to kit protocol and pass through 0.1 μm low-protein binding syringe filter to remove any aggregates. 22. Decant 0.05% Tween-PBS wash solution, add 80 μL/well of anti-His detection antibody solution into designated wells, and incubate for 1 h at RT (protected from light). 23. Wash plate(s) two times with 0.05% Tween-PBS wash solution. 24. Prepare tertiary solution according to kit protocol and pass through 0.1 μm low protein binding syringe filter to remove any aggregates. 25. Wash plates(s) twice with distilled water. 26. Remove protective underdrain and place plate face down on vacuum manifold. Completely fill the backside of the plate with distilled water and apply vacuum to draw water through the membrane (“back to front”) (see Note 39). 27. Allow plate to dry completely, protected from light (see Note 40). 28. Scan and count plate(s) with suitable analyzer equipped with the appropriate detection channel (see Note 18).

4

Notes 1. As enzyme-linked immunospot (ELISPOT) and FluoroSpot assays differ only in the modality of detecting secretory footprints of cells on membranes, we collectively refer to both as ImmunoSpot® assays. In the former, the detection antibody is tagged to enable the engagement of an enzymatic reaction that results in the local precipitation of a substrate visible under

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white light. In the latter, the plate-bound detection antibodies are visualized via fluorescent tags using appropriate excitation and emission wavelengths. Data provided in the chapter in this volume by Yao et al. [23] establish that ELISPOT and FluoroSpot assays have equal sensitivity for detecting antibodysecreting cell (ASC)-derived secretory footprints. 2. In proliferating germinal center B cells (GCB), somatic hypermutation (SHM) introduces random mutations into the variable regions of the B cell receptors (BCR) responsible for antibody binding. As a result, subclones endowed with improved, or decreased, BCR binding affinity for the antigen are generated. GCB subclones that possess an increased BCR affinity for antigen are positively selected to proliferate and undergo further rounds of SHM to further improve BCR affinity. Collectively, the positive selection of GCB subclones with improved binding for antigen underlies the progressive affinity maturation of the ensuing antibody response. 3. B cell antigen receptors (BCR) are the membrane-anchored forms of secreted immunoglobulin (Ig) molecules (commonly referred to as antibodies). Surface BCR are encoded by the same Ig class/subclass and possess an identical antigen specificity to the antibody that a B cell will secrete once it differentiates into an ASC, such as a plasma cell (PC). 4. As the concentration of antigen declines with time, only those germinal center B (GCB) cells with the highest affinity will continue to successfully compete for antigen binding, and only the latter will continue to proliferate and undergo progressive SHM to eventually differentiate along the PC lineage. 5. The half-life of different Ig classes, and IgG subclasses, in serum is variable and relatively short in vivo. The half-life of IgG1, IgG2, and IgG4 in humans is 21–28 days, whereas for IgG3 it is ~1 week [32]. For IgA and IgM, their half-lives are even shorter (3–7 days) [33, 34] and IgE has the shortest halflife in serum, ~2–3 days [35]. 6. The extended half-life of serum IgG in vivo is dependent on its interactions with FcRn [36, 37] which protects it from catabolism and enables IgG recycling. Likewise, the in vivo half-life of other antibody classes such as IgA, IgE, and IgM are also greatly extended when in association with Ig-binding surface receptors [35, 38]. 7. IgE secreted by B cells can bind to high-affinity Fc receptors (FcεRI) expressed on mast cells and basophils and endow these cells with antigen-recognition properties. IgA antibodies are secreted to mucosal surfaces. 8. Traditional B cell ELISPOT assays have been performed by direct coating of the assay membrane with the antigen of

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interest. However, many (in fact, most) antigens do not adsorb sufficiently to the membrane to enable reliable detection of ASC-derived secretory footprints. We have overcome this limitation by introducing an affinity coating approach for achieving high-density antigen absorption to the assay membrane [11]. 9. These samples were collected early in the COVID-19 pandemic when the prototype Wuhan-Hu-1 strain of the SARS-CoV2 virus was circulating, and the testing was done with Spike and Nucleocapsid (NCAP) antigens that corresponded to the original Wuhan-Hu-1 strain. Thus, homotypic B cell memory was studied. 10. While ImmunoSpot® enables measurement of a single ASC within a bulk population of cells, flow cytometry falls short of detecting antigen-specific cells when they are present as very low-frequency events. In ImmunoSpot® assays, there is no inherent lower limit of detection. If, e.g., 3 million PBMC are plated at 3 × 105 PBMC across 10 replicate wells, 1 in 3 million is the detection limit, etc. Importantly, owing to increased Poisson noise occurring with such low-frequency measurements, the number of replicate wells evaluated needs to be increased accordingly to obtain accurate low-frequency measurements. As shown in Fig. 3, antigen-specific Bmem quite frequently occur in low frequencies. 11. To establish the frequency of antigen-specific ASC among all ASC-producing IgM/IgG/IgA/IgE or all four IgG subclasses (IgG1-IgG4) requires ~2 million PBMC (that can be isolated from 2 to 3 mL of blood) when done according to our protocols (see also Note 23). To obtain the same information by flow cytometry, up to ten-fold more cells would be required [39, 40]. 12. For details of four-color B cell ImmunoSpot® detection, see the chapter by Yao et al. in this volume [23]. Flow cytometry does not reliably reveal the class/subclass of Ig produced by the individual B cell because surface BCR expression can be highly variable and this is an underappreciated complexity of probe staining. In particular, in the case of IgG+ ASC, they express little if any surface BCR and this undermines assessment of their antigen specificity and subclass usage using traditional surface staining approaches. Consequently, fixation and intracellular staining are required to define the IgG subclass usage of these cells (a procedure that results in substantial cell loss in the sample). 13. Each investigator in our laboratory can routinely test, in a single experiment, 10–20 PBMC samples for reactivity against a panel of antigens, assessing the frequency of ASC producing

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each of the Ig classes and IgG subclasses (see also the chapter by Yao et al. in this volume [23]). With additional logistical refinements, this throughput is readily scalable upward. 14. For most antigens we have investigated thus far (SARS-CoV2 NCAP and receptor binding domain (RBD) proteins, the HCMV gH pentamer complex, EBV EBNA1, and OVA), we have been unsuccessful in establishing reliable B cell ImmunoSpot® assays when following the classic protocol in which the antigen is directly absorbed to the membrane irrespective of how high an antigen concentration was used for coating. In contrast, our recent introduction of the affinity coating strategy [11] generated pristine results with each of these antigens. We succeeded with direct coating only with recombinant hemagglutinin proteins of seasonal influenza viruses or the fulllength trimeric Spike protein of SARS-CoV-2, albeit having to use very high antigen coating concentrations (exceeding 20 μg/mL) to achieve high-quality secretory footprint formation. 15. In addition to detecting antigen-specific, B cell-derived secretory footprints on affinity-coated plates, as described here, it is also possible to detect such cells via the inverted ImmunoSpot® approach. In the latter, the Ig secretory footprint is captured irrespective of the ASC’s specificity (e.g., by coating the membrane with an anti-IgG capture antibody) and the antigenspecific B cells’ secretory footprints among all those retained on the membrane are detected by the capture of labeled antigen. Such inverted assays work perfectly when the frequency of antigen-specific B cell footprints is high among all secretory footprints captured, but are challenging for the detection of low-frequency events. Inverted assays are described in detail in the chapter by Becza et al. in this volume [12]. While direct assays permit the capture of all Ig classes or subclasses produced by antigen-specific ASC via four-color ImmunoSpot® detection (see the chapter by Yao et al. in this volume [23]), the inverted assay detects only the selected Ig class, e.g., IgG+ ASC. 16. B cell ImmunoSpot® data can be expressed as spot-forming units (SFU) per cell input per well to determine the frequency of antigen-specific cells. However, owing to considerable interindividual variations in the abundance of total Ig class/subclass producing ASC in test samples following polyclonal stimulation, and the prevalence of IgG [6], it is better to report data as the frequency of ASC producing a given Ig class/subclass among all ASC producing that Ig class/subclass. 17. ASC secrete Ig in an undirected fashion into 3D space. The antibody released toward the membrane will be captured as a secretory footprint; however, the remainder of the secreted

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antibody will diffuse away from the cell and is diluted into the culture supernatant. As the concentration of diffused antibodies increases in the culture medium, these antibodies are captured on the membrane distantly from the source ASC, increasing the background signal in the assay and undermining the resolution of individual secretory footprints. Such an elevated background in an ImmunoSpot® assay is termed an ELISA effect that can obscure the accurate counting, or even detection of secretory footprints [6]. 18. The chapter by Karulin et al. in this volume [10] introduces machine learning-based spot-forming unit (SFU) analysis that can partially compensate for ELISA effects and SFU crowding, thus extending the linear range of accurate quantification for cell numbers plated per well and SFU detected. 19. When performing such serial dilution experiments, using 4 replicate wells per cell input plated, and linear regression of the SFU frequency through extrapolation from the mean of these 4 replicates, that calculated frequency was very close to the frequencies established by relying on just one replicate well per cell input dilution (see the chapter by Yao et al. in this volume [23], and N. Becza, manuscript in preparation). Thus, serial dilutions involving single wells for each cell dilution, progressing in a 1 + 1 (two-fold) dilution series is a valid option for establishing accurate SFU frequencies and greatly reduces the cell numbers and reagents required. 20. We failed to detect SARS-CoV-2 Spike or NCAP antigenspecific, class-switched (IgG or IgA) ASC in individuals collected prior to the COVID-19 pandemic that by definition would be immunologically naive to the SARS-CoV-2 virus [6]. However, IgM+ ASC reactivity against these antigens in the pre-COVID-19 donor cohort population was ambiguous because the current polyclonal stimulation approach also results in the differentiation of naive IgM+ B cells, which may express broadly reactive specificities and can generate secretory footprints in negative control wells. In contrast, all subjects with PCR-verified infection, harbored S and N antigen-specific IgG+ ASC. These findings are consistent with the notion that clonal expansions to detectable frequency levels, classswitching, and affinity maturation (all features of Bmem) are needed for detecting antigen-specific B cells in ImmunoSpot® assays. Therefore, it appears that detection of class-switched, antigen-specific ASC in ImmunoSpot® assays implies prior in vivo priming of Bmem with (cognate or cross-reactive) specificity for the test antigen.

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21. Thus far, we have not seen evidence for class-switching of Bmem during short-term in vitro polyclonal stimulation using R848 plus IL-2. 22. So far, having studied “healthy volunteers” exclusively, we only detect IgE+ ASC following in vitro stimulation with agonistic anti-CD40 mAb plus IL-4 and IL-21, but not after stimulation with R848 plus IL-2 [41]. Thus, our detection system enables identification of IgE+ ASC, but whether IgE+ Bmem exist in vivo, or whether they switch de novo to IgE in vivo or in vitro is controversial [42, 43]. 23. If special protocols are followed, PBMC can be frozen without impairing the B cells’ functionality ([30] and N. Becza manuscript in preparation). Thus, by freezing B cells of a sample in several aliquots, the same PBMC can be tested repeatedly, reproducing the results of the previous experiment with high accuracy [6], or for extending those studies, e.g., for assessing the affinity distribution of the antigen-specific Bmem repertoire. Of note, when planning the numbers of PBMC to be frozen per cryovial, as a rule of thumb, one can anticipate recovery of ~50% of PBMC initially frozen after these cells are thawed and have undergone 5 days of in vitro polyclonal stimulation to promote terminal differentiation of resting Bmem into ASC. It is also important to note that any number of PBMC between 1 and 10 million can be frozen per cryovial permitting the optimization of PBMC utilization when planning experiments (N. Becza, manuscript in preparation). 24. While the potential of B cell ImmunoSpot® assays to establish affinity distributions is embedded in the nature of the assay, this potential has not been realized yet. The chapter by Becza et al. in this volume [12] is the first step in this direction. To progress along these lines is one of the major focuses in our laboratory. 25. It can be assumed that after vaccination with the prototype Wuhan-Hu-1 strain Spike antigen, in addition to T-cell immunity directed against shared antigenic determinants of Spike variants, heterotype-specific, cross-reactive B cell memory for the receptor binding domain (RBD) mediates the partial protection seen toward newly emerging antigenic variants of this virus. Such SARS-CoV-2 variants evade the first wall (reflected by serum antibodies) and are thus capable of causing infections, but the engagement of cross-reactive Bmem (the second wall) can still convey protection from severe disease through rapidly generating a quasi-secondary neutralizing antibody response against the mutated RBD of the new variant. 26. In our ongoing studies, we find that ~40% Wuhan-Hu-1 infection-elicited Bmem are cross-reactive with the Omicron (BA.1) variant (Fig. 5 and Liu et al., work in progress).

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27. Our introduction of the PVDF membrane to T-cell ELISPOT assays [28], with its by far superior capture antibody adsorption properties [44], has been key for improving our ability to detect secretory footprints to the point needed for transforming ImmunoSpot® into the robust T-cell monitoring platform it has become for detecting rare – even very rare – antigenspecific T cells ex vivo, in freshly isolated PBMC or other lymphoid cell material. We refer to Figure 1 in [45] to appreciate the difference in assay performance using the PVDF membrane vs. the previously used mixed cellulose ester membrane. 28. The shape of secretory footprints (spot morphologies) produced by T cells follows simple rules since the capture antibody’s (i.e., an anti-cytokine-specific mAb) affinity for the analyte to be detected is high and fixed. Consequently, only the quantity of analyte (cytokine) produced by the T cell will define the morphology of the resulting secretory footprint [16]. Predictable (log normal, [46]) spot sizes permit objective automated size gating [47]. However, ASC-derived secretory footprints in antigen-coated wells are primarily defined by the affinity of the secreted antibody for the membrane-bound antigen. Thus, the formation of ASC-derived secretory footprints follows a different set of rules, and hence requires a fundamentally different analytical approach. The chapter contributed by Karulin et al. in this volume [10] introduces a software solution suited for accurate, high-throughput, and high-content B cell ImmunoSpot® analysis. 29. Thawing of cryopreserved cells causes a fraction of the cells (up to 30%) to die, and the DNA released from such cells can cause clumping of the thawed cell material. This cell clumping can be reduced, if not completely eliminated, by including an immunologically neutral endonuclease, Benzonase. Ready-touse Benzonase-containing serum-free wash solutions are available: CTL anti-aggregate Wash™ 20X solution. 30. A suitable assay medium for use in B cell ImmunoSpot® is RPMI 1640 with 10% FCS, 2 mM L-glutamine, 100 U/mL penicillin, 100 μg/mL streptomycin, 8 mM HEPES, and 50 μM 2-mercaptoethanol. 31. The volume of in vitro stimulation cultures can be scaled up or down accordingly, but we recommend keeping the cell density of PBMC at approximately 1–2 × 106 cells/mL. Smaller in vitro stimulation cultures can be initiated in 48- or 24-well plates with a final volume of 1 or 2 mL, respectively. Be sure to fill empty wells in tissue culture plates with sterile PBS to avoid dehydration of cell cultures. For larger volumes, use appropriate tissue culture flasks, upright or laying, with the culture medium between 0.5 and 1 cm in height.

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32. Kit is suited for detecting either antigen-specific ASC that differentiated in vivo, or antigen-specific Bmem that have been polyclonally stimulated in vitro to promote their transition to ASC. Each kit contains anti-His capture antibody, Ig classspecific (IgA, IgE, IgG, and IgM) detection reagents, diluent buffers, low autofluorescence PVDF-membrane plates, and polyclonal B cell activator (B-Poly-S). Alternatively, IgG classspecific (IgG1, IgG2, IgG3, and IgG4) detection reagents can be substituted in the context of such four-color B cell ImmunoSpot® assays. 33. Traditional B cell ELISPOT assays have been performed by direct coating of the assay membrane with the antigen of interest. However, many (in fact, most) antigens do not adsorb sufficiently to the membrane to enable reliable detection of ASC-derived secretory footprints. We have overcome this limitation by introducing an affinity coating approach for achieving high-density antigen absorption to the assay membrane [11]. 34. We recommend optimizing the concentration of His-tagged protein(s) used for affinity capture coating. A concentration of 10 μg/mL His-tagged protein has yielded well-formed secretory footprints for most antigens, but increased concentrations of the anti-His affinity capture antibody and/or His-tagged protein may be required to achieve optimal assay performance [11]. 35. Kit is suited for detecting either antigen-specific ASC that differentiated in vivo, or antigen-specific Bmem that have been polyclonally stimulated in vitro to promote their transition to ASC. Each kit contains pan anti-human IgG capture antibody, anti-His detection reagents, diluent buffers, low autofluorescence PVDF-membrane plates, and polyclonal B cell activator (B-Poly-S). 36. The optimal concentration of affinity (His)-tagged antigen probe used for detection of all antigen-specific secretory footprints (e.g., SFU), low- or high-affinity alike, should be determined empirically. 37. If the entire plate will not be coated with the anti-His affinity capture antibody solution, the remainder of the EtOH prewet wells should receive 80 μL/well of PBS. 38. Plate washes may also be performed manually. For automated washing, the pin height and flow rate should be customized to avoid damaging the assay membranes, which is the case for the CTL 405LSR plate washer. 39. Lowering nonspecific background staining, and reduction of “hot spots” in the center of the assay wells, is achieved through performing the “back to front” water filtration technique.

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40. To completely dry plates, blot assay plate(s) on paper towels to remove residual water before either placing them in a running laminar flow hood at a 45° angle for >20 min or placing face down on paper towels for >2 h in a dark drawer/cabinet. Do not dry assay plates at temperatures exceeding 37 °C as this may cause the membrane to warp or crack. Fluorescent spots may not be readily visible while the membrane is still wet and the background fluorescence may be elevated. Scan and count plates only after membranes have dried completely.

Acknowledgments We wish to thank the R&D and the Software Development teams at CTL for their continued support and technological innovation that made our B cell ImmunoSpot® endeavor possible. We thank Drs. Alexey Y. Karulin and Graham Pawelec for in-depth discussions of the subject matter, and Diana Roen for carefully proofreading the manuscript, and also Gregory Kovacs for his support in the generation of graphic illustrations. All efforts were funded from CTL’s research budget. Conflicts of Interest P.V.L. is Founder, President, and CEO of CTL, a company that specializes in immune monitoring by ImmunoSpot®. Z.L., N.B., A.V.V., and G.A.K are employees of CTL. J.B. was a summer intern at CTL when he developed the tetanus toxoid heavy chain (TTHc)-specific B cell ImmunoSpot® assay and contributed the frequency measurements of TTHc-specific Bmem cells incorporated in Fig. 3.

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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 Tracking Circulating HLA-Specific IgG-Producing Memory B Cells with the B-Cell ImmunoSpot Assay Delphine Kervella, Sebastiaan Heidt, Robert Fairchild, Stephen Todryk, and Oriol Bestard Abstract Donor-specific antibodies (DSA) against human leukocyte antigen (HLA) molecules are a major risk factor for rejection of transplanted organs (in antibody-mediated rejection [ABMR]), particularly in patients who have prior sensitization or receive insufficient immunosuppression through minimization or noncompliance. These DSA are measured routinely in the serum of patients prior to transplantation mainly using bead-based technologies or cell-based assays. However, the absence of detectable serum DSA does not always reflect the absence of sensitization or histologically defined ABMR, and so it has been proposed that the detection and measurement of memory B cells capable of secreting antibodies against donor HLA antigens could be carried out using B-cell ImmunoSpot, to better inform the degree of immune sensitization of transplant patients prior to as well as after transplantation. Such an assay is described here. Key words ImmunoSpot, B cells, memory, Transplantation, HLA

Abbreviations Ab ABMR DSA ELISPOT FluoroSpot HLA IgG IL ImmunoSpot mBC PBMC PBS RPMI RT

antibody antibody-mediated rejection donor-specific antibody enzyme-linked immunospot assay fluorescent-spot assay human leukocyte antigen immunoglobulin interleukin cover term for ELISPOT and FluoroSpot memory B cell peripheral blood mononuclear cells phosphate-buffered saline Roswell Park Memorial Institute medium room temperature

Alexander E. Kalyuzhny (ed.), Handbook of ELISPOT: Methods and Protocols, Methods in Molecular Biology, vol. 2768, https://doi.org/10.1007/978-1-0716-3690-9_12, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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Introduction A major cause of organ transplant rejection (the kidney being the focus in this paper) is the reaction of circulating serum antibodies against mismatched HLA (human leukocyte antigen) molecules of the donor [1]. Such alloreactive antibodies may be preexistent in the circulation due to previous sensitization events such as a previous transplant, transfusion, or pregnancy, and are particularly avid at causing rapid acute antibody-mediated rejection (ABMR) when at high titers [1, 2]. These antibodies have the ability to fix complement and give rise to characteristic ABMR histology. Acute ABMR is usually caused by high titers of IgG1 or IgG3 cytotoxic antibodies that bind complement, but other subclasses and lower titers are also involved in chronic ABMR and graft loss [3]. In the case of an unsensitized recipient at the time of transplant, HLA matching guides clinicians for donor compatibility [2], and donor-specific antibodies (DSA) to mismatched HLA molecules can potentially develop subsequently. Development of such de novo DSA depends on the adjuvant effect of the organ (whether it is damaged, e.g., by ischemia) [4], and may be involved more in chronic transplant dysfunction rather than acute rejection. Treatment of ABMR is not standardized but includes antibody and B-cell depletion [5]. Extensive characterization of HLA antibody specificities pretransplant provides clinicians with a measure of the risk of rejection of the transplant. Historically, DSA were measured by their ability to bind and sensitize donor target lymphocytes for complementdependent lysis. These days single HLA bead testing, using technology such as Luminex, is used for determining HLA specificity of circulating antibodies in patient sera. Notably, despite known sensitization events before transplantation, and even at the time of actual histological ABMR occurring after transplant, serum DSA are not always detectable or their levels may not correlate with rejection. There is a strong case to be made for an assay to be used, which can detect memory B cells capable of rapidly secreting DSA and causing ABMR, and a major candidate is the B-cell ELISPOT assay. Memory B cells persist following immunization, even once the sensitizing antigen has diminished and antibody-secreting cells have contracted and are rapidly reactivated by signals such as their cognate antigen and mitogens. The role of the B-cell effector pathway of adaptive immunity in solid organ transplantation (SOT), has gained significant attention over the last years [6– 11]. An issue with measuring B-cells secreting DSA is the large number of potential HLA molecules that could be recognized and that need to be included as target antigens in the assay on a per-patient basis. Recently-developed prototypic ImmunoSpot assays for measuring circulating memory B-cells-secreting

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Table 1 Different formats used to detect HLA-specific memory B cells using ELISPOT/FluoroSpot assays Antigens

Detection systems

References

Cell lysates from HLA-typed cells, stabilized for MHC class II

Karahan Capture: goat anti-human IgG (Jackson Labs, et al. Baltimore) [8, 9] Detection: mouse anti-mouse IgG2b for MHC class II; mouse anti-human beta-2-microglubulin for MHC class I TMB substrate

Biotin-labeled HLA-monomers

Coating: streptavidin was coated to ELISPOT plates to enable biotin-HLA capture as the coated antigen target. Detection: HRP-conjugated goat anti-human IgG TMB substrate

Heidt et al. [6, 8]

Multimerized HLA monomers, fluorescently labeled

Capture: anti-human IgG monoclonal antibody, clone MT145, (Mabtech, Stockholm, Sweden); Detection: anti-human IgG antibody conjugated to fluorochrome (Merck, Darmstadt, Germany)

Lu´cia et al. [7, 10, 11]

antibodies against HLA molecules are described, which comprise variations in target antigens to make them more pragmatic (Table 1) [6–11].

2

Materials

2.1 Isolation and Culture of Human PBMC

1. Heparinized blood from a patient 2. Ficoll-Hypaque solution, or Lymphoprep 3. RPMI-1640 medium supplemented with standard 10% fetal calf serum, penicillin/streptomycin and L-glutamine (further referred to as complete medium), or IMDM, similarly supplemented 4. Stimuli used in various concentrations and combinations: Resiquimod TLR7/8 agonist (R848), IL-2, IL-10, IL-21, and anti-CD40 antibody. 5. CO2 incubator at 37 °C 6. PBS buffer

2.2 ImmunoSpot Assay (See Table for Lab-Specific Details)

1. Transparent, white or black ELISPOT plates (Merck, Darmstadt, Germany; or Millipore; or CTL) 2. Coating antibody or coating antigen 3. Detection antibody 4. Fluorescently-labeled HLA multimers (Pure Protein, L.L.C., Oklahoma City, OK, USA)

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5. Enhancer (AID-Diagnostika GmbH, Strasberg, Germany) 6. FluoroSpot plate analyzer 7. ImmunoSpot kits for the detection of B cells, and analyzers for analyzing plates and counting spots based on features, including size and intensity, can be purchased from companies, including CTL (Shaker Heights, Ohio) and Mabtech (Nacka Strand, Sweden).

3

Methods All steps before and during peripheral blood mononuclear cells (PBMC) isolations and stimulation should be performed under sterile conditions (in laminar flow hood; see Note 1).

3.1 PBMC Stimulation

1. Resuspend freshly isolated or thawed PBMC in culture medium and count cells (see Note 2). 2. Centrifuge cells at 500g for 8 min. 3. Resuspend cells in culture medium supplemented with cytokines (either R848 + IL-2, or more complete R848 + α-CD40 + IL-2 + IL-21 + IL-10), at 1.5 × 106 cells per mL in 15 mL tubes with a filter-vented cap or similar. 4. Incubate at 37 °C, 5% CO2 for 6 days.

3.2 ImmunoSpot Assay (Focusing on Methods in Publications: [7–9]) 3.2.1 Plate Coating (Day 5), See Fig. 1 for Diagrammatic Representation (See Note 3) 3.2.2 Plate Seeding (Day 6)

1. Prepare the anti-human IgG antibody: dilute at 15 μg/mL in sterile PBS. 2. Prewet the ELISPOT plate with 35% ethanol (30 μL per well) for up to 2 min. 3. Wash plate 5 times with sterile water (200 μL per well). The membrane should not dry during this process. 4. Add the coating antibody solution (100 μL per well). 5. Incubate the plate overnight at 4 °C (for up to 5 days). 1. Wash plate 5 times with sterile PBS to remove excess of coating antibody (200 μL per well). 2. Block plate by adding 200 μL of complete medium per well, for at least 1 h at room temperature (RT). 3. Centrifuge cells at 500 g for 8 min. The supernatants of cell cultures can be kept for further analysis (i.e., contains IgG produced by ASC that can be further characterized by ELISA or Luminex). 4. Resuspend cells in complete medium and count.

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Fig. 1 Illustration of the B-cell ImmunoSpot in transplantation. (a) PBMCs are cultured with TLR agonist to expand and differentiate all B cells, including antigen-specific B cells. (b) (1) ELISPOT plate, precoated with capture antibody for all IgG isotypes, has the expanded B cells added to it (2). During incubation, all secreted antibodies are captured (3). The captured antibodies are then probed with fluorescent-labeled multimerized HLA molecules (4). Spots, representing a specific ASC are revealed upon analysis with an analyzer instrument

5. Seed cells in complete medium (see Note 4). Given the low frequency of HLA-specific B cells, seed a large number of cells in each well (450,000). For the detection of polyclonal IgG production, seed lower numbers of cells (45,000 and 4500 cells); each condition duplicated. 6. Incubate for 20 h at 37 °C 5% CO2. 3.2.3

Detection (Day 7)

Detection of polyclonal IgG-secreting cells: prepare secondary antibody (anti-IgG antibody conjugated to fluorochrome). Detection of HLA-specific Ab-producing cells: prepare pure protein, L.L.C. HLA tetramers (0.5 μg per well, diluted in 100 μL PBS). Note that a combination of tetramers of different specificities, coupled to different fluorochromes, is possible in the same well, with no more than three per well. Target HLA tetramers related to donor–recipient HLA mismatches as well as self-HLA antigens should be also used as controls. 1. Wash plate five times with PBS. 2. Add 100 μL of anti-IgG secondary antibody (polyclonal) and 100 μL of HLA-tetramer solution to the respective wells. 3. Incubate at RT for 2 h (protected from light). 4. Wash plate five times with PBS. 5. Add 100 μL of enhancer in each well. 6. Incubate for 15 min at RT.

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Fig. 2 Multicolor FluoroSpot assay results. (a) Positive FluoroSpots in two different sensitized individuals, for each specificity and combined detection. (b) Detection of polyclonal IgG-producing cells (right panel) ascertains for correct proliferation and differentiation of B cells. (c) No detection of spots when using selfHLA tetramers

7. Dry plate and read plate with a FluoroSpot reader/analyzer. Spots are detected by fluorescence and counted in an automated ELISPOT reader. 3.2.4

Data Analysis

Data can be analyzed with the different parameters collected during data acquisition (number of spots, spots fluorescence intensity, area, and activity) using the software associated with the given analyzer (see Note 5). An example of data acquired is presented in Fig. 2. The detection of polyclonal IgG-producing cells is an internal control of the test to ascertain global proliferation and differentiation of B cells during the first phase. Results from polyclonal wells can also be used to normalize results for HLA-specific detection (i.e., a ratio or %). Negative controls for each tetramer are obtained by using selfHLA and/or individuals without HLA sensitization. This assay can be adapted using different detection techniques, as described in Table 1 (see Note 6).

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An example of the results of a FluoroSpot assay using fluorescently labeled HLA-multimers is shown in Fig. 2. Responses are shown following culture to expand and differentiate mBCs. The role of B-cell alloimmune memory in solid organ transplantation (SOT) has gained significant interest in the last decade due to the preponderance of antibody-mediated rejection as the main cause of immune-mediated allograft loss [13, 14], the progressively high numbers of highly sensitized patients on waiting lists worldwide because of previous transplants, pregnancies, or blood transfusions, which reduce their chances to receive an HLA-compatible organ [15], and importantly, because of the development of a number of new assays capable of measuring this effector pathway at the cellular level [16]. HLA-specific B cells may even enhance alloreactive T-cell responses [12]. Importantly, the implementation of sensitive immune assays measuring anti-HLA Ab in the sera has helped to significantly improve both pre- and post-transplant alloimmune risk stratification and diagnosis of antibody-mediated graft lesions [17]. However, while quantification of the humoral alloimmune response may be broadly achieved by measuring serum HLA antibodies, this approach does not provide the full picture of the B-cell alloimmune response. Notably, B-cell alloimmunity also involves bone marrow (BM)-residing long-lived plasma cells (LLPC), which continuously secrete alloantibodies, and the memory B-cell (mBC) pool that are located in secondary lymphoid organs and constantly recirculate in the bloodstream. Such mBC may rapidly switch to become antibody-secreting cells (ASC) and produce high-affinity antigenspecific IgG antibodies (DSA) if they re-encounter a previously recognized alloantigen [18]. While BM-residing LLPC are technically difficult to be systematically evaluated in clinical practice, HLA-specific mBC circulating in peripheral blood can be readily characterized with a number of novel assays such as the B-cell ELISPOT/FluoroSpot assay (ImmunoSpot). A major characteristic of this assay is its capacity to detect at the single-cell level frequencies of circulating mBC capable of switching to plasmablast-like phenotype cells after polyclonal stimulation, and producing IgG antibodies specific against HLA antigens, which may be detected using distinct HLA antigens as detection matrix and read with an ELISPOT/FluoroSpot reader. Such functionality cannot be determined using flow cytometry with fluorescently labeled HLA molecules. This ImmunoSpot assay allows the detection at the single-cell level of HLA-specific IgG produced by ASC that have polyclonally proliferated in vitro from circulating HLA-specific mBC [10]. While the limitations of this technique are the requirement of a FluoroSpot reader and the time needed for the assay (7 days), it enables the enumeration of HLA-specific mBCs circulating in peripheral blood, especially those at very low frequencies.

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This technique could be applied in different clinical settings [15]. In highly sensitized patients, it could further refine the immunization status of these patients and eventually adapt desensitization strategies to the presence or absence of donor(HLA)-specific mBC [15]. In addition, after transplantation, the assessment of circulating donor (HLA)-specific mBC frequencies could reveal the presence of de novo memory B-cell activation, even in the absence of serum antibodies, and thus establish guided therapeutic strategies. Finally, excluding the development of donor (HLA)-specific mBC could be of high value for novel trials evaluating the efficacy of new therapies primarily targeting alloreactive B cells as well as for those therapeutic strategies aiming at immunosuppression minimization. Finally, although these techniques have been mainly used in kidney transplantation, they could be also useful for other solid organ transplants in which B-cell alloimmunity also plays a crucial role. 3.3

4

Conclusion

The HLA-specific B-cell FluoroSpot may be a useful assay to further refine current immune-risk stratification, which is exclusively based on serum anti-HLA antibodies, to ultimately better characterize the alloimmune risk of transplant recipients.

Notes Several considerations must be made in order to successfully carry out the ImmunoSpot assay, particularly the adherence to exacting standards. 1. Blood must be processed to high standards (standard operating procedures [SOPs]), and then the resulting PBMCs frozen and cryo-stored appropriately. 2. Careful thawing is required to achieve good yield and viability. This needs to be checked using a suitable method/device. The input of accurately determined viable cell numbers into ImmunoSpot is crucial. 3. ELISPOT plate coating must be optimized through titration when being established in a new laboratory. Likewise when using custom detection reagents. Kits come already optimized. 4. It is advisable to titrate cells across wells to observe a titratable effect and to resolve the true number of reactive cells when such cells are in high frequency. 5. Counting parameters for resulting spots need to be diligently set to acquire data at sufficient sensitivity and consistency. 6. Companies such as CTL provide comprehensive methods, SOPs, and technical support to enable the robust success of their B-cell ImmunoSpot kits.

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Disclosures The HLA-specific B-cell FluoroSpot assay based on fluorescently labeled multimerized HLA monomers is currently commercialized by Pure HLA Protein, LLC, who owns the full license. ST is a consultant for CTL. References 1. Bestard O, Thaunat O, Bellini MI et al (2022) Alloimmune risk stratification for kidney transplant rejection. Transpl Int 35:10138 2. Meneghini M, Crespo E, Niemann M et al (2021) Donor/recipient HLA molecular mismatch scores predict primary humoral and cellular alloimmunity in kidney transplantation. Front Immunol 11:623276 3. Lefaucheur C, Viglietti D, Bentlejewski C et al (2016) IgG donor-specific anti-human HLA antibody subclasses and kidney allograft antibody-mediated injury. J Am Soc Nephrol 27(1):293–304 4. Gorbacheva V, Fan R, Beavers A et al (2019) Anti-donor MHC Class II alloantibody induces glomerular injury in mouse renal allografts subjected to prolonged cold ischemia. Am Soc Nephrol 30(12):2413–2425 5. Shiu KY, Stringer D, McLaughlin L et al (2020) Effect of optimized immunosuppression (including rituximab) on anti-donor alloresponses in patients with chronically rejecting renal allografts. Front Immunol 11:79 6. Heidt S, Roelen DL, de Vaal YJ et al (2012) A novel ELISPOT assay to quantify HLA-specific B cells in HLA-immunized individuals. Am J Transplant 12(6):1469–1478 7. Lu´cia M, Luque S, Crespo E et al (2015) Preformed circulating HLA-specific memory B cells predict high risk of humoral rejection in kidney transplantation. Kidney Int 88(4): 874–887 8. Karahan GE, de Vaal YJ, Roelen DL et al (2015) Quantification of HLA class II-specific memory B cells in HLA-sensitized individuals. Hum Immunol 76(2–3):129–136 9. Karahan GE, de Vaal YJH, Krop J et al (2017) A memory B cell crossmatch assay for quantification of donor-specific memory B cells in the peripheral blood of HLA-immunized individuals. Am J Transplant 17(10):2617–2626 10. Luque S, Lu´cia M, Crespo E et al (2018) A multicolour HLA-specific B-cell FluoroSpot assay to functionally track circulating

HLA-specific memory B cells. J Immunol Methods 462:23–33 11. Luque S, Lu´cia M, Melilli E et al (2019) Value of monitoring circulating donor-reactive memory B cells to characterize antibody-mediated rejection after kidney transplantation. Am J Transplant 19(2):368–380 12. Burton H, McLaughlin L, Shiu KY et al (2022) The phenotype of HLA-binding B cells from sensitized kidney transplant recipients correlates with clinically prognostic patterns of interferon-γ production against purified HLA proteins. Kidney Int 102(2):355–369 13. Sellare´s J, de Freitas DG, Mengel M et al (2012) Understanding the causes of kidney transplant failure: the dominant role of antibody-mediated rejection and nonadherence. Am J Transplant 12(2):388–399 14. Stringer D, Gardner L, Shaw O et al (2023) Optimized immunosuppression to prevent graft failure in renal transplant recipients with HLA antibodies (OuTSMART): a randomised controlled trial. eClinicalMedicine 56. Available from: https://www.thelancet.com/ journals/eclinm/article/PIIS2589-5370(22) 00548-X/fulltext. [cited 2023 May 9] 15. Bestard O, Couzi L, Crespo M et al (2021) Stratifying the humoral risk of candidates to a solid organ transplantation: a proposal of the ENGAGE working group. Transpl Int 34(6): 1005–1018 16. Tambur AR, Bestard O, Campbell P et al (2023) Sensitization in transplantation: assessment of Risk 2022 working group meeting report. Am J Transplant 23(1):133–149 17. Schinstock CA, Askar M, Bagnasco SM et al (2021) A 2020 Banff Antibody-mediated Injury Working Group examination of international practices for diagnosing antibodymediated rejection in kidney transplantation – a cohort study. Transpl Int 34(3):488–498 18. Tarlinton D, Good-Jacobson K (2013) Diversity among memory B cells: origin, consequences, and utility. Science 341(6151): 1205–1211

Chapter 13 Assessing the Affinity Spectrum of the Antigen-Specific B Cell Repertoire via ImmunoSpot® Noe´mi Becza, Zhigang Liu, Jack Chepke, Xing-Huang Gao, Paul V. Lehmann, and Greg A. Kirchenbaum Abstract The affinity distribution of the antigen-specific memory B cell (Bmem) repertoire in the body is a critical variable that defines an individual’s ability to rapidly generate high-affinity protective antibody specificities. Detailed measurement of antibody affinity so far has largely been confined to studies of monoclonal antibodies (mAbs) and are laborious since each individual mAb needs to be evaluated in isolation. Here, we introduce two variants of the B cell ImmunoSpot® assay that are suitable for simultaneously assessing the affinity distribution of hundreds of individual B cells within a test sample at single-cell resolution using relatively little labor and with high-throughput capacity. First, we experimentally validated that both ImmunoSpot® assay variants are suitable for establishing functional affinity hierarchies using B cell hybridoma lines as model antibody-secreting cells (ASC), each producing mAb with known affinity for a defined antigen. We then leveraged both ImmunoSpot® variants for characterizing the affinity distribution of SARS-CoV-2 Spike-specific ASC in PBMC following COVID-19 mRNA vaccination. Such ImmunoSpot® assays promise to offer tremendous value for future B cell immune monitoring efforts, owing to their ease of implementation, applicability to essentially any antigenic system, economy of PBMC utilization, highthroughput capacity, and suitability for regulated testing. Key words ELISPOT, FluoroSpot, B lymphocytes, Immune monitoring, Antibodies, Antibody titers, Immune memory, Functional affinity

1

Introduction

1.1 Definition of Affinity, Avidity, and Functional Affinity

Antibodies, also called immunoglobulins (Ig), can specifically bind to nearly any type of molecule (collectively termed an antigen), whereby the antigen-binding domain located in the hypervariable region of the antibody molecule (called the paratope) associates with a defined region of the antigen referred to as an epitope. The specificity and strength of antigen-antibody interactions depends

Paul V. Lehmann and Greg A. Kirchenbaum contributed equally and share senior authorship. Alexander E. Kalyuzhny (ed.), Handbook of ELISPOT: Methods and Protocols, Methods in Molecular Biology, vol. 2768, https://doi.org/10.1007/978-1-0716-3690-9_13, © The Author(s) 2024

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on the presence of complementary structures on both surfaces in three-dimensional space (“lock and key principle”) with the summation of attractive/repulsive forces contributing to the net binding strength. The binding forces involved in antibody-antigen interactions include (a) pairs of oppositely charged molecular groups, (b) hydrophobic regions capable of attracting each other, (c) coordinated hydrogen bonds, and (d) van der Walls-type interactions. As all four binding forces are reversible, the binding of antibodies to antigens is reversible too. Consequently, there is a constant “on-off” flickering of the paratope-epitope association that follows a second-order biochemical reaction. The association(Kon) and dissociation- (Koff) constants of the paratope-epitope interaction jointly determine the equilibrium constant (Kd) that defines “affinity” (see Note 1). The affinity of antibodies elicited during the course of an immune response can range from nearly undetectable (e.g., Kd < 10-4 M), to extremely high (e.g., Kd > 10-10) [1–8]. Consequently, a difference of greater than six orders of magnitude in antibody concentration can be required for low vs. high-affinity antibodies to attain the same level of epitope coverage. Thus, while all such “specific” antibodies can result in “specific” antigenbinding if they are present in sufficiently high concentration, it would take one million times more antibody molecules for a low-affinity antibody compared to a high-affinity antibody to achieve the same density of antigen binding! Therefore, the affinity of an antibody is a critical factor determining its capacity to contribute towards protective immune reactions. Affinity will suffice for defining the binding strength of an antibody to an antigen when only a single arm (so-called Fab fragment) of the antibody can interact with the antigen. This is the typical scenario when antibodies (immobilized or not) interact with individual antigen molecules in solution. This is because, on most protein antigens, the epitopes are singular unique structures, i.e., there are no other epitopes present on the same antigen molecule for the other arm of the antibody to associate with. Avidity defines the net binding strength when a single antibody molecule (that possesses 2–10 Fab fragments or arms) can attach to multiple epitopes that are structurally connected. This is the case when repetitive epitopes occur on the same antigen molecule and are within the reach of the antibody’s arms, or when separate, but physically linked, antigen molecules are sufficiently close in proximity that the antibody can associate with more than one antigen molecule simultaneously (e.g., when two membrane-anchored antigen molecules are in close proximity to each other). The net binding strength attained from such multivalent binding (avidity) is exponentially higher than that resulting from monovalent binding (affinity). This is because, as mentioned above, the interaction of the antibody’s paratope with the antigen’s epitope is a reversible

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“on” and “off” flickering; during the “off” state, in case of a monovalent interaction, the reaction partners can diffuse away. If, however, more than one arm of the same antibody molecule is simultaneously involved in unsynchronized “on/off” flickering, the time window in which two or more arms are dissociated is greatly reduced, and the ligand(s) could separate and diffuse away. To define affinity, single arms of antibodies (so-called Fab fragments) need to be studied; however, these do not occur naturally. As all naturally occurring antibodies in humans (and other commonly used animal models) have at least two arms (IgG, IgE, and monomeric IgA have two; IgM and dimeric IgA have 10 and 4, respectively), the occurrence of monovalent or multivalent binding (i.e., whether affinity or avidity applies), depends on the special circumstances of the antigen encounter. In terms of immunobiological consequences, however, the overall attachment strength of the antibody molecule with the antigen is fundamental for host defense. The latter is pragmatically called functional affinity. Since, in this chapter, we will also be analyzing the binding of bivalent antibodies to antigens, we adopt the term functional affinity, when appropriate, hereafter. 1.2 Biological Benefits of Antibodies with High Functional Affinity

A critical effector function of antibodies is their ability to bind and neutralize antigens (e.g., toxins or viruses) by preventing their association with cells of the host. Importantly, for this process to occur effectively, these antibodies need to possess a functional affinity for the antigen that surpasses that of the toxin/virus for their endogenous host receptor(s). Thus, the higher the functional affinity of these antibodies (e.g., the lower the Kd), the lower the concentration required for them to exert their neutralizing effector function. Moreover, another effector function of antibodies with high functional affinity is their ability to cross-link soluble antigens and generate immune complexes facilitating the effective elimination of the antigen by phagocytes. Activation of the complement system, another effector function initiated by antibodies, depends on the cross-linking of at least two of the six “arms” of the C1q molecule [9–11]; which is the first component that initiates the classical complement cascade. For this to occur, two antibody molecules need to be bound simultaneously, in close vicinity, so C1q can bind to both at once. As antibodies with high functional affinity bind more stably, even when they occur at low concentrations, they outperform antibodies with lower functional affinity in their capacity to initiate complement activation. The same applies when antibodies bind to antigen and label the latter for FcR-mediated elimination via phagocytosis (opsonization) or destruction (antibody-dependent cellular cytotoxicity, ADCC). For all these effector functions, the concentration of “specific” antibodies in bodily fluids is critical for their efficacy. As antibody

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levels tend to decline with time after an immune response, only those antibodies at the high-level end of the functional affinity spectrum will effectively contribute to host defense. 1.3 Affinity Maturation of the Antibody Response

Owing to the fundamental role that antibody functional affinity plays in immune protection, the B cell system has evolved means to maximize the affinity of antibodies deployed in the course of an immune response. This starts with the recombination of V(D)J gene segments within the IgH/IgL loci during early B cell ontogeny, a process capable of generating an estimated 1014 unique B cell receptors (BCR) [12] (see Note 2). As B cells are limited to the expression of a single BCR (with rare exceptions), each B cell is endowed with a unique antigen-binding specificity. Importantly, any given specificity occurs at a very low frequency among naive B (and T cells) and such cells continuously recirculate through secondary lymphoid tissues where they may encounter their cognate antigen (see Note 3). Both naive B and T cells are efficiently retained in secondary lymphoid tissues during the onset of a primary immune response when they first encounter “their” antigen, and, driven by (affinity-based) antigen receptor triggering, these cells are induced to proliferate. In this way, clonal expansions occur resulting in an increase in the frequency of antigen-specific B and T cells among all lymphocytes in the body. The activated T and B cells also acquire specialized effector functions, i.e., T cells become Th1/Th2/Th17/Tfh21 cells, each of which is capable of secreting a distinct cytokine signature, and B cells undergo Ig class switching that is characteristic of antigen-experienced B cells: they switch from IgM-expressing naive B cells to downstream Ig classes such as IgG or IgA, and within the IgG producers, to subclasses IgG1/ IgG2/IgG3/IgG4. Of note, the process of Ig class switching is governed by instructive signals provided primarily by T helper cells (see Note 4). During the primary immune response, the proliferating antigen-stimulated B cells also undergo an additional fundamental process aimed at improving the efficacy of the ensuing antibody response: the acquisition of somatic hypermutations (SHM) in the IgH and IgL encoding the antigen-binding variable regions of their BCR. Initially, naive B cells with adequate affinity for the antigen become stimulated; these cells can then acquire SHM as they undergo additional rounds of proliferation within the germinal center reaction. Their progeny, therefore, will consist of subclones that have BCR with a spectrum of affinities for the eliciting antigen: some with higher, and others with lower affinity. Of these daughter cells, only those with the highest affinity for the antigen continue to participate in the germinal center reaction and undergo further rounds of proliferation and acquisition of additional SHM. This process of positive selection for high-affinity subclones (and negligence of daughter cells with lower affinity) continues throughout

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the primary immune response and becomes more and more stringent as the antigen becomes gradually eliminated as a consequence of the ensuing successful immune response. This process of affinity maturation through the acquisition of SHM can also be re-initiated at a later time point if the antigen is reencountered, providing the cellular basis for why booster immunizations can progressively raise the affinity of the elicited antibody response. Whether in the context of a primary or secondary (recall) immune response, the B cell progeny endowed with the highest affinity BCR for the eliciting antigen will differentiate into plasma cells (PC) that secrete antibodies with identical specificity of the BCR expressed at the time of their disengagement from the germinal center reaction. If such antibody-secreting cells (ASC) settle into the bone marrow or other suitable niches, they can become long-lived PC (see Note 5). Relevant to this chapter, the affinity distribution of B cells for any given antigen is expected to be variable among human subjects, dependent on the dose and duration of antigen persistence during the primary immune response (that dictates the initial affinity-based positive selection process) and on the timing, dose, and duration of secondary, and possibly subsequent, antigen encounters, thus triggering further rounds of affinity-based selection of the antigenspecific B cell repertoire. Based on past efforts generating monoclonal antibody (mAb)-secreting B cell hybridomas, experience supports that the more booster shots that are given, the higher the chance of isolating a clone that secretes antibodies with very high affinity. 1.4 Measuring Antibody Functional Affinity in the Serum vs. the B Cells Themselves

It is important to understand the fundamental differences between immune protection mediated by the B cell system via the firstvs. second walls of adaptive humoral defense. Antibodies already present in serum and other bodily fluids can instantly bind to antigens as soon as they attempt to enter the body. This first wall of defense can prevent reinfections and it can be readily assessed by serum antibody measurements. Serum antibodies, however, are relatively short-lived molecules (see Note 6) and their continued presence depends on constant replenishment by PC. While PC are potentially long-lived, their lifespans are heterogenous and likely fall on a continuum [13, 14]. During the recent COVID-19 pandemic, we were reminded how rapidly following natural infection, or after vaccination, the induced specific serum antibody levels can decline [15, 16]. We also frequently learned that the detection of memory B cells (Bmem) is far more reliable for revealing whether an infection has occurred than measuring serum antibodies [17]. The second wall of B cell-mediated protection is conferred through the reactivation of Bmem. If the first wall of adaptive humoral defense fails, and a (re-)infection occurs, antigen-specific Bmem (and memory T cells) can rapidly engage in secondary immune responses. These lymphocytes can mount a stronger and

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faster counterattack against the offending pathogen because they are present in greatly increased numbers in the body compared to the numbers of naive B (and T) cells in the pre-immune repertoire. Moreover, many of these pre-existing memory cells have already undergone differentiation into effector lineages (Th1/Th2/Th17, etc. for T cells, IgG subclass, or IgA-switched B cells). Additionally, the B cell affinity maturation process re-engages during secondary immune responses, starting, however, from the elevated levels established following the prior antigen encounter(s). Studying the first wall (existing antigen-specific serum antibodies) and the second wall (the antigen-specific Bmem repertoire in the blood, see Note 7) of adaptive humoral immunity therefore provides fundamentally different information. The former provides a low-resolution and fading image of the past as it still applies to the integrity of the first wall, and the latter permits to gain a highresolution image of the second wall, thus assessing the immune potential present in case of a subsequent antigen encounter. Highresolution in this context means being able to assess individual B cells within the entire antigen-specific repertoire, the type and quantity of antibody they produce, and, relevant for this chapter, the functional affinity distributions of the individual B cells. 1.5 Measuring Functional Affinity by B Cell ImmunoSpot®

Being able to assess the functional affinity distribution of the antigen-specific Bmem compartment in an individual requires defining the functional affinity for hundreds (the more the better) of individual B cells to obtain a representative picture. Doing so by the standard approaches, either through surface plasmon resonance (SPR) or biolayer interferometry (BLI) measurements of mAbs, would be an effort too involved for immune monitoring purposes (see Note 8). We introduce here modifications of the ImmunoSpot® assay that promise to fill this gap. The principle of one of the underlying ImmunoSpot® variants suited for this purpose, the inverted assay, is shown in Fig. 1a. In brief, the membrane is coated with an antibody suited for capturing the Ig produced by ASC irrespective of their antigen specificity or affinity: in case of testing human PBMC, the capture reagents typically would be xenogeneic anti-human IgG, A, or E-specific antibodies, or a pan Ig-specific (anti-kappa + anti-lambda light chain) antibody (see Notes 9 and 10). Onto this lawn of Ig capture reagent, the cells are plated (see Note 11). During the time in which the ASC resides on the membrane, their secreted antibodies will be captured in close proximity to the respective ASC in the form of a secretory footprint (often also referred to as a “spot forming unit,” SFU) (see Note 12). After a brief period of cell culture during which the secreted Ig analyte is captured, the cells are removed from the plate (see Note 13), and the antigen probe is added to replicate wells containing the same number of ASC, but decreasing

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Fig. 1 Schematic representation of antigen-specific inverted or direct ImmunoSpot® assays. The PVDF membrane on the bottom of a 96-well plate is densely coated (a) with a pan anti-Ig(G) class-specific (in this example IgG) capture antibody that binds with high-affinity to secreted Ig(G) irrespective of antigen specificity or (b) with an antigen. The PBMC are plated on the resulting lawn of (a) pan anti-Ig(G) class-specific capture antibody, and individual ASC producing the particular Ig class will generate a secretory footprint or (b) antigen and individual ASC-producing antibody with sufficient affinity to the antigen will generate a secretory footprint. After removal of the cells, (A) the affinity-tagged (in this example His) antigen is added at a sufficient concentration to be retained by antigen-specific secretory footprints generated by ASC producing low- or highaffinity antibody or (b) antigen-bound antibody is visualized by biotinylated human Ig class/subclass (in this example IgG) specific detection antibodies. (a) Antigen-specific secretory footprints are then visualized using a biotinylated anti-affinity tag detection reagent; which is revealed by the addition of a fluorescently-conjugated streptavidin (FluoroSpot, as shown) or via an enzymatic reaction (ELISPOT, not shown). (b) Antigen-specific secretory footprints are revealed by the addition of a fluorescently-conjugated streptavidin (FluoroSpot, as shown) or via an enzymatic reaction (ELISPOT, not shown). (a, b) Counting the spot-forming units (SFU) per well reveals the number of antigen-specific ASC within the PBMC plated

(graded) concentrations of antigen. When the antigen probe is added in excess, all secretory footprints retain the antigen probe: ASC with low- and high-affinity will be revealed alike. As the concentration of antigen probe becomes limiting, however, increasingly only the high-affinity secretory footprints (SFUs) will capture a sufficient quantity of the antigen probe for their detection. For such affinity measurements, the cells need to be plated at a predetermined “Goldilocks” number that, being assay dependent, is between 50 and 200 SFUs/well (being at the upper end of the linear range between cell numbers plated and SFU counted, so that as many individual ASC can be assessed per well as possible, yet without ASC crowding interfering with the image analysis, see also Note 14). Testing a fixed number of replicate wells for each antigen concentration thus permits to assessment of the affinity spectrum within the same number (ideally ≥300) of antigen-specific ASC for each antigen concentration. The percentage of SFU lost with each successive reduction in antigen probe concentration reveals the affinity thresholds for the ASC subpopulations lost, respectively. A detailed protocol for such B cell affinity measurements is provided below.

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Fig. 2 Assessing the affinity of Granzyme B (GzB)-specific mAb by inverted ImmunoSpot® assay vs. surface plasmon resonance testing. In columns 1–4, the results of an inverted ImmunoSpot® assay are shown. Onto a lawn of anti-murine Igκ capture antibody, B cell hybridoma clones 12.4 and 30.1, each of which secretes GzB-specific mAb, and a third hybridoma clone that secretes a SARS-CoV-2 Spike-specific mAb, were plated at 200 cells per well, in the rows specified. His-tagged recombinant human GzB (rhGzB) was added in decreasing concentrations, as shown, and the captured rhGzB probe was detected via the His-tag. In the column on the right, replicate wells seeded with the B cell hybridomas, respectively, were detected using an anti-murine IgG detection capture antibody to verify the presence of IgG+ secretory footprints. KD values of the GzB-specific hybridomas for rhGzB, as established by surface plasmon resonance testing, are indicated on the right. ND not determined 1.6 B Cell Hybridoma Studies Validating ImmunoSpot® Affinity Measurements

While performing B cell affinity measurements using the inverted ImmunoSpot® approach outlined above seems simple and intuitive, this strategy has not yet been introduced. To experimentally validate this approach, we tested B cell hybridomas with established affinities for defined antigens. An example is shown in Fig. 2 using a pair of anti-human Granzyme B (GzB) hybridomas we generated during our efforts to raise mAbs against this protein. Among 7 such hybridomas, two clones secreted mAb at opposing ends of the antigen-retention spectrum when tested using an inverted ImmunoSpot® approach: GzB12.4 and GzB30.1. These two clones, along with an additional SARS-CoV-2 Spike-specific control hybridoma (generously provided by Giuseppe A. Sautto), were all plated at ~200 cells per well. Visualizing the secretory footprints with an anti-murine IgG detection antibody showed that most cells within each of the three hybridoma clones were capable of generating a secretory footprint and that the per-cell IgG productivity rates were comparable across the three (the SFU sizes and fluorescence intensities were similar between individual ASC within each hybridoma line and between the three B cell hybridoma clones (Fig. 2). When recombinant His-tagged human GzB (rhGzB) was added at a concentration of 500 ng/mL, the secretory footprints of both

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GzB12.4 and GzB30.1 bound the antigen revealing ~200 SFU, whereas the footprints of the control hybridoma did not capture rhGzB. When the concentration of rhGzB was decreased in a 1:5 dilution series, footprints of GzB12.4 continued to capture this antigen down to a concentration of 4 ng/mL, whereas the secretory footprints of clone GzB30.1 were no longer discernable when the rhGzB probe was added at concentrations lower than 100 ng/ mL. By ImmunoSpot®, clone GzB12.4 was identified as secreting mAb with high affinity for rhGzB, whereas clone GzB30.1 secreted mAb that appeared to possess a substantially lower affinity for this molecule. To verify by an independent method that this was indeed the case, we purified mAb from GzB12.4 and GzB30.1 and established their functional affinity for rhGzB using surface plasmon resonance (Biacore): a KD value of 7.0 × 10-11 was calculated for mAb GzB12.4, while the KD value of mAb GzB30.1 was 2.3 × 10-8. Thus, both techniques concurred that clone GzB12.4 secreted mAb with a substantially higher functional affinity for rhGzB than the mAb produced by clone GzB30.1. Supplementary Figures 1 and 2 (see Note 15) provide further examples in which inverted ImmunoSpot® assays provided confirmatory results for hybridoma pairs secreting mAb with differences in functional affinity for alternative antigens; influenza hemagglutinin (H1) or SARSCoV-2 Spike, respectively. Another semiquantitative approach for assessing the affinity of individual ASC via ImmunoSpot® is based on the direct assay (Fig. 1b), studying the SFU morphologies on antigen-coated wells, in particular when coated with graded antigen densities. In direct B cell ImmunoSpot® assays, the membrane is either coated with the antigen itself, or adapter molecules are used for attaching the antigen to the membrane (see Note 16). As such, only ASC which produce an antibody with sufficient functional affinity for the membrane-associated antigen will be capable of generating a secretory footprint. Next to the per cell quantity of secreted antibody, the antibodies’ functional affinity for the antigen defines the size and density characteristics of the resulting secretory footprint: ASC that produce high-affinity antibodies will create crisp and more intense spots, whereas ASC-producing antibodies with reduced functional affinity yield more diffuse and fainter/sparse footprints [18]. The difference between secretory footprints generated by high- or low-affinity ASC will be further accentuated when the bonus effect of multivalent binding is negated by reducing the antigen-coating density. At low antigen-coating densities, antibodies can only bind using a single “arm,” i.e., the binding is defined by affinity alone; if the antigen is coated densely onto the assay membrane, however, the antibody can attach with both arms to neighboring antigen molecules and now avidity can amplify the antibody’s functional affinity. Figure 3 illustrates the spot morphologies observed when murine B cell hybridomas secreting mAb with

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Fig. 3 Direct ImmunoSpot® assay leveraging limiting antigen coating reveals differential mAb functional affinity for influenza hemagglutinin. The specified murine B cell hybridomas, each secreting monoclonal antibody (mAb) reactive with recombinant hemagglutinin (rHA) protein (COBRA P1), however, with previously defined differential affinity [19], were plated at the Goldilocks numbers (100 cells per well) into wells that had been pre-coated with decreasing concentrations of rHA protein, as specified. The captured secretory footprints were visualized using anti-murine IgG1-specific detection antibodies. Secretory footprints from each of the B cell hybridoma lines were readily apparent in wells coated with a high concentration of rHA. However, the number of secretory footprints and their density declined at different rates for the two clones when hybridoma cells were seeded into wells coated with decreasing concentrations of rHA. Based on the results of this experiment, the mAb secreted by the 1-B7.2 B cell hybridoma line has an increased functional affinity for the coated rHA protein compared to the mAb secreted by 2-A10.4, confirming the previously published affinity hierarchy established by Sautto et al. [19]

differential functional affinities for an influenza hemagglutinin (H1) were evaluated for secretory footprint formation on membranes in which the antigen-coating density was progressively limited. 1.7 B Cell Affinity Distribution Measurements in PBMC with Inverted ImmunoSpot® Tests

After having experimentally validated the suitability of both ImmunoSpot® assay variants for B cell affinity measurements using hybridomas as model ASC, we set out to translate these tests for studies of antigen-specific ASC in human PBMC. As described above, B cells endowed with a wide affinity spectrum for the eliciting antigen participate in the primary immune response. However, progressively the engaged BCR repertoire is selected for those with increasingly high affinity, particularly in cases of prolonged or repetitive antigen exposure. Every individual’s Bmem repertoire at a given time point is therefore a variable defining that person’s capacity to engage in an effective anamnestic, adaptive humoral defense

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reaction. One critical variable for evaluation is the clonal size of the antigen-specific Bmem compartment, as revealed by their frequency in PBMC. A second critical variable for assessment is the antibody class/subclass usage of antigen-specific Bmem since this will offer insight into what types of antibodies will be secreted acutely following antigen reencounter. Third, but not least, the frequency of high-affinity Bmem capable of producing the desired Ig class/subclass will define the efficacy of the anamnestic antibody response elicited in case of antigen reencounter. Importantly, all three of these parameters can readily be assessed using tailored B cell ImmunoSpot® assays. As PBMC can be frozen without losing B cell functionality (see Note 11), we recommend undertaking high-resolution B cell ImmunoSpot® testing using aliquots of the same sample and a tiered approach. In the first experiment, the frequency of antigenspecific Bmem-derived ASC is established by performing serial dilutions of the cell input to define the Goldilocks number by linear regression (see Notes 14 and 17). Of note, this first experiment only requires ~2 × 106 cells per antigen to define the frequency of antigen-specific ASC secreting each Ig class/IgG subclass, respectively (see Note 18 and see the chapter by Yao et al. in this volume [20] that is dedicated to such measurements). In a subsequent experiment, thawing an additional aliquot(s) of the PBMC sample, the affinity distribution of the antigen-specific ASC is established at the Goldilocks number, testing each antigen concentration in the same number of replicate wells. The number of PBMC needed for this second experiment depends on the Goldilocks number, the number of replicate wells needed to obtain a total of minimally 300 SFU, and the concentration range of antigen being evaluated. It can be calculated with high accuracy once the Goldilocks number has been established. In the example provided in Fig. 4, as detailed below, the number of PBMC needed to perform an inverted ImmunoSpot® assay encompassing 4 concentrations of SARSCoV-2 Spike (receptor-binding domain, RBD) antigen probe, each evaluated in 10 replicate wells with ~50 SFU/well (thus assessing the affinity distribution within ~500 IgG+ Bmem of this specificity) was 4 × 106 PBMC. In Fig. 4, we present typical results obtained using the inverted ImmunoSpot® assay to assess the affinity distribution of antigenspecific ASC present within a human PBMC sample using limiting quantities of the antigen probe for their detection. In this case, cryopreserved PBMC collected from a healthy human donor 7 days following a second COVID-19 mRNA vaccination were tested. In the first experiment, the Goldilocks number for the SARS-CoV2 RBD-specific IgG+ ASC was determined for this PBMC sample (as described above) at 50 SFU/1 × 105 PBMC per well. In the initial test, a saturating concentration of the RBD probe was used and enabled the detection of all antigen-specific IgG+ ASC;

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Fig. 4 Measuring B cell affinity distribution in PBMC with inverted ImmunoSpot® assays using limiting quantities of the antigen probe for detection. Cryopreserved PBMC collected from a healthy human donor 7 days following a second COVID-19 mRNA vaccination were tested. The PBMC were seeded into an inverted assay at 1 × 105 PBMC per well, which in a previous experiment was determined to result in ~50 SARS-CoV2 Spike (RBD)-specific IgG+ spot-forming units (SFU), a number suitable for studying individual secretory footprints. The His-tagged RBD probe was added to 10 replicate wells at the specified concentrations, followed by the detection of the His-tagged, plate-bound antigen. (a–d) Representative well images are shown for each RBD probe concentration. (e) Standard bar graph representation of cumulative SFU counts at each probe concentration. (f) Composite bar representation of the same data with color code specifying spot numbers detected at each probe concentration. (g) Intensity of spots detected at each RBD probe concentration was stratified into ten intensity groupings (denoted using grayscale) and the percentage of SFU residing in each “bin” is reflected by the width of the corresponding section. (h) Flow cytometry standard (FCS)-type intensity by size scatter plot analysis of the cumulative spots detected using 100 ng/mL or 12.5 ng/mL of the RBD probe, respectively

low- and high-affinity alike. Thawing a second aliquot of these cryopreserved cells, the PBMC were plated into 10 replicate wells for each of the four RBD probe concentrations tested at the Goldilocks number. In Fig. 4a–d, representative well images are shown for each RBD probe concentration. A reduction in SFU numbers is readily seen visually as the RBD probe concentration decreases. The corresponding SFU numbers are shown in a conventional bar diagram in Fig. 4e; most of the expected cumulative ~500 SFU were detected at 100 ng RBD/mL. Figure 4f shows these results in a composite bar diagram format that is better suited for comparing affinity distributions between different PBMC samples, e.g., to visualize affinity maturation for a subject’s B cell response with repeat immunizations, or for representing such data for cohorts.

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Figure 4 also illustrates that, next to the decrease in SFU, the fluorescence intensity of the individual secretory footprints changes as the RBD probe concentration decreases. Footprints of highaffinity ASC continue to stain bright as the RBD probe concentration declines, but ASC whose threshold of binding capability is approximated at the given probe concentration become fainter. Such transitions can be graphically captured. In Fig. 4g, the intensity of spots detected at each RBD probe concentration is stratified into ten intensity groupings, and the percentage of SFU residing in each “bin” is reflected by the width of the corresponding section. Flow cytometry standard (FCS)-type representation of ImmunoSpot® data also permits such further high-content data analysis. Figure 4h shows an FCS-type scatter plot representation (fluorescence intensity vs. size) of spot morphologies, and the difference seen between the highest and lowest RBD probe concentrations. Inverted ImmunoSpot® assays are particularly well-suited for measuring B cell affinity distributions when the frequency of antigen-specific ASC producing the Ig class/subclass to be captured on the membrane (e.g., IgG) is relatively high within all ASC present in the sample producing that particular Ig class/subclass (see Note 19). This is often the case when studying plasmablasts that transiently circulate in the blood acutely (5–9 days) after initiation of a B cell response (see Note 20). If the frequency of antigen-specific ASC vs. total ASC is low in a given sample, direct ImmunoSpot® assays offer an alternative approach for measuring the affinity distribution of Bmem-derived ASC. 1.8 B Cell Affinity Distribution Measurements in PBMC with Direct ImmunoSpot® Tests

Direct ImmunoSpot® assays inherently reveal information about ASC affinity. Following basic rules of antigen–antibody binding, ASC that produce high-affinity antibodies will leave dense and sharp secretory footprints on membrane-bound antigens, while ASC-producing antibodies with lower functional affinity for the membrane-associated antigen will form faint and diffuse spots [18]. Performing direct ImmunoSpot® assays under conditions when the membrane is coated with graded antigen densities helps to further enhance such affinity studies. Typical results for these types of tests done on human PBMC are shown in Fig. 5, in this case using PBMC from a COVID-19 mRNA vaccine recipient. Like for the inverted ImmunoSpot® assay, here too the cells were plated into all replicate wells at an assay-specific, Goldilocks number (3 × 104 PBMC/well) needed to achieve 50 Spike-specific SFU/well. A direct ImmunoSpot® assay was performed in which, however, the Spike protein coating density was graded. In Fig. 5a–d, representative well images are shown for each coating concentration of Spike protein. Here too, a reduction in SFU numbers is seen as the coating density was decreased. The corresponding SFU numbers are shown in a conventional bar diagram in Fig. 5e, and a composite bar diagram format in Fig. 5f.

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Fig. 5 ImmunoSpot® enables the evaluation of ASC affinity through limiting antigen coating. Cryopreserved PBMC collected from a healthy human following a second COVID-19 mRNA vaccination were tested. The PBMC were seeded into the assay at 3 × 104 PBMC per well, which in a previous experiment was determined to result in ~50 SARS-CoV-2 Spike-specific IgG+ spot-forming units (SFU), a number suitable for studying individual secretory footprints. Assay wells were initially coated with anti-His capture antibody, followed by affinity capture of Spike protein at 10, 5, 2.5, or 1 μg/mL, in 20 replicate wells, respectively. (a–d) Representative well images are shown for each Spike coating concentration. (e) Standard bar graph representation of cumulative SFU counts at each Spike coating concentration. (f) Composite bar representation of the same data with color code specifying spot numbers detected at each Spike coating concentration. (g) Flow cytometry standard (FCS)-type intensity by size scatter plot analysis of the cumulative spots detected in wells coated with either 10 or 2.5 μg/mL of Spike protein, respectively

In this chapter, we present data demonstrating that B cell ImmunoSpot® assays are capable of distinguishing between model ASC (murine B cell hybridomas) known to secrete mAb with variable functional affinity for defined antigens. Further, through characterizing the B cell response elicited in COVID-19 mRNA vaccine recipients, we highlight the utility of antigen-specific, inverted, and direct ImmunoSpot® assays for characterizing the functional affinity distribution of the antigen-specific B cells present in PBMC. In the following Materials and Methods section, we provide detailed protocols for performing such inverted and direct ImmunoSpot® assays. Here we also refer to a chapter by Yao et al. [21] in this volume that describes defining of Goldilocks number in detail, and to a chapter by Karulin et al. [29] on highcontent image analysis of B cell-derived SFU. The underlying B cell biology is elaborated in greater detail in a chapter by Lehmann et al. [18], also in this volume.

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Materials

2.1 Thawing of Cryopreserved PBMC

1. Class II biosafety cabinet (BSC) 2. Cryopreserved PBMC sample(s) (see Note 11). 3. 70% (v/v) ethanol (EtOH) 4. DNase-containing washing medium (pre-warmed to 37 °C) (see Note 21). 5. Complete B cell medium (BCM) (pre-warmed to 37 °C) (see Note 22).

2.2 In Vitro Polyclonal Stimulation of B Cells in PBMC 2.3 Antigen-Specific, Single-Color Inverted FluoroSpot Assay

1. B-Poly-S 2. 25 cm2 sterile culture flask 3. Humidified incubator set at 37 °C, 5% CO2 1. Commercially available, single-color human IgG Inverted (His) B cell kit (see Note 23). 2. His-tagged recombinant protein (see Note 24). 3. 190 proof (95%) EtOH 4. Cell culture-grade water. 5. Reagent reservoir(s) (sterile). 6. 0.05% Tween-PBS wash solution 7. 0.1 μm low-protein binding syringe filter 8. Plate washer. 9. Vacuum manifold. 10. ImmunoSpot® S6 Ultimate 4 LED Analyzer, or a suitable instrument equipped with the appropriate detection channels, running CTL’s ImmunoSpot® UV.

2.4 Antigen-Specific FluoroSpot Assay (Affinity Capture Coating)

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1. Commercially available, single-color human IgG affinity capture (His) B cell kit (see Note 25). 2. His-tagged recombinant protein (see Note 26).

Methods

3.1 Thawing of Cryopreserved PBMC (Sterile Conditions)

1. Place cryovial(s) into a 37 °C bead bath, or better, glass bead bath, for 8 min to thaw. 2. Remove cryovial(s) and wipe with 70% EtOH inside the BSC before unscrewing the cap(s). 3. Using a sterile pipette, transfer contents of cryovial(s) into a labeled conical tube (if applicable, up to 5 vials of the same donor’s cell material can be pooled in one conical tube).

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4. Rinse each of the cryovials with 1 mL of warm anti-aggregate solution. Add the warm rinse solution to the conical tube dropwise while swirling the tube to ensure its adequate mixing with the cells in the thawing medium. 5. Double the volume of the cell suspension by dropwise addition of warm anti-aggregate solution while swirling the tube to ensure adequate mixing of the cells and thawing medium. 6. Continue doubling the volume of the cell suspension by dropwise addition of warm anti-aggregate solution while swirling the tube until the cryopreserved cell material has been diluted ten-fold. If multiple cryovials are pooled, calculate using 1 mL of cryopreserved cell suspension + 9 mL of anti-aggregate solution to determine the necessary final resuspension volume. 7. Centrifuge balanced tubes at 330 × g for 10 min with the centrifuge brake on, non-frigerated. 8. Decant supernatant and resuspend the cell pellet(s) using pre-warmed B cell medium (BCM) to achieve a cell density of ~2–5 × 106 cells/mL. You may estimate this number assuming a typical recovery of 70–80% of the frozen PBMC. 9. Pipet 15 μL of live/dead cell counting dye onto a piece of parafilm to form a droplet. 10. Remove 15 μL of cell suspension and combine with a droplet of live/dead cell counting dye. Pipet up and down three to five times to mix the sample while avoiding the formation of bubbles. 11. Transfer 15 μL of the cell and dye suspension into each chamber of a hemacytometer. 12. Determine live cell count and viability using CTL’s live/dead cell counting suite. 13. Increase volume of cell suspension(s) with additional sterile PBS and centrifuge balanced tubes at 330 × g for 10 min with a centrifuge, non-refrigerated, brake on. 14. Decant supernatant and gently resuspend the cell pellet(s) using pre-warmed BCM at a cell density of ~2–4 × 106 cells/mL. 3.2 In Vitro Polyclonal Stimulation of B Cells in PBMC (Sterile Conditions)

1. Dilute CTL’s B-Poly-S polyclonal stimulation reagent 1:500 into prewarmed BCM to achieve a final concentration of 2X. Into labeled sterile culture vessels, add 50% vol of BCM containing 2X concentration of B-Poly-S. 2. Add the same volume of cell suspension at ~2–4 × 106 cells/ mL to achieve a final culture at ~1–2 × 106 cells/mL with 1X potency of CTL’s B-Poly-S polyclonal stimulation reagent (see Note 27). 3. Transfer culture vessels (flasks or plates) into a humidified incubator set at 37 °C, 5% CO2 for 4–6 days (96–144 h).

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1. One day before plating cells (Day 1), prepare 70% EtOH and anti-human IgG capture antibody solutions. 2. Remove underdrain and pipet 15 μL of 70% EtOH solution into the center of each well (or designated wells) of the assay plate. Immediately after the addition of the 70% EtOH solution to the entire plate (or designated wells), add 180 μL/well of PBS (see Note 30). Decant and wash wells again with 180 μL/well of PBS. 3. Decant the assay plate, replace underdrain, and immediately add 80 μL/well of the anti-human IgG capture antibody solution into each well (or designated wells) of the low autofluorescence PVDF-membrane plate provided with the kit (see Note 31). 4. Incubate the plate overnight at 4 °C in a humified chamber. 5. On the day of the assay (Day 0), decant the assay plate and wash wells with 180 μL/well of PBS. Next, decant the plate and add 150 μL/well of pre-warmed BCM to block the plate (≥1 h at RT). 6. If using PBMC following polyclonal activation in vitro, collect the cell suspension(s) and transfer it into labeled conical tube(s). Wash culture vessel(s) interior to recover remaining cells with sterile PBS and transfer to the corresponding conical tube(s). Increase the volume to fill the conical tube with additional PBS and then centrifuge balanced tubes at 330 × g for 10 min with the centrifuge brake on. Alternatively, thaw previously cryopreserved PBMC as detailed above if prior in vitro stimulation is not required to elicit antigen-specific ASC activity in the sample(s). 7. Decant supernatant and resuspend the cell pellet(s) using pre-warmed BCM to achieve a cell density of approximately 2–5 × 106 cells/mL. 8. Pipet 15 μL of live/dead cell counting dye onto a piece of parafilm to form a droplet. 9. Remove 15 μL of cell suspension and combine with a droplet of live/dead cell counting dye. Pipet up and down three to five times to mix the sample while avoiding the formation of bubbles. 10. Transfer 15 μL of the cell and dye suspension into each chamber of a hemacytometer. 11. Determine live cell count and viability using CTL’s live/dead cell counting suite. 12. Increase volume of cell suspension(s) with additional sterile PBS and centrifuge balanced tubes at 330 × g for 10 min with centrifuge brake on (see Note 32).

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13. Decant supernatant and resuspend the cell pellet(s) using pre-warmed BCM at the pre-determined cell density to achieve the so-called “Goldilocks” cell input of 50 SFU/well (see Notes 19, 28, and 29). 14. Decant the BCM used for blocking the assay plate and replace it with 100 μL/well of pre-warmed BCM to block the plate. 15. Into a reagent reservoir, add cell suspension adjusted to a density of 500 SFU/mL (see Note 14). Using a multichannel pipet, transfer 100 μL of the cell suspension(s) into designated wells of the assay plate. Gently tap the edges of the ImmunoSpot® assay plate to ensure equal distribution of the cell input. 16. Incubate cells in the assay plate for 16–18 h at 37 °C, 5% CO2 (see Note 33). 17. After completion of the assay incubation period, remove the plate and decant cells. Wash the plate two times with PBS (200 μL/well), followed by two additional washing steps with 0.05% Tween-PBS wash solution (see Note 34). 18. Prepare His-tagged antigen probe solution at limiting concentrations (see Note 35) according to kit protocol and pass through a 0.1 μm low protein binding syringe filter to remove any protein aggregates. 19. Decant 0.05% Tween-PBS wash solution, add 80 μL/well of the various His-tagged antigen probe solutions (containing limiting/graded quantities of antigen probe) into designated wells, and incubate for 2 h at RT (protected from light). 20. Wash plate(s) two times with 0.05% Tween-PBS wash solution. 21. Prepare anti-His detection antibody solution according to kit protocol and pass through a 0.1 μm low-protein binding syringe filter to remove any aggregates. 22. Decant 0.05% Tween-PBS wash solution, add 80 μL/well of anti-His detection antibody solution into designated wells, and incubate for 1 h at RT (protected from light). 23. Wash plate(s) two times with 0.05% Tween-PBS wash solution. 24. Prepare tertiary solution according to kit protocol and pass through a 0.1 μm low-protein binding syringe filter to remove any aggregates. 25. Wash plates(s) twice with distilled water. 26. Remove the protective underdrain and place the plate face down on the vacuum manifold. Completely fill the backside of the plate with distilled water and apply a vacuum to draw water through the membrane (“back to front”) (see Note 36).

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27. Allow the plate to dry completely, protected from light (see Note 37). 28. Scan and count plate(s) with a suitable analyzer equipped with the appropriate detection channel (see Note 38). 3.4 Antigen-Specific (Affinity Capture Coating) FluoroSpot Assay (Limiting Antigen-Coating Concentrations)

1. Two days before plating cells (Day 2), prepare 70% EtOH and anti-His affinity capture antibody solutions. 2. Remove underdrain and pipet 15 μL of 70% EtOH solution into the center of each well (or designated wells) of the assay plate. Immediately after the addition of the 70% EtOH solution to the entire plate (or designated wells), add 180 μL/well of PBS (see Note 30). Decant and wash wells again with 180 μL/well of PBS. 3. Decant the assay plate, replace underdrain, and immediately add 80 μL/well of the anti-His affinity capture antibody solution into each well (or designated wells) of the low autofluorescence PVDF-membrane plate provided with the kit (see Note 39). 4. Incubate the plate overnight at 4 °C in a humified chamber. 5. The following day (Day 1) dilute the His-tagged protein (s) into diluent B (provided with the kit) at pre-determined limiting/graded concentrations (see Note 26). 6. Decant the assay plate and wash wells with 180 μL/well of PBS. Immediately, add 80 μL/well of the corresponding His-tagged protein coating solutions into the designated wells. 7. Incubate the plate overnight at 4 °C in a humidified chamber. 8. On the day of the assay (Day 0), decant the assay plate and wash wells with 180 μL/well of PBS. Next, decant the plate and add 150 μL/well of pre-warmed BCM to block the plate (≥1 h at RT). 9. If using PBMC following polyclonal activation in vitro, collect the cell suspension(s) and transfer it into labeled conical tube (s) (see Note 27). Wash culture vessel(s) interior with sterile PBMC and also transfer into the corresponding conical tube (s). Increase volu vitrome to fill up tube(s) with additional PBS and then centrifuge balanced tubes at 330× g for 10 min with centrifuge brake on. Alternatively, follow the procedures detailed above to obtain freshly isolated PBMC, or to thaw PBMC that were previously cryopreserved, if prior in vitro stimulation is not required to elicit antigen-specific ASC activity in the sample(s). 10. Decant supernatant and resuspend the cell pellet(s) using pre-warmed BCM to achieve a cell density of approximately 2–5 × 106 cells/mL.

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11. Pipet 15 μL of live/dead cell counting dye onto a piece of parafilm to form a droplet. 12. Remove 15 μL of cell suspension and combine with a droplet of live/dead cell counting dye. Pipet up and down three to five times to mix the sample while avoiding the formation of bubbles. 13. Transfer 15 μL of the cell and dye suspension into each chamber of a hemacytometer. 14. Determine live cell count and viability using CTL’s live/dead cell counting suite. 15. Increase volume of cell suspension(s) with additional sterile PBS and centrifuge balanced tubes at 330× g for 10 min with centrifuge brake on (see Note 32). 16. Decant supernatant and resuspend the cell pellet(s) using pre-warmed BCM at the pre-determined cell density to achieve the so-called “Goldilocks” cell input of 50 SFU/well. 17. Decant the BCM used for blocking the assay plate and replace it with 100 μL/well of pre-warmed BCM to block the plate. 18. Into a reagent reservoir, add cell suspension adjusted to a density of 500 SFU/mL (see Note 18). Using a multichannel pipet, transfer 100 μL of the cell suspension(s) into designated wells of the assay plate. Gently tap the edges of the ImmunoSpot® assay plate to ensure equal distribution of the cell input. 19. Incubate cells in the assay plate for 16–18 h at 37 °C, 5% CO2 (see Note 33). 20. After completion of the assay incubation period, remove the plate and decant cells. Wash the plate two times with PBS (200 μL/well), followed by two additional washing steps with 0.05% Tween-PBS wash solution (see Note 34). 21. Prepare detection antibody solution by following kit protocol and pass through a 0.1 μm low protein binding syringe filter to remove any aggregates. 22. Decant 0.05% Tween-PBS wash solution, add 80 μL/well of detection antibody solution into designated wells, and incubate for 2 h at RT (protected from light). 23. Wash plate(s) two times with 0.05% Tween-PBS wash solution. 24. Prepare a tertiary solution by following kit protocol and pass through a 0.1 μm low protein binding syringe filter to remove any aggregates. 25. Decant 0.05% Tween-PBS wash solution, add 80 μL/well of tertiary solution into designated wells, and incubate for 1 h at RT (protected from light). 26. Wash plates(s) twice with distilled water.

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27. Remove the protective underdrain and place the plate face down on the vacuum manifold. Completely fill the backside of the plate with distilled water and apply a vacuum to draw water through the membrane (“back to front”) (see Note 36). 28. Allow the plate to dry completely, protected from light (see Note 37). 29. Scan and count plate(s) with a suitable analyzer equipped with the appropriate detection channels (see Note 38).

4

Notes 1. The unit in which antibody affinity is measured is molarity (M). It defines the concentration of soluble antibody required to attain 50% of the maximal antigen binding capacity, e.g., by surface plasmon resonance (SPR) or biolayer interferometry (BLI). 2. There are 129 functional V (variable), 27 D (diversity), and 9 J (joining) gene segments available within the human IgH locus located on chromosome 14 [21]. In concert with the large number of V and J gene segments present within the Igκ (chromosome 2) and Igλ (chromosome 22) loci, respectively, plus the addition and deletion of nucleotides at the junctional borders of the recombined gene segments, and the pairing of IgH and IgL proteins to generate a functional B cell receptor (BCR), it is estimated that up 1014 different BCR specificities can be created through combinatorial diversity. As the adult human body has ~1012 lymphocytes, of which 1–10% in the blood are B cells, one can estimate that when an antigen is first encountered, there are very few naive B cells in the entire body that are endowed with a specific BCR. 3. An immune response requires that B and T cells encounter antigens in a secondary lymphoid tissue, such as a draining lymph node (LN) or the spleen. Naive B and T cells continuously recirculate among secondary lymphoid tissues, arriving there via the bloodstream and entering via the “lymph node homing receptors” they express. After searching the secondary lymphoid tissue for the presence of “their” antigen, if such is absent, they return to the blood via the afferent lymph vessels to continue their surveillance of other secondary lymphoid tissues, as well. Antigens are transported to the regional LN (s) that drain these areas of the body. The antigens arrive in that LN(s) either with the efferent lymph fluid or via dendritic cells actively transporting them. In this way, the three essential components of an efficient immune response (antigen, antigen-specific B cells, and antigen-specific T helper cells) are all

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brought together in the LN(s). The architecture of the LN is also specialized in that it provides the ideal microenvironment for the ensuing immune response to evolve. 4. Naive T cells undergoing a primary immune response in secondary lymphoid tissues are sensitive to the type of cytokine/ chemokine microenvironment created by cells of the innate immune system, as the latter become activated by pattern receptor-perception of the antigen. In this way, through different local microenvironments, Th1, Th2, Th17, and other type of T cell lineages arise. The process of Ig class switch recombination, in turn, is dictated by the type of cytokines these “helper” T cells secrete in response to recognizing cognate peptides presented by antigen-specific B cells. Engaging the “right” class/subclass of T and B cell responses is critical for mounting an effective antigen clearance/control without inciting collateral immunopathology. 5. While still not fully elucidated, plasma cells require specific stimuli provided by stromal and other accessory cells for their longevity and long-term residence in specialized survival niches [13, 22]. 6. The half-life of IgG1, IgG2, and IgG4 in humans is 21–28 days, and it is 7 days for IgG3 [23]. The half-life of IgA and IgM is even shorter (3–7 days) [24, 25]. IgE has a halflife in serum of 2–3 days [26]. 7. While antibody-secreting plasma cells are tissue-resident in the bone marrow or other specialized survival niches, memory B cells (Bmem) recirculates among secondary lymphoid tissues, just as naive B cells do (see also Note 3). Returning to the bloodstream via efferent lymph vessels, the Bmem are carried with the bloodstream where they egress into remote secondary lymphoid tissues and survey for their cognate antigen. By sampling blood, one can therefore gain insights into the Bmem compartment present in the body. 8. Defining the affinity distribution of, e.g., 300 antigen-specific B cells in an individual by SPR (or BLI) would mean having to either establish 300 antigen-specific B cell hybridoma clones, or generate 300 paired IgH/IgL sequences and suitable expression constructs, to express and purify these 300 monoclonal antibodies (mAb), and then to test the affinity of each of the 300 mAbs, one by one. Compare this effort and cost to the ease with which information on the affinity distribution of antigenspecific Bmem/ASC can be obtained via the ImmunoSpot® approach introduced here. 9. To the choice of capture antibody: when plasmablast populations elicited acutely (days 5–9) following induction of a B cell response are being studied, most of the spontaneously

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producing ASC in the blood will be antigen-specific. In such a case, the use of a pan Ig-specific (anti-kappa + anti-lambda light chain) capture antibody is justified and enables assessment of all classes and subclasses of antigen-specific antibodies, including IgM. In this case, IgM+ secretory footprints are inferred to originate from antigen-specific B cells engaging in a primary immune response. Furthermore, such inverted assays performed using pan Ig-specific capture conditions can be multiplexed with detection reagents which enable resolving the Ig class/subclass usage of distinct antigen-specific secretory footprints. 10. When inverted ImmunoSpot® assays are performed with in vitro polyclonally stimulated PBMC, however, third-party IgA and IgM-producing B cells are also induced in high numbers and readily compete for “real estate” on membranes coated with pan Ig capture antibodies. Moreover, the presence of IgM-derived secretory footprints generated by naive B cells possessing broad or highly cross-reactive specificities also hinders the specificity of such inverted assays. To overcome these obstacles, we recommend the usage of Ig class-specific capture antibodies; such as the anti-human IgG capture antibody shown in Fig. 1. In this way, only in vivo class-switched B memory cells are being interrogated in the ImmunoSpot® test. 11. Peripheral blood mononuclear cells (PBMC) can be cryopreserved without loss of spontaneous, or polyclonal stimulationinduced ASC activity ([27] and Becza et al., manuscript in preparation). This permits “direct ex vivo” batch testing of samples irrespective of the time-point of their actual collection. By freezing the cells in aliquots, assay results can be reproduced with high accuracy [17], and/or follow-up experiments can be performed building on the previous test result that establishes the “Goldilocks number” of antigen-specific ASC in a given PBMC sample, as described in Note 14. It is important for planning cryopreservation of PBMC to know that any number of PBMC can be frozen per vial between 1 and 10 million PBMC, in each case recovering at thawing about 90% viable and fully functional B cells, and that after 5 days of in vitro polyclonal B cell stimulation about 50% of these cells will be recovered as viable cells for testing (Becza et al., manuscript in preparation). 12. In the inverted assay, as shown in Fig. 1a, the capture of antibodies produced by ASCs occurs at a fixed high affinity, that of the anti-IgG capture antibody for the ASC-derived IgG. Only the net amount of IgG produced per ASC defines the size and density of the resulting secretory footprint. The affinity of the antibody captured in the footprint will be revealed, however, when the antigen is added at various concentrations.

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13. The B cells survive the ImmunoSpot® assay unharmed and can potentially be re-used in subsequent functional assays, for cloning, or analysis via means of molecular biology. 14. In the initial test, it is suggested to plate PBMC (or other single-cell suspensions) in serial dilution over a wide range of cell numbers per well because the frequencies of antigen-specific ASC can span orders of magnitudes between individuals [17] and ASC-producing different classes and subclasses of antigen-specific (and total) Ig also spans a wide range within an individual, and in the human population. In this way, not only the frequency of ASC can be reliably established by linear regression analysis [28] within the entire cell material tested but the Goldilocks number for secretory footprints can be defined as the maximal number of cells that can be plated while still being able to discern clearly individual secretory footprints. This number for an optimized, antigen-specific (SARS-CoV-2 Spike RBD) inverted assay is 50 (maximally 100) SFU/well using anti-IgG specific capture antibodies and Bmem-derived ASC following in vitro polyclonal stimulation of PBMC [17]. Once the Goldilocks number has been established, in a second experiment, the cell number(s) can be chosen accordingly for creating replicates containing ≥300 individual ASC-derived footprints per antigen concentration for studying the functional affinity distribution. 15. Due to spacing limitations precluding their inclusion, two additional supplementary figures for this chapter can be found at https://immunospot.com/supplfigs-affinity 16. Absorption of biomolecules to the hydrophobic PVDF membrane can occur with variable efficiency and the coating concentration required for reliable detection of antigen-specific ASC may be prohibitive; and even when raising the coating antigen concentration “indefinitely,” no, or no satisfactory antigen coating may be achieved. Affinity coating readily circumvents this problem: it is illustrated in Fig. 4 of the chapter by Lehmann et al. in this volume [29]. 17. A similar linear regression approach can also be used for defining the Goldilocks number of in vivo-differentiated ASC such as circulating plasmablasts or bone marrow/tissue-resident plasma cells. 18. To establish the frequency, and Ig class or IgG subclass usage, of Bmem-derived ASC for one antigen using a direct ImmunoSpot® assay approach, a typical serial dilution experiment requires (with a safety margin) 2 × 106 PBMC that can be obtained from 2 mL of blood, as follows: one starts the PBMC dilution series with 2 × 105 PBMC per well to permit the detection of relatively low frequency Bmem-derived ASC as well, and progresses with a 1 + 1 (two-fold) serial dilution of

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the cells. Importantly, frequency assessments done with single wells per cell dilution provide frequency assessments with essentially the same accuracy as serial dilutions done with 4 replicate wells (see Yao et al. in this volume [21]). Inclusion of negative control wells seeded with 2 × 105 and 5 × 104 PBMC is also recommended but not required; cell material permitting. Thus, ~6.5 × 105 PBMC will be needed on day 5 from in vitro polyclonal stimulation cultures, for which 1.3 million (for safety margin 2 × 106) freshly isolated PBMC need to be cryopreserved accommodating the 50% cell recovery after in vitro stimulation. The cell numbers recovered on day 5 from such in vitro cultures even permit to establish the frequency of all IgA-, IgG- and IgM-producing B cells in that sample; for such measurements, the serial dilution typically starts with an input of 2 × 104 cells per well. Note the efficiency of PBMC utilization for such B cell ImmunoSpot® assays. 19. Owing to polyclonal stimulation of Bmem to trigger their terminal differentiation, a large majority of IgG+ ASC will not be antigen-specific yet will compete for “real estate” on the lawn of anti-IgG capture reagent used for coating. Consequently, inverted assays aimed at studying lower frequency ASC specificities are directly limited by the maximal number of total IgG+ ASC that can be input into a single well while still maintaining the ability to resolve individual antigen-specific secretory footprints. 20. The plasmablasts that can be detected circulating in the blood 5–9 days after onset of a B cell response are trafficking from the draining lymph nodes to the bone marrow where they may become resident as plasma cells. 21. Thawing of cryopreserved cells causes a fraction of the cells (up to 20%) to die, and the DNA released from such cells can cause clumping of the thawed cell material. This cell clumping can be reduced, if not completely eliminated, by including an immunologically neutral endonuclease, Benzonase. Ready-touse Benzonase-containing, serum-free wash solutions are available: CTL Anti-Aggregate Wash™ 20X Solution. 22. A suitable assay medium for use in B cell ImmunoSpot® is RPMI 1640 with 10% FCS, 2 mM L-glutamine, 100 U/mL penicillin, 100 μg/mL streptomycin, 8 mM HEPES, and 50 μM 2-mercaptoethanol. 23. Kit is suited for detecting either antigen-specific ASC that differentiated in vivo, or antigen-specific Bmem that have been polyclonally stimulated in vitro to promote their transition to ASC. Each kit contains an anti-human IgG-specific capture antibody, anti-His detection reagents, diluent buffers, low autofluorescence PVDF-membrane plates, and polyclonal B cell activator (B-Poly-S).

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24. The optimal concentration of affinity (His)-tagged antigen probe used for detection of all antigen-specific secretory footprints (e.g., SFU), low- or high-affinity alike, should be determined empirically. Likewise, prior optimization of limiting antigen probe concentrations used to resolve ASC with differential functional affinity is highly recommended. 25. Kit is suited for detecting either antigen-specific antibodysecreting cells (ASC) that differentiated in vivo, or antigenspecific Bmem that have been polyclonally stimulated in vitro to promote their transition to ASC. Each kit contains anti-His capture antibodies, detection reagents, diluent buffers, low autofluorescence PVDF-membrane plates, and polyclonal B cell activator (B-Poly-S). 26. For most His-tagged proteins, high-density coating of the assay membrane can be achieved via affinity capture of a 10 μg/mL solution. Lower protein coating concentrations reduce the density of membrane-associated antigens and concomitantly increase the minimal functional affinity of an antigen-specific ASC to generate a detectable secretory footprint. 27. The volume of in vitro stimulation cultures can be scaled up or down but we recommend keeping the cell density of PBMC at ~1–2 × 106 cells/mL. If larger numbers of in vitro stimulated PBMC are required for downstream ImmunoSpot® assays, two or more 25 cm2 flasks can be set up per donor or larger 75 cm2 culture flasks can be used with a final volume of 20–25 mL. Likewise, smaller in vitro stimulation cultures can be initiated in 24-well plates with a final volume of 2 mL. Be sure to fill empty wells in tissue culture plates with sterile PBS to avoid dehydration of cell cultures. 28. Prior to performing an inverted ImmunoSpot® assay using limiting quantities of antigen detection probe, it is recommended to first determine the Goldilocks cell input to achieve ~50 SFU/well using an aliquot of cryopreserved cell material (see also Note 11). 29. In such instances when the population of antigen-specific ASC is low among all ASC, we recommend increasing the number of replicate wells and seeding at lower cell inputs. Moreover, to conserve on cell material required, increasing the fold dilution of the antigen probe and/or testing only at pre-determined concentrations are both valid options. 30. Activation of the PVDF membrane with 70% EtOH is instantaneous and can be seen visually as a graying of the membrane. It is important to be sure that the EtOH solution has spread across the entire membrane before adding the first wash of PBS. If needed, tapping the plate can promote contact of the

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EtOH solution with the PVDF membrane. We recommend only pre-wetting one plate at a time with 70% EtOH to ensure that the contact time is ≤1 min; longer contact times may promote leaking of the membrane and result in suboptimal assay performance. 31. If the entire plate will not be coated with the anti-human IgG capture antibody solution, the remainder of the EtOH pre-wet wells should receive 80 μL/well of PBS. 32. Contaminating antibody in the cell suspension(s) is efficiently captured by the anti-Ig capture antibody and may also result in elevated membrane staining that interferes with accurate enumeration of individual antigen-specific ASC in inverted ImmunoSpot® assays. 33. Shorter B cell ImmunoSpot® assay incubation times are suggested if using an enzymatic-based detection approach to avoid merging of spots and/or elevated membrane background staining. 34. Plate washes may also be performed manually. For automated washing, the pin height and flow rate should be customized to avoid damaging the assay membranes, which is the case for the CTL 405LSR plate washer. 35. Antigen probe concentrations can be generated through serial dilution (two-fold, three-fold, or five-fold) or by directly adding fixed concentrations of antigen probe into the diluent provided with the ImmunoSpot® assay kit. 36. Optimal removal of fibers and other debris, along with the reduction of “hot spots” in the center of the assay wells, is achieved through performing the “back to front” water filtration technique. 37. To completely dry plates, blot assay plate(s) on paper towels to remove residual water before either placing them in a running laminar flow hood at a 45° angle for >20 min or placing them face down on paper towels for >2 h. in a dark drawer/cabinet. Do not dry assay plates at temperatures exceeding 37 °C as this may cause the membrane to warp or crack. Fluorescent spots may not be readily visible while the membrane is still wet and the background fluorescence may be elevated. Scan and count plates only after membranes have dried completely. 38. The chapter by Karulin et al. in this volume [28] introduces machine learning-based spot-forming unit (SFU) analysis that can partially compensate for ELISA effects and SFU crowding, thus extending the linear range of accurate quantification for cell numbers plated per well and SFU detected.

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Acknowledgments We wish to thank the R&D and the Software Development teams at CTL for their continued support and technological innovation that made our B cell ImmunoSpot® endeavor possible, in particular acknowledging Dr. Alexey Karulin’s long-term contributions to this field. We also thank Dr. Graham Pawelec and Diana Roen for carefully proofreading the manuscript and providing constructive feedback. Lastly, we thank Gregory Kovacs for his support in the generation of graphic illustrations. All efforts were funded from CTL’s research budget. Conflicts of Interest P.V.L. is Founder, President, and CEO of CTL, a company that specializes in immune monitoring by ImmunoSpot®. N.B., Z.L., J.C., and G.A.K. are employees of CTL. The data X.G. contributed to this chapter (Fig. 2) were generated during his employment at CTL.

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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 14 Using PBMCs in a Multiplex FluoroSpot Assay for Detection of Innate Immune Response-Modulating Impurities (IIRMIs) Bartek Makower and Niklas Ahlborg Abstract The ELISA-based monocyte activation test (MAT) facilitates the replacement of the rabbit pyrogen test (RPT) for the detection of Innate Immune Response-Modulating Impurities (IIRMIs) in injectable drugs by activation of monocytes in human peripheral blood mononuclear cells (PBMCs). We describe the use of a triple-color IL-1β/IL-6/TNF-α FluoroSpot assay as a sensitive tool for quantification of the frequencies of IIRMI-activated monocytes as well as determination of the relative amount of pyrogenic cytokine (s) produced by each activated cell. Key words Cytokine, ELISpot, FluoroSpot, Pyrogens, Endotoxin, Monocyte activation test

1

Introduction Cytokine FluoroSpot [1] is used to simultaneously analyze the secretion of multiple cytokines by single cells. It is based on the principle of ELISpot, a powerful tool often used for the analysis of disease- and vaccine-induced T-cell responses but, limited to analysis of one cytokine at a time. In a FluoroSpot assay, in contrast to ELISpot, different fluorophores are used for the detection of separate cytokines. As a result, differently colored spots are obtained, each representing the cytokine footprint of single cells producing either single or multiple cytokines. The comprehensive assessment of multiple cytokines, along with the concurrent examination of co-expressing cells and the relative quantification of cytokine secretion [2], yields a more intricate and informative dataset than the conventional analysis focused on single cytokines. This multifaceted approach proves particularly valuable in investigating the immune responses of T cells to infectious diseases and vaccines [3]. However, the cytokine FluoroSpot is not limited to the analysis of T cells but can be used to study, in principle, any cell-secreting cytokines. It can also be applied for in vitro studies where monocytes or other

Alexander E. Kalyuzhny (ed.), Handbook of ELISPOT: Methods and Protocols, Methods in Molecular Biology, vol. 2768, https://doi.org/10.1007/978-1-0716-3690-9_14, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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cells are used to measure the impact of drugs or drug contaminants such as pyrogens. Pyrogens are substances that may induce systemic inflammation resulting in fever and hypotension upon administration of the drug. Pyrogenic substances can come from various sources including microbiological organisms and manufacturing processes. These substances can be found in equipment, packaging materials, and raw materials used in the production of medical devices and drugs. Their presence in sterile preparations suggests the presence of bacterial contamination. The MAT assay is an alternative to the RPT and the bacterial endotoxin test (BET) to detect and quantify pyrogenic contaminants in pharmaceutical products intended for parenteral use [4]. The MAT assay detects both endotoxin and non-endotoxin pyrogens and is based on the production of cytokines by monocytes in response to pyrogen triggering. The produced cytokines are quantified using ELISA. The source of monocytes for the test can be PBMCs, whole blood, or cells of a monocytic cell line. The use of pooled, frozen PBMCs from a large cell preparation batch of four healthy blood donors, secures high sensitivity, good reproducibility, and low variability [5]. The MAT has advantages over the RPT and BET, such as its high sensitivity for all types of pyrogens and its quantitative nature. Monocytes are a heterogeneous group of cells with varying morphology and functions. In humans, three distinct subsets have been identified based on their expression of the cell surface markers CD14 and CD16. However, the relationship between these subsets and the production of cytokines has largely been studied by measurement of cytokine levels using ELISA, thus making it difficult to determine the functional profiles at the single-cell level. By using FluoroSpot, human monocytes from healthy volunteers could be divided into several subgroups based on their secretion of cytokines such as IL-1β, IL-6, TNF-α, GM-CSF, and MIP-1β [6]. This approach revealed an intricate relationship between the different subsets of monocytes and their secretion of cytokines in response to TLR-2 or TLR-4 stimulation. The innate immune system uses pattern recognition receptors (PRRs) to detect harmful substances and stress signals. Toll-like receptors (TLRs) are the main family of PRRs that are involved in this process. When activated, TLRs initiate signaling cascades that lead to the production of pro-inflammatory cytokines, reactive oxygen species, and chemokines. TLRs are expressed in various cells, including monocytes, macrophages, and dendritic cells. There are at least ten different TLRs in mammals, each of which recognizes different ligands. TLR2, for example, responds to peptidoglycans from Gram-positive bacteria, while TLR4 is activated by endotoxin in Gram-negative bacteria. The process of manufacturing biopharmaceuticals often generates a mixture of

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the desired therapeutic protein and host cell proteins (HCPs) with immunostimulatory properties (IIRMIs). Purification steps are intended to minimize the presence of these impurities, but some may still be present in the final drug product. These trace contaminants can affect drug efficacy by causing adverse events, such as the recruitment of immune cells to the site of inoculation, activation of antigen-presenting cells, and inhibition of regulatory T cells. This can increase the risk of unwanted immune responses, such as the development of anti-drug antibodies (ADA), which can compromise the clinical effectiveness of the drug. Many TLR ligands act as adjuvants, enhancing antigen uptake and presentation, T-cell activation, and antibody production. It is essential to detect, quantify, and remove these undesired substances to acceptable levels in the final drug product. Monocytes in PBMCs express high levels of TLRs and other PRRs and can therefore be used as a sensitive tool for detecting clinically relevant IIRMIs. A recent study [7] identified cytokine secretion from healthy human donor PBMC as a sensitive method for the in vitro monitoring of innate immune responses to ten common IIRMIs using a 16-plex cytokine assay as a readout. The authors found that IIRMIs activated a broad and often overlapping spectrum of cytokines. Specifically, IL-6 and TNF-α are produced by monocytes and T cells and are responsible for pyrogenicity; IL-1β is produced by monocytes and DCs and is responsible for the inflammation, fever, and activation of specific subsets of lymphocytes (i.e., IL-1β promotes TH17 differentiation). The study identified three signature cytokines induced by individual IIRMIs. For example, IL-6 was a signature cytokine for Poly (I:C) TLR3, CLO75 (TLR8), and ODN (TLR9) while IL-1β was a signature cytokine for Flagellin (TLR5) and LPS (TLR4). TNF-α was induced by all IIRMI tested except ODN (TLR9) and Poly (I:C). Using the FluoroSpot assay captures the full spectrum of the response and identification of polyfunctional monocytes contributes information about the potential in vivo function of these cell populations [8].

2

Materials

2.1 Instrumentation and Software 2.2

FluoroSpot Kits

1. IRIS FluoroSpot reader (Mabtech) 2. Apex™ software using the RAWspot™ algorithm (Mabtech) FluoroSpot Plus kit: Human IL-1β/IL-6/TNF-α kit #FSP121309-2 (Mabtech, Nacka Strand, Sweden). The kit components are listed below: Pre-coated transparent, flat-bottomed, 96-well plates with polyvinylidene difluoride (PVDF) membranes designed for low autofluorescence (IPFL; Millipore, Cork, Ireland). The plates are

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Fig. 1 Trippel fluorospot for the detection of Flagellin-stimulated PBMCs secreting IL-1β and/or IL-6 and TNFα. Human PBMCs (25,000 cells/well) were stimulated with 2.5 ng/mL Flagellin for 16 h. (a) Raw images from the IRIS reader showing IL-1β (green), IL-6 (yellow), and TNF-α (red) spots after stimulation. (b) Representation of monocyte populations producing one, two, or three cytokines simultaneously identified by the RAWspot™ algorithm in the Apex software. (c) Average RSV showed for cells producing one, two, or three cytokines simultaneously

coated with monoclonal antibodies (mAbs) MT175 (antihuman IL-1β), 13A5 (anti-human IL-6), and MT25C5 (antihuman TNF-α). Detection mAbs: anti-human IL-1β (7P10), BAM, anti-human IL-6 (39C3), biotin and anti-human TNF-α (MT20D9), WASP. Fluorophore conjugates: anti-BAM mAb, 490, SA-550, and antiWASP mAb, 640. Buffer/solution: FluoroSpot enhancer used to improve fluorescent signal (Fig. 1). 2.3

Buffers

Blocking reagent after coating: 10% fetal calf serum (FCS) in phosphate-buffered saline (PBS), pH 7.4, or in RPMI 1640.

2.4

Media

Cell culture medium: RPMI 1640 (with 2 mM L-glutamine) + 10% FCS (see Note 1), 10 mM HEPES, and 100 μg/mL penicillin +100 μg/mL streptomycin.

2.5

Cells

Cryopreserved, pooled human PBMCs from four healthy blood donors.

2.6

Stimuli (Fig. 2)

Endotoxin (LPS) TLR4 agonist (Sigma/ Merck), Endotoxin standard BRP, #E0150000 EDQM, European Pharmacopoeia (EP). Heat killed Staphylococcus aureus (HKSA), a Gram-positive extra-cellular growing bacterium. TLR2 agonist (Invivogen, Cat: tlrl-hksa)

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Fig. 2 Titration of LPS (TLR4), Flagellin (TLR5), and HKSA (TLR2) in trippel FluoroSpot. Spot forming units (SFU) for the total IL-1β, IL-6, and TNF-α responses are shown

Flagellin (Salmonella typhimurium) from Gram-negative bacteria. TLR5 agonist (Invivogen, Cat: tlrl-stfla). Pam3CSK4 (Pam3CysSerLys4), a synthetic triacylated lipopeptide TLR2/TLR1 agonist (Invivogen. Cat: tlrlpms). R848 (Resiquimod), TLR7/TLR8 Agonist (Mabtech, Sweden).

3

Methods The protocol describes trippel staining for the detection of human IL-1β, IL-6, and TNF-α. PVDF plates are pre-coated with mAbs to IL-1β, IL-6, and TNF-α. Cells ± stimuli are added, and secreted IL-1β, IL-6, and TNF-α are captured by the specific mAbs. After cell removal, spots are detected in two steps. First, a mixture of mAb anti-IL-1β-BAM, anti-IL-6-biotin, and anti-TNF-α-WASP is added (see Note 4). Second, spots are detected by adding a mixture of mAb anti-BAM-green fluorophore (for IL-1β) SA-550 yellow fluorescence (for IL-6), and mAb anti-WASP-red fluorophore (for TNF-α).

3.1 Trippel-Color FluoroSpot

Thawing of Cells (Sterile Conditions) Before you begin: heat a water bath to 37 °C. Pre-heat the cell culture medium to be used for step 2. The medium used in later washing steps can be at room temperature. 1. Transfer the cryotube rapidly from the frozen storage to a 37 °C water bath and thaw the cells until only a small ice crystal remains (see Note 6). 2. Slowly add 0.5–1 mL of the pre-heated cell culture medium to the cryotube, resuspend, and transfer the cells from the cryotube to a 15 mL, sterile polypropylene centrifuge tube. Rinse the cryotube with 1 mL of cell culture medium and transfer it to the 15 mL tube. Fill the tube with a cell culture medium. 3. Wash by centrifugation for 10 min at 300× g.

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4. Discard the supernatant and tap the tube to make the pellet less compact. Resuspend the cells in 1 mL of cell culture medium by slowly pipetting up and down. Add cell culture medium to a total volume of 15 mL. 5. Centrifuge for 10 min at 300× g. 6. Discard the supernatant and resuspend the pellet in 1 mL of cell culture medium. Add additional cell culture medium (e.g., 5 mL depending on expected cell number and desired cell concentration). 7. Place the cells at 37 °C in a cell incubator with a humidified, 5% carbon dioxide atmosphere for 1 h. Leave the cap slightly open. Utilize this 1 h of downtime to wash and block your FluoroSpot plate. Preparation of the Pre-coated Plates (Sterile Conditions) 1. Remove the plate from the sealed package and wash the plate three times with sterile PBS pH 7.4 (200 μL/well). 2. Block/condition the plate by adding a cell incubation medium containing 10% FCS (200 μL/well). Incubate for at least 30 min at room temperature. Adding Stimuli to the Plates (Sterile Conditions) 1. Remove the blocking solution and add 100 μL stimuli diluted in cell culture medium/well (see Note 2). Counting and Dilution of Cells (Sterile Conditions) 1. After incubation (cell resting), resuspend the cells and let any aggregated cell debris sediment (takes around 1 min). Then carefully transfer the cell suspension, without the debris, to a fresh 15 mL tube. Add a maximum of two cryotubes to each 15 mL tube (or around 30–40 million cells). 2. Count the cells and determine cell viability. Counting can be done with an automated cell counter or trypan blue staining using a microscope. Because only living cells will be able to secrete the analyte, make sure to exclude dead and, if possible, apoptotic cells from the cell count. Dilute the cells to 250,000 viable PBMCs/mL. If the cell concentration turns out to be lower than required, centrifuge the cells again, resuspend, and dilute to the desired volume. Incubation of Cells in the Plates (Sterile Conditions) 1. Add 100 μL (25,000) viable PBMCs/well to a washed and blocked FluoroSpot plate containing 100 μL stimuli/well (see preparation of the pre-coated plates and adding stimuli to the plates) or only cell culture medium as a negative control.

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2. Incubate the plate overnight at +37 °C in a humidified atmosphere with 5% carbon dioxide (see Note 7). Avoid moving the plate during the incubation time (see Note 8). Detection of Spots 1. Discard the cells and wash the plate five times with PBS in a plate washer or by hand with 200 μL/well (see Note 3). 2. In the same tube, dilute the detection mAbs for IL-1β, IL-6, and TNF-α labeled with BAM biotin and WASP, respectively, in PBS containing 0.1% bovine serum albumin (BSA) following the dilution recommendations in the datasheet, add 100 μL/ well, and leave the plate for 2 h at room temperature. 3. Wash the plate as in step 1 above. 4. Dilute secondary detection reagents, i.e., mAb anti-BAM labeled with green fluorophore streptavidin (SA) labeled with yellow and anti-WASP labeled with red in the same tube to their respective working concentrations in PBS–0.1% BSA, following the dilution recommendations in the datasheet. Add 100 μL/well and leave the plate for 1 h at room temperature. 5. Wash the plate as in step 1 above, empty it by flicking the plate, and thereafter add FluoroSpot enhancer solution for 5–15 min at room temperature (50 μL/well). Empty by flicking the plate to remove the FluoroSpot enhancer. Do not wash. It is important that the plate is not tapped against paper towels since dust particles may interfere with the assay. Remove the soft plastic underdrain and let the plate dry in the dark before analysis. Further storage of the dry plate should be at room temperature in the dark. The plate should be completely dry before analysis. Analysis of Spots 1. To facilitate the analysis process, we recommend utilizing an automated FluoroSpot reader such as the Mabtech IRIS, equipped with FITC, Cy3, and Cy5 filters. In the resulting images, green spots indicate cells producing IL-1β, yellow spots indicate cells producing IL-6, and red spots indicate cells producing TNF-α. By overlaying the IL-1β, IL-6, and TNF-α spot images from the same well in a computerized system, we can identify double-stained or triple-stained spots based on the relative positions of the green, yellow, and red spots. The RSV (see Note 5) can be determined for each spot using the Apex software, the RSV data can be exported from each well for further analysis (Fig. 1).

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2. Fluorescent spots, like any fluorescent signal, can diminish over time if exposed to excessive light. While the fluorescence on dry plates has been observed to be remarkably stable, allowing plates to be stored for months, it is advisable to analyze the plate within 1 week of development for optimal results. This is because the membrane, particularly in green staining, may undergo slight brightening over time. 3. To prevent any potential crossover between filters, it is recommended to include wells in the assay where only one detection system has been employed. When analyzing these wells using filters specific to other fluorophores, the resulting image should ideally show no observable spots. This practice helps ensure accurate and reliable detection without any interference or contamination between the different detection systems.

4

Notes 1. Serum in cell medium: A serum pre-evaluated for cell culture and that yields low spontaneous cytokine secretion should be used in the medium; FCS is recommended. 2. Preparation of stimuli: LPS 5 mL of sterile H2O is added to the endotoxin standard BRP vial (1.2 μg endotoxin) equal to 2000 EU/mL. The recommended working concentration of LPS is between 0.01 and 1 EU/mL. HKSA. 1 mL of sterile endotoxin-free water is added to 1010 lyophilized cells of HKSA. The recommended working concentration of HKSA is between 0.06 × 106 and 8 × 106 cells/mL. Flagellin Dilute Flagellin to 0.5 mg/mL in sterile H2O. The recommended working concentration of Flagellin is between 0.8 ng/ mL and 100 ng/mL. Pam3CSK4. Dilute Pam3CSK4 to 2.0 mg/mL in sterile H2O. The recommended working concentration of Pam3CSK4 is between 0.3 ng/mL and 20 ng/mL. R848 (Resiquimod). Dilute R848 to 1.0 mg/mL in sterile H2O. The recommended working concentration of R848 is between 15 ng/mL and 1000 ng/mL. 3. Plate washing: washing of plates can be done using a multichannel device. In washing steps not requiring sterile

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conditions, a regular ELISA plate washer with a washing head adjusted to ELISPOT/fluorospot plates can be used. 4. FluoroSpot is based on a fluorescent detection system. Most analytes (except for immunoglobulins) are detected in a two-step fashion, where the first detection step is performed using a combination of biotinylated and tag-labeled detection antibodies. The tags BAM and WASP and the high-affinity anti-tag antibodies are proprietary to Mabtech and have been developed to work optimally in FluoroSpot. BAM and WASP are short peptides with the sequences: DAEFRHDSGY (BAM) and PDYRPYDWASPDYRD (WASP). 5. RSV can only be compared between spots of the same color, i.e., IL-1β RSV cannot be compared with IL-6 RSV since different detection systems and fluorophores may differ in sensitivity. 6. If you intend to thaw multiple cryotubes, only thaw a few at a time. The freezing medium contains DMSO which is toxic to cells so proceed to step 5 as soon as possible. 7. Wrap the plate in aluminum foil or use sterile breathable film. Most incubators maintain an optimal level of humidity, but it is good practice to reduce the risk of evaporation of the cell culture medium. 8. Place the plates next to – not on top of – each other. Do not stack plates in the incubator. Stacking plates can lead to uneven conditions between the plates (e.g., temperature, CO2 level, evaporation rate), which may affect the cells and thereby the spot numbers. References 1. Gazagne A, Claret E, Wijdenes J et al (2003) A Fluorospot assay to detect single T lymphocytes simultaneously producing multiple cytokines. J Immunol Methods 283:91–98 2. Del Aguila Pla P, Jalden J (2018) Cell detection by functional inverse diffusion and non-negative group sparsity—part I: modeling and inverse problems. IEEE Trans Signal Process 66:5407– 5421 3. Sandberg JT, Varnaite R, Christ W et al (2021) SARS-CoV-2-specific humoral and cellular immunity persists through 9 months irrespective of COVID-19 severity at hospitalisation. Clin Transl Immunol 10:1–18 4. Poole S, Thorpe R, Meager A et al (1988) Detection of pyrogen by cytokine release. Lancet 8577:130

5. Solati S, Aarden L, Zeerleder S et al (2015) An improved monocyte activation test using cryopreserved pooled human mononuclear cells. Innate Immun 21:677–684 6. Smedman C, Ernemar T, Gudmundsdotter L et al (2011) FluoroSpot analysis of TLR-activated monocytes reveals several distinct cytokine-secreting subpopulations. Scand J Immunol 75:249–258 7. Holley CK, Cedrone E, Donohue D et al (2021) An in vitro assessment of Immunostimulatory responses to ten model innate immune response modulating impurities (IIRMIs) and peptide drug product, Teriparatide. Molecules 26:1–20 8. Boyette LB, Macedo C, Hadi K et al (2017) Phenotype, function, and differentiation potential of human monocyte subsets. PLoS One 12: 1–20

Chapter 15 Four-Color ImmunoSpot® Assays Requiring Only 1–3 mL of Blood Permit Precise Frequency Measurements of Antigen-Specific B Cells-Secreting Immunoglobulins of All Four Classes and Subclasses Lingling Yao, Noe´mi Becza, Andrea Maul-Pavicic, Jack Chepke, Greg A. Kirchenbaum, and Paul V. Lehmann Abstract The B lymphocyte response can encompass four immunoglobulin (Ig) classes and four IgG subclasses, each contributing fundamentally different effector functions. Production of the appropriate Ig class/subclass is critical for both successful host defense and avoidance of immunopathology. The assessment of an antigenspecific B cell response, including its magnitude and Ig class/subclass composition, is most often confined to the antibodies present in serum and other biological fluids and neglects monitoring of the memory B cell (Bmem) compartment capable of mounting a faster and more efficient antibody response following antigen reencounter. Here, we describe how the frequency and Ig class and IgG subclass use of an antigen-specific Bmem repertoire can be determined with relatively little labor and cost, requiring only 8 × 105 freshly isolated peripheral blood mononuclear cells (PBMC), or if additional cryopreservation and polyclonal stimulation is necessary, 3 × 106 PBMC per antigen. To experimentally validate such cell saving assays, we have documented that frequency measurements of antibody-secreting cells (ASC) yield results indistinguishable from those of enzymatic (ELISPOT) or fluorescent (FluoroSpot) versions of the ImmunoSpot® assay, including when the latter are detected in alternative fluorescent channels. Moreover, we have shown that frequency calculations that are based on linear regression analysis of serial PBMC dilutions using a single well per dilution step are as accurate as those performed using replicate wells. Collectively, our data highlight the capacity of multiplexed B cell FluoroSpot assays in conjunction with serial dilutions to significantly reduce the PBMC requirement for detailed assessment of antigen-specific B cells. The protocols presented here allow GLP-compliant high-throughput measurements which should help to introduce high-dimensional Bmem characterization into the standard immune monitoring repertoire. Key words ELISPOT, FluoroSpot, B cells, Immune monitoring, Immune memory, Ig class, IgG subclass, High throughput, Antibody-secreting cell

Alexander E. Kalyuzhny (ed.), Handbook of ELISPOT: Methods and Protocols, Methods in Molecular Biology, vol. 2768, https://doi.org/10.1007/978-1-0716-3690-9_15, © The Author(s) 2024

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Introduction One might ask why bother with detecting antigen-specific memory B cells (Bmem), fragile live cells that need to be processed within a short time window after they have been collected from the body when simple serum antibody measurements provide the soughtafter information? The answer is simple: Bmem measurements can provide insights into immune responsiveness that serum antibodies cannot. Because another chapter in this volume [1] is dedicated to this issue, here we will just touch on the major points. Antigenspecific plasma cells (PC) and Bmem both arise during an immune response triggered by antigen encounter, but the generation of these two daughter cell lineages follows different fate-decision pathways. Precursors of both cell types, germinal center B cells (GCB), undergo somatic hypermutations (SHM) that result in the generation of subclones with slightly modified B cell antigen receptors (BCR). From this repertoire of daughter cells, subclones that have an increased affinity for the antigen are positively selected to undergo further rounds of proliferation and SHM, and eventually differentiate into PC. Contrary to the previously held notion, PC are not necessarily long-lived (see Note 1) and neither are the Ig molecules they secrete (see Note 2). In contrast, GCB progeny endowed with lower affinity BCR for the antigen can still join the long-lived Bmem compartment. Hence, PC and Bmem fulfill different roles in maintaining host immune defense. The antibodies produced by PC constitute the first wall of acquired humoral immune defense [2]. They provide immediate protection by preventing the re-entry of the antigen, or, if it enters, by neutralizing it and/or facilitating its elimination by phagocytes via immune complex formation (precipitation), opsonization, and complement fixation. As evidenced during the recent COVID-19 pandemic, and previously with seasonal influenza, the first wall of adaptive humoral defense may fail to prevent (re)-infection when antibody titers drop below protective levels, or upon emergence of viral escape mutants capable of evading the neutralizing activity of antibodies elicited by the original (homotype) virus strain. In such cases, Bmem provide the second wall of adaptive humoral host defense [2]. Owing to their increased frequencies compared to antigen-specific naive B cells, and having already switched from IgM to the expression of specialized Ig classes and subclasses (see Note 3), Bmem not only mediate a faster and more efficient “secondary” antibody response against the same (homotypic) virus but also against antigenically-related viral escape mutants (heterotypes). This is because, within the antigen-specific B cell repertoire that was clonally expanded by the homotypic virus during the primary response, there will be Bmem that joined the memory compartment with mutated BCR that have affinity for the heterotype as well.

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Because such cross-reactive Bmem occur in increased frequencies compared to naive B cells and have already undergone Ig class switching, this population is poised to engage in a quasi-secondary B cell response if infection with a variant virus occurs. From the above, it follows that measurements of serum antibodies provide insights only about the fading first wall of immune protection. Measurements of Bmem, in contrast, provide insights into the cellular basis of long-term immunity. Through measuring the frequency of antigen-specific Bmem within all peripheral blood mononuclear cells (PBMC) (see Note 4), the magnitude of the existing memory compartment within an individual can be directly quantified. Such information sheds light on the vigor of future secondary antibody responses upon antigen encounter. Moreover, by establishing the Ig class/subclass utilization of the Bmem compartment, one can also predict the type of antibody that will be produced upon antigen encounter (see Note 5). There are few techniques capable of detecting rare antigenspecific Bmem while also providing information regarding their relative abundance, Ig class/subclass usage, and functional affinity (see Note 6). B cell ImmunoSpot® assays are ideally suited for this purpose as they enable the detection of Ig molecules secreted by individual antibody-secreting cells (ASC). While resting Bmem do not secrete antibodies, such cells can readily be differentiated into ASC following a simple in vitro stimulation protocol (see Note 7). The ImmunoSpot® assay principle for detecting ASC, irrespective of their antigen specificity, is described in Fig. 1a, and its variant for detecting antigen-specific ASC in Fig. 1b. In this chapter, we share our expertise on how to best establish the frequency of antigen-specific, Bmem-derived ASC in human PBMC, including their Ig class and subclass usage, and how to do so with the least labor, and the lowest number of PBMC possible (see Note 4). The type of testing described here is also essential for determining the so-called “Goldilocks” number of PBMC to be seeded into subsequent ImmunoSpot® assays aimed at evaluating the affinity distribution present among antigen-specific ASC (see the chapter by Bezca et al. in this volume, [3]), or the crossreactivity of homotype antigen-primed ASC with heterotypic antigens (see the chapter by Lehmann et al., also in this volume [1]). Owing to the requirement to detect individual ASC-derived secretory footprints to accurately determine the frequency of antigen-specific B cells, an ImmunoSpot® assay-related challenge to overcome is that Bmem-derived ASC producing different classes and subclasses of Ig span orders of magnitude [4]. Importantly, this problem is readily overcome by seeding PBMC (or other single-cell suspensions) in serial dilutions. For establishing the frequencies of antigen-specific ASC following in vitro polyclonal stimulation of PBMC, we recommend starting at 2 × 105 PBMC per well and progressing in a 1 + 1 (two-fold dilution series) down

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Fig. 1 Schematic representation of (a) the pan (total), or (b) the antigen-specific, direct ImmunoSpot® assays (the latter, using the affinity capture coating variant). For (a), the PVDF membrane on the bottom of a 96-well plate is densely coated with a pan anti-Ig capture antibody that will bind the ASC-secreted Ig(G) with highaffinity irrespective of the ASC’s antigen specificity. In (b), the membrane is coated first with an anti-affinity tag-specific antibody (in this example anti-His) that captures the (His)-tagged antigen with high affinity. In this way, a dense coating of the membrane with the antigen is accomplished. As the next step in both assay variants, the PBMC containing the ASC are plated. In (a), an ASC-secreted Ig(G) antibody is captured around each ASC that is secreting Ig(G), and results in the formation of individual secretory footprints. In (b), the antibody produced by antigen-specific ASC is captured only on the lawn of the antigen. The subsequent steps are similar for both assay variants. After removal of the cells, the membrane-associated antibody is visualized using biotinylated anti-human Ig class/subclass-specific detection antibody reagents that subsequently are revealed by the addition of a fluorescently conjugated streptavidin (FluoroSpot, as shown) or via an enzymatic reaction (ELISPOT, not shown). Counting the spot-forming units (SFU) per well reveals the number of (a) total Ig (G) or (b) antigen-specific, Ig(G)-producing ASC within the PBMC plated. The spot morphologies in (b) also provide insights into the functional affinities of the antibody secreted by the individual ASC for the antigen, a topic covered in detail in the chapter by Becza et al., in this issue [3]

the 96-well plate to generate 8 additional data points. Similarly, for establishing the frequency of all ASC-producing IgM, IgG, or IgA, irrespective of their antigen-specificity, we recommend starting at 2 × 104 and performing a similar two-fold dilution series down the 96-well plate for 8 points of cell titration (see Note 10). Four-color ImmunoSpot® assays are suited to generate maximal data while saving on cells (see Note 11). Figures 2 and 3 show that such fluorescence-based tests detect the secretory footprints of individual Bmem with the same efficacy as single-color enzymatic assays. As can be seen for higher cell numbers in Fig. 3, the confluence of secretory footprints and the resulting ELISA effect interfere with the accurate recognition and counting of individual spotforming units (SFU). At SFU counts lower than 100 SFU per well, however, a close to perfect linear relationship exists between the number of PBMC seeded per well and the number of SFU per well, from which, by linear regression, the frequency of SFU within all PBMC plated can be accurately extrapolated. A fully automated software module built into the ImmunoSpot® software permits the identification of the linear range of SFU counts, the calculation of means of replicates, and the frequency extrapolation (see the chapter by Karulin et al. on this issue in this volume [5]).

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Fig. 2 Single-color ELISPOT and FluoroSpot assays have similar sensitivities for detecting ASC-derived secretory footprints. Peripheral blood mononuclear cells (PBMC) were polyclonally stimulated with B-Poly-S (or B-Poly-SE for detection of IgE+ ASC in panel D) for 5 days in vitro (see Notes 7 and 10), washed, and seeded into single-color ELISPOT or FluoroSpot assays in parallel, detecting the immunoglobulin (Ig) class/ subclass specified in panels A–J, respectively; the type of assay shown in Fig. 1a was performed. Since antibody-secreting cells (ASC) producing the different Ig classes or subclasses occur in vastly different frequencies in PBMC following in vitro polyclonal stimulation (as detailed in the text), the assays were performed by plating PBMC in a 1 + 1 serial dilution series, starting at 2 × 105, in 4 replicates per cell dilution. Images of representative wells are shown for the dilution step in which the individual secretory footprints were clearly discernable; between 50 and 70 SFU/well. The cell inputs for panels A-J were: 781, 391, 3125, 200,000, 781, 6250, 6250, 25,000, 3125, and 6250 per well, respectively. The number of secretory footprints (spot forming units, SFU) measured using ImmunoSpot® software is reported in the left upper corner of the corresponding well images, and means ± SD of the four replicate wells are specified under each pair of wells. Statistical analysis (unpaired Student’s t-test) indicated no significant difference (“ns”) between the ELISPOT and corresponding FluoroSpot counts between the replicates for each condition. The presented results are representative of 4 independent experiments using different PBMC donors, leading to the same conclusion

Our in-depth studies of such regression analysis-based frequency calculations showed that performing serial dilution experiments involving replicates offers only a negligible advantage in precision over performing the assay with single wells (Fig. 4 and Becza et al., manuscript in preparation). Doing such serial dilutionbased frequency measurements in four-color reduces the required number of PBMC by four-fold compared to performing 4 independent single-color assays, and doing so using a single well serial dilution approach permits another four-fold reduction in cell material compared to testing each cell input in quadruplicate (as was done in the data presented in Figs. 2 and 3). Figure 5 depicts the results obtained from a typical serial dilution-based four-color ImmunoSpot® test detecting all four antibody classes (IgM, IgG, IgA, and IgE) of SARS-CoV-2 Spikespecific ASC in a convalescent individual with PCR-verified infection. Similarly, the IgG subclass usage of Spike-specific ASC was determined in parallel using an IgG1/IgG2/IgG3/IgG4 fourcolor assay. The abundance of all ASC in the test sample producing each Ig class or IgG subclass was also established to permit

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Fig. 3 Single-color ELISPOT and multi-color FluoroSpot assays have similar sensitivities for detecting ASC-derived secretory footprints. Peripheral blood mononuclear cells (PBMC) were polyclonally stimulated with B-Poly-S for 5 days in vitro, washed, and then seeded into single-color enzymatic (SCE) ELISPOT, singlecolor FluoroSpot (SCF), three-color FluoroSpot (TCF), or four- (quadruple) color FluoroSpot (QCF) assays in parallel, diluting the cell inputs two-fold per well, detecting pan IgG+ (a, b) or IgM+ ASC (c, d) in four replicate wells, irrespective of antigen specificity; the type of assay illustrated in Fig. 1a was performed. In panels (a) and (c), representative images (one of the four replicates tested) depicting secretory footprints visualized using anti-IgG or anti-IgM detection reagents are shown, respectively, with the corresponding cell input specified. ELISA effects are evident at high cell inputs. In panels (b) and (d), the spot-forming unit (SFU) counts for the four replicate wells for each condition are shown as means ± SD (y-axis) at the corresponding cell inputs (x-axis). The detection modalities are color-coded, as specified, including the matching trend lines calculated through linear regression analysis also denoting R2 values

calculation of the frequency of antigen-specific ASC among all ASC, for each Ig class and subclass (see Notes 12 and 13). As seen for this individual, and consistent with the majority of convalescent COVID-19 donors we tested thus far (Kirchenbaum, unpublished observation), the SARS-CoV-2 Spike-specific Bmem primarily secrete IgG, and IgG1 in particular. Notably, although our IgA class-specific detection system works perfectly well for the detection of total IgA+ ASC activity (see Fig. 2c), very few Spikespecific IgA+ Bmem-derived ASC were detectable in this individual. Moreover, a small population of Spike-specific IgG3+ Bmem-derived ASC was also detectable in this donor. Importantly, for the precise enumeration of the latter very rare antigen-specific ASC, higher cell

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Fig. 4 Linear regression analysis performed using single wells in serial dilution experiments yields similar accuracy for determining the frequency of ASC compared to that calculated from the mean of 4 replicate wells at each cell input. Peripheral blood mononuclear cells (PBMC) were polyclonally stimulated with B-Poly-S for 5 days in vitro, washed, and then serially diluted two-fold, with four replicate wells for each cell input, in a four-color FluoroSpot assay to determine the frequency of antibody-secreting cells (ASC) producing IgA (a), IgM (b), or IgG (c). The type of assay shown in Fig. 1a was performed, detecting ASC irrespective of their antigen specificity. Red symbols denote means ± SD of SFU counts in the four replicate wells, at each of the cell inputs, respectively; the solid red trend line was calculated by linear regression analysis of these means with the R2 value specified. The respective frequencies as calculated by extrapolating the linear regression line to a 106 PBMC input are given in parentheses. The color-coded “Cell titration 1-4” data with the corresponding dashed trend lines, R2 values, and frequencies were obtained by independent single-well analysis of the four replicates. The results are representative of 5 independent experiments using different PBMC donors leading to the same conclusion

inputs would be required, inputs that would be too high for determining the frequency of antigen-specific ASC producing IgG/IgG1 (highlighting the importance of testing samples across multiple cell inputs through serial dilutions and the value of multicolor analysis). Of note, the large number of IgM+ SFU detected in Spike antigen-coated wells does not appear to be “specific” since comparable numbers of SFU were also present in negative control wells. Such IgM+ SFU likely originate from naive B cells possessing broadly reactive BCR specificities, and which differentiated into ASC following in vitro polyclonal stimulation. This lack of “specificity” exhibited by IgM+ ASC following polyclonal stimulation of human PBMC has also been reported previously [6] and reiterates the importance of including negative controls in such B cell ImmunoSpot® assays. In contrast, while IgG+ Spike- or NCAP-antigenspecific ASC were absent in all PBMC collected in the pre-COVID era, abundant such IgG+ ASC were detected in individuals with PCR-verified SARS-CoV-2 infection [4]. Therefore, unlike IgM, detecting antigen-specific IgG (and IgG subclass)-producing ASC in PBMC following short-term in vitro polyclonal stimulation signifies in vivo primed and class-switched memory B cells. Antigenspecific, IgM+ ASC can be detected, however, when PBMC are studied acutely (5–9 days) after induction of a primary B cell response. Performing the above test required 8 × 105 PBMC to be seeded into the antigen-specific assay, plus we added an optional pan Ig class/subclass assay requiring an additional 8 × 104 PBMC (see

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Fig. 5 Four-color ImmunoSpot® assays permit assessment of SARS-CoV-2 Spike-specific ASC frequency, plus the frequency of all ASC producing different Ig classes and IgG subclasses using 8.8 × 105 PBMC (see Note 4). Cryopreserved peripheral blood mononuclear cells (PBMC) from a convalescent donor with PCR-verified SARS-CoV-2 infection were polyclonally stimulated with B-Poly-S for 5 days in vitro, washed, and then evaluated in four-color ImmunoSpot® assays. (a) Recommended plate layout for determining ASC frequencies for each Ig class and IgG subclass using a serial dilution approach: columns 1 and 2 measure pan (total) Ig class and subclass usage are illustrated in Fig. 1a; columns 3 and 4 for measuring the antigen-specific ASC. (b) Spike-specific SFU counts were measured using the four-color Ig class detection system over the entire PBMC range plated, and (d) for the IgG subclass system with Ig class/subclass specified by color. Note deviation from linearity at high cell inputs/SFU counts. (c) Trend line for Spike-specific IgG+ ASC calculated by linear regression analysis of the wells containing SFU < 100/well with the R2 value; in parenthesis the extrapolated frequency per 106 PBMC. (d) The results for IgG1+ ASC are represented in (e)

Fig. 5a and Note 4). As PBMC can be cryopreserved without loss of B cell functionality [7], samples can be run in batches instead of testing them one by one as soon as the blood is drawn (see Note 13). During freeze-thawing up to 30% of the cells may be lost, but the functionality of the recovered B cells will be unaltered compared to freshly isolated PBMC ([7] and Becza et al., manuscript in preparation). If an additional polyclonal stimulation is needed prior to the actual ImmunoSpot® test to convert resting Bmem into ASC, approximately 50% of the frozen PBMC will be recovered after thawing and performing the 5 day in vitro polyclonal stimulation protocol (N. Becza, manuscript in preparation). Therefore, only 8.8 × 105 PBMC are needed for the type of test shown in Fig. 5, and allowing for a safety margin in cell recovery, we suggest that 2–3 × 106 PBMC should be cryopreserved per aliquot to perform such a test. Furthermore, we recommend freezing several aliquots of PBMC (or other single-cell suspensions) to permit subsequent tests using higher cell inputs and/or additional replicates in scenarios where antigen-specific ASC frequencies are very low (see Note

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15). Furthermore, the availability of additional aliquots of cell material permits further in-depth characterization of affinity distributions or heterotypic cross-reactivity within the Bmem-derived ASC repertoire (see Notes 16 and 17) and references [1, 3]. In the following, we provide detailed protocols for cryopreservation of PBMC to maintain their full functionality, subsequent polyclonal stimulation of these PBMC to differentiate resting Bmem into ASC, and four-color ImmunoSpot® assays for defining the Ig class and IgG subclass usage of antigen-specific ASC.

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Materials

2.1 Isolation and Cryopreservation of PBMC from Whole Blood

1. Class II biosafety cabinet (BSC) 2. Green vacutainer tubes containing sodium heparin 3. Lymphoprep™ 4. 15 or 50 mL conical tubes 5. Centrifuge capable of spinning tubes at 800 × g (temperature set to 25 °C) 6. Sterile transfer pipette 7. Ca2+, Mg2+-free phosphate-buffered saline (PBS), pH 7.2 (room temperature) 8. Parafilm 9. CTL-LDC™ counting kit 10. CTL-Cryo™ ABC media kit 11. ImmunoSpot® S6 Ultimate 4 LED analyzer, or a suitable instrument equipped with the appropriate detection channels, running CTL’s live/dead cell counting suite software 12. DMSO 13. Bead bath (set to 37 °C) 14. 1.8 mL Cryovials (with internal thread and silicone washer seal) 15. Mr. Frosty™ freezing container, or controlled rate freezer. 16. Isopropyl alcohol 17. -80 °C freezer 18. Liquid nitrogen tank

2.2 Thawing of Cryopreserved PBMC

1. Cryopreserved PBMC sample(s) (see Note 14) 2. 70% EtOH 3. DNase-containing washing medium (pre-warmed to 37 °C) (see Note 18) 4. Complete B cell medium (BCM) (pre-warmed to 37 °C) (see Note 19)

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2.3 In Vitro Polyclonal Stimulation of B Cells in PBMC 2.4 Single-Color ELISPOT Assay

1. B-Poly-S 2. Tissue culture plate (48 or 24-well) 25 cm2 sterile culture flask 3. Humidified incubator set at 37 °C, 5% CO2 1. Commercially available, single-color human Ig class (IgA, IgE, IgG, or IgM) or subclass (IgA1, IgA2, IgG1, IgG2, IgG3, or IgG4) ELISPOT kit (see Note 20) 2. 190 proof (95%) EtOH 3. Cell culture-grade water 4. 96-well, round bottom dilution plate 5. 0.05% Tween-PBS wash solution 6. 0.1 μm low-protein binding syringe filter 7. Plate washer 8. ImmunoSpot® S6 Ultimate 4 LED analyzer, or a suitable instrument equipped with the appropriate detection channels, running CTL’s ImmunoSpot® UV

2.5 Single-, Three- or Four-Color FluoroSpot Assays

Commercially available, single-color human Ig class (IgA, IgE, IgG, or IgM) or subclass (IgA1, IgA2, IgG1, IgG2, IgG3, or IgG4) FluoroSpot kit (see Note 21). 1. Commercially available, three-color human Ig class (IgA, IgG, and IgM) FluoroSpot kit (see Note 21) 2. Commercially available, four-color human Ig class (IgA, IgE, IgG, and IgM) FluoroSpot kit (see Note 21) 3. Vacuum manifold

2.6 Four-Color Antigen-Specific FluoroSpot Assay (Affinity Capture Coating)

1. Commercially available, four-color human Ig class (IgA, IgE, IgG, and IgM) affinity capture (His) FluoroSpot kit (see Note 22) 2. Commercially available, four-color human IgG subclass affinity capture (His) FluoroSpot kit (see Note 22) 3. The His-tagged recombinant protein (see Notes 23 and 24)

3

Methods

3.1 Isolation and Cryopreservation of PBMC from Whole Blood (Sterile Conditions)

1. Obtain blood samples according to IRB-approved protocol. Keep blood at ambient temperature, do not refrigerate. 2. Keeping donors’ material separate, inside a class II BSC, pool each donor’s blood into labeled conical tubes. Rinse each vacutainer tube with PBS at ambient temperature and combine it with whole blood.

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3. Measure blood volume and dilute 1:1 with PBS. If the volume of whole blood to be processed is ≥20 mL, transfer half of the volume into another labeled 50 mL tube prior to diluting with PBS. 4. Layer diluted blood at ambient temperature slowly over Lymphoprep™, also at ambient temperature taking care not to disrupt the interface. 5. Centrifuge balanced tubes at 800 × g for 20 min with the centrifuge brake off, non-refrigerated. 6. Identify the buffy coat at the interface between Lymphoprep™ and diluted plasma layers. Carefully remove the cells at this interface and transfer them to a fresh conical tube (see Note 25). 7. Wash the harvested PBMC by adding additional PBS at RT and pellet cells by spinning at 330 × g for 10 min with centrifuge brake on, non-refrigerated. 8. Decant supernatant and resuspend cell pellet(s) using PBS at RT. If appropriate, pool cell pellets from a single donor into one conical tube. Centrifuge balanced tubes at 300 × g for 15 min with centrifuge brake on, non-refrigerated. 9. Decant supernatant and resuspend the cell pellet(s) using PBS to achieve a cell density of ~2–5 × 106 cells/mL. 10. Pipet 15 μL of live/dead cell counting dye onto a piece of parafilm to form a droplet. 11. Remove 15 μL of cell suspension and combine with a droplet of live/dead cell counting dye. Pipet up and down three to five times to mix the sample while avoiding the formation of bubbles. 12. Transfer 15 μL of the cell and dye suspension into each chamber of a hemacytometer. 13. Determine live cell count and viability using CTL’s live/dead cell counting suite. 14. Increase volume of cell suspension(s) with additional PBS and centrifuge balanced tubes at 330× g for 10 min with centrifuge brake on, non-refrigerated. 15. Decant supernatant and gently resuspend the cell pellet (s) using pre-warmed CTL-Cryo™ medium at a cell density of 6–10 × 106 cells/mL to generate aliquots containing 3–5 × 106 PBMC/vial, respectively. 16. Double the volume of the sample(s) by dropwise addition of pre-warmed CTL-Cryo™ medium containing 20% v/v DMSO while gently swirling the tube to ensure adequate mixing.

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17. Immediately transfer sample(s) into labeled 1.2 mL cryovials and place into Mr. Frosty™ freezing containers and transfer into -80 °C freezer (see Note 26), or use a controlled rate freezer. 3.2 Thawing of Cryopreserved PBMC (Sterile Conditions)

1. Place cryovial(s) into a 37 °C bead bath, or better, glass bead bath, for 8 min to thaw. 2. Remove cryovial(s) and wipe with 70% EtOH inside the BSC before unscrewing the cap(s). 3. Using a sterile pipette, transfer contents of cryovial(s) into a labeled conical tube (if applicable, up to 5 vials of the same donor’s cell material can be pooled in one conical tube). 4. Rinse each of the cryovials with 1 mL of warm anti-aggregate solution. Add the warm rinse solution to the conical tube dropwise while swirling the tube to ensure its adequate mixing with the cells in the thawing medium. 5. Double the volume of the cell suspension by dropwise addition of warm anti-aggregate solution while swirling the tube to ensure adequate mixing of the cells and thawing medium. 6. Continue doubling the volume of the cell suspension by dropwise addition of warm anti-aggregate solution while swirling the tube until the cryopreserved cell material has been diluted ten-fold. If multiple cryovials are pooled, calculate using 1 mL of cryopreserved cell suspension + 9 mL of anti-aggregate solution to determine the necessary final resuspension volume. 7. Centrifuge balanced tubes at 330 × g for 10 min with the centrifuge brake on, non-frigerated. 8. Decant supernatant and resuspend the cell pellet(s) using pre-warmed B cell medium (BCM) to achieve a cell density of ~2–5 × 106 cells/mL. You may estimate this number assuming a typical recovery of 70–80% of the frozen PBMC. 9. Pipet 15 μL of live/dead cell counting dye onto a piece of parafilm to form a droplet. 10. Remove 15 μL of cell suspension and combine with a droplet of live/dead cell counting dye. Pipet up and down three to five times to mix the sample while avoiding the formation of bubbles. 11. Transfer 15 μL of the cell and dye suspension into each chamber of a hemacytometer. 12. Determine live cell count and viability using CTL’s live/dead cell counting suite. 13. Increase volume of cell suspension(s) with additional sterile PBS and centrifuge-balanced tubes at 330 × g for 10 min with a centrifuge, non-refrigerated, brake on.

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14. Decant supernatant and gently resuspend the cell pellet (s) using pre-warmed BCM at a cell density of ~2–4 × 106 cells/mL. 3.3 Polyclonal In Vitro Stimulation of B Cells in PBMC (Sterile Conditions)

1. Dilute CTL’s B-Poly-S polyclonal stimulation reagent 1:500 into prewarmed BCM to achieve a final concentration of 2X. Into labeled sterile culture vessels add 50% vol of BCM containing 2X concentration of B-Poly-S. 2. Add the same volume of cell suspension at ~2–4 × 106 cells/ mL to achieve a final culture at ~1–2 × 106 cells/mL with 1X potency of CTL’s B-Poly-S polyclonal stimulation reagent (see Note 27). 3. Transfer culture vessels (flasks or plates) into a humidified incubator set at 37 °C, 5% CO2 for 4–6 days (96–144 h).

3.4 Four-Color, Antigen-Specific FluoroSpot Assay (Affinity Capture Coating)

1. Two days before plating cells (Day 2), prepare 70% EtOH and anti-His affinity capture antibody solutions. 2. Remove underdrain and pipet 15 μL of 70% EtOH solution into the center of each well (or designated wells) of the assay plate. Immediately after the addition of the 70% EtOH solution to the entire plate (or designated wells) add 180 μL/well of PBS. Decant and wash wells again with 180 μL/well of PBS. 3. Decant the assay plate, replace underdrain, and immediately add 80 μL/well of the anti-His affinity capture antibody solution into each well (or designated wells, Note 28) of the low autofluorescence PVDF-membrane plate provided with the kit. 4. Incubate the plate overnight at 4 °C in a humified chamber. 5. The following day (Day 1) dilute the His-tagged protein (s) into diluent A (provided with the kit) to the previously determined optimal concentration (see Note 24). 6. Decant the assay plate and wash wells with 180 μL/well of warm PBS. Immediately, add 80 μL/well of the corresponding His-tagged protein-coating solution(s) into the designated wells (see Note 29). 7. Incubate the plate overnight at 4 °C in a humidified chamber. 8. On the day of the assay (Day 0), decant the assay plate and wash wells with 180 μL/well of warm PBS. Next, decant the plate and add 150 μL/well of pre-warmed BCM to block the plate (≥1 h at RT). 9. If using PBMC following in vitro polyclonal activation, collect the cell suspension(s) and transfer it into labeled conical tube (s). Keep the cells warm during processing. Wash the culture vessel’s interior with sterile warm PBS to collect residual PBMC and transfer it into the corresponding conical tube(s). Increase the volume to fill the tube with additional warm PBS

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and then centrifuge balanced tubes at 330 × g for 10 min non-refrigerated, centrifuge with the brake on. Alternatively, follow the procedures detailed above to obtain freshly isolated PBMC, or to thaw PBMC that were previously cryopreserved, if prior in vitro stimulation is not required to elicit antigenspecific ASC activity in the sample(s). 10. Decant supernatant and resuspend the cell pellet(s) using pre-warmed BCM to achieve a cell density of ~2–5 × 106 cells/mL (the cell number recovered at this point can be estimated to be 50% of the number of cells frozen). 11. Pipet 15 μL of live/dead cell counting dye onto a piece of parafilm to form a droplet. 12. Remove 15 μL of cell suspension and combine with a droplet of live/dead cell counting dye. Pipet up and down three to five times to mix the sample while avoiding the formation of bubbles. 13. Transfer 15 μL of the cell and dye suspension into each chamber of a hemacytometer. 14. Determine live cell count and viability using CTL’s live/dead cell counting suite. 15. Increase volume of cell suspension(s) with additional sterile warm PBS and centrifuge balanced tubes at 330 × g for 10 min with centrifuge brake on, non-refrigerated (see Note 30). 16. Decant supernatant and resuspend the cell pellet(s) using pre-warmed BCM at 2 × 106 PBMC/mL (see Note 31). 17. Decant the BCM used for blocking the ImmunoSpot® assay plate and replace it with 100 μL/well of pre-warmed BCM. 18. Prepare PBMC serial dilution series in a round bottom 96-well polystyrene plate to match the plate layout shown in Fig. 5a (see Note 32). For this, we recommend the following procedure. Into the round bottom 96 well dilution plate, add 10 μL of pre-warmed BCM into all wells, except for row A of columns 1 and 2 for the pan-Ig assay and columns 3 and 4 for the antigen-specific assay. Into wells A3 and A4 add 200 μL/well each of the 2 × 106 PBMC stock (for the pan-Ig assays), dilute 40 μL of the 2 × 106 PBMC stock with 360 μL of warm BCM to obtain 400 μL of cell suspension at 2 × 105 PBMC/mL, of which plate 200 μL each into wells A1 and A2. Using a multichannel pipettor, perform a two-fold dilution series of the PBMC by transferring 100 μL from each row to the next, diluting the cells by gently aspirating and ejecting twice at each dilution step. Once the cell dilution in the round bottom plate is complete, using a multichannel pipettor and fresh tips, transfer 100 μL of the serially diluted cells from the dilution plate into the actual ImmunoSpot® test plate.

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19. Incubate cells in the ImmunoSpot® assay plate for 16–18 h at 37 °C, 5% CO2. 20. After completion of the assay incubation period, decant (or reutilize) cells and wash the plate two times with warm PBS (200 μL/well), followed by two additional washing steps with 0.05% Tween-PBS wash solution (see Note 33). 21. Prepare anti-Ig class/subclass-specific detection antibody solution(s) according to kit protocol and pass through a 0.1 μm low-protein binding syringe filter to remove any protein aggregates. 22. Decant 0.05% Tween-PBS wash solution, add 80 μL/well of the anti-Ig class/subclass-specific detection antibody solution into designated wells, and incubate for 2 h at RT (protected from light). 23. Wash plate(s) two times with 0.05% Tween-PBS wash solution. 24. Prepare a tertiary solution by following kit protocol and pass through a 0.1 μm low-protein binding syringe filter to remove any aggregates. 25. Decant 0.05% Tween-PBS wash solution, add 80 μL/well of tertiary solution into designated wells, and incubate for 1 h at RT (protected from light). 26. Wash plates(s) twice with distilled water. 27. Remove the protective underdrain and place the plate face down on the vacuum manifold. Completely fill the backside of the plate with distilled water and apply a vacuum to draw water through the membrane (“back to front”) (see Note 34). 28. Allow the plate to dry completely, protected from light (see Note 38). 29. Scan and count plate(s) with a suitable analyzer equipped with the appropriate detection channels (see Note 35).

4

Notes 1. While plasma cells (PC) elicited during the primary immune response can secrete large amounts of antibodies, their lifespans are heterogeneous and likely fall on a continuum [8, 9]. 2. The half-life of different Ig classes, and IgG subclasses, is variable and relatively short in vivo. The half-life of IgG1, IgG2, and IgG4 in humans is 21–28 days, whereas for IgG3 it is ~1 week [10]. For IgA and IgM, their half-lives are even shorter (3–7 days) [11, 12] and IgE has the shortest half-life in serum, ~2–3 days [13].

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3. A hallmark of immunological memory is the rapid increase in the level of class-switched, antigen-specific IgG and IgA. 4. To establish the frequency, and Ig class or IgG subclass usage of Bmem-derived ASC for one antigen using a direct ImmunoSpot® assay approach, a typical serial dilution experiment requires 8.8 × 105 PBMC to be seeded into the assay – see the plate layout shown in Fig. 5a. Therefore, if freshly isolated PBMC are to be tested on day 5–9 post onset of the B cell response when spontaneously Ig-secreting, antigen-specific plasmablasts are present in the blood, 1 mL of blood should suffice to complete such a test. If the PBMC from such blood is to be cryopreserved before testing, 1.5–2 × 106 PBMC should be frozen to obtain, with a safety margin, 8.8 × 105 viable and fully functional PBMC after thawing [7]. If resting memory B cells in PBMC are to be assessed, however, the PBMC need first to be subject to a 5 day in vitro stimulation culture [15], after which about 50% of the (fresh or thawed) PBMC are recovered (Becza et al., manuscript in preparation). Thus, for working with thawed PBMC and after polyclonal stimulation, freezing 3 (with added safety, 4) × 106 PBMC is required to end up with 8.8 × 105 PBMC on the day of the test. Any number of freshly isolated PBMC between 1 and 10 × 106 per vial can be cryopreserved recovering the proportional number of fully functional PBMC after thawing (Becza et al., manuscript in preparation). 5. Stimulating optimal Ig class usage during an infection or following vaccination is vital to successful host defense and the avoidance of collateral immune-mediated pathology (reviewed in [14]). 6. Flow cytometry does not reliably reveal the class/subclass of Ig produced by the individual B cell because surface BCR expression can be highly variable and this is an underappreciated complexity of probe staining. In particular, in the case of IgG+ ASC, they express little if any surface BCR and this undermines the assessment of their antigen specificity and subclass usage by traditional surface staining approaches. Consequently, fixation and intracellular staining are required to define the IgG subclass usage of these cells (a procedure that results in substantial cell loss in the sample). 7. Memory B cells exist in a quiescent state in the absence of recent antigen encounter and do not secrete their individual BCR as soluble antibodies. To overcome this obstacle for detecting them in ImmunoSpot® assays, in vitro polyclonal stimulation protocols can be used to trigger the antigenindependent activation of resting memory B cells into ASC [15].

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8. Spontaneous ASC activity can also be evaluated directly ex vivo (e.g. plasmablasts that occur in PBMC 5–9 days after onset of a B cell response, or PC residing in the bone marrow). 9. If testing PBMC in the absence of prior stimulation (e.g. to measure plasmablasts), or other samples for spontaneous ASC activity directly ex vivo, the optimal cell inputs for establishing the frequency of all ASC-producing IgM, IgG, and IgA will be much higher. 10. IgE+ ASC are quite rare and consequently, ImmunoSpot® assays that aim to determine their relative frequency require high initial starting cell inputs. Despite testing numerous human PBMC samples (>50) of healthy, non-allergic individuals following in vitro stimulation with CTL’s B-Poly-S, we have not detected IgE+ ASC activity in any of these samples thus far. IgE+ ASC can be detected in PBMC of such individuals, however, following polyclonal stimulation with IL-21 in conjunction with anti-CD40 and IL-4 [15] that mimics T cell help (suggesting class switching to IgE during the in vitro cell culture period using this particular polyclonal stimulation protocol). 11. Cell material is most often the limiting component for immune monitoring. 12. B cell ImmunoSpot® data can be expressed as SFU per cell input per well to determine the frequency of antigen-specific cells. However, owing to the variable abundance of pan IgG+ ASC in test samples following polyclonal stimulation, some prefer to report data as the frequency of antigen-specific B cells secreting a given Ig class/subclass among all B cells secreting that Ig class/subclass. 13. Each investigator in our laboratory can routinely test, following the protocol outlined in this chapter, in a single experiment, 10–20 PBMC samples for reactivity against a panel of antigens, assessing the frequency of ASC producing each of the Ig classes and IgG subclasses. With additional logistical refinements, this throughput is readily upward scalable. 14. If a special protocol is followed, PBMC can be frozen without impairing the B cells’ functionality ([7] and N. Becza manuscript in preparation). Thus, by freezing B cells of a sample in several aliquots, the same PBMC can be tested repeatedly, reproducing the results of the previous experiment with high accuracy [4], or extending those studies. Of note, when planning the numbers of PBMC to be frozen per cryovial, as a rule of thumb, one can anticipate recovery of ~50% of PBMC initially frozen after these cells are thawed and have undergone 5 days of polyclonal stimulation to promote terminal differentiation of resting Bmem into ASC (see also Note 4). It is also

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important to know that any number of PBMC between 1 and 10 × 106 can be frozen per cryovial permitting the optimization of PBMC utilization when planning experiments (N. Becza, manuscript in preparation). 15. In ImmunoSpot® assays, there is no inherent lower limit of detection. If, e.g., 3 × 106 PBMC are plated at 3 × 105 PBMC across 10 replicate wells, 1 in 3 × 106 is the detection limit, etc. Importantly, however, owing to increased Poisson noise occurring with such low-frequency measurements, the number of replicate wells evaluated needs to be increased accordingly to obtain accurate low-frequency measurements. 16. The so-called “Goldilocks” number is defined as the maximal number of cells that can be plated in an assay well while still being able to discern clearly individual secretory footprints derived from antigen-specific ASC. As it is assay-dependent, it needs to be experimentally established, but 50 SFU/well is a safe estimate. 17. Once the so-called “Goldilocks” number has been established, using a serial dilution approach as illustrated in Fig. 5, in a second experiment, the cell number(s) can be chosen accordingly for generating replicates containing ≥300 individual ASC-derived footprints for studying the functional affinity distribution (see also the chapter in this volume by Becza et al. [3]). Obviously, the higher the assay-specific Goldilocks number, the fewer replicates are needed to obtain ≥300 cumulative secretory footprints. 18. Thawing of cryopreserved cells causes a fraction of the cells (up to 30%) to die, and the DNA released from such cells can cause clumping of the thawed cell material. This cell clumping can be reduced, if not completely eliminated, by including an immunologically neutral endonuclease, Benzonase. Ready-touse Benzonase-containing, serum-free wash solutions are available: CTL anti-aggregate Wash™ 20X solution. 19. A suitable assay medium for use in B cell ImmunoSpot® is RPMI 1640 with 10% FCS, 2 mM L-glutamine, 100 U/mL penicillin, 100 μg/mL streptomycin, 8 mM HEPES, and 50 μM 2-mercaptoethanol. 20. Kit is suited for detecting all antibody-secreting cells (ASC) producing a given Ig class or Ig subclass, irrespective of antigen-specificity, that differentiated in vivo, or following an in vitro polyclonal stimulation protocol to promote their transition to ASC. Each kit contains pan anti-Ig capture antibody, Ig class or Ig subclass-specific detection reagents, diluent buffers, PVDF-membrane plates, development substrate solutions, and polyclonal B cell activator (B-Poly-S or B-Poly-

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SE). Of note, B-Poly-SE is capable of stimulating ASC that produce one of the distinct Ig classes/subclasses; including IgE (see also Note 10). 21. Kit is suited for detecting all antibody-secreting cells (ASC) producing a given Ig class or Ig subclass, irrespective of antigen-specificity, that differentiated in vivo, or following an in vitro polyclonal stimulation protocol to promote their transition to ASC. Each kit contains pan anti-Ig capture antibody, Ig class or Ig subclass-specific detection reagents, diluent buffers, low autofluorescence PVDF-membrane plates, and polyclonal B cell activator (B-Poly-S or B-Poly-SE). Importantly, Ig class-specific and/or Ig subclass-specific detection reagents can be combined to generate multiplexed detection systems enabling two-, three, or four-color B cell ImmunoSpot® assays. 22. Kit is suited for detecting either antigen-specific antibodysecreting cells (ASC) that differentiated in vivo, or antigenspecific Bmem that have been polyclonally stimulated in vitro to promote their transition to ASC. Each kit contains anti-His capture antibody, Ig class-specific (IgA, IgE, IgG, and IgM) detection reagents, diluent buffers, low autofluorescence PVDF-membrane plates, and polyclonal B cell activator (B-Poly-S). Alternatively, IgG class-specific (IgG1, IgG2, IgG3, and IgG4) detection reagents can be substituted in the context of such four-color B cell ImmunoSpot® assays. 23. Traditional B cell ELISPOT assays have been performed by direct coating of the assay membrane with the antigen of interest. However, many (in fact, most) antigens do not adsorb sufficiently to the membrane to enable reliable detection of ASC-derived secretory footprints. We have overcome this limitation by introducing an affinity coating approach for achieving high-density antigen absorption to the assay membrane [16]. 24. We recommend optimizing the concentration of His-tagged protein(s) used for affinity capture coating. A concentration of 10 μg/mL His-tagged protein has yielded well-formed secretory footprints for most antigens, but increased concentrations of the anti-His affinity capture antibody and/or His-tagged protein may be required to achieve optimal assay performance [16]. 25. Take care to collect as little Lymphoprep™ as possible. At this point, the interphase of two conical tubes can be combined into one tube. If the proportion of Lymphoprep™ is too high (≥ 10% v/v), significant cell loss may occur. 26. A cooling rate of 1 °C/min is optimal for cell cryopreservation. Be sure to fill the lower compartment of the Mr. Frosty™ Freezing container with 100% isopropyl alcohol and replace it after five cryopreservation cycles. After approximately 20 h,

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and no more than 2 days, transfer cryopreserved PBMC from the -80 °C freezer into liquid nitrogen for long-term storage. 27. The volume of in vitro stimulation cultures can be scaled up or down accordingly, but we recommend keeping the cell density of PBMC at approximately 1–2 × 106 cells/mL. If larger numbers of in vitro stimulated PBMC are required for downstream ImmunoSpot® assays, tissue culture flasks should be used; store flat or standing such the height of the cell suspension in the flask is between 0.5 and 1 cm. Smaller in vitro stimulation cultures can be initiated in 48- or 24-well plates with a final volume of 1–2 mL, respectively. Be sure to fill empty wells in tissue culture plates with sterile PBS to avoid dehydration of cell cultures. 28. If the entire plate will not be coated with the anti-His affinity capture antibody solution, the remainder of the EtOH pre-wet wells should receive 80 μL/well of PBS. 29. If performing both an antigen-specific affinity capture and total ASC assay on the same plate, we recommend adding the pan anti-Ig capture antibody at this stage. 30. If the cells are not washed thoroughly, contaminating antibodies in the cell suspension(s) can compete for binding of the affinity captured antigen and may also result in elevated membrane staining that interferes with accurate enumeration of individual antigen-specific ASC. 31. This starting cell input was used to generate the data presented in Figs. 2, 3, 4 and 5 and to highlight the vastly different frequencies of ASC that produce each Ig class or subclass following in vitro polyclonal stimulation. 32. Do NOT do serial dilutions in the actual ImmunoSpot® plates because the pipet tips can easily damage the membrane. For multiple PBMC, these cell transfers can readily be done simultaneously using a multichannel pipettor. 33. Plate washes may also be performed manually. For automated washing, the pin height and flow rate should be customized to avoid damaging the assay membranes, which is the case for the CTL 405LSR plate washer. 34. Optimal removal of background staining, fibers, and other debris, along with reduction of “hot spots” in the center of the assay wells, is achieved through performing the “back to front” water filtration technique. 35. The chapter by Karulin et al. in this volume [5] introduces artificial intelligence-based SFU analysis that can partially compensate for ELISA effects and SFU crowding, thus extending the linear range of accurate quantification for cell numbers plated per well and SFU detected.

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Acknowledgments We wish to thank the R&D and the Software Development teams at CTL for their continued support and technological innovation that made our B cell ImmunoSpot® endeavor possible. We thank Drs. Alexey Y. Karulin and Graham Pawelec for in-depth discussions of the subject matter, Diana Roen for carefully proofreading the manuscript, and Gregory Kovacs for his support in the generation of graphic illustrations. All efforts were funded from CTL’s research budget. Conflicts of Interest P.V.L. is Founder, President, and CEO of CTL, a company that specializes in immune monitoring by ImmunoSpot®. L.Y., N.B., A.M.P., J.C., and G.A.K. are employees of CTL.

References 1. Lehmann PV, Becza N, Liu Z et al (2023) Monitoring memory B cells by next generation ImmunoSpot® provides insights into humoral immunity that measurements of circulating antibodies do not reveal. Methods Mol Biol 2. Akkaya M, Kwak K, Pierce SK (2020) B cell memory: building two walls of protection against pathogens. Nat Rev Immunol 20(4): 229–238. https://doi.org/10.1038/s41577019-0244-2 3. Becza N, Liu Z, Chepke J et al (2023) Assessing the affinity spectrum of the antigenspecific B cell repertoire in freshly isolated cell material via ImmunoSpot®. Methods Mol Biol 4. Wolf C, Koppert S, Becza N et al (2022) Antibody levels poorly reflect on the frequency of memory B cells generated following SARSCoV-2, seasonal influenza, or EBV infection. Cell 11(22). https://doi.org/10.3390/ cells11223662 5. Karulin AY, Megyesi Z, Kirchenbaum GA et al (2023) Artificial intelligence-based counting algorithm enables accurate and detailed analysis of the broad spectrum of spot morphologies observed in antigen-specific B cell EliSpot and FluoroSpot assays. Methods Mol Biol 6. Bisceglia H, Barrier J, Ruiz J et al (2023) A FluoroSpot B assay for the detection of IgA and IgG SARS-CoV-2 spike-specific memory B cells: Optimization and qualification for use in COVID-19 vaccine trials. J Immunol Methods 515:113457. https://doi.org/10.1016/j. jim.2023.113457

7. Fecher P, Caspell R, Naeem V et al (2018) B cells and B cell blasts withstand cryopreservation while retaining their functionality for producing antibody. Cell 7(6). https://doi.org/ 10.3390/cells7060050 8. Lightman SM, Utley A, Lee KP (2019) Survival of long-lived plasma cells (LLPC): piecing together the puzzle. Front Immunol 10:965. https://doi.org/10.3389/fimmu.2019. 00965 9. Robinson MJ, Ding Z, Dowling MR et al (2023) Intrinsically determined turnover underlies broad heterogeneity in plasma-cell lifespan. Immunity 56(7):1596–1612. e1594. https://doi.org/10.1016/j.immuni.2023. 04.015 10. Morell A, Terry WD, Waldmann TA (1970) Metabolic properties of IgG subclasses in man. J Clin Invest 49(4):673–680. https:// doi.org/10.1172/JCI106279 11. Blandino R, Baumgarth N (2019) Secreted IgM: New tricks for an old molecule. J Leukoc Biol 106(5):1021–1034. https://doi. org/10.1002/JLB.3RI0519-161R 12. van Tetering G, Evers M, Chan C et al (2020) Fc engineering strategies to advance IgA antibodies as therapeutic agents. Antibodies 9(4). https://doi.org/10.3390/antib9040070 13. Normansell R, Walker S, Milan SJ et al (2014) Omalizumab for asthma in adults and children. Cochrane Database Syst Rev 1:CD003559. h t t p s : // d o i . o r g / 1 0 . 1 0 0 2 / 1 4 6 5 1 8 5 8 . CD003559.pub4

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14. Webb NE, Bernshtein B, Alter G (2021) Tissues: the unexplored frontier of antibody mediated immunity. Curr Opin Virol 47:52– 67. https://doi.org/10.1016/j.coviro.2021. 01.001 15. Franke F, Kirchenbaum GA, Kuerten S et al (2020) IL-21 in conjunction with anti-CD40 and IL-4 constitutes a potent polyclonal B cell

stimulator for monitoring antigen-specific memory B cells. Cell 9(2). https://doi.org/ 10.3390/cells9020433 16. Koppert S, Wolf C, Becza N et al (2021) Affinity tag coating enables reliable detection of antigen-specific B cells in Immunospot assays. Cell 10(8). https://doi.org/10.3390/ cells10081843

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 16 High-Plex ELISPOT: FOLISPOT Based on Fluorescence Detection and DNA Complementary Pairing Mi Liu Abstract Enzyme-linked immunospot (ELISPOT) is one of the most important methods to measure the number of specific cells by detecting protein secretion at a single-cell level. However, traditional ELISPOT based on enzyme-substrate color development can only detect one target. Therefore, scientists developed multipletarget ELISPOT based on enzyme-substrate coloring. Besides, FluoroSPOT that can detect 2–4 fluorescent signals are developed. Nevertheless, the maximum detection targets of multiple-target ELISPOT and FluoroSPOT are around 4, and the signal amplification system can be further optimized. Fluorescencebased oligo-linked immunospot (FOLISPOT), which utilized DNA-barcoded antibodies to provide a highly multiplexed method with signal amplification, was developed to detect multiple targets simultaneously. In this method, multiple targets can be detected in one round and multiple rounds of detection can be conducted, and thus a large number of targets can be detected. Besides, signal amplification is achieved by DNA complementary pairing and modular orthogonal DNA concatemers, and thus cells secreting limited amounts of proteins can be detected. According to the studies, FOLISPOT can detect more spots than ELISPOT and can detect targets that are undetectable by ELISPOT. Furthermore, FOLISPOT can be utilized to detect more than 6 targets, by allowing sequential detection of multiple targets in one round and sequential detection in multiple rounds. Key words FOLISPOT, DNA complementary pairing, DNA concatemers, ELISPOT, Fluorescencebased oligo-linked immunospot

1

Introduction Currently, immunoassay plays an important role in the analysis of many common samples and cells. There are many kinds of immunoassay methods, such as precipitation reaction, agglutination reaction, and immunolabeling technology. ELISPOT is one of the most widely used methods for the quantitative determination of specific cells, such as cells that can secrete IFN- γ [1–3]. It can be used to

Reprinted (adapted) with permission from Anal. Chem. 2022, 94, 24, 8704–8714, Publication Date: June 1, 2022, https://doi.org/10.1021/acs.analchem.2c01087, Copyright, 2022, American Chemical Society. Alexander E. Kalyuzhny (ed.), Handbook of ELISPOT: Methods and Protocols, Methods in Molecular Biology, vol. 2768, https://doi.org/10.1007/978-1-0716-3690-9_16, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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quantitatively measure the secretion of antigens or antibodies in a medium to detect activated cells at the single-cell level. Thanks to the single-cell level measurements, ELISPOT is often used to measure the number of cells secreting a specific protein, such information is crucial for evaluating the disease or treatment of cancer, tuberculosis, and other diseases. Although traditional ELISPOT has many advantages in the field, their techniques also have some limitations when detecting the lower abundance targets or simultaneously detecting multiplex proteins in a single detection. Therefore, in order to meet some special needs, it is particularly important to develop a high-sensitivity detection system with a signal amplification function and high-plex detection capacity. To overcome these limitations faced by traditional ELISPOT, multi-color FluoroSPOT assay and multiple-target ELISPOT based on enzyme-substrate coloring were developed to simultaneously detect several cytokines or antibodies secreted by T cells or B cells [2, 4–7]. By detecting each cytokine with a specific fluorophore (or enzyme-substrate system) and analyzing spots of different colors through a fluorophore-specific filtering system, thus cells that produce single or multiple cytokines can be investigated. However, the color types of fluorophores are limited and this method can only detect up to about four targets at the same time, and cannot do multiple-round measurements, which is not enough for multiplexing capabilities [2, 4, 8–12]. Therefore, a new method fluorescence-based oligo-linked immunospot (FOLISPOT), that can simultaneously detect multiple targets and contains an amplification method to support high sensitivity, was developed [10]. The method was designed to be easily accessible, utilizing a simple and robust workflow compatible with commonly available imaging platforms and cost-effective and readily available reagents. FOLISPOT achieves the function of signal amplification and simultaneous detection of multiple targets in a single detection by DNA barcode antibodies and DNA complementary pairing signal amplification system (which replace the enzyme–substrate). In addition, due to the binding between DNA strands can be designed to be stable at the specific conditions and can be washed off at other conditions, FOLISPOT can function multiple-round detections, with each round detecting several targets. Thus, the number of detection targets is maximized (Fig. 1 [10]). In summary, FOLISPOT can be used to detect the secretion frequency of multiple proteins at the single-cell level at one time, and FOLISPOT are rapid, signal-amplified, highly multiplexed method that overcame current key challenges and could detect multiple signals simultaneously. The straightforward material preparations and protocols, along with compatibility with common equipment, make this method readily adoptable [10].

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Fig. 1 Detection of cell-secreted antigens by FOLISPOT. (a) Schematic diagram of detecting 4 targets simultaneously by FOLISPOT with single stage amplification system. (b) Schematic diagram of detecting 6 targets simultaneously by FOLISPOT with single stage amplification system. (c) Schematic diagram of detecting 6 targets simultaneously by FOLISPOT with dual-stage amplification system [10]

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Materials 1. Unmodified, dye-labeled, oligonucleotides,

and

thiol-modified

DNA

2. Bst LF polymerase 3. dNTP set solution 4. Tris 5. Boric acid 6. Tween-20 7. Agarose 8. Sodium chloride 9. 1 kb plus DNA ladder 10. GenElute 11. RPMI medium 12. Ficoll Histopaque 13. Maleimide-PEG-succinimidyl ester crosslinkers 14. Phytohemagglutinin (PHA-P) 15. Lipopolysaccharides 16. Ethylenediaminetetraacetic acid disodium salt dihydrate 17. Trifluoroacetic acid 18. Magnesium chloride 19. DMF 20. Amicon Ultra Centrifugal Filter (100 kDa MWCO) 21. Zaba spin desalting column (7000 MWCO) 22. NAP-5 columns 23. 1 M Tris-HCL (pH = 8.0) 24. 0.5 M EDTA (pH = 8.0) 25. Sodium carbonate-sodium bicarbonate buffer (0.05 mol/L, pH = 9.6) 26. Deionized formamide (see Note 1) 27. Acetonitrile 28. Gel Red 29. Fetal bovine serum 30. Penicillin–streptomycin solution 31. Washing buffer: (a) 1xPBS (b) 0.05%(vol/vol) tween-20 32. Blocking buffer:

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(a) 1xPBS (b) 1%BSA 33. Hybridization buffer: (a) 2 × SSC (b) 10% Dextran sulfate (c) 10% Formamide (deionized) (d) 10 mM MgCl2 (e) 1 mM EDTA 34. Hybridization washing buffer: (a) 2 × SSC (b) 10% Formamide (deionized) (c) 10 mM MgCl2 (d) 1 Mm EDTA 35. Fluorescent hybridization buffer: (a) 1 mM Tris-HCl (b) 500 mM NaCl (c) 10 mM MgCl2 (d) 1 mM EDTA 36. Dissociation buffer: (a) 1 × PBS (b) 50% Formamide

3

Methods

3.1 Antibody–DNA Conjugation and Purification

1. DNA oligonucleotides were activated as follows: 250 uM 5′-thiol-modified DNA oligonucleotides were activated by 100 mM DTT for 2 h at room temperature in the dark. 2. The sample was purified by using NAP5 columns. 3. Antibodies were concentrated to 2 mg/mL by using 0.5 mL 100 kDa Amicon Ultra Filters and then were allowed to react with maleimide-PEG2-succinimidyl ester crosslinkers for 2 h at 4 °C. 4. Antibodies were then purified by using 0.5 mL 7 kDa Zeba desalting columns. 5. Activated DNA oligonucleotides were incubated with modified antibodies (20:1, DNA: antibody molar ratio) overnight at 4 °C. 6. The conjugated antibodies were then washed by using PBS in 0.5 mL 100 kDa Amicon Ultra Filters ten times to remove non-reacted DNA oligonucleotides.

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Table 1 DNA docking strand sequences used for antibody conjugation DNA docking strand index

DNA sequence

DNA docking strand 1

AAATTCCTCTACCACCTACATCAC

DNA docking strand 2

TATTTAGTGTTCGAATAGTTCGATCTAG

DNA docking strand 3

AATTCTATGACACCGCCACGCCCTATATCCTCGCAATAACCC

DNA docking strand 4

GTTTCCTATATTTAGCGTCCGTGTCGTTCTCCCGCGCAACAG

DNA docking strand 5

GAATCCAAATCGTCCGTTAGCAGTGAACCGTACCGTCTCAAT

DNA docking strand 6

CAAGTCTAGTCCAAGGTCCGTCAAGGTCTCCGACGCATACTC

DNA docking strand 7

AGTTCCTGTAGTATCCCGTCGCATAGTCGTACATTCACCGTC

*The list of bridge sequences used for conjugation that is referred in reference [10] is provided in Table 1. 3.2 Characterization of DNA-Conjugated Antibodies by MALDITOF [10]

1. The DNA-modified antibody (DNA-Ab, 3 mg/mL) was done buffer exchange and transferred to Milli-Q water by using 7 kDA Zeba spin columns. 2. A matrix solution was prepared by dissolving sinapinic acid (1 mg) in acetonitrile (50 μL) and water with 0.1% trifluoroacetic acid (50 μL). 2 μL of the DNA–antibody solution was deposited onto the MALDI plate and then mixed with 2 μL of MALDI matrix. 3. The plate was allowed to dry at room temperature for approximately 1–2 h before analysis. The MALDI-TOF mass data were collected using the AB SCIEX 4800 MALDI-TOF/TOF analyzer.

3.3 Preparation of Concatemers

1. The reaction was conducted in 1 mL PBS with final concentrations of 10 mM MgSO4, 80 units/mL of Bst LF polymerase, 600 μM each of dATP/dCTP/dTTP, 10 μM hairpin, and 10 μM primer. The mixture was incubated for 9 h at 37 °C, followed by heating to inactivate the polymerase at 80 °C. 2. Concatemers were stored at -20 °C for the following experiments. *Primer sequences and details of the conditions are listed in Tables 2 and 3.

3.4 Verification and Purification of Concatemers

1. The lengths of concatemers were evaluated by diluting 1 μL of the sample with 9 μL of 10 × loading buffer. The diluted samples were then run on 1% agarose gels for 20 min on the E-gel apparatus and imaged with the Gel Red channel on a SYSTEM GelDoc XR+ IMAGELA scanner.

p.4

p.5

p.6

p.7

p.8

p.9

C.4

C.5

C.6

C.7

C.8

C.9

ATGATGATG-TATGATGATG-TATGATGATG-TTTTTTTCTACCATTTTCTACC

TAGGTTTAT-TTAGGTTTAT-TTAGGTTTAT-TTTTTCATTTACATTCATTTAC

GACGGTGAATGTACGACTATGCGACGGGATACTACAGGAACTTTCCAATAATA-A-CCAATAATA

GAGTATGCGTCGGAGACCTTGACGGACCTTGGACTAGACTTGTTCAATCAAAA-A-CAATCAAAA

ATTGAGACGGTACGGTTCACTGCTAACGGACGATTTGGATTCTT-TTCATTTAC-ATTCATTTAC

CTGTTGCGCGGGAGAACGACACGGACGCTAAATATAGGAAACTT-ACAACTTAAC

GGGTTATTGCGAGGATATAGGGCGTGGCGGTGTCATAGAATTTTAATACTCTC

GTTAAGTTG-TGTTAAGTTG-TGTTAAGTTG-TTTCCTTCTATTACCTTCTATT

p.3

C.3

CTAGATCGAACTATTCGAACACTAAATATT-CATCATCAT

C.11 p.11

p.2

C.2

GTGATGTAGGTGGTAGAGGAATTT-TTATAAACCTA

GAGAGTATT-TGAGAGTATT-TGAGAGTATT-TTTTCCTTTTATATCCTTTTAT

p.1

C.1

Primer sequence

C.10 p.10

Primer ID

ID

ATTTTCTACCGGGCCTTTTGGCCCGGT AGAAAATGGTAGAAAA/3InvdT/

ATTCATTTACGGGCCTTTTGGCCCGTA AATGAATGTAAATGAA/3InvdT/

ACCAATAATAGGGCCTTTTGGCCCTATT ATTGGTTATTATTGG/3InvdT/

ACAATCAAAAGGGCCTTTTGGCCCTTT TGATTGTTTTTGATTG/3InvdT/

ATTCATTTACGGGCCTTTTGGCCCGTA AATGAATGTAAATGAA/3InvdT/

ACAACTTAACGGGCCTTTTGGCCCGTT AAGTTGTGTTAAGTTG/3InvdT/

AAATACTCTCGGGCCTTTTGGCCCGAG AGTATTTGAGAGTATT/3InvdT/

ACATCATCATGGGCCTTTTGGCCCATG ATGATGTATGATGATG/3InvdT/

AATAAACCTAGGGCCTTTTGGCCCTAG GTTTATTTAGGTTTAT/3InvdT/

Hairpin sequence

(continued)

h.11.11 ATCCTTTTATGGGCCTTTTGGCCCATAA AAGGATATAAAAGGA/3InvdT/

h.10.10 ACCAATAATAGGGCCTTTTGGCCCTAT TATTGGTTATTATTGG/3InvdT/

h.9.9

h.8.8

h.7.7

h.6.6

h.5.5

h.4.4

h.3.3

h.2.2

h.1.1

Hairpin ID

Table 2 The information of primers and hairpins used in preparing concatemers by primer exchange reaction (PER)

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TTTTGATTG-TTTTTGATTG-TTTTTGATTGTTTTCATTACTTATCATTACTT-ATCATTACTT

TATTATTGG-T-TATTATTGG-TTATTATTGG-TTTTTCTTACTCATTCTTACTC-ATTCTTACTC

C.13 p.13

C.14 p.14

Primer sequence

GTAAATGAA-TGTAAATGAA-TGTAAATGAA-TTTTTATTCACTATTATTCACT-ATTATTCACT

Primer ID

C.12 p.12

ID

Table 2 (continued)

Hairpin sequence

h.14.14 ATTCTTACTCGGGCCTTTTGGCCCGA GTAAGAATGAGTAAGAA/3InvdT/

h.13.13 ATCATTACTTGGGCCTTTTGGCC CAAGTAATGATAAGTAATGA/3InvdT/

h.12.12 ATTATTCACTGGGCCTTTTGGCCCAGT GAA TAATAGTGAATAA/3InvdT/

Hairpin ID

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Table 3 DNA amplification concatemer sequences ID

DNA amplification strand sequence (contains multiple repeating units)

C.1

GTGATGTAGGTGGTAGAGGAATTT-TT-(ATAAACCTA-A)nATAAACCTA-A(40≤n≤60)

C.2

CTAGATCGAACTATTCGAACACTAAATA-TT-(CATCATCAT-A)nCATCATCAT-A (40≤n≤60)

C.3

GGGTTATTGCGAGGATATAGGGCGTGGCGGTGTCATAGAATTTT- (AATACTCTC-A)nAATACTCTC-A(40≤n≤60)

C.4

CTGTTGCGCGGGAGAACGACACGGACGCTAAATATAGGAAACTT-A-(CAACTTAAC-A) n-CAACTTAAC-A(40≤n≤60)

C.5

ATTGAGACGGTACGGTTCACTGCTAACGGACGATTTGGATTCTT- -TTCATTTAC-A(TTCATTTAC-A)n-TTCATTTAC-A (40≤n≤60)

C.6

GAGTATGCGTCGGAGACCTTGACGGACCTTGGACTAGACTTGTT- CAATCAAAA-A(CAATCAAAA-A)n-CAATCAAAA-A(40≤n≤60)

C.7

GACGGTGAATGTACGACTATGCGACGGGATACTACAGGAACTTT- CCAATAATA-A(CCAATAATA-A)n-CCAATAATA-A(40≤n≤60)

C.8

TAGGTTTAT-T-TAGGTTTAT-T-TAGGTTTAT-TTT-TTCATTTACA- (TTCATTTAC-A)nTTCATTTAC-A(40≤n≤60)

C.9

ATGATGATG-T-ATGATGATG-T-ATGATGATG-TTTTTTTCTACC-A- (TTTTCTACC-A)nTTTTCTACC-A(40≤n≤60)

C.10 GAGAGTATT-T-GAGAGTATT-T-GAGAGTATT-TTTTCCTTTTAT-A- (TCCTTTTAT-A)nTCCTTTTAT-A(40≤n≤60) C.11 GTTAAGTTG-T-GTTAAGTTG-T-GTTAAGTTG-TTT-CCTTCTATTA- (CCTTCTATT-A)nCCTTCTATT-A( 40≤n≤60) C.12 GTAAATGAA-T-GTAAATGAA-T-GTAAATGAA-TTTTTATTCACT-A-TTATTCACT-A(TTATTCACT-A)n-TTATTCACTA (40≤n≤60) C.13 TTTTGATTG-T-TTTTGATTG-T-TTTTGATTGTTT-TCATTACTT-ATCATTACTT-A(TCATTACTT-A)n-TCATTACTT-A(40≤n≤60) C.14 TATTATTGG-T-TATTATTGG-T-TATTATTGG-TTT-TTCTTACTCA-TTCTTACTC-A(TTCTTACTC-A)n-TTCTTACTC-A (40≤n≤60)

2. In addition, concatemers were purified by using a GenElute kit and lyophilized to a solid state on a CentriVap. *The amplification concatemers longer than 160 bp were prepared by primer exchange reaction (PER), and concatemers shorter than 160 bp were prepared by chemical synthesis. 3.5 Detection Strands

Detection strands are 20 mer DNA oligonucleotides with fluorophores on the 5′ end and inverted dT modification on the 3′ end. Detection strands were designed to bind the dimers of the primer unit sequence to achieve stable but easily reversible binding that is necessary for DNA exchange imaging.

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Table 4 DNA strands used for detection (see Note 3) ID

Detection of DNA strand sequence

i.1*

Fluorophore-TT-TAGGTTTAT-T-TAGGTTTAT-T

i.2*

Fluorophore-TT-ATGATGATG-T-ATGATGATG-T

ai.3*

Fluorophore-TT-GAGAGTATT-T-GAGAGTATT-T

i.4*

Fluorophore-TT-GTTAAGTTG-T-GTTAAGTTG-T

i.5*

Fluorophore-TT-GTAAATGAA-T-GTAAATGAA-T

i.6*

Fluorophore-TT-TTTTGATTG-T-TTTTGATTG-T

i.7*

Fluorophore-TT-TATTATTGG-T-TATTATTGG-T

i.8*

Fluorophore-TT-GTAAATGAA-T-GTAAATGAA-T

i.9*

Fluorophore-TT-GGTAGAAAA-T-GGTAGAAAA-T

i.10*

Fluorophore-TT-ATAAAAGGA-T-ATAAAAGGA-T

i.11*

Fluorophore-TT-AATGAAAGA-T-AATGAAAGA-T

i.12*

Fluorophore-TT-AGTGAATAA-T-AGTGAATAA-T

i.13*

Fluorophore-TT-AAGTAATGA-T-AAGTAATGA-T

i.14*

Fluorophore-TT-GAGTAAGAA-T-GAGTAAGAA-T

The format of the detection strands sequences is: 5′-fluorophore-tt-primer*-t-primer*-t-3′. They were with 5′-fluorophore (FAM or Texas Red). They are named as i.primer ID*and the sequences are listed in Table 4. 3.6

Cell Culture

PBMCs were grown in RPMI-1640 supplemented with 10%(vol/vol) fetal bovine serum and 1% (vol/vol) penicillin/streptomycin solution under 37 °C and 5% CO2.

3.7 Signal Amplification

150 nM primary concatemer was added into wells and incubated overnight at 37 °C in a solution containing 10% formamide,10% dextran sulfate, 10 mM MgCl2, and 1 mM EDTA. All primary and secondary concatemers were added into wells and incubated overnight at 37 °C in a solution containing 10% formamide,10% dextran sulfate, 10 mM MgCl2, and 1 mM EDTA. *The mechanism of signal amplification by DNA-modified antibodies is shown in Fig. 2 and the concatemer sequences for each target are given in Tables 3 and 5.

3.8 Fluorophore Hybridization and Dehybridization

Detection strands were hybridized at 1 μM in 1 mM Tris-HCl (supplemented with 500 mM NaCl, 10 mM MgCl2, and 1 mM EDTA) for 1 h at room temperature. Detection strands were removed by incubating at room temperature in PBS containing

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Fig. 2 Mechanism of signal amplification by DNA-modified antibodies. (a) Scheme of signal amplification by a single-stage amplification system. Step 1: synthesis of primary DNA amplification strands by primer exchange reaction (PER); step 2: binding of amplification strands with DNA-conjugated antibodies before the detection; step 3: addition of DNA detection strands to perform the measurements. (b) Scheme of signal amplification by a dual-stage amplification system. Step 1: synthesis of secondary DNA amplification strands by primer exchange reaction (PER); step 2: binding of secondary amplification strands with primary DNA amplification strands during the measurement; step 3: addition of DNA detection strands to perform the measurements [10]

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Table 5 Antibodies used in six-target multiplexed FOLISPOT experiments, DNA docking strand sequences, respective amplification concatemers, and detection sequence

Target

DNA sequence (conjugate to detection antibodies)

DNA sequence (conjugate to detection antibodies)

DNA sequence (conjugate to detection antibodies)

Il-2

DNA docking strand 7

C.7 + C.14

i.14*

TGF- β

DNA docking strand 2

C.2 + C.9

i.9*

IFN- γ

DNA docking strand 3

C.3 + C.10

i.10*

TNF- α

DNA docking strand 4

C.4 + C.11

i.11*

IL-10

DNA docking strand 5

C.5 + C.12

i.12*

IL-6

DNA docking strand 6

C.6 + C.13

i.13*

50% formamide for 10 min, followed by two times washing with PBS at room temperature. A new round of detection hybridization was performed as above. Fluorescence intensity was measured by using an instrument (such as a Cytation 5 spectrophotometer). 3.9 Detection of Multiple Targets by FOLISPOT

In this example, FOLISPOT with primary signal amplification system and FOLISPOT with dual-stage amplification system were applied to simultaneously analyze 4 or 6 cytokine secreting (IL-2, IL-10, TGF-β1, IL-6, IFN-γ, and TNF-α) in PBMC samples. The fluorescent probe modified on DNA detection chains for IL-2, IL-10, and IFN-γ were FAM, and the fluorescent probe modified on DNA detection chains for IL-6, TGF-β1, and TNF-α were Texas red (see Note 2). In addition, three groups of different cell densities were set up for comparison in this experiment, one was 2 × 105 HPBMC/well, one was 5 × 104 and the other group was 1 × 104 HPBMC/well. The detailed flow chart of this experiment is shown in Figs. 3 and 4. 1. Mix six different capture antibodies (anti-human IL-2 capture antibody/anti-human TGF-β capture antibody/anti-human IFN-γ capture antibody/anti-human IL-10 capture antibody/anti- Human TNF-α capture antibody/anti-human IL-6 capture antibody) and dilute the concentration of each capture antibody to 10 μg/mL in sterile PBS, and coat the mixture on a 96-well plate by incubating overnight at 4 °C. 2. Decant capture antibody solution, wash off unbound antibodies with 200 μL sterile PBS 3 times. Block membrane with 200 μL cell medium (RPMI-1640, 10% fetal bovine serum, 1% nonessential amino acids, penicillin, streptomycin, and glutamine) for at least 2 h at 37 °C. 3. Decant blocking medium. Add 100 μL PBMCs into each well at a final concentration of 1x105, 5x105, and 2x106 cells/mL and incubate overnight at 37 °C, 5% CO2.

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Fig. 3 Simultaneous detection of four cell-secreted antigens by FOLISPOT with a single-stage amplification system. Step (1): coating captures antibody. Step (2): incubating cells with a stimulant after blocking. The antigens secreted by the activated cells will be captured by the coated capture antibody, and a circle full of captured antigens will form only around the activated cells. Step (3): DNA-barcoded antibodies were added to bind with antigens secreted by stimulated cells. Step (4): addition of DNA amplification strands. Step (5): addition of DNA detection strands to bind with amplification strands. The detection strands will form a spot at the site of activated cells, and the images with fluorescence spots can be captured by the machine to perform the first-round recording. Step (6): washing away the first-round detection strands and addition of the secondround detection strands, followed by second-round imaging and recording. After the measurement, the images were analyzed and the spot numbers were counted by software

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Fig. 4 Schematic diagram of detection of 6 cell-secreted antigens (3 detection rounds) by FOLISPOT with dualstage amplification systems. Step (1): the capture antibodies are coated onto a plate; step (2) the plate is blocked with culture medium, and then cells are added to the plate and incubated with stimulants; step

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4. Decant cells and wash the plate six times with washing buffer. Mix six kinds of DNA-conjugated detection antibodies (antihuman IL-2 detection antibody/antihuman TGF-β detection antibody/anti-human IFN-γ detection antibody/anti-human IL-10 detection antibody/ anti-human TNF-α detection antibody/anti-human IL-6 detection antibody) and dilute the concentration of each DNA-conjugated detection antibody to 2 μg/mL with blocking buffer, followed by adding the mixture into the corresponding wells and incubating at room temperature for 2 h. 5. Wash three times with washing buffer, and then wash once with hybridization washing buffer. Mix six kinds of primary DNA concatemers (C.7, C.2, C.3, C.4, C.5, and C.6) and dilute the concentration of each primary DNA concatemers to 150 nM with hybridization buffer, followed by adding 100 μL per well to the plate and incubating at 37 °C overnight. 6. Wash three times with hybridization washing buffer, and then mix and add 100 μL per well 6 kinds of secondary DNA concatemers (C.14, C.9, C.10, C.11, C.12, and C.13 at 150 nM) to the plate, followed by incubating at 37 °C for 5 h (note: the primary signal amplification group is not processed). 7. Wash three times with hybridization washing buffer, then wash with fluorescent hybridization buffer once, followed by adding the first round DNA detection strands i.14* and i.9* (1 μM) and reacting at room temperature for 1 h. 8. Decant unbound free DNA detection strands, and then capture the images by utilizing the detection machines, such as Cytation 5 Cell Imaging Multi-Mode Reader (BioTek Instruments, INC.). 9. Remove the fluorescent hybridization buffer, add the dissociation buffer, and react at room temperature for 20 min. 10. Wash two times with PBS buffer, then wash two times with fluorescent hybridization buffer, add the second round of DNA detection strands i.12* and i.13* (1 μM), and react at room temperature for 1 h. ä Fig. 4 (continued) (3) after incubation, cells are removed and the plate is washed, followed by adding DNA-barcoded detection antibodies (see Note 4); step (4) primary amplification strands are added; step (5) secondary amplification strands are added; step (6) detection strands are added and the signals are measured for the first round; step (7) the first round detection strands are washes away and the second round detection strands are added, followed by the second round signal measurement; step (8) the second round detection strands are washes away and the third round detection strands are added, followed by the third round signal measurement [10]

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11. Decant unbound free DNA detection strands, and then capture the images by utilizing the detection machines, such as Cytation 5 Cell Imaging Multi-Mode Reader (BioTek Instruments, INC.). 12. Remove the fluorescent hybridization buffer, add the dissociation buffer, and react at room temperature for 20 min. 13. Wash two times with PBS buffer, then wash two times with fluorescent hybridization buffer, add the third round of DNA detection strands i.10* and i.11* (1 μM), and react at room temperature for 1 h. 14. Decant unbound free DNA detector, and then capture the images by utilizing the detection machines, such as Cytation 5 Cell Imaging Multi-Mode Reader (BioTek Instruments, INC.). 3.10 Detection of Antigens by Traditional ELISPOT

1. Prepare the PVDF membrane 96-well ELISPOT plates (Millipore) by soaking them in 35% ethanol for 1 min. 2. Wash thoroughly with PBS to remove any residual ethanol. 3. Add capture antibody (10 μg/mL in PBS) to the prepared ELISPOT plate and incubate overnight at 4 °C. 4. Wash the plate three times with 200 μL/well of sterile PBS, and then add 200 μL/well of RPMI 1640 medium (10% FBS), followed by incubating for 2 h at 37 °C in the incubator. 5. Remove the RPMI 1640 medium (10% FBS) and add 100 μL PBMCs in RPMI 1640 medium (10% FBS), together with 20 μM PHA and LPS, to different wells in ELISPOT plate, followed by incubating overnight at 37 °C with 5% CO2. 6. Wash the plate six times with 200 μL/well of sterile 0.01% PBST and add 100 μL biotinylated detection antibody in PBS with 0.5%BSA (2 μg/mL) into the plate, followed by incubating at 37 °C for 2 h. 7. Wash plate six times with 200 μl/well of 0.01% PBST and add 100 μL Streptavidin-ALP into each well followed by incubating at room temperature for 45 min. 8. Wash the plate three times with 200 μL/well of PBST and then wash the plate three times with PBS, followed by adding 100 μL BCIP/NBT plus and incubating at dark for 10 min. 9. Remove BCIP/NBT plus and wash the plate with water, followed by drying the plate in the dark. 10. Capture the images and count the spot number by detection machines, such as the CTS reader system.

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3.11 Detection of Four Targets by FOLISPOT with a Single-Stage Amplification System

In the FOLISPOT detection with a single-stage amplification system, the DNA-conjugated detection antibodies can bind with amplification concatemers, followed by fluorescent signal detection. Two detection rounds were conducted, and, in each detecting round, two different detection dyes were utilized. After the measurement of fluorescent intensity in each round, detection strands were washed away and detection strands for the next round were added (Figs. 5 and 6) for next round detection. The switch of different round detections was achieved by detection strand exchange. In this FOLISPOT detection, four different kinds of cells secreted cytokine antigens, IL-10, TGF-β1, IFN-γ, and TNF-α, were analyzed simultaneously and only the primary amplification system was applied. In the first detection round, cells secreting TGF-β or IL-10 were detectable, while in the second round, cells secreting TNF-α or IFN-γ were detectable. Meanwhile, cells simultaneously secreting TGF-β and IL-10 were measured and cells simultaneously secreting TNF-α and IFN-γ were detectable. Both in wells with a low number of cells (2 × 105 peripheral blood mononuclear cells, PBMCs) and with a high number of cells (4 × 105 PBMCs), clear and strong signals were detectable.

3.12 Detection of Six Targets with FOLISPOT

To further elevate the detection sensitivity of FOLISPOT, cells secreting 6 cytokines (IL-10, TGF-β1, IFN-γ, TNF-α, IL-2, and IL-6) were detected simultaneously by FOLISPOT with a dualstage signal amplification system (Fig. 7). Compared FOLISPOT with a single-stage amplification system (Fig. 7), FOLISPOT with a dual-stage amplification system increased the detection sensitivity of every cytokine (IL-10, TGF-β1, IFN-γ, TNF-α, IL-2, or IL-6). Meanwhile, FOLISPOT with a dual-stage amplification system could detect around 135 cells (2 × 105 PBMCs/well) that are secreting both IL-10 and IL-6. These illustrated that FOLISPOT with a dual-stage amplification system could be applied to measure cells that are secreting multiple antigens and especially be applied to detect weakly activated cells secreting fewer proteins.

3.13 Comparison of FOLISPOT with Traditional ELISPOT

The detection sensitivity of FOLISPOT and ELISPOT was investigated by detecting the same cytokines secreted by activated T cells, and the spot number in the FOLISPOT assay is higher than the spot number in ELISPOT assay (Figs. 8 and 9). These data illustrated that FOLISPOT could be applied to detect signals that are not strong enough to be detected by ELISPOT.

3.14

Widely used traditional ELISPOT can only detect one single target at one time, therefore, FOLISPOT, which can simultaneously detect multiple targets in one sample, could improve the detection accuracy and capacity. FOLISPOT could reduce the repeated work, materials, and sample demanded for tests. The signal amplification system in FOLISPOT is controllable by adjusting DNA

Conclusion

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Fig. 5 Analysis of cytokine-secreted cells by FOLISPOT with single and dual-stage amplification systems. (a) Measurement of cytokine-secreted cells by FOLISPOT at the condition of 2 × 105 HPBMC/well. (b) Measurement of cytokine-secreted cells by FOLISPOT at the condition of 5 × 104 HPBMC/well [10]

Fig. 6 Measurement of cytokine-secreting cells by FOLISPOT with a single-stage amplification system. (a) Image of an example well measured by FOLISPOT with a single-stage amplification system. (b) Results of measurement of cytokine-secreting cells by FOLISPOT with a single-stage amplification system. (c) Number of cytokine-secreting cells measured by FOLISPOT under different cell densities. (d) In the case of 2 × 105

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concatemers. By adjusting the signal amplification system, we potentially could detect some proteins that we mistakenly thought were not secreted. According to previous studies [10], FOLISPOT can simultaneously detect at least six targets in one text. In addition, a better detection sensitivity than traditional ELISPOT in detecting cytokines was observed when utilizing FOLISPOT. According to the above description, FOLISPOT is applicable under special situations. For instance, when the amount of cytokines that target cells secrete is too little and traditional ELISPOT is unable to detect such a small amount of cytokine, FOLISPOT can be used to detect cells only secreting small amounts of cytokines thanks to FOLISPOT is more sensitive. In addition, when scientists need to identify the number of target cells that are simultaneously secreting several different kinds of cytokines, FOLISPOT can be utilized. Such as, when scientists explore more details of some antigenspecific T cells in type 1 diabetes or cancer secreting different cytokines upon stimulation by specific antigens, FOLISPOT could be very helpful.

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Notes 1. Washing buffer applied in each detection round. The concentration of formamide (deionized) in the washing buffer can be altered according to the DNA strands that need to be washed off in every washing round before taking images. 2. Washing time in each detection round The incubation time of the washing buffer can be altered according to the DNA strands that need to be washed off in every washing round before taking images. 3. Length of DNA strands applied to connect different strands The length of DNA strands for connecting different DNA strands needs to be altered according to the need for binding strength. If they need to bind tightly and need to be washed off in later rounds, the length can be longer; if they need to bind loosely and need to be washed off in easier rounds, the length can be shorter.

ä Fig. 6 (continued) HPBMCs/well, the number of cells that secrete TNF-α, the number of cells that secrete IFNγ, and the number of cells that simultaneously secrete both TNF-α and IFN-γ detected by FOLIPSOT. (e) In the case of 4 × 105 HPBMCs/well, the number of cells that secrete TGF-β, the number of cells that secrete IL-10, and the number of cells that simultaneously secrete both TGF-β and IL-10 detected by FOLIPSOT [10]

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Fig. 7 Analysis of cytokine-secreted cells by FOLISPOT with single and dual-stage amplification systems. (a) Measurement of cytokine-secreted cells by FOLISPOT at the condition of 1 × 104 HPBMC/well. (b) Measurement of cytokine-secreted cells by FOLISPOT at the control condition of no stimulant [10]

Fig. 8 Measurement of cytokine-secreting cells by FOLISPOT with a single- or dual-stage amplification system. (a) Comparison of measurement of TGF-β-secreting cells by traditional ELISPOT, FOLISPOT with a single-stage amplification system, and FOLISPOT with a dual-stage amplification system. (b) Comparison of measurement of TNF-α-secreting cells by traditional ELISPOT, FOLISPOT with a single-stage amplification system, or FOLISPOT with a dual-stage amplification system. (c) Comparison of measurement of cytokinesecreting cells by FOLISPOT with a single- or dual-stage amplification system at different cell densities. (d) Results of measurement of TGF-β-secreting cells by traditional ELISPOT. (e) Results of measurement of TGF-β-secreting and IL-2-secreting cells by FOLISPOT with a single- or dual-stage amplification system in the case of 2 × 105 HPBMCs/well. (f) Results of measurement of TGF-β-secreting and IL-2-secreting cells by FOLISPOT with a single- or dual-stage amplification system in the case of 5 × 104 HPBMCs/well. (g) Results of measurement of TGF-β-secreting and IL-2-secreting cells by FOLISPOT with a single- or dual-stage amplification system in the case of 1 × 104 HPBMCs/well [10]

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4. Incubation of detection DNA strands for next round detection The incubation time of detection DNA strands after washing off the last round of detection DNA strands needs to be altered according to the need for binding strength and the length of DNA strands. References 1. Udomkarnjananun S, Kerr SJ, Francke MI et al (2021) A systematic review and meta-analysis of enzyme-linked immunosorbent spot (ELISPOT) assay for BK polyomavirus immune response monitoring after kidney transplantation. J Clin Virol 140:104848 2. Janetzki S (2015) Immune monitoring technology primer: the enzyme-linked immunospot (Elispot) and Fluorospot assay. J Immunother Cancer 3:30 3. Blume J, Kostler J, Weissert R (2015) Benefit of ELISpot in early diagnosis of tuberculous meningoencephalitis: Case report and literature review. eNeurologicalSci 1(3–4):51–53 4. Okamoto Y, Abe T, Niwa T et al (1998) Development of a dual color enzyme-linked immunospot assay for simultaneous detection of murine T helper type 1- and T helper type 2-cells. Immunopharmacology 39(2):107–116 5. Axelsson B (2022) Detection and enumeration of cytokine-secreting cells by FluoroSpot. Methods Mol Biol 2386:81–99 6. Bisceglia H, Barrier J, Ruiz J et al (2023) A FluoroSpot B assay for the detection of IgA and IgG SARS-CoV-2 spike-specific memory B cells: optimization and qualification for use in COVID-19 vaccine trials. J Immunol Methods 515:113457

7. Danielzik T, Koldehoff M, Buttkereit U et al (2018) Sensitive detection of rare antigenspecific T cells directed against Wilms’ tumor 1 by FluoroSpot assay. Leuk Lymphoma 59(2): 490–492 8. Dillenbeck T, Gelius E, Fohlstedt J et al (2014) Triple cytokine FluoroSpot analysis of human antigen-specific IFN-gamma, IL-17A and IL-22 responses. Cell 3(4):1116–1130 9. Luque S, Lucia M, Crespo E et al (2018) A multicolour HLA-specific B-cell FluoroSpot assay to functionally track circulating HLA-specific memory B cells. J Immunol Methods 462:23–33 10. Ma J, Peng Z, Ma L et al (2022) A multipletarget simultaneous detection method for immunosorbent assay and immunospot assay. Anal Chem 94(24):8704–8714 11. Nilsson A, Hobinger A, Jahnmatz P et al (2022) Four-parameter FluoroSpot assay reveals that the varicella zoster virus elicits a robust memory T cell IL-10 response throughout childhood. J Virol 96(22):e0131022 12. Udomkarnjananun S, Kerr SJ, Townamchai N et al (2021) Donor-specific ELISPOT assay for predicting acute rejection and allograft function after kidney transplantation: a systematic review and meta-analysis. Clin Biochem 94:1– 11

Chapter 17 Triple-Color FluoroSpot Analysis of Polyfunctional Antigen-Specific T Cells by Quantification of Spot-Forming Units and Relative Spot Volumes Niklas Ahlborg, Christian Smedman, and Bartek Makower Abstract Switching from ELISpot to FluoroSpot enables the analysis of spot-forming units representing cells producing different cytokines as well as the frequencies of spots derived from cells co-secreting multiple cytokines. Due to the fluorescent read-out signal, sophisticated reader instruments can also measure the relative spot volume, making it possible to differentiate between spots generated by cells secreting different levels of one or more cytokines. Here we describe how triple FluoroSpot assays can be used to define polyfunctional T cells secreting multiple cytokines and how different T-cell populations can differ in the levels of cytokines they secrete. Key words Cytokine, ELISpot, FluoroSpot, Polyfunctional T cells

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Introduction ELISpot is a sensitive method for the detection of cytokine secretion by antigen-specific T cells, but the method is limited to the analysis of a single cytokine at a time. Based on the same principle as ELISpot but utilizing several capture and detection antibodies with different fluorophores, FluoroSpot enables the detection of multiple cytokines simultaneously. Detection of different cytokines with specific fluorophores also makes it possible to determine if spots are derived from cells secreting single or multiple cytokines, thus facilitating the quantification of polyfunctional T cells. Polyfunctional antigen-specific T cells, secreting multiple cytokines, are of relevance for the control of various pathogens and have higher protective efficacy after vaccination (1–4). Another factor that influences the functionality of T cells is the level of cytokine they produce. Several studies have indicated that polyfunctional T cells are more efficient in terms of protective capacity because of their ability to secrete higher levels of cytokines, compared to T cells

Alexander E. Kalyuzhny (ed.), Handbook of ELISPOT: Methods and Protocols, Methods in Molecular Biology, vol. 2768, https://doi.org/10.1007/978-1-0716-3690-9_17, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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Fig. 1 Example of FluoroSpot analysis of human polyfunctional T cells using spot-forming units (SFU) and relative spot volume (RSV) as analytical parameters. Human peripheral mononuclear cells (PBMCs) from patients with suspected latent tuberculosis (n = 13) were stimulated with peptide pools including T-cell epitopes from Epstein–Barr virus (EBV) and Cytomegalovirus (CMV). FluoroSpot was used to detect spots derived from antigen-specific T cells secreting IFN-ɣ, IL-2, and/or TNF-α. The graphs display IFN-ɣ secretion derived from cells secreting only IFN-ɣ, IFN-ɣ, and IL-2, IFN-ɣ, and TNF-α or all three cytokines. Background spot counts in wells with unstimulated PBMCs were low and subtracted. A) IFN-ɣ SFU were the highest for cells secreting only IFN-ɣ or IFN-ɣ and TNF-α. B) In contrast, when measuring RSV for IFN-ɣ, IFN-ɣ RSV was the highest for cells secreting IFN-ɣ and IL-2 or all three cytokines. Note that the (a) graph has a logarithmic Y-axis and (b) a linear Y-axis

displaying a more limited cytokine repertoire (4–7). To assess the involvement of T cells in protective immunity, it is thus important to not only if T cells are able to produce multiple cytokines but also the magnitude of production. Notably, a recent study used triple color FluoroSpot and showed that antigen-specific T cells secreting a single cytokine can be more prevalent than polyfunctional T cells, but that the polyfunctional T cells secrete higher levels of certain cytokines (7). A similar example of how the measurement of SFU and RSV can yield contrasting results is shown in Fig. 1. In addition to defining cytokine co-secretion, analysis of FluoroSpot assays with advanced readers makes it possible to define the relative spot volume (RSV) that reflects the level of cytokine (s) secreted by single cells. To achieve such an analysis in a reliable manner, the FluoroSpot reader must first define the spot center accurately. This is a prerequisite for defining if spots of different colors, representing different cytokines, are derived from a single cell or from two cells in close proximity to each other. Second, to facilitate a reliable measure of the RSV, the reader should define the volume of the spot using three-dimensional raw spot data, i.e., it should define the spot volume by signal processing rather than analyzing a compressed camera image.

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This is achieved by using a totally novel approach to spot recognition, whereby RAWspot™ technology encapsulates the true physics of spot-formation into a mathematical model, predicting each responding cell localization and their level of secretion based on how well the model matches the final outcome as observed in the uncompressed camera image. Since spots take on distinctly different diffusion patterns based on the rate of cytokine release during the actual assay, the mathematical model in RAWspot™ can elegantly employ a temporal dimension to easily understand even the most complicated clusters of spot formation. In essence, the high detection sensitivity of the FluoroSpot assay, facilitating detection of cytokine-secreting antigen-specific T cells, combined with the method’s capacity of defining cytokine co-expression and the relative levels of cytokines secreted, makes FluoroSpot an ideal assay for studies of polyfunctional T cells.

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Material 1. Transparent 96-well plates with low autofluorescence polyvinylidene difluoride (PVDF) membranes (IPFL; MerckMillipore, Cork, Ireland). 2. Blocking buffer to be used after coating: 10% fetal calf serum (HyClone Laboratories Inc., Logan, UT, USA) in phosphatebuffered saline (PBS), pH 7.4. 3. Human PBMCs suspended in culture medium; RPMI 1640 supplemented with glutamine (2 mM), HEPES (10 mM), and penicillin/streptomycin (diluted 1:100; all from Life Technologies Ltd., Paisley, UK) and 10% FCS (HyClone). 4. Monoclonal antibodies (mAbs) for coating: anti-human IFN-ɣ mAb 1D1K, anti-human IL-2 mAb MT2A91/2C95, and antihuman TNF-α mAb MT25C3 (Mabtech, Nacka Strand, Sweden). 5. MAbs for the primary detection step: anti-human IFN-ɣ mAb 7-B6–1 labeled with peptide tag BAM, anti-human IL-2 mAb MT8G10 labeled with biotin, and anti-human-TNF-α mAb MT20D9 labeled with peptide tag WASP (Mabtech). 6. MAbs for the secondary detection step: anti-BAM mAb labeled with 490 nm fluorophore, streptavidin (SA) labeled with 550 nm fluorophore, and anti-WASP mAbs labeled with 640 nm fluorophore (Mabtech). 7. MAbs for cellular activation: anti-CD3 mAb CD3–2 and antihuman CD28 mAb CD28-A (Mabtech). 8. Peptide pools for activation of T cells. Two peptide pools based on selected peptide epitopes for CD4+ and CD8+ T cells from EBV and CMV proteins (Mabtech).

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9. Fluorescence enhancer used to strengthen the fluorescent signal (Mabtech). 10. Analysis of spots is performed with the ELISpot/FluoroSpot reader Mabtech IRIS (Mabtech) equipped with automated calibration, APEX™ software, LED lights, and narrow band filters for fluorophores with excitation/emission at 380/430, 490/510, 550/570, and 640/660 nm.

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Methods A triple-color FluoroSpot assay is used to detect cytokine secretion by antigen-specific T cells. FluoroSpot plates are coated with capture mAbs specific for human IFN-ɣ, IL-2, and TNF-α. Human PBMCs with or without stimulus are added and incubated, and cytokines secreted by activated T cells are captured by the respective capture mAbs. The cells are removed and captured cytokine is detected in two steps. In the first step, detection mAbs specific for the three cytokines are used. The detection mAbs are labeled with different tags. In the second step, three anti-tag detection reagents labeled with different fluorophores are used. Spots of three different colors are identified using a FluoroSpot reader. The spots are footprints of cells that have secreted one or several of the cytokines. The reader analysis will quantify the number of spots for each color, identify co-localized spots derived from co-expressing cells, and will define the relative amount of cytokines secreted by different cells.

3.1 Triple-Color FluoroSpot

EtOH Treatment and Coating of the Plates (Sterile Conditions) 1. Pre-wet the membrane of a FluoroSpot plate with 15 μL/well of 35% EtOH for a maximum of 1 min. Wash the plate with 200 μL of sterile water five times. Make sure that the membrane does not dry after the treatment (see Note 1). 2. Mix the three coating mAbs for IFN-ɣ, IL-2, and TNF-α in sterile PBS to a final concentration of 15 μg/mL of each mAb. Add 100 μL of the mixed mAb solution to each well and incubate overnight at +4–8 C (see Note 1). 3. Empty the plate and wash it five times with sterile PBS. Add 200 μL/well cell culture medium with 10% FCS to condition/ block the membranes for 30 min at room temperature (see Note 2). Remove the blocking solution just prior to adding the cell suspension. Incubation of PBMCs in the Plates (Sterile Conditions) 4. PBMCs are incubated in a cell culture medium only, with stimuli (in this case a mix of two peptide pools from EBV and CMV) or with a positive control (anti-CD3), preferably in

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triplicates for each condition. To compensate for the capture of IL-2, co-stimulatory anti-CD28 mAb is added to all wells. Either mix all components before adding them to the wells or suspend the components separately and add them separately to the wells. If components are added sequentially, always add the PBMCs last. The final volume should be 100–150 μL/well (see Notes 3–5). 5. Incubate the plate at +37 °C in a humified incubator with 5% CO2. Avoid evaporation by wrapping the plate in aluminum foil and do not move the plate during incubation. Incubate the cells for 18–24 h (see Note 6). Detection of Spots 6. Discard the cells by emptying the plate and wash them five times with PBS (see Note 7). 7. In the same vial, dilute the three primary detection mAbs for IFN-ɣ, IL-2, and TNF-α to their respective working concentrations according to the manufacturer’s instruction, in PBS containing 0.1% BSA. Add 100 μL/well and incubate the plate for 2 h at room temperature. 8. Wash the plate five times with PBS. 9. In the same vial, dilute the three tag-specific secondary detection reagents to their respective working concentrations in accordance with the manufacturer’s instructions, in PBS containing 0.1% BSA. Add 100 μL/well and incubate the plate for 2 h at room temperature. 10. Wash the plate five times with PBS. Empty the plate, add fluorescence enhancer at 50 μL/well, and leave the plate for 10 min at room temperature. Remove the fluorescence enhancer by decanting and flicking the plate. Do not wash. Avoid tapping the plate against tissue paper since dust particles can interfere with the analysis. Leave the plate to dry. To speed up the drying, remove the soft plastic underdrain on the backside of the plate and dry in a laminar hood. Plates can be stored dark at room temperature. 3.2

Analysis of Spots

1. The plate should be completely dry before analysis. The reader has to be equipped with filters adapted for the fluorophores used (as described in Subheading 2). 2. Read the plate using Mabtech default settings for each filter. The brightness and contrast of the image can be adjusted afterward without affecting SFU or RSV data. The analysis is based on uncompressed HDR images and analyzed by the RAWspot™ algorithm using a mathematical model for determining spot centers and the volume of spots (see Note 8).

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3. Depending on background levels in unstimulated controls, make adjustments to intensity and size. Counted spots are updated immediately in all wells without the need for a re-count.

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Notes 1. EtOH treatment of plates. It is essential to not let the wells dry out after the EtOH treatment. Should this happen, it is necessary to repeat the EtOH treatment. For the same reason, after EtOH treatment, keep the water from the last wash in the wells until the mAb solution has been prepared and the coating procedure is ready to be initiated. More detailed information about the EtOH treatment is presented in a video available at: https://www.mabtech.com/knowledge-hub/step-step-guideelispot. To avoid the necessity of EtOH treat and coat plates, an option is to use a human IFN-ɣ/IL-2/TNF-α FluoroSpot kit from Mabtech that contains precoated plates and all necessary reagents. 2. Blocking step. The cell culture medium used for the cells should be used in the blocking step. As an alternative to using a cell culture medium supplemented with FCS, it is possible to use a serum-free medium such as AIM-V for the cell culture. If so, the serum-free medium should be used also for the blocking step. 3. Cell culture medium. If a cell culture medium supplemented with FCS is used, it is important to evaluate the FCS batch in pilot tests. Ideally, an FCS batch that yields low background in wells with unstimulated cells and strong results in wells with antigen-stimulated cells or positive controls should be used. 4. Cell stimulation and incubation. Anti-CD28 mAb (final concentration 1 ug/mL) is added to the PBMCs since the capture of IL-2, by the IL-2 capture antibodies, can have a negative impact on the production of other cytokines. Anti-CD28 mAb provides a co-stimulatory signal to antigen-specific T cells that compensates for the depleted IL-2 but does not activate T cells by itself. Anti-CD3 mAb (a final concentration of 100 ng/mL) is included as a positive control and will activate both CD4+ and CD8+ T cells provided the PBMCs have good viability. If frozen PBMCs are used, the viability should be >85% after thawing for good results. Add PBMCs premixed with stimulus or if components are added separately, add PBMCs last. If PBMCs are dispensed first and stimulus after, cells may be pushed to the edges of the wells.

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5. Peptide pools. The EBV peptide pool consists of 100 peptides derived from 22 EBV proteins. 66 peptides are MHC class I-restricted epitopes, and 34 peptides are MHC class II-restricted epitopes. The CMV peptide pool consists of 42 peptides derived from CMV proteins. Twenty-eight peptides are MHC class I-restricted epitopes and 14 are MHC class II-restricted epitopes. The lyophilized peptide pools are dissolved in 40 μL DMSO followed by the addition of 85 μL PBS whereafter they are aliquoted and stored at -20 °C or used directly. Both CMV and EBV are persistent viruses with a high prevalence in the population. The peptide pools cover a wide variety of HLA types, and most individuals will respond to HLA-matched peptides from one or both peptide pools. 6. Cell incubation time. The production of different cytokines by antigen-stimulated T cells displays different kinetics. IFN-ɣ, IL-2, and TNF-α can be measured in FluoroSpot after an overnight incubation whereas some other cytokines (e.g., IL-4, IL-5, IL-21, and IL-22) may need night 2 days incubation. When combining different cytokines in a triple FluoroSpot assay it is important to consider the kinetics of all three cytokines. It is important to use a defined incubation time in a series of experiments since the RSV is based on the detection of accumulated cytokine. Hence, longer incubation times may increase the RSV. 7. Non-sterile washes. An ELISA washer can be used provided the washer head has been adapted for ELISpot/FluoroSpot plates. Do not include Tween or other detergents in the buffer. 8. RSV can only be compared between spots of the same color i.e., IFN-ɣ RSV cannot be compared with IL-2 RSV since different detection systems and fluorophores may differ in sensitivity.

Acknowledgments The authors thank Tyler Sandberg for the critical reading of the manuscript. References 1. Sun Y, Santra S, Schmitz JE et al (2008) Magnitude and quality of vaccine-elicited T cell responses in the control of immunodeficiency virus replication in Rhesus monkeys. J Virol 82: 8812–8819 2. Betts MR, Nason MC, West SM et al (2006) HIV nonprogressors preferentially maintain highly functional HIV-specific CD8+ T cells. Blood 107:4781–4789

3. Almeida JR, Price DA, Papagno L et al (2007) Superior control of HIV-1 replication by CD8+ T cells is reflected by their avidity, polyfunctionality, and clonal turnover. J Exp Med 204:2473– 2485 4. Darrah PA, Patel DT, De Luca PM et al (2007) Multifunctional TH1 cells define a correlate of vaccine-mediated protection against Leishmania

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major. Nat Med 3(7):843–850. https://doi. org/10.1038/nm1592 5. Kannanganat S, Ibegbu C, Chennareddi L et al (2007) Multiple-cytokine-producing antiviral CD4 T cells are functionally superior to singlecytokine-producing cells. J Virol 81:8468–8476 6. Precopio ML, Betts MR, Parrino J et al (2007) Immunization with vaccinia virus induces

polyfunctional and phenotypically distinctive CD8+ T cell responses. J Exp Med 204:1405– 1416 7. Sandberg JT, Varnaite˙ R, Christ W et al (2021) SARS-CoV-2-specific humoral and cellular immunity persists through 9 months irrespective of COVID-19 severity at hospitalization. Clin Transl Immunol 10(7):e1306. https://doi. org/10.1002/cti2.1306

Chapter 18 Performance and Stability of New Class of Fetal Bovine Sera (FBS) and Its Lyophilized Form in ELISpot and FluoroSpot Assays: Applications for Monitoring the Immune Response in Vaccine, and Cell and Gene Immunotherapy in Clinical Trials Zhinous Hosseini, Christopher J. Groves, Penny Anders, Kristen Cave, Madelyn Krunkosky, Brandi Chappell, Sofie Pattyn, Devin Davis, Sylvia Janetzki, and Elizabeth Reap Abstract Interferon-gamma (IFNγ) ELISpot and FluoroSpot are widely used assays to detect functional cell responses in immunotherapy clinical studies. Recognized for their importance in vaccine development studies to quantitate immune responses, these assays have more recently risen to the forefront in cell and gene therapy as well as cancer immunotherapy fields where responses against cancer neoantigens are not easily detectable above assay background. Here, we test a new class of fetal bovine serum (FBS), CultraPure FBS, in ex vivo ELISpot and FluoroSpot assays and cultured FluoroSpot assays following in vitro expansion. Several CultraPure FBS lots that have been specially formulated through the process of lyophilization (lyo-FBS) were compared to liquid CultraPure FBS. We stimulated human PBMCs with antigen-specific peptide pools diluted in media supplemented with liquid CultraPure FBS or lyo-FBS and found equivalent cytokine production with negligible to no assay background with both liquid and lyo-FBS formats. Moreover, the lyo-FBS showed lot-to-lot consistency and 90-day refrigerated (4 °C) stability in both ex vivo direct and in vitro cultured assays. In addition, we present here a method using lyo-FBS for the expansion of low-frequency antigen-specific T cells, mimicking the low frequency seen with cancer neoantigens by utilizing a cultured FluoroSpot assay. Our results demonstrate the presence of Granzyme B, interferon-gamma (IFNγ), and tumor necrosis factor (TNF) production by antigen-specific polyfunctional T cells following a 9-day culture using media supplemented with lyo-FBS. Key words ELISpot, FluoroSpot, Serum, lyophilized FBS, Cellular Immune Responses, Immune monitoring, HCMV pp65, Cell and gene therapy, vaccines, Cancer immunotherapy

Alexander E. Kalyuzhny (ed.), Handbook of ELISPOT: Methods and Protocols, Methods in Molecular Biology, vol. 2768, https://doi.org/10.1007/978-1-0716-3690-9_18, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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Introduction ELISpot and FluoroSpot are important assays used in both research and clinical studies for the evaluation of single-cell functional immune responses, especially secretion of IFNγ [1, 2]. Determining the functionality of rare or low-frequency cells in a clinical sample is a highly useful endpoint for vaccine studies [3], cell and gene therapy development [4], and detecting neoantigen responses in cancers [5]. These assays have become a mainstay in both pharmaceutical research and clinical studies and require high sensitivity where positive responses over background are an important factor in resolving actual responses from noise. Therefore, keeping background reactivity levels of tested samples to a minimum in ELISpot and FluoroSpot assays is a necessity and a challenge for rare cell detection [6]. Although ELISpots and FluoroSpots are powerful research techniques, these assays have historically been plagued with high assay background and variability, despite major improvements in antibody specificity and plate reading technology [8, 9]. To employ these assays successfully in clinical trials, extensive and timeconsuming optimizations must occur [5, 6]. A major contributor to background and variability in ELISpot and FluoroSpot assays is conventional FBS, which is a necessary ingredient in most cell culture media [7]. Conventional FBS is used in liquid form, while it is stored and shipped frozen. It is often derived from various collection sites and under varying harvest and storage conditions prior to pooling and bottling, resulting in lot-to-lot inconsistency [10]. Therefore, each new lot of FBS used in ELISpot or FluoroSpot assays must be evaluated for low background while maintaining a nutrient-rich environment supportive of functional cellular responses. Identifying a particular vendor and a specific lot of FBS with favorable characteristics is essential to providing high-quality reproducible results with these assays [10, 11]. After an FBS vendor/lot is qualified, the specific lot is typically purchased in bulk and must be stored under temperature-controlled conditions for the duration of the study, which can last a year or more. The challenges related to this process for monitoring clinical studies include lot qualification, lot expiration, storage issues, and the possibility of running out of a specific lot of FBS. Therefore, having a reliably stable source of FBS that results in low background, supporting cellular responses, minimal lot-to-lot variability, ambient temperature long-term storage, and ease of use would be a considerable improvement to the immune monitoring field. Recently, Berrong et al. [11] tested and validated a new class of FBS (liquid CultraPure FBS) for use in ELISpot assays that overcome the historical challenges of using traditional FBS. This new

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class of FBS has been pre-validated for improved performance, enhanced lot-to-lot consistency, exhibits low background, and is supportive of a robust cellular response in ELISpot assays [11]. Liquid CultraPure FBS is also available in a lyophilized format, removing nearly 95% of the water content in a proprietary process that preserves more of the native growth factors present in serum while eliminating the hassles of storage at freezing temperatures during its manufacture, delivery, and storage. Traditional liquid-FBS may undergo up to four freeze-thaw cycles prior to its use. These drastic temperature changes are widely known to reduce the performance of liquid-FBS. Lyophilized CultraPure FBS eliminates the freezethaw cycles and cold chain constraints, significantly lowers the total cost of shipping, and dramatically cuts the use of plastics leading to a reduced carbon footprint and improved social and environmental impact. This recently developed format of FBS also represents a striking improvement in convenience and ease of use for laboratory bench scientists. We wondered whether these findings would extend to direct ELISpot and FluoroSpot assays and cultured FluoroSpot assays, where high background is often seen hindering the detection of potentially important new tumor and vaccine targets. We hypothesized that lyophilized CultraPure FBS (lyo-FBS) would have similar performance characteristics to liquid CultraPure FBS in the ELISpot and FluoroSpot assays, To determine the functional cellular responses supported by lyo-FBS media, we performed a cultured FluoroSpot assay from a pre-screened donor whose PBMCs contain only few HCMV pp65 antigen-specific cells. These cells were cultured and expanded over 9 days with peptide pools and cytokines, which markedly improved the detection of rare cells with low background, suggesting equivalent performance characteristics of this new class of lyo-FBS to liquid CultraPure FBS.

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Materials Human IFN-γ FluoroSpot kit (MabTech) Sterile PBS, calcium, and magnesium free Sterile RPMI Medium 1640 1 M HEPES MEM NEAA Penicillin–streptomycin solution 100X Lyophilized CultraPure FBS (Lyo-FBS) (Seraworx) CultraPure FBS (Seraworx) CEFTA control peptide pool – see Note 1 for peptide spec info.

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HCMV pp65 peptide pool – see Note 2 for peptide spec info. Pepmix actin peptide pool – see Note 3 for peptide spec info. Phytohemagglutinin-L (PHA-L) (Invitrogen) Trypan Blue 0.4%, 0.85% NaCl Cryopreserved PBMCs (≥ to 90% viability) isolated from healthy human donors and prepared from purchased leukopaks after overnight (o/n) ship. Complete media: RPMI 1640, 10% FBS, 1X penicillin–streptomycin solution, 25 mM HEPES, 1X NEAA Laminar flow hood Cell culture incubator set to 5% CO2 and 37 °C Mabtech IRIS FluoroSpot/ELISpot plate reader BioRad TC20-automated cell counter or equivalent

3

Methods

3.1 For Cultured ELISpot or FluoroSpot Continue with These Procedures. If Not Performing a Cultured ELISpot or FluoroSpot Proceed to Subheading 3.2

1. Prepare the lyo-FBS by creating a 4.50% stock solution with purified or deionized water. See Notes 3 and 4. 2. Prepare sterile filtered complete media and pre-warm in 37 °C water bath. 3. Thaw cryopreserved PBMCs, wash 1× with complete media, and count cells. See Note 5 for thawing instructions. 4. Resuspend cells in complete media to obtain a concentration of viable 4 × 106 cells/mL. 5. Add IL-7 to the cell suspension for a final concentration of 5 ng/mL. 6. Add 1 mL of cell suspension, 4 × 106 cells/well, to each well of a 24-well plate in triplicate. 7. Add peptide pool positive (CEFTA) and negative (actin) controls as well as assay-specific peptide pool (HCMV pp65) to designated wells per plate map so that the final concentration is 2 μg/mL for each pool. 8. Incubate at 5% CO2 and 37 °C o/n. 9. The next day, carefully remove 0.5 mL media and replace it with new 0.5 mL warm complete media containing IL-7 at a final concentration of 5 ng/mL. 10. Incubate at 5% CO2 and 37 °C for 72 h. 11. Carefully remove 0.5 mL of medium and replace it with new 0.5 mL warm complete media containing IL-7 (a final concentration of 5 ng/mL) and IL-2 (a final concentration of 4 ng/ mL).

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12. Incubate at 5% CO2 and 37 °C for 72 h. 13. Harvest the cells by pooling them from the same treatments in each well into a 50 mL conical tube. Rinse each well with 1 mL of complete media and add to the tube. 14. Centrifuge at 300× g for 10 min at room temperature. 15. Resuspend the pellet in 1 mL of warm complete media per culture well with a maximum of 5 mL per 50 mL conical tube. 16. Rest cells o/n in the incubator (5% CO2, 37 °C) with the caps slightly loosened for adequate ventilation. 17. For a direct, ex vivo assay control, thaw cryopreserved PBMCs and rest o/n followed by seeding onto FluoroSpot plate and peptide stimulation. 18. Proceed to Subheading 3.3. 3.2 For Direct (Ex Vivo) ELISpot/ FluoroSpot Assay, Thaw Cryopreserved PBMCs and Rest Overnight

1. Prepare the lyo-FBS by creating a 4.50% stock solution with purified or deionized water. See Notes 4 and 5. 2. Prepare a plate layout and a step-by-step protocol that includes all calculations and volumes of reagents. 3. Prepare complete media using each type of sterile-filtered serum to be evaluated followed by sterile filtering of the media after the addition of FBS and other supplements. Warm in 37 °C water bath. 4. Thaw cryopreserved PBMCs. See Note 6 for the protocol 5. Add PBMCs to a 50 mL conical tube at a concentration of approximately 10x106 in 5 mL of complete media specific for each condition. 6. Rest and incubate cells at 37 °C, 5% CO2 o/n. See Note 7.

3.3 Preparation of Precoated Plates and Seeding of Stimulated PBMC

1. Remove precoated plates from the kit and wash 3X with sterile PBS (200 μL per well). 2. Block plates by adding 200 μL per well of complete medium. Block each condition with the specific serum that has either CultraPure liquid FBS or lyo-FBS according to the plate layout and incubate for 2 h at room temperature. 3. Remove cells from the incubator. Centrifuge at 230× g for 10 min and decant supernatant. 4. Resuspend cells in appropriate media (serum-specific media) so that the concentration of viable cell suspension is 5 × 106 cells/ mL. 5. Prepare the peptide pools at 2X concentration, prepare PHA at 20 μg/mL, and HCMV pp65 at 4 μg/mL in appropriate complete media. The final concentrations will be 10 μg/mL for PHA and 2 μg/mL for HCMV pp65. See Notes 8 and 9.

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6. Next, add 50 μLs of PBMC cell suspension (in serum-specific media) to the wells according to plate layout in triplicate. 7. Transfer the plate to an incubator for o/n incubation. See Note 10. 8. Finally, follow the instructions provided in the Mabtech kit for signal development and detection.

4

Notes 1. CEFTA: The CEFTA peptide pool induces secretion of e.g., IFN-γ, IL-2, IL-5, and IL-13 from human CD4 T cells. It consists of 35 MHC class II-restricted peptides from human HCMV pp65, EBV, influenza virus, tetanus toxin, and adenovirus 5. 2. HCMV pp65 peptide pool: Pool of 138 peptides derived from a peptide scan (15mers with 11 aa overlap) through 65 kDa phosphoprotein (pp65) (Swiss-Prot ID: P06725) of human Cytomegalovirus (HCMV pp65) – strain AD169. 3. Pepmix actin: Control pool of 92 peptides derived from a peptide scan (15mers with 11 aa overlap) through actin, alpha skeletal muscle (Swiss-Prot ID: P68133) of Homo sapiens (human). 4. Reconstitution of the lyo-FBS. Vortex until homogenous for about 2 min. If lyo-FBS particles are not dissolved after 2 min of vertexing, incubate in a 37 °C water bath for 5 min and vortex again. 5. It is important to sterile filter the reconstituted lyo-FBS. Use a 0.45 μm filter with hydrophilic PVDF membrane, low extractables, and low adsorption membranes. The reconstituted solution may now be used as equivalent to conventional, liquid FBS in media. 6. Thaw cells carefully following below procedure: (a) Thaw the PBMC cryovial in a 37 °C water bath until a small piece of ice is visible. (b) Transfer to a 15 mL falcon tube containing 9 mL of 10% FBS complete media. (c) Centrifuge at 300× g for 5 min, decant supernatant, resuspend in media, and count cells using an automated/manual chamber cell counter. (d) To get 4 × 106 cells/mL (for each well of 24-well plate) ! viable cell count = mL media to add to the cell pellet. 4x106 (e) Spin cell suspension, decant media, and resuspend in volume media were calculated in the previous step.

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7. Loosen the cap on 50 mL conical tube to allow for ventilation or use tubes with venting caps. Extended resting of cells (preferably >16 h) can reset cells to a tissue-like state with increased responsiveness without expanding the number of TCR-specific T cells [12–15]. 8. Prepare 20% excess volume of peptide pool solutions to account for pipetting loss for each peptide pool. 9. Using a reagent reservoir and a multi-channel pipette, add 50 μL of either serum-specific media or peptide pool made in serum-specific media to the wells according to the plate layout. 10. Transfer the plate to the incubator without jiggling the plate. Do not open or move the incubator during this incubation step. Incubators should be isolated and preferably with motion stabilizing or shock-absorbing pads. Incubators located on benchtops next to centrifuges are not recommended. 11. Plates were sent to ZellNet Consulting, Inc. (Fort Lee, NJ) for spot enumeration using the IRIS reader (Mabtech, Nacka Strand, Sweden) employing next-generation plate evaluation with RAWSpot™ technology. 12. The response status to antigens was assessed using the distribution-free resampling (DFR) [16, 17]. A free statistical online tool for running DFR testing is available at https:// rundfr.fredhutch.org/. For all experiments in this chapter, pre-screened donor PBMCs with low, medium, and high immune responses to HCMV pp65 measured by IFNγ secretion were used to assess the performance of liquid CultraPure FBS and lyo-FBS in ELISpot and FluoroSpot assays. To this end, we evaluated assay background levels and immune responses in assays performed with liquid CultraPure FBS compared to lyo-FBS. We also compared lot-to-lot performance consistency, stability, and expansion of antigen-specific cells. PBMCs from three human donors, run in triplicate, show similar IFNγ responses to HCMV pp65 stimulation with three different lots of liquid CultraPure FBS and lyo-FBS in a FluoroSpot assay (Fig. 1). Backgrounds lower than 2 SFU/2.5e5 PBMC were seen in all but one out of nine controls (media alone and cells with media alone) of lyo-FBS lot 1. ELISpot results from a donor PBMC using media made with three lots of freshly prepared liquid CultraPure FBS and lyo-FBS (done in triplicate) show low background for controls and expected HCMV pp65 responses as measured by IFNγ (Fig. 2a). Ninety-day 4 °C storage of these lots supplemented with liquid CultraPure or lyo-FBS show comparable results to the freshly prepared lots using the same PBMC donor (Fig. 2b). All lots and FBS formulations exhibit negligible to no background in media only control wells. Cells and media background for fresh liquid CultraPure FBS is

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Fig. 1 Three lots of liquid CultraPure FBS and lyo-FBS show similar IFNγ responses to HCMV pp65 in a FluoroSpot assay. In this experiment, three human PBMC donors representing low to barely detectable, medium, and high HCMV pp65 responses were employed to assess background levels and responses to antigens for all three lots of liquid CultraPure and lyo-FBS. In all FBS lots, media-only and cells with mediaonly controls showed very low background, in all but one case lower than 2 SFU/2.5e5 PBMC. For all FBS lots, the low responder donor displayed similar antigen-specific responses to the HCMV pp65 peptide pool in media containing either CultraPure or lyo-FBS. (Fig. 1a). Across all FBS lots and formulations, the medium responder donor showed similar backgrounds and antigen-specific responses to media containing either CultraPure or lyo-FBS. While responses were biologically similar in all groups, Lots 1 and 2 exhibited statistically significant differences by two-way ANOVA with liquid FBS slightly higher than lyo-FBS (Fig. 1b). The high-responder donor demonstrated comparable antigen-specific responses across all three lots and formulations of FBS. Lots 1 and 2 exhibited statistically significant differences by two-way ANOVA with liquid FBS slightly higher than lyo-FBS (Fig. 1c)

5 ± 3 SFU/2.5e5 when averaged, and lyo-FBS lots are fewer than 2 ± 1 SFU/2.5e5 PBMCs. All FBS formats and lots produce a robust response “signal” via spot counts above 400 SFU/2.5e5 PBMCs on average. Therefore, liquid CultraPure FBS and lyo-FBS show high signal-to-noise ratios in ELISpot assays and are stable when stored at 4 °C for up to 90 days. We show in Fig. 1c that three lots of liquid CultraPure FBS and lyo-FBS have low background and similar IFNγ responses in FluoroSpot assay (Fig. 1c is reproduced in Fig. 3a again for comparison). We extend these findings and show that media supplemented with either liquid CultraPure FBS or lyo-FBS stored at 4 °C for 90 days demonstrates similar low background responses and expected HCVM pp65 IFNγ responses and remains stable for 90 days in a FluoroSpot assay.

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Fig. 2 An ELISpot assay shows three lots of liquid CultraPure FBS and lyo-FBS have the low background and similar IFNγ responses and remain stable for 90 days when stored at 4 °C. All lots and formulations of FBS exhibit low to negligible background with media only. Cells and media controls are below 5 SFU/2.5e5 PBMCs when averaged from triplicate replicates, and lyo-FBS lots are fewer than 2 SFU/2.5e5 PBMCs when averaged. All three lots and formulations of FBS have similar HCMV pp65 IFNγ responses using PBMCs from the same donor. Lot 1 shows a reduction in HCMV pp65 IFNγ responses with the lyo-FBS lot compared to responses to the liquid CultraPure lot by two-way ANOVA (Fig. 2a). Cells and media controls for 4 °C stored FBS formulations and lots are similar to the results for freshly prepared lot/formulations. Liquid CultraPure lots 1 and 2 have backgrounds below 5 SFU/2.5e5 PBMCs when averaged, and lyo-FBS lots are fewer than 2 SFU/2.5e5 PBMCs when averaged

We developed a multiplex cultured FluoroSpot assay protocol to evaluate the capacity for rare immune cell expansion in the presence of appropriate growth-promoting cytokines and peptide pools using three lots of lyo-FBS supplemented media [18]. We utilize the HCMV pp65 low donor PBMC exhibiting a barely detectable IFNγ response identified in Fig. 1a (±3 SFU/2.5e5 PBMCs per triplicate) to compare the responses after culture with peptide antigen and cytokines for 9 days. We see a significant increase in HCMV pp65 IFNγ responses, in triplicate, after expansion showing 400 ± 30 SFU/2.5e5 PBMCs on average while maintaining a low background (Fig. 4a) compared to before expansion (±3 SFU/2.5e5 PBMCs). Since our method was successful in expanding IFNγ T cells, we wanted to determine the quality of cells expanded. In addition to the HCMV pp65 IFNγ-secreting cells, we show the frequency of six additional populations which are identifiable following culture with the triple-analyte assay in a pie chart (Fig. 4b). As expected, the highest post-culture responses come from IFNγ and TNF single positive cells. Seven response populations are detected from the cultured PBMCs, where we observe three monofunctional populations that secrete IFNγ, TNF, or Granzyme B, and 4 polyfunctional populations that secrete two or

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Fig. 3 Three lots of liquid CultraPure FBS and lyo-FBS have low background and similar HCMV pp65 IFNγ responses via a FluoroSpot assay and remain stable for 90 days when stored at 4 °C. For comparison, the data from Fig. 1c are shown again in Fig. 3a. Media containing all lots and formulations of FBS which was stored at 4 °C for 90 days, demonstrating similar HCMV pp65 IFNγ responses and remains stable for 90 days (Fig. 3b). Backgrounds on both controls for 4 °C stored FBS lots and formulations are similar to the results for freshly prepared lots. All three fresh FBS and lyo-FBS lots stored for 90 days have similar HCMV pp65 IFNγ responses using the same donor PBMCs

Fig. 4 In vitro expansion of antigen-specific HCMV pp65 cells enables the detection of polyfunctional T cells in a 9-day cultured FluoroSpot Assay. The cultured IFNγ response from the low donor PBMC done in triplicate expanded to 400 SFU/2.5e5 PBMCs shows a significant increase in antigen-specific cells (Fig. 4a). Figure 4b is a pie chart that demonstrates the frequency of the seven populations that resolve following culture from the triple-analyte assay where the highest post-culture responses come from IFNy and TNFα monospecific T cells. Figure 4c shows the number of SFU of the seven resolvable response populations

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Fig. 5 Images of a triple analyte FluoroSpot assay reveal triple-positive polyfunctional T cells. Images of 9-day in vitro culture results obtained using the Mabtech IRIS plate reader reveal distinct single-color populations of IFNγ (green), TNF (red), and Granzyme B (yellow) (Fig. 5a). Figure 5b (lower left) shows an overlay of all three image filters from the same well showing polyfunctional cells (amber). Pepmix actin negative control peptide pool background showing a single spot (orange) as an additional control that was run alongside cells and media (Fig. 5b, upper right)

three of the analytes (Fig. 4c). As expected, triple-positive populations are infrequent, but still clearly detectable in this cultured FluoroSpot assay. We demonstrate that by performing in vitro cultured expansion of low-responding donor PBMCs using lyo-FBS supplemented media, antigen, and cytokines, we are able to detect the expansion of antigen-specific T cells. Example FluoroSpot images from the Mabtech IRIS plate reader of the 9-day in vitro culture from the same well reveal IFNγ (green), TNF (red), and Granzyme B (yellow) signals (Fig. 5a). The same cell in each filter image is indicated with an arrow. Images from the same well are overlaid in Fig. 5b and show that one polyfunctional cell is secreting all three cytokines (amber, shown with a blue arrow). Pepmix actin control showing a single spot (orange) was run as an additional control in addition to our normal cells and media controls (Fig. 5b). The in vitro cultured FluoroSpot assay demonstrates that lyo-FBS can be used with negligible to low background and supports the determination of polyfunctional T cells. Together, our data show that lyo-FBS can be used as an alternative or substitute to liquid CultraPure FBS in ELISpot and FluroSpot assays achieving exquisite sensitivity. Due to the importance of the right sera for any immune monitoring in both pre-clinical and clinical assays, Q2 solutions have developed a protocol for ongoing testing to continuously validate newly released liquid CultraPure FBS and lyo-FBS lots. Such prevalidated FBS in both liquid and lyophilized formats can be recommended for use with confidence in direct ELISpot and FluoroSpot

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assays and cultured FluoroSpot assays for improved performance, standardization, optimization, and reproducibility. We expect these results to positively impact the field of immune monitoring.

Acknowledgments We thank Drs. Patrick Hurban and Patrice Hugo, Q2 solutions, for critical review of this chapter. References 1. Czerkinsky CC, Nilsson LA, Nygren H et al (1983) A solid-phase enzyme-linked immunospot (ELISPOT) assay for enumeration of specific antibody-secreting cells. J Immunol Methods 65:109–121 2. Janetzki S, Rueger M, Dillenbeck T (2014) Stepping up-ELISpot: multi-level analysis in FluoroSpot assays. Cell 3(4):1102–1115 3. Reap EA, Dryga SA, Morris J et al (2007) Cellular and humoral immune responses to alphavirus replicon vaccines expressing cytomegalovirus pp65, IE1, and gB proteins. Clin Vaccine Immunol 14(6):748–755 4. Patton KS, Harrison MT, Long BR et al (2021) Monitoring cell-mediated immune responses in AAV gene therapy clinical trials using a validated IFN- ELISpot method. Mol Ther Methods Clin Dev 22:183–195 5. Arnaud M, Chiffelle J, Genolet R et al (2022) Sensitive identification of neoantigens and cognate TCRs in human solid tumors. Nat Biotechnol 40:656–660 6. Kalyuzhny A, Stark S (2001) A simple method to reduce the background and improve well-towell reproducibility of staining in ELISPOT assays. J Immunol Methods 257(1–2):93–97 7. Janetzki S, Price L, Britten CM et al (2010) Performance of serum-supplemented and serum-free media in IFNγ Elispot Assays for human T cells. Cancer Immunol Immunother 59(4):609–618 8. Janetzki S, Price L, Schroeder H et al (2015) Guidelines for the automated evaluation of Elispot assays. Nat Protoc 10(7):1098–1115 9. Janetzki S, Panageas B-PL et al (2008) Results and harmonization guidelines from two largescale international Elispot proficiency panels conducted by the Cancer Vaccine Consortium (CVC/SVI). Cancer Immunol Immunother 57:303–315 10. Van der Valk J, Bieback K, Buta et al (2017) Fetal bovine serum (FBS): past – present –

future, alternatives to animal experimentation. ALTEX 35(1):99–118 11. Berong M, Ferrari PC et al (2023) Validation of the performance and suitability of a new class of FBS optimized for use in single-cell functional assays. J Immunol Methods 515:113452 12. Romer PS, Berr S, Avota E et al (2011) Preculture of PBMCs at high cell density increases sensitivity of t-cell responses, revealing cytokine release by cd28 super agonist tgn1412. Blood 118(26):6772–6782 13. Wegner J, Hackenberg S, Scholz CJ et al (2015) High-density preculture of PBMCs restores defective sensitivity of circulating cd8 t cells to virus- and tumor-derived antigens. Blood 126(2):185–194 14. Kutscher S, Dembek CJ, Deckert S et al (2013) Overnight resting of PBMCs changes functional signatures of antigen specific T cell responses: impact for immune monitoring within clinical trials. PLoS One 8(10):e76215 15. Santos R, Buying A, Sabri N et al (2014) Improvement of IFNγ elispot performance following overnight resting of frozen PBMCs samples confirmed through rigorous statistical analysis. Cells 4(1):1–1 16. Moodie Z, Price L, Gouttefangeas C et al (2010) Response definition criteria for Eli spot assays revisited. Cancer Immunol Immunother 59(10):1489–1501 17. Moodie Z, Price L, Janetzki S, Britten CM (2012) Response determination criteria for Eli spot: toward a standard that can be applied across laboratories. Methods Mol Biol 792: 185–196 18. Somogyi E, Csiszovski Z, Molna´r L et al (2021) A peptide vaccine candidate tailored to individuals’ genetics mimics the multi-targeted T cell immunity of COVID-19 convalescent subjects. Front Genet 12:1–19

INDEX A Accuracy..........................................3–5, 10, 81, 106–109, 111, 194, 221, 233, 235, 257, 267, 289 Activated paper................................................................ 19 Affinity ........................................... 60–62, 64, 65, 78, 83, 168–170, 173, 175, 178, 179, 182, 183, 190, 192, 194–196, 211–237, 252–254, 259, 269 Affinity capture coating .........................70, 82, 173, 175, 184–187, 196, 225, 228, 254, 260, 263, 269 Affinity maturation....................................... 62, 179, 190, 193, 214–216, 222 Agilent Quanteon™ .......................................... 91, 92, 94 AI-based ImmunoSpot® ...........................................59–83 Antibodies ....................9, 15, 55, 60, 87, 105, 120, 136, 154, 167, 202, 211, 243, 252, 274, 297, 306 Antibody considerations ...........................................20–21 Antibody-DNA conjugation ........................................ 277 Antibody evaluation ........................................................ 20 Antibody-secreting cell (ASC) ...................60–65, 67–70, 78–83, 170–177, 179–183, 186, 188, 190–194, 196, 202, 204, 205, 207, 215–219, 221, 223, 224, 227, 228, 232–237, 253–259, 264, 266–270 Antibody titers ......... 117, 154, 167, 168, 175, 178, 252 Antigen dilutions.................................120, 125, 128, 130 Antigen-specific .................................. 2, 9, 29–47, 59–83, 106, 113, 118, 154, 155, 159, 161, 164, 165, 168, 170–181, 183–188, 191–196, 205, 207, 214–217, 219, 221, 223–225, 227, 228, 230, 232–237, 252–270, 292, 297–303, 311, 312, 314, 315 Antigen stocks ...................................................... 119, 121 Antigen titers........................................................ 171, 181 Anti-tuberculosis treatment (ATT) ................................ 52 Apex™ ........................................................................... 243 Artificial intelligence (AI) ...................................... 65, 270 Automatic scanning and counting ................................. 77

B Bacille Calmette-Guerin (BCG)..................................... 53 Background .......................................... 2, 3, 7, 10–12, 18, 20, 21, 23, 30, 45, 67, 68, 78, 79, 82, 83, 103, 127, 129–131, 137, 138, 148, 155, 161, 193, 196, 197, 237, 270, 298, 302, 306, 307, 311–315

B-cell ELISPOT ..............................................59–83, 154, 162, 168, 190, 196, 202, 207, 269 B cells .........2, 21, 60, 98, 118, 154, 167, 202, 214, 252 in vitro stimulation.................................................. 263 polyclonal in vitro stimulation................................ 263 B lymphocytes ............................................................... 214 Bromochloroindoyl phosphate/nitro blue tetrazolium (BCIP/NBT)........................ 55, 56, 96, 148, 287

C Cancer immunotherapy ............................................51–56 Capping ........................................ 6, 7, 11, 31, 32, 34, 35 Cats ....................................................................... 136–151 CD4 T-cells ............................................................ 38, 310 CD40 ............................................................................. 203 CD8 T-cell.......................................................... 37, 38, 60 Cell and gene therapy ................................................... 306 Cellular immune response ............................................ 118 Cellular immunity ......................................................... 118 Control peptides ...................................4, 6, 10, 307, 315 Control pools ................................................... 43–45, 310 Counting algorithm ..................................................59–83 COVID-19 ......................... 63, 117, 118, 141, 143, 154, 155, 170–172, 181, 191, 193, 221, 222, 224, 256 COVID-19 vaccination ....................................... 142, 176 Cross-contamination....................................................... 35 CTL-LDC™......................................................... 183, 259 CultraPure .................................. 306, 307, 309, 311–315 Cytokine ............................................ 12, 52, 60, 87, 113, 119, 139, 177, 204, 214, 241, 274, 297, 307 Cytokine mRNAs ................................139, 142, 144, 148 Cytomegalovirus (CMV) ..................................10, 44, 45, 298–300, 303, 310

D Deletion peptides ............................................... 11, 31–35 Depreciation ..............................................................46–47 Destaining ....................................................................... 20 Detection ........................9, 17, 33, 52, 60, 87, 105, 121, 139, 159, 167, 203, 215, 241, 253, 274, 297, 306 limit of blank ............................................................... 4 upper and lower limit.................................................. 4 Detection strands ....................... 281–283, 285, 287–289

Alexander E. Kalyuzhny (ed.), Handbook of ELISPOT: Methods and Protocols, Methods in Molecular Biology, vol. 2768, https://doi.org/10.1007/978-1-0716-3690-9, © The Editor(s) (if applicable) and The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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Diffusion blotting .....................................................22, 23 Direct B-cell .................................................................... 63 DNA ...............................15, 20, 80, 150, 177, 195, 235, 268, 274, 276, 278, 281, 283–285, 287, 288 DNA amplification ...................................... 281, 283, 285 DNA complementary pairing .............................. 273–296 DNA concatemers................................................ 287, 289 DNA oligonucleotides ................................ 276, 277, 281 DNA strands........................................274, 282, 291, 296 Dual-stage amplification .....................283, 284, 289, 294

E Electroblotting ..........................................................22–25 ELISPOT........................1, 20, 30, 51, 59, 88, 105, 120, 136, 154, 181, 203, 241, 260, 273, 297, 306 accuracy........................................................................ 2 cross contamination .................................................... 6 limit of detection......................................................... 3 limit of quantitation .................................................... 3 linearity ........................................................................ 3 neoepitopes.................................................................. 6 precision....................................................................... 2 range ............................................................................ 3 robustness .................................................................... 3 ruggedness................................................................... 3 single-color ......................................70, 255, 256, 260 specificity ..................................................................... 2 validation ........................................................... v, 1–12 Endotoxin..................................... 35, 138, 242, 244, 246 Enhancer........................................................................ 204 Epidemiological studies .................................................. 53 Epitope mapping...................................31, 33, 40, 43, 60 Epstein-Barr virus (EBV) ............................ 44, 170, 172, 174, 192, 298–300, 303, 310 Ex vivo .............................60, 78, 80, 195, 233, 267, 309

F Feline coronavirus ................................................ 136–151 Feline coronavirus serotypes 1 (FCoV1) ........... 136–138, 140–147, 149, 151 Feline coronavirus serotypes 2 (FCoV2) ........... 136, 138, 140–144, 146, 147, 149, 151 Feline Cytokine ELISpot.............................................. 147 Fetal Bovine Sera (FBS)............................. 138, 139, 142, 148, 287, 306–315 Ficoll-Paque.......................................................... 119, 122 Flagellin ................................................................ 243–246 Flow cytometry ................................64, 66, 87–103, 173, 191, 207, 222, 223, 266 Flu .................................................................................... 44 Fluorenylmethoxycarbonyl (Fmoc) ..................... 6, 7, 11, 30, 31, 34, 35

Fluorescence enhancer ................................................ 204, 300, 301 Fluorescence-based oligo-linked immunospot (FOLISPOT) ............................................ 273–296 Fluorophore hybridization and dehybridization......... 282 FluoroSpot..................................... 59–83, 105, 113, 168, 175, 184, 185, 188, 190, 203, 206, 207, 217, 225, 227, 228, 241–249, 254–257, 260, 263, 274 affinity capture coating ........................................... 228 four-color........................................................ 185–187 limiting antigen probe ............................................ 227 Four-color........................................................68, 70, 184, 196, 254, 255, 257–260, 263, 269 Functional affinity ................................ 62, 174–176, 211, 213–216, 219–221, 224, 234, 236, 253, 254, 268

G Gram-positive extra-cellular growing bacterium......... 244

H HCMV pp65 ........................................................ 307–314 Heat killed Staphylococcus aureus (HKSA).......... 244–246 High-pressure liquid chromatography (HPLC)................................................... 31, 33–36 High throughput ....................................... 17, 18, 60, 69, 106, 108, 118, 119, 173, 177, 178, 195 HLA-specific IgG producing B cells ................... 202–207 Human leukocyte antigen (HLA)..................35, 44, 202, 203, 205–208, 303 Humoral immunity ......................................167–197, 216 Hybridoma ........................ 178, 215, 218–220, 224, 232

I IFN-γ ELISPOT .................................118–121, 126, 128 Ig class ...................................................... 68, 70, 80, 173, 175–179, 183, 184, 190, 192, 214, 217, 221, 223, 232–234, 252–256, 258–260, 264, 266–270 IgG isotypes................................................................... 205 IgG subclass........... 68, 70, 80, 177, 183, 184, 190–192, 216, 221, 234, 255, 257–260, 264, 266, 267 IL-1β ....................................................242–245, 247, 249 Immobilization..........................................................18, 24 ImmobilonTM .................................................................. 18 Immune memory ............................................................ 54 Immune monitoring ....................... 6, 33, 36, 38, 41, 60, 106, 173, 178, 181, 183, 216, 267, 306, 315, 316 Immune suppression.................................................12, 54 Immunopathology ................................................. 53, 232 Immunosorbant ..................................................... 20, 155 ImmunoSpot® .............................................. 60, 106, 157, 168, 183, 202, 216, 253

HANDBOOK antigen-specific .................................................... 75–77 automatic scanning and counting ............................ 76 direct B-cell ......................................................... 73–74 inverted .............. 70, 75–77, 83, 176, 179, 181, 183, 184, 186, 192, 217–219, 221–224, 233, 236, 237 Pan-Ig .................................................................. 70–72 Infection ........................... 12, 42, 51–53, 61, 62, 67, 80, 118, 136–138, 143, 144, 146, 154, 161, 170, 172, 174, 177, 181, 193, 194, 215, 253, 255, 257, 258, 266 Influenza A (Flu)............................................................. 44 Interferon-gamma (IFN-γ) ............................52, 53, 118, 120, 121, 128, 130, 136–151, 284, 287, 289, 292, 307, 310 Interleukin-2 (IL-2)......................................80, 136–144, 147, 156, 161, 194, 203, 204, 284, 287, 289, 298–303, 308, 310 Interleukin-6 (IL-6)............................. 87–103, 242–245, 247, 249, 284, 287, 289 Interleukin-10 (IL-10) .................................87–103, 203, 204, 284, 287, 289, 292 Interleukin-21 (IL-21) ..............194, 203, 204, 267, 303 In vitro stimulation .................................... 186, 188, 194, 195, 227, 228, 235, 236, 253, 264, 267, 270 IRIS FluoroSpot................................................... 243, 308 IRIS reader ........................................................... 244, 311

L Latent tuberculosis infection (LTBI)............................. 52 Limiting antigen probe........................................ 227, 236 Limit of blank (LOB) ...............................................3, 4, 9 Limit of detection (LOD) ..............................3, 7, 11, 76, 173, 191, 268 Limit of quantitation ........................................................ 3 Linearity........................................ 3, 4, 8, 12, 69, 79, 258 Lipopolysaccharide (LPS) ................................. 88–91, 93, 95–99, 101, 129, 243–246, 287 Lyophilization ...........................................................34, 35 Lyophilized FBS............................................................ 307

M Mabtech..................................... 155–157, 161, 163, 164, 203, 204, 243, 245, 247, 249, 299–302, 307, 308, 310, 311, 315 MALDI .......................................................................... 278 MALDI-TOF ................................................................ 278 Memory ......................................... 32, 54, 59–63, 67, 68, 76, 81, 141, 153–165, 167–197, 202–207, 215, 216, 233, 252, 253, 257, 266 Monocyte activation test (MAT).................................. 242 Monocytes .....................63, 89, 92, 94, 97, 98, 241–244 mRNA extraction .......................................................... 140 Mycobacterium tuberculosis (MTB) .......................51, 52

OF

ELISPOT: METHODS

AND

PROTOCOLS Index 319

N Neoepitopes....................................................................... 6 Nitrocellulose ............................................................15, 17 Nitrocellulose membranes ................................. 17–18, 23 Nylon membranes .....................................................19–20

O Oligonucleotides ......................................... 276, 277, 281

P Pam3CSK4 .................................................................... 245 Pan-Ig ................................ 61–63, 70–72, 233, 257, 264 Peptide ..................................................6, 19, 29, 56, 106, 122, 169, 232, 249, 299, 308 depreciation ......................................................... 45–47 storage.................................................................. 45–47 Peptide analysis and purification .................................... 31 Peptide impurities .....................................................33, 35 Peptide pool .....................................6, 10, 11, 29–47, 57, 118, 298–300, 303, 307–313, 315 Peptide synthesis .................................................. 7, 11, 30 PepTivator® ........................................119, 121, 122, 126 Peripheral blood mononuclear cells (PBMCs).........4, 65, 89, 106, 118, 136, 156, 172, 204, 216, 242, 253, 282, 299, 307 cryopreservartion ..........................108, 233, 259, 260 culture ............................................................. 139, 203 RNA isolation.......................................................... 149 Phytohemagglutinin (PHA) ...................... 119, 121, 122, 126, 130, 138, 144, 147, 151, 287, 309 Phytohemagglutinin-L (PHA-L) ................................. 308 Polyacrylamide gels ...................................................15, 23 Polyclonal in vitro stimulation ............................. 81, 194, 225, 234, 263 Polyfunctional T cells..........................297–299, 314, 315 Polymerase chain reaction ............................................ 150 Polyvinylidene difluoride (PVDF).................... 15, 18–19, 21, 23, 60, 78, 81, 82, 138, 147, 175, 195, 217, 234, 236, 237, 243, 245, 254, 287, 299, 310 Precision ................................... 3–5, 8, 9, 66–69, 80, 255 Primer exchange reaction (PER)................ 279, 281, 283 Protein spanning ................................................ 10, 38–41 Protein transfer.............................................16–20, 22, 23 Pyrogens ........................................................................ 242

R R848 (resiquimod) ...................................... 80, 155, 156, 161, 194, 203, 204, 245, 246 Range .............3–5, 7–10, 12, 17, 33, 37, 38, 40, 45, 62, 63, 65, 67–69, 87, 130, 146, 155, 174, 176–179, 193, 212, 217, 221, 234, 237, 254, 258, 270 RAWspot™........................................................... 243, 244

HANDBOOK OF ELISPOT: METHODS AND PROTOCOLS

320 Index

Reagent Tracker™ .......................................107, 109–111 Receptor binding domain (RBD) ...................... 136, 138, 141–145, 147, 149, 151, 154, 155, 181, 192, 194, 221–223, 234 Resiquimod ................................................................... 203 Respiratory-syncytial virus (RSV) ..................44, 45, 244, 247, 249, 298, 301, 303 Reverse transcriptase ..................................................... 150 Reverse Transcription and PCR of cDNA................... 140 RNA isolation................................................................ 149 Robustness...........................................3, 4, 8, 68, 81, 106 RPMI 1640 ............................................ 90, 91, 119–123, 125, 126, 139, 156, 157, 195, 203, 235, 244, 268, 282, 284, 287, 299, 308 Ruggedness........................................................................ 3

S SARS-CoV-2 (SCoV2) ................. 39, 41, 42, 44, 63–66, 117–119, 121, 122, 126, 136–138, 140–145, 147, 149, 151, 153–165, 170–172, 174, 181, 191–194, 218, 219, 221, 222, 224, 234, 255–258 Spike RBD-His recombinant protein .................... 156 Spike S1-His recombinant protein......................... 156 Spike S1 NTD-His & AVI recombinant protein .. 156 Spike S2 ECD-His recombinant protein ............... 156 Wuhan-Hu-1 proteins ............................................ 157 Semi-dry transfer............................................................. 24 Serum.................................. 18, 19, 52, 56, 60, 100, 136, 138, 139, 167–170, 172, 175, 179, 181, 190, 194, 202, 203, 207, 208, 215, 216, 232, 246, 252, 253, 264, 276, 282, 284, 299, 307, 309 Signal amplification .................................... 274, 282–284, 287, 289, 292 Simple diffusion ........................................................21, 22 Single-stage amplification .......................... 275, 283, 285, 289, 291, 294 Sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE)........................21, 22 Specificity ....................................... 2, 3, 6, 10, 11, 15–25, 37, 52, 61, 64, 70–72, 82, 170, 174–178, 181, 190–193, 202, 205, 206, 211, 214–217, 221, 230, 233, 235, 253, 254, 256, 257, 266, 306 Spike RBD-His recombinant protein .......................... 156 Spike S1-HIS recombinant protein.............................. 156 Spike S1 NTD-His & AVI recombinant protein ........ 156 Spike S2 ECD-His recombinant protein ..................... 156 Standard operating procedures (SOPs) ................. 2, 4, 5, 7–10, 12, 36, 207

Storage ........................................... 16, 17, 34, 36, 45–47, 124, 159, 245, 247, 270, 306, 307, 311 Strand sequences .................................278, 281, 282, 284 SytoxTM Blue Dead Cell Stain........................................ 90

T T-cell activation assays .................................................... 30 T cells .................................... 2, 38, 52, 60, 90, 106, 118, 136, 154, 172, 207, 232, 241, 267, 298, 311 Tetramers HLA ......................................................................... 206 Tetramers TLR agonist Tetramethylbenzidine (TMB) .................... 157, 164, 203 TFA cleavage ................................................................... 34 TGF-β1 ................................................................. 284, 289 TLR agonist................................................................... 205 TLR2 agonist ................................................................ 244 TNF-α ......................................................... 242–245, 247, 284, 287, 289, 292, 298–303 Transplantation ........................................... 202, 205, 208 Treatment efficacy ........................................................... 54 Trifluoroacetate (TFA) ...................................... 31, 34, 35 Triple-color FluoroSpot....................................... 298, 300 Trippel-color.................................................................. 245 Trypan blue ......................... 90, 123, 156, 159, 246, 308 Tuberculin skin test (TST) .......................................52, 53 Tuberculosis (TB) ................................... 51–56, 274, 298 diagnosis .............................................................. 51–53 2019-nCoV ................................................. 119, 121, 122

U Ultimate Analyzer ........................................................... 76 Upper and lower limit .................................................. 3, 4

V Vaccine development ........................................................53, 55 Vacuum blotting .......................................................23, 24

W Western blotting........................................................15–25 Wet transfer ..................................................................... 24 Wuhan-Hu-1 proteins .................................................. 157

Z Zetabind (ZB) ................................................................. 19