Vaccine Design: Methods and Protocols, Volume 1. Vaccines for Human Diseases (Methods in Molecular Biology, 2410) 1071618830, 9781071618837

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Methods in Molecular Biology 2410

Sunil Thomas Editor

Vaccine Design Methods and Protocols, Volume 1: Vaccines for Human Diseases Second Edition

METHODS

IN

MOLECULAR BIOLOGY

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

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

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

Vaccine Design Methods and Protocols, Volume 1: Vaccines for Human Diseases Second Edition

Edited by

Sunil Thomas Lankenau Institute for Medical Research, Wynnewood, PA, USA

Editor Sunil Thomas Lankenau Institute for Medical Research Wynnewood, PA, USA

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-1883-7 ISBN 978-1-0716-1884-4 (eBook) https://doi.org/10.1007/978-1-0716-1884-4 © The Editor(s) (if applicable) and The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022, corrected publication 2022 This work is subject to copyright. All rights are reserved by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, 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.

Dedication The healthcare and frontline workers who worked tirelessly taking care of COVID-19 patients. Researchers who studied diligently the biology of SARS-CoV-2 and developed vaccines to protect against COVID-19.

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Preface A healthy society should not have just one voice.—Li Wenliang (1986–2020) (the first physician to recognize the outbreak of COVID-19 in Wuhan, China)

Vaccinations have greatly reduced the burden of infectious diseases. Aggressive vaccination strategies have helped eradicate smallpox in humans and rinderpest, a serious disease of cattle. Vaccination has greatly reduced many pediatric infectious diseases. Vaccines not only protect the immunized but can also reduce disease among unimmunized individuals in the community through “herd protection.” Vaccines have also led to increased production of fish and farm animals, thereby improving food security. The development of vaccines has improved our understanding of immunology and the principles of immunity. This has led to the research and development of vaccines for cancer and neurodegenerative diseases. The world’s health and economy deteriorated since the first report of COVID-19 in China in December 2019. The pandemic has resulted in a huge interest in the development of vaccines. Even the skeptics were clamoring for early development of vaccines. Generally, vaccines take around 10–15 years to reach the clinic. Advances in the knowledge of molecular biology, immunology, and bioinformatics have led to the development of mRNA and adenovirus vector vaccines that are more efficacious than conventional vaccines. Collaboration at multiple levels led to the development and quick employment of COVID19 vaccines in the clinic within a year of the observation of the disease, making it the quickest vaccines ever to be developed and deployed. In 2016, we published the first edition of the book Vaccine Design: Methods and Protocols. Volume 1: Vaccines for Human Diseases and Volume 2: Vaccines for Veterinary Diseases. The books were a tremendous success. The Methods in Molecular Biology series Vaccine Design: Methods and Protocols, Second Edition, contains 87 chapters in three volumes. Volume 1: Vaccines for Human Diseases, has an introductory section on future challenges for vaccinologists, the immunological mechanism of vaccines, and trends in vaccinology. The design of human vaccines for viral, bacterial, fungal, and parasitic diseases as well as vaccines for tumors is also described in this volume. Volume 2: Vaccines for Veterinary Diseases includes vaccines for farm animals and fishes. Volume 3: Resources for Vaccine Development includes chapters on vaccine adjuvants, vaccine vectors and production, vaccine delivery systems, vaccine bioinformatics, vaccine regulation, and intellectual property. It has been 225 years since Edward Jenner vaccinated his first patient in 1796 to protect against smallpox. This book is a tribute to the pioneering effort of his work. The job of publishing the second edition of the book Vaccine Design: Methods and Protocols was assigned at a tough time. Most of the universities were closed due to COVID-19 immediately after I took up the assignment. Several of the authors, their collaborators, and families were infected with the virus while contributing to the book. Nevertheless, the authors completed their chapters within the stipulated time. I am extremely grateful to the authors for completing the task in spite of the hardship faced while contributing to the books. My sincere thanks to all the authors for contributing to Vaccine Design: Methods and Protocols (Edition 2); Volume 1: Vaccines for Human Diseases; Volume 2: Vaccines for Veterinary

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Diseases; and Volume 3: Resources for Vaccine Development. I would also like to thank the series editor of Methods in Molecular Biology™, Prof. John M. Walker, for giving me the opportunity to edit this book. My profound thanks to my parents Thomas and Thresy, wife Jyothi for the encouragement and support, and also our twins Teresa and Thomas for patiently waiting for me while preparing the book. Working on the book was not an excuse for staying away from the laboratory. I made sure that my children were told about new exciting data generated in the laboratory and the advances in science published daily before bedtime. Wynnewood, PA, USA

Sunil Thomas

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

PART I

VACCINES: INTRODUCTION

1 Challenges for Vaccinologists in the First Half of the Twenty-First Century . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sunil Thomas, Ann Abraham, Patrick J. Callaghan, and Rino Rappuoli 2 Principles in Immunology for the Design and Development of Vaccines . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Claudius U. Meyer and Fred Zepp 3 Revisiting the Principles of Designing a Vaccine . . . . . . . . . . . . . . . . . . . . . . . . . . . . Shubhranshu Zutshi, Sunil Kumar, Prashant Chauhan, and Bhaskar Saha 4 Status of COVID-19 Pandemic Before the Administration of Vaccine . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sunil Thomas

PART II

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TRENDS IN VACCINOLOGY

5 mRNA Vaccines to Protect Against Diseases. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 111 Sunil Thomas and Ann Abraham 6 Artificial Intelligence in Vaccine and Drug Design . . . . . . . . . . . . . . . . . . . . . . . . . . 131 Sunil Thomas, Ann Abraham, Jeremy Baldwin, Sakshi Piplani, and Nikolai Petrovsky

PART III

VACCINES FOR HUMAN VIRAL DISEASES

7 Vaccines Targeting Numerous Coronavirus Antigens, Ensuring Broader Global Population Coverage: Multi-epitope and Multi-patch Vaccines . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 149 Sukrit Srivastava, Spyros D. Chatziefthymiou, and Michael Kolbe 8 Use of Micro-Computed Tomography to Visualize and Quantify COVID-19 Vaccine Efficiency in Free-Breathing Hamsters . . . . . . . . . . . . . . . . . . 177 Laura Seldeslachts, Christopher Cawthorne, Suzanne F. Kaptein, Robbert Boudewijns, Hendrik Jan Thibaut, Lorena Sanchez Felipe, Sapna Sharma, Georg Schramm, Birgit Weynand, Kai Dallmeier, and Greetje Vande Velde

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9 Design of Replication-Competent VSV- and Ervebo-Vectored Vaccines Against SARS-CoV-2. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Qixing Liu, Zhe Ding, Jiaming Lan, and Gary Wong 10 CRISPR Engineering of Bacteriophage T4 to Design Vaccines Against SARS-CoV-2 and Emerging Pathogens . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jingen Zhu, Neeti Ananthaswamy, Swati Jain, Himanshu Batra, Wei-Chun Tang, and Venigalla B. Rao 11 Techniques for Developing and Assessing Immune Responses Induced by Synthetic DNA Vaccines for Emerging Infectious Diseases . . . . . . . . Ziyang Xu, Michelle Ho, Devivasha Bordoloi, Sagar Kudchodkar, Makan Khoshnejad, Leila Giron, Faraz Zaidi, Moonsup Jeong, Christine C. Roberts, Young K. Park, Joel Maslow, Mohamed Abdel-Mohsen, and Kar Muthumani 12 Towards Determining the Epitopes of the Structural Proteins of SARS-CoV-2 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sunil Thomas 13 Development, Production, and Characterization of Hepatitis B Subviral Envelope Particles as a Third-Generation Vaccine . . . . . . . . . . . . . . . . . . . Juan Manuel Battagliotti, Diego Fontana, Marina Etcheverrigaray, Ricardo Kratje, and Claudio Prieto 14 Generation of CpG-Recoded Zika Virus Vaccine Candidates . . . . . . . . . . . . . . . . . Ivan Trus, Daniel Udenze, and Uladzimir Karniychuk

PART IV 15

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VACCINES FOR HUMAN BACTERIAL DISEASES

Salmonella Uptake into Gut-Associated Lymphoid Tissues: Implications for Targeted Mucosal Vaccine Design and Delivery. . . . . . . . . . . . . . Angelene F. Richards, Fernando J. Torres-Velez, and Nicholas J. Mantis Development of Human Recombinant Leptospirosis Vaccines . . . . . . . . . . . . . . . . Natasha Rodrigues de Oliveira, Thaı´s Larre´ Oliveira, Se´rgio Jorge, and Odir Antoˆnio Dellagostin Induction of T Cell Responses by Vaccination of a Streptococcus pneumoniae Whole-Cell Vaccine . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Emily M. Roy, Fan Zhang, Richard Malley, and Ying-Jie Lu Development of a Bacterial Nanoparticle Vaccine Against Escherichia coli. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Melibea Berzosa, Yadira Pastor, Carlos Gamazo, and Juan Manuel Irache Construction of Novel Live Genetically Modified BCG Vaccine Candidates Using Recombineering Tools . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Mario Alberto Flores-Valdez and Michel de Jesu´s Aceves-Sa´nchez

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An Update on Tuberculosis Vaccines . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 387 Radha Gopalaswamy and Selvakumar Subbian Structure-Based Design of Diagnostics and Vaccines for Lyme Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 411 Sunil Thomas Development of a SONIX Vaccine to Protect Against Ehrlichiosis . . . . . . . . . . . . 423 Sunil Thomas

PART V 23

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VACCINES FOR HUMAN PARASITIC DISEASES

Development of the Antileishmanial Vaccine . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sunil Kumar, Shubhranshu Zutshi, Mukesh Kumar Jha, Prashant Chauhan, and Bhaskar Saha In Silico Design of Recombinant Chimera T Cell Peptide Epitope Vaccines for Visceral Leishmaniasis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Amanda Sanchez Machado, Vivian Tamietti Martins, Maria Victoria Humbert, Myron Christodoulides, and Eduardo Antonio Ferraz Coelho Preclinical Assessment of the Immunogenicity of Experimental Leishmania Vaccines . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Vivian Tamietti Martins, Amanda Sanchez Machado, Maria Victoria Humbert, Myron Christodoulides, and Eduardo Antonio Ferraz Coelho Production of Oral Vaccines Based on Virus-Like Particles Pseudotyped with Protozoan-Surface Proteins. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Lucı´a Lara Rupil, Marianela del Carmen Serradell, and Hugo Daniel Luja´n A Fast-Track Phenotypic Characterization of Plasmodium falciparum Vaccine Antigens through Lyse-Reseal Erythrocytes Mediated Delivery (LyRED) of RNA Interference for Targeted Translational Repression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Malabika Chakrabarti, Swati Garg, Akshay Munjal, Sweta Karan, Soumya Pati, Lalit C. Garg, and Shailja Singh Plasmodium falciparum Antigen Expression in Leishmania Parasite: A Way Forward for Live Attenuated Vaccine Development . . . . . . . . . . . . . . . . . . . Akriti Srivastava, Swati Garg, Sweta Karan, Shikha Kaushik, Anand Ranganathan, Soumya Pati, Lalit C. Garg, and Shailja Singh Molecular Characterization of a Vector-Based Candidate Antigen Using the 30 -RACE and Genome Walking Methods and In Silico Analysis of the Correspondent Protein for Vaccine Design and Development . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Abbasali Raz, Mahdieh Manafi, and Mahdokht Ilbeigi Khamseh Nejad In Vitro Culture of Plasmodium falciparum for the Production of Mature Gametocytes for Performing Standard Membrane Feeding Assay and Infection of Anopheles spp.. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Abbasali Raz and Mahdokht Ilbeigi Khamseh Nejad

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Plasmodium berghei Infection in BALB/c Mice Model as an Animal Model for Malaria Disease Research . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 589 Abbasali Raz Standard Membrane Feeding Assay for Malaria Transmission Studies. . . . . . . . . . 597 Abbasali Raz, Jafar J. Sani, and Hemn Yousefi

PART VI 33

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Generation of Tumor Targeted Dendritic Cell Vaccines with Improved Immunogenic and Migratory Phenotype . . . . . . . . . . . . . . . . . . . . . . . . . 609 Adam M. Swartz, Kelly M. Hotchkiss, Smita K. Nair, John H. Sampson, and Kristen A. Batich Monocytes as a Cellular Vaccine Platform to Induce Antitumor Immunity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 627 Min-Nung Huang, Vincent M. D’Anniballe, and Michael D. Gunn Beyond Sequencing: Prioritizing and Delivering Neoantigens for Cancer Vaccines . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 649 Alexander S. Roesler and Karen S. Anderson

PART VII 36

DEVELOPMENT OF CANCER VACCINES

VACCINES FOR ALLERGY

Proteomics for Development of Food Allergy Vaccines . . . . . . . . . . . . . . . . . . . . . . 673 Monica Carrera and Susana Magada´n

PART VIII

VACCINES FOR TOXINS

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Estimating Vaccine Potency Using Antibody-Based Competition Assays . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 693 Jennifer Doering, Greta Van Slyke, Oreola Donini, and Nicholas J. Mantis Correction to: Status of COVID-19 Pandemic Before the Administration of Vaccine . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C1 Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Contributors MOHAMED ABDEL-MOHSEN • Vaccine & Immunotherapy Center, The Wistar Institute, Philadelphia, PA, USA ANN ABRAHAM • Lankenau Institute for Medical Research, Wynnewood, PA, USA MICHEL DE JESU´S ACEVES-SA´NCHEZ • Centro de Investigacion y Asistencia en Tecnologı´a y disen˜o del Estado de Jalisco, A.C. Biotecnologı´a Me´dica y Farmace´utica. Av. Normalistas 800, Col. Colinas de la Normal, Guadalajara, Jalisco, Mexico NEETI ANANTHASWAMY • Department of Biology, The Catholic University of America, Washington, DC, USA KAREN S. ANDERSON • Mayo Clinic, Scottsdale, AZ, USA; Center for Personalized Diagnostics, Biodesign Institute, Arizona State University, Tempe, AZ, USA JEREMY BALDWIN • Vaxine Pty Ltd, Adelaide, SA, Australia KRISTEN A. BATICH • Department of Neurosurgery, Duke University Medical Center, Durham, NC, USA; The Preston Robert Tisch Brain Tumor Center, Duke University Medical Center, Durham, NC, USA HIMANSHU BATRA • Department of Biology, The Catholic University of America, Washington, DC, USA JUAN MANUEL BATTAGLIOTTI • UNL, CONICET, FBCB (School of Biochemistry and Biological Sciences), CBL (Biotechnological Center of Litoral), Cell Culture Laboratory, Ciudad Universitaria, Santa Fe, Argentina MELIBEA BERZOSA • Department of Microbiology and Parasitology, Institute of Tropical Health, University of Navarra, Pamplona, Spain DEVIVASHA BORDOLOI • Vaccine & Immunotherapy Center, The Wistar Institute, Philadelphia, PA, USA ROBBERT BOUDEWIJNS • Virology and Chemotherapy, Molecular Vaccinology & Vaccine Discovery, KU Leuven Department of Microbiology, Immunology and Transplantation, Rega Institute, Leuven, Belgium PATRICK J. CALLAGHAN • Lankenau Institute for Medical Research, Wynnewood, PA, USA MO´NICA CARRERA • Department of Food Technology, Spanish National Research Council (CSIC), Institute of Marine Research (IIM), Pontevedra, Spain CHRISTOPHER CAWTHORNE • KU Leuven Department of Imaging and Pathology, Nuclear Medicine and Molecular Imaging, Leuven, Belgium MALABIKA CHAKRABARTI • Special Centre for Molecular Medicine, Jawaharlal Nehru University, New Delhi, India SPYROS D. CHATZIEFTHYMIOU • Deutsches Elektronen-Synchrotron (DESY), Hamburg, Germany; Department of Structural Infection Biology, Center for Structural Systems Biology (CSSB), Helmholtz-Center for Infection Research (HZI), Hamburg, Germany PRASHANT CHAUHAN • National Centre for Cell Science, Ganeshkhind, Pune, Maharashtra, India MYRON CHRISTODOULIDES • Neisseria Research Group, Molecular Microbiology, School of Clinical and Experimental Sciences, University of Southampton Faculty of Medicine, Southampton General Hospital, Southampton, England, UK

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EDUARDO ANTONIO FERRAZ COELHO • Programa de Pos-Graduac¸a˜o em Cieˆncias da Sau´de: Infectologia e Medicina Tropical, Faculdade de Medicina, Universidade Federal de Minas Gerais, Belo Horizonte, Minas Gerais, Brazil VINCENT M. D’ANNIBALLE • Department of Medicine, Duke University Medical Center, Durham, NC, USA KAI DALLMEIER • Virology and Chemotherapy, Molecular Vaccinology & Vaccine Discovery, KU Leuven Department of Microbiology, Immunology and Transplantation, Rega Institute, Leuven, Belgium NATASHA RODRIGUES DE OLIVEIRA • Universidade Federal de Pelotas, Centro de Desenvolvimento Tecnologico, Campus Universita´rio s/n, Pelotas, RS, Brazil ODIR ANTOˆNIO DELLAGOSTIN • Universidade Federal de Pelotas, Centro de Desenvolvimento Tecnologico, Campus Universita´rio s/n, Pelotas, RS, Brazil ZHE DING • Institut Pasteur of Shanghai, Chinese Academy of Sciences, Shanghai, China JENNIFER DOERING • Division of Infectious Diseases, Wadsworth Center, New York State Department of Health, Albany, NY, USA OREOLA DONINI • Soligenix, Inc, Princeton, NJ, USA MARINA ETCHEVERRIGARAY • UNL, CONICET, FBCB (School of Biochemistry and Biological Sciences), CBL (Biotechnological Center of Litoral), Cell Culture Laboratory, Ciudad Universitaria, Santa Fe, Argentina MARIO ALBERTO FLORES-VALDEZ • Centro de Investigacion y Asistencia en Tecnologı´a y disen˜o del Estado de Jalisco, A.C. Biotecnologı´a Me´dica y Farmace´utica. Av. Normalistas 800, Col. Colinas de la Normal, Guadalajara, Jalisco, Mexico DIEGO FONTANA • UNL, CONICET, FBCB (School of Biochemistry and Biological Sciences), CBL (Biotechnological Center of Litoral), Cell Culture Laboratory, Ciudad Universitaria, Santa Fe, Argentina; UNL, FBCB (School of Biochemistry and Biological Sciences), CBL (Biotechnological Center of Litoral), Biotechnological Development Laboratory, Ciudad Universitaria, Santa Fe, Argentina CARLOS GAMAZO • Department of Microbiology and Parasitology, Institute of Tropical Health, University of Navarra, Pamplona, Spain LALIT C. GARG • Gene Regulation Laboratory, National Institute of Immunology, New Delhi, India SWATI GARG • Special Centre for Molecular Medicine, Jawaharlal Nehru University, New Delhi, India LEILA GIRON • Vaccine & Immunotherapy Center, The Wistar Institute, Philadelphia, PA, USA RADHA GOPALASWAMY • Department of Bacteriology, ICMR-National Institute for Research in Tuberculosis, Chennai, Tamil Nadu, India MICHAEL D. GUNN • Department of Medicine, Duke University Medical Center, Durham, NC, USA MICHELLE HO • Vaccine & Immunotherapy Center, The Wistar Institute, Philadelphia, PA, USA KELLY M. HOTCHKISS • Department of Neurosurgery, Duke University Medical Center, Durham, NC, USA MIN-NUNG HUANG • Department of Medicine, Duke University Medical Center, Durham, NC, USA MARIA VICTORIA HUMBERT • Neisseria Research Group, Molecular Microbiology, School of Clinical and Experimental Sciences, University of Southampton Faculty of Medicine, Southampton General Hospital, Southampton, England, UK

Contributors

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MAHDOKHT ILBEIGI KHAMSEH NEJAD • Malaria and Vector Research Group (MVRG), Biotechnology Research Center (BRC), Pasteur Institute of Iran, Tehran, Iran JUAN MANUEL IRACHE • Department of Technology and Pharmaceutical Chemistry, University of Navarra, Pamplona, Spain SWATI JAIN • Department of Biology, The Catholic University of America, Washington, DC, USA MOONSUP JEONG • GeneOne Life Science Inc., Seoul, South Korea MUKESH KUMAR JHA • National Centre for Cell Science, Ganeshkhind, Pune, Maharashtra, India; Department of Microbiology and Immunology, Columbia University, New York, NY, USA SE´RGIO JORGE • Universidade Federal de Pelotas, Faculdade de Medicina Veterina´ria, Campus Universita´rio s/n, Pelotas, RS, Brazil SUZANNE F. KAPTEIN • Virology and Chemotherapy, Molecular Vaccinology & Vaccine Discovery, KU Leuven Department of Microbiology, Immunology and Transplantation, Rega Institute, Leuven, Belgium SWETA KARAN • Gene Regulation Laboratory, National Institute of Immunology, New Delhi, India ULADZIMIR KARNIYCHUK • Vaccine and Infectious Disease Organization-International Vaccine Centre (VIDO-InterVac), University of Saskatchewan, Saskatoon, SK, Canada SHIKHA KAUSHIK • Special Centre for Molecular Medicine, Jawaharlal Nehru University, New Delhi, India MAKAN KHOSHNEJAD • Vaccine & Immunotherapy Center, The Wistar Institute, Philadelphia, PA, USA MICHAEL KOLBE • Department of Structural Infection Biology, Center for Structural Systems Biology (CSSB), Helmholtz-Center for Infection Research (HZI), Hamburg, Germany; MIN-Faculty University Hamburg, Hamburg, Germany RICARDO KRATJE • UNL, CONICET, FBCB (School of Biochemistry and Biological Sciences), CBL (Biotechnological Center of Litoral), Cell Culture Laboratory, Ciudad Universitaria, Santa Fe, Argentina SAGAR KUDCHODKAR • GeneOne Life Science Inc., Seoul, South Korea SUNIL KUMAR • National Centre for Cell Science, Ganeshkhind, Pune, Maharashtra, India JIAMING LAN • Institut Pasteur of Shanghai, Chinese Academy of Sciences, Shanghai, China QIXING LIU • Institut Pasteur of Shanghai, Chinese Academy of Sciences, Shanghai, China YING-JIE LU • Division of Infectious Diseases, Boston Children’s Hospital, Harvard Medical School, Boston, MA, USA HUGO DANIEL LUJA´N • Centro de Investigacion y Desarrollo en Inmunologı´a y Enfermedades Infecciosas (CIDIE), Consejo Nacional de Investigaciones Cientı´ficas y Te´cnicas (CONICET)/Universidad Catolica de Cordoba (UCC), Cordoba, Argentina; Facultad de Ciencias de la Salud, Universidad Catolica de Cordoba (UCC), Cordoba, Argentina AMANDA SANCHEZ MACHADO • Programa de Pos-Graduac¸a˜o em Cieˆncias da Sau´de: Infectologia e Medicina Tropical, Faculdade de Medicina, Universidade Federal de Minas Gerais, Belo Horizonte, Minas Gerais, Brazil SUSANA MAGADA´N • Biomedical Research Center (CINBIO), Universidade de Vigo, Immunology, Pontevedra, Spain RICHARD MALLEY • Division of Infectious Diseases, Boston Children’s Hospital, Harvard Medical School, Boston, MA, USA MAHDIEH MANAFI • Malaria and Vector Research Group (MVRG), Biotechnology Research Center (BRC), Pasteur Institute of Iran, Tehran, Iran

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NICHOLAS J. MANTIS • Department of Biomedical Sciences, University at Albany School of Public Health, Albany, NY, USA; Division of Infectious Diseases, Wadsworth Center, New York State Department of Health, Albany, NY, USA VIVIAN TAMIETTI MARTINS • Programa de Pos-Graduac¸a˜o em Cieˆncias da Sau´de: Infectologia e Medicina Tropical, Faculdade de Medicina, Universidade Federal de Minas Gerais, Belo Horizonte, Minas Gerais, Brazil JOEL MASLOW • GeneOne Life Science Inc., Seoul, South Korea CLAUDIUS U. MEYER • Department of Pediatrics, University Medical Center Mainz, Mainz, Germany AKSHAY MUNJAL • Special Centre for Molecular Medicine, Jawaharlal Nehru University, New Delhi, India KAR MUTHUMANI • Vaccine & Immunotherapy Center, The Wistar Institute, Philadelphia, PA, USA; GeneOne Life Science Inc., Seoul, South Korea SMITA K. NAIR • Division of Surgical Sciences, Department of Surgery, Duke University Medical Center, Durham, NC, USA; The Preston Robert Tisch Brain Tumor Center, Duke University Medical Center, Durham, NC, USA THAI´S LARRE´ OLIVEIRA • Universidade Federal de Pelotas, Centro de Desenvolvimento Tecnologico, Campus Universita´rio s/n, Pelotas, RS, Brazil YOUNG K. PARK • GeneOne Life Science Inc., Seoul, South Korea YADIRA PASTOR • Department of Microbiology and Parasitology, Institute of Tropical Health, University of Navarra, Pamplona, Spain SOUMYA PATI • Department of Life Science, School of Natural Sciences, Shiv Nadar University, Gautam Buddh Nagar, Uttar Pradesh, India NIKOLAI PETROVSKY • Vaxine Pty Ltd, Adelaide, SA, Australia; College of Medicine and Public Health, Flinders University, Adelaide, SA, Australia SAKSHI PIPLANI • Vaxine Pty Ltd, Adelaide, SA, Australia; College of Medicine and Public Health, Flinders University, Adelaide, SA, Australia CLAUDIO PRIETO • UNL, FBCB (School of Biochemistry and Biological Sciences), CBL (Biotechnological Center of Litoral), Biotechnological Development Laboratory, Ciudad Universitaria, Santa Fe, Argentina ANAND RANGANATHAN • Special Centre for Molecular Medicine, Jawaharlal Nehru University, New Delhi, India VENIGALLA B. RAO • Department of Biology, The Catholic University of America, Washington, DC, USA RINO RAPPUOLI • GSK Vaccines, Siena, Italy ABBASALI RAZ • Malaria and Vector Research Group (MVRG), Biotechnology Research Center (BRC), Pasteur Institute of Iran, Tehran, Iran ANGELENE F. RICHARDS • Department of Biomedical Sciences, University at Albany School of Public Health, Albany, NY, USA; Division of Infectious Diseases, Wadsworth Center, New York State Department of Health, Albany, NY, USA CHRISTINE C. ROBERTS • GeneOne Life Science Inc., Seoul, South Korea ALEXANDER S. ROESLER • School of Medicine, Duke University, Durham, NC, USA; Mayo Clinic, Scottsdale, AZ, USA EMILY M. ROY • Division of Infectious Diseases, Boston Children’s Hospital, Harvard Medical School, Boston, MA, USA LUCI´A LARA RUPIL • Centro de Investigacion y Desarrollo en Inmunologı´a y Enfermedades Infecciosas (CIDIE), Consejo Nacional de Investigaciones Cientı´ficas y Te´cnicas

Contributors

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(CONICET)/Universidad Catolica de Cordoba (UCC), Cordoba, Argentina; Facultad de Ciencias de la Salud, Universidad Catolica de Cordoba (UCC), Cordoba, Argentina BHASKAR SAHA • National Centre for Cell Science, Ganeshkhind, Pune, Maharashtra, India; Trident Academy of Creative Technology, Bhubaneswar, Odisha, India JOHN H. SAMPSON • Department of Neurosurgery, Duke University Medical Center, Durham, NC, USA; The Preston Robert Tisch Brain Tumor Center, Duke University Medical Center, Durham, NC, USA LORENA SANCHEZ FELIPE • Virology and Chemotherapy, Molecular Vaccinology & Vaccine Discovery, KU Leuven Department of Microbiology, Immunology and Transplantation, Rega Institute, Leuven, Belgium JAFAR J. SANI • Malaria and Vector Research Group (MVRG), Biotechnology Research Center (BRC), Pasteur Institute of Iran, Tehran, Iran GEORG SCHRAMM • KU Leuven Department of Imaging and Pathology, Nuclear Medicine and Molecular Imaging, Leuven, Belgium LAURA SELDESLACHTS • KU Leuven Department of Imaging and Pathology, Biomedical MRI/MoSAIC, Leuven, Belgium MARIANELA DEL CARMEN SERRADELL • Centro de Investigacion y Desarrollo en Inmunologı´a y Enfermedades Infecciosas (CIDIE), Consejo Nacional de Investigaciones Cientı´ficas y Te´ cnicas (CONICET)/Universidad Catolica de Cordoba (UCC), Cordoba, Argentina; Laboratorio de Parasitologı´a y Micologı´a, Departamento de Bioquı´mica Clı´nica, Facultad de Ciencias Quı´micas, Universidad Nacional de Cordoba (UNC). Haya de la Torre y Medina Allende, Ciudad Universitaria, Cordoba, Argentina SAPNA SHARMA • Virology and Chemotherapy, Molecular Vaccinology & Vaccine Discovery, KU Leuven Department of Microbiology, Immunology and Transplantation, Rega Institute, Leuven, Belgium SHAILJA SINGH • Special Centre for Molecular Medicine, Jawaharlal Nehru University, New Delhi, India AKRITI SRIVASTAVA • Department of Life Science, School of Natural Sciences, Shiv Nadar University, Dadri, UP, India SUKRIT SRIVASTAVA • Infection Biology Group, Indian Foundation for Fundamental Research, Raebareli, Uttar Pradesh, India SELVAKUMAR SUBBIAN • The Public Health Research Institute Center at New Jersey Medical School, Rutgers University, Newark, NJ, USA ADAM M. SWARTZ • Division of Surgical Sciences, Department of Surgery, Duke University Medical Center, Durham, NC, USA WEI-CHUN TANG • Department of Biology, The Catholic University of America, Washington, DC, USA HENDRIK JAN THIBAUT • Virology and Chemotherapy, Molecular Vaccinology & Vaccine Discovery, KU Leuven Department of Microbiology, Immunology and Transplantation, Rega Institute, Leuven, Belgium; Translational Platform Virology and Chemotherapy (TPVC), KU Leuven Department of Microbiology, Immunology and Transplantation, Rega Institute, Leuven, Belgium SUNIL THOMAS • Lankenau Institute for Medical Research, Wynnewood, PA, USA FERNANDO J. TORRES-VELEZ • Division of Infectious Diseases, Wadsworth Center, New York State Department of Health, Albany, NY, USA IVAN TRUS • Vaccine and Infectious Disease Organization-International Vaccine Centre (VIDO-InterVac), University of Saskatchewan, Saskatoon, SK, Canada

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Contributors

DANIEL UDENZE • Vaccine and Infectious Disease Organization-International Vaccine Centre (VIDO-InterVac), University of Saskatchewan, Saskatoon, SK, Canada GRETA VAN SLYKE • Division of Infectious Diseases, Wadsworth Center, New York State Department of Health, Albany, NY, USA GREETJE VANDE VELDE • KU Leuven Department of Imaging and Pathology, Biomedical MRI/MoSAIC, Leuven, Belgium BIRGIT WEYNAND • KU Leuven Department of Imaging and Pathology, Division of Translational Cell and Tissue Research, Leuven, Belgium GARY WONG • Institut Pasteur of Shanghai, Chinese Academy of Sciences, Shanghai, China; De´partement de microbiologie-infectiologie et d’immunologie, Universite´ Laval, Que´bec, QC, Canada ZIYANG XU • Vaccine & Immunotherapy Center, The Wistar Institute, Philadelphia, PA, USA HEMN YOUSEFI • Malaria and Vector Research Group (MVRG), Biotechnology Research Center (BRC), Pasteur Institute of Iran, Tehran, Iran FARAZ ZAIDI • Vaccine & Immunotherapy Center, The Wistar Institute, Philadelphia, PA, USA FRED ZEPP • Department of Pediatrics, University Medical Center Mainz, Mainz, Germany FAN ZHANG • Division of Infectious Diseases, Boston Children’s Hospital, Harvard Medical School, Boston, MA, USA JINGEN ZHU • Department of Biology, The Catholic University of America, Washington, DC, USA SHUBHRANSHU ZUTSHI • National Centre for Cell Science, Ganeshkhind, Pune, Maharashtra, India

Part I Vaccines: Introduction

Chapter 1 Challenges for Vaccinologists in the First Half of the Twenty-First Century Sunil Thomas, Ann Abraham, Patrick J. Callaghan, and Rino Rappuoli Abstract The COVID-19 pandemic of 2020–2021 has highlighted the importance of vaccines and vaccination in human health. The pandemic has resulted in social distancing, travel restrictions, decreased trade, high unemployment, commodity price decline, and financial stress that has impacted the global economy. Since December 2020, a massive vaccination campaign is undergoing in every country on the planet to protect against SARS-CoV-2. Vaccination is the cheapest health-care interventions that can save more lives than any other drugs or therapies. Some of the common diseases of the twentieth century including smallpox and polio are seldom reported due to intense vaccination programs that eradicated it. Smallpox is completely eradicated globally; whereas, polio is confined to only a couple of countries. Vaccination has not only improved the health of man but also improved food security by preventing diseases in farm animals and aquacultured fish. Awareness of the principles of immunology and novel vaccines has led to effective vaccination strategies. Climate change could lead to generation of new strains of infectious microorganisms that would require development of novel vaccines. Recent years have seen the increase in incidence of braineating amoeba and flesh-eating bacteria (necrotizing fasciitis). There are no vaccines for these diseases. Though vaccination programs have eradicated several diseases and increased the quality of life, there are several diseases that have no effective vaccines. Currently there are no vaccines for cancer, neurodegenerative diseases, autoimmune diseases, as well as infectious diseases like tuberculosis, AIDS, and parasitic diseases including malaria. Spontaneous evolution of pathogenic microorganisms may lead to pandemics that impact the health of not only humanity but also other animals. Hence, the challenge to vaccinologists is the development of novel vaccines and vaccination strategies within limited time period and using minimum resources. In addition, the vaccine developed should be administered globally within a short duration so as to prevent generation of pathogenic variants more lethal than the parent strain. Key words Climate change, Infectious disease, Vaccine, COVID-19, Vaccine challenge

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Introduction Infectious diseases are disorders caused by microorganisms including bacteria, viruses, fungi, or parasites that are passed, directly or indirectly, between people or due to exposure from infected animals. Infectious diseases are a leading cause of death worldwide, particularly in low-income countries, especially in young children.

Sunil Thomas (ed.), Vaccine Design: Methods and Protocols, Volume 1: Vaccines for Human Diseases, Methods in Molecular Biology, vol. 2410, https://doi.org/10.1007/978-1-0716-1884-4_1, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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The current pandemic COVID-19 caused by SARS-CoV-2 (started in December 2019) has led to millions of infections and deaths within 1 year of its occurrence. The incidence of death due to COVID-19 was more in developed countries compared to developing countries. Millions of deaths, travel restrictions, decreased trade, high unemployment, and financial stress have impacted the global economy; having an effective vaccine earlier would have resulted in a different scenario. Vaccinologists raced against time to develop and test COVID-19 vaccines; by December 2020, they were ready to be administered globally. The importance of vaccine in the health and well-being of the society was confirmed during the COVID-19 pandemic. Vaccines have revolutionized healthcare since the mid-twentieth century. Though less than 1% of microbial species are human pathogens, infectious diseases are still major cause of human morbidity and mortality, accounting for tens of millions of deaths each year [1, 2]. The number of deaths inflicted by microorganisms and the number of individuals maimed from infectious disease would actually be much higher in a world without vaccinations [3, 4]. Millions of deaths are prevented annually due to the reduced incidence of infectious diseases such as measles, polio, diphtheria, pertussis, tetanus, and whooping cough [4–8]. Undoubtedly, vaccines are among the most important scientific and public health achievements since their discovery over 200 years ago [3]. Despite the usage and popularity of vaccines in developed countries, one-third of all deaths in several developing countries occur in children, and infectious diseases are the leading cause of these deaths [9]. Hence, there is a need for mass vaccination programs benefitting children of these countries to prevent curable infectious disease. The COVID-19 pandemic caused by SARS-CoV-2 has opened new challenges to the vaccinologists. There is a need to quickly develop novel vaccines against new and emerging diseases with limited resources. Several diseases including AIDS, malaria, that are yet to have vaccines require new strategies for vaccine development. This chapter states the challenges that will be faced by vaccinologists in the first half of the twenty-first century. 1.1 Change and Emerging Infectious Diseases

Climate change will be the greatest threat to humanity in the twenty-first century. Climate change is associated with increase in temperature, changes in weather patterns, increase in sea level, frequency of hurricanes and cyclones, increase or decrease in rain fall patterns depending on the region, forest fires, expansion of desertification, salinization of cultivable lands, decrease in snow, and destruction of glaciers and ice shelves. Climate change will impact food security, availability of drinking water, flooding of coastal areas, air pollution, and may also lead

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to mass migration. Lack of fresh water would lead to use of contaminated water that may lead to waterborne diseases. Changes in the weather and temperature have an impact on insect vectors. Increase in temperature leads to increase in mosquitos thereby increasing the range of diseases like malaria, dengue, and Zika virus. The population of ticks is controlled during winter season; however, a warm winter season would not regulate the tick population [10], thereby increasing incidence of Lyme disease, rickettsiosis, and ehrlichiosis. Changes in rain fall patterns especially in the desert regions that are breeding grounds for locust could lead to increase in locust population that threatens the global food security. The recent locust swarms (year 2020) extended from East Africa to India [11] and devastated crops in the region. Hunger and malnutrition weaken the immune system; people subjected to starvation or on low nutrient diet are more prone to infectious diseases [12, 13]. 1.2 Development of Vaccines to Protect against COVID-19

Severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) is responsible for the disease COVID-19 that has decimated the health and economy of our planet. The virus causes the disease not only in people but also in companion and wild animals. People with diabetes are at risk of the disease. As yet we do not know why the virus has been highly successful in causing the pandemic within 3 months of its first report [14, 15]. The disease was first reported in Wuhan, China, in December 2019 [16]. As of March 2021, 116 million people are infected with COVID-19 with 2.5 million deaths. The unofficial figures may be many times higher. COVID-19 is regarded as a respiratory disease that manifests with fever, cough, shortness of breath or difficulty breathing, chills, muscle pain, headache, sore throat, and loss of taste and smell. Other symptoms include diarrhea, nausea, and vomiting. Many patients with the COVID-19 are asymptomatic but are able to transmit the virus to others [14, 15]. The prolonged pandemic has resulted in social distancing, travel restrictions, decreased trade, high unemployment, commodity price decline, and financial stress that has impacted the global economy. COVID-19 disease is caused by the severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2), a member of the betacoronavirus genus. The disease has resulted in a mortality of 0.5–8.0%. Several factors influenced the death rate in people with COVID-19. Age, health, and behavior of the population impacted the death rate due to COVID-19. Old people and people with underlying diseases such as diabetes, lung diseases (due to smoking), liver disease, cardiovascular disease, and obesity are more prone to death due to COVID19. As yet, there are no effective drugs available for treatment of the disease [14]. The major structural proteins of SARS-CoV-2 are spike (S), membrane (M), envelope (E), and the nucleocapsid (N) proteins [14]. The spike protein of SARS-CoV-2 uses the host angiotensin-

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converting enzyme 2 (ACE2) as the entry receptor. Hence, the research community was interested in studying the S protein for vaccine development. There are several variants of COVID-19 vaccines. The vaccines developed by companies are based on mRNA, adenovirus vector, or live attenuated. The efficiency of the vaccines ranges from 50% to 95%. The mRNA vaccines were found to be more efficient than conventional vaccines. Vaccines were administered beginning December 2020. Most countries did not strictly enforce social distancing. This has resulted in quick spread of the virus and its mutation. Viruses generally acquire mutations over time, giving rise to new variants. Several variants of the SARS-COV-2 are reported recently. Lineage B.1.1.7 is also known as the UK variant. B.1.1.7 variant may be associated with an increased risk of death compared with other variants. This variant has a mutation in the receptor binding domain (RBD) of the spike protein at position 501, where the amino acid asparagine (N) has been replaced with tyrosine (Y). The 501Y.V2 variant (lineage B.1.351) is the South African strain. This variant has multiple mutations in the spike protein, including K417N, E484K, and N501Y. Some of the vaccines may not be protected by this variant. P.1 is a close relative of the B.1.351 lineage and is the Brazil variant. This variant has 17 unique mutations, including three in the receptor binding domain of the spike protein. Lineage B.1.429 is the California variant (CAL.20C). The S protein L452R mutation is within a known receptor binding domain that has been found to be resistant to certain S protein monoclonal antibodies [17]. The vaccines currently developed do not protect against all the variants. Hence, the challenge is to develop vaccines that could protect against all the variants of SARS-CoV-2. The COVID-19 vaccine was developed during lockdown conditions with minimum support. The Universities and laboratories were under lockdown during the vaccine development. Companies had to use their resources to achieve their goals. Pfizer BioNTech and Moderna had to use a never tested vaccine technology— mRNA—to develop an effective COVID-19 vaccine. The mRNA vaccine was successful because there was prior knowledge on the use of the strategy for overexpression of proteins and use of technology to protect against diseases [18–21]. Companies and research organizations engaged in basic and translational research, collaborate effectively and have a team that could perform tasks with minimum support and at minimum time has a cutting edge in developing new and innovative vaccines. 1.3 Development of Vaccines to Protect against HIV

Acquired immunodeficiency syndrome (AIDS) caused by human immunodeficiency virus (HIV) has caused millions of infections and deaths since early 1980s. Despite the countless efforts of research, there is yet to develop a vaccine or drug that can

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successfully prevent or eradicate the disease. It is a continual raging pandemic that is affecting millions of people globally. Approximately, 38 million people are living with HIV around the globe [22]. Tiredness and fatigue are common problems among people with HIV. Other symptoms of HIV include lack of energy, sleep disturbance, anxiety, and depression [23]. AIDS costs billions of dollars in lost earnings, largely as a result of the deaths of hundreds of thousands of workers that could have been prevented with effective treatment. Lack of energy and fatigue in people with AIDS contribute to loss of income. The structure of the HIV virus is uniquely complex due to its mutability and genetic diversity, which in turn adds to the challenge of inducing an immune response in the host. HIV-1, is an RNA virus, has been estimated to have the highest mutation rate of any biological product [24, 25]. This introduction of antigenic variation limits recognition of the virus by the immune system, and this issue is compounded by antibodies having minimal accessibility and specificity for native envelope proteins on virus’ surface [26]. Further, infected cells that express HIV have reduced expression of MHC class I on their surface, causing these infected cells to be resistance to cytolysis, effectively providing a hideout for the virus [26]. As yet there are no vaccines to protect against HIV. There is an urgent need to develop a vaccine against the life-threatening disease.

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Development of Vaccines for Flaviviruses The Flaviviridae family consists of 70 different viruses that are arthropod-borne viruses and transmitted by mosquitoes or ticks. The mosquitoes-transmitted viruses include yellow fever, dengue fever, Japanese encephalitis, West Nile virus, and Zika virus. The flaviviruses transmitted by ticks are responsible for encephalitis and hemorrhagic diseases and include tick-borne encephalitis (TBE), Kyasanur Forest disease (KFD) and Alkhurma disease, and Omsk hemorrhagic fever [27]. The major flaviviruses of economic importance include the West Nile virus (WNV), dengue, and Zika virus, and none of them have vaccines for humans. The West Nile virus is known to infect mosquitoes, birds, horses, and humans. It is a member of the Flavivirus family and is spread through infected mosquitoes [28]. Despite the significant health risk, including three million infections in the United States, there are no vaccines for the disease. Approximately 80% of cases are asymptomatic, which makes it difficult to diagnose patients who are infected [29]. Several vaccine candidates are in various stages of clinical development; however, demonstrating efficacy and the unpredictability of WNV outbreaks pose a challenge for securing an effective vaccine. ChimeriVAX-

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WN02 is a promising candidate with  96% seroconversion rates observed during phase II of clinical trials. Another phase II clinical trial observed a  92% seroconversion rates in all tested doses for 50 year olds and older. Other phase I clinical trials include WNV-DENV chimeric vaccine, a naked DNA vaccine, and WN-80E, which have been completed. These candidates have shown promising immunogenic properties in humans; however, a major hurdle is the cost-effectiveness of developing a vaccine for WNV. Due to the low incidence rates and inconsistent activity, it would not be beneficial for pharmaceutical companies to introduce WNV routine immunization [30]. The West Nile virus is increasing in vulnerable populations; therefore, it is essential for vaccinologists to reconsider the cost-effectiveness and determine the importance of immunizing humans. Zika virus emerged slowly yet made an outstanding impact in the lives of many. The virus was first found in a Rhesus monkey and in mosquitoes in the Zika forest (Uganda) in 1947 and 1948. In 1952, the first human case was reported [28]. It is primarily transmitted by mosquitoes. The Zika virus caught the world’s attention in 2007 when it caused an epidemic on Yap Island, Micronesia. Later in 2013–2014, large epidemics arose in French Polynesia and other regions of the South Pacific. In May 2015, the World Health Organization reported the first local transmission of Zika virus in the Region of the Americas (Americas), with autochthonous cases identified in Brazil. In December, the Ministry of Health estimated that 440,000–1,300,000 suspected cases of Zika virus (ZIKV) disease had occurred in Brazil in 2015. By January 20, 2016, locally transmitted cases had been reported to the Pan American Health Organization from Puerto Rico and 19 other countries or territories in the Americas [31]. ZIKV infection is linked with activation of Guillian-Barre´ syndrome (GBS) in adults infected with the virus and microcephaly in infants following maternal infection [32]. Association between ZIKV infection and microcephaly via vertical transmission of virus from pregnant mother to her fetus was illustrated based on an increased incidence of microcephaly possibly pertaining to ZIKV outbreak, presence of ZIKV in microcephalic fetal brain as well as the presence of viral RNA in the amniotic fluid of pregnant women [33–35]. Zika-related birth defects are also linked to pregnancy losses, which include miscarriages and abortions (CDC) [32]. Once infected, the disease symptoms include fever, rash, headache, joint pain, red eyes, and muscle pain. The World Health Organization declared the Zika virus (ZIKV) as a Public Health Emergency of International Concern in 2016 [36]. The virus has infected approximately 1.5 million people. ZIKV stems from two genetic lineages (African and Asian) with 35,000 children, ages 2–16 years in 10 dengue-endemic countries in Asia and the Americas [38]. Dengvaxia is efficacious in seropositive individuals, but increases the risk for severe dengue in seronegative persons about 2 years after administration of the first dose. In a phase IIb trial in Thailand, 2.8% of children vaccinated at ages 2–5 years and 1.4% of children vaccinated at age 9 or above were hospitalized for breakthrough dengue infections [39]. For countries considering the introduction of Dengvaxia, WHO recommends a prevaccination screening strategy whereby only persons with evidence of a past dengue infection would be vaccinated [40]. Dengvaxia, lacking DENV nonstructural protein antigens, does not protect seronegatives because it failed to raise a competent

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T-cell response and/or antibodies to NS1 [39]. Dengvaxia was used in the Philippines to vaccinate 9–10-year-old school children, living in areas highly endemic for dengue. After about 830,000 had received at least 1–3 recommended doses, risks of enhanced disease in dengue-naı¨ve vaccinees were reported [41]. Several types of dengue vaccines are in the developing stages, and some are under preclinical and clinical evaluation [30]. Takeda’s tetravalent dengue vaccine candidate (TAK-003) is based on a liveattenuated DENV-2 virus that provides the genetic backbone for all four of the vaccine viruses. TAK-003 demonstrated continued benefit independent of baseline serostatus in reducing dengue with some decline in efficacy during the second year [42]. Biswal et al. (2020) [43] demonstrated that TAK-003 has an acceptable safety profile in healthy children aged 4–16 years and is efficacious in the prevention of symptomatic dengue disease in both individuals who are dengue-naive and those previously exposed. Efficacy varied against individual serotypes, with an overall efficacy of 66% in individuals who were dengue-naive and 76% in those who were preexposed. In addition, TAK-003 reduced the number of dengue cases that were hospitalized by 90% along with an 86% reduction in dengue hemorrhagic fever. These data represent a major step forward in the development of an effective and safe dengue vaccine for use in people of all ages, irrespective of previous dengue exposure at the time of vaccination. The immunogenicity and protective efficacy of a candidate tetravalent dengue virus purified inactivated vaccine (TDENV PIV) formulated with alum or an Adjuvant System (AS01, AS03, or AS04) was evaluated in a 0, 1-month vaccination schedule in rhesus macaques. When tested in a nonhuman primate infection model, adjuvanted TDENV PIV vaccine formulations showed an acceptable safety profile and were highly immunogenic, with all formulations inducing robust and persistent neutralizing antibody responses against each of the four DENV serotypes [44]. However, currently the only available WHO-approved dengue vaccine is Dengvaxia.

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Development of Vaccines for Norovirus Norovirus, also known as Norwalk virus, is responsible for gastrointestinal distress or commonly known as the “stomach flu.” The first recorded outbreak of the norovirus was in Norwalk, Ohio in 1968, where 50% of the elementary students were infected. Symptoms of the virus are vomiting, diarrhea, and a low fever [45]. Noroviruses are the leading cause for gastroenteritis spanning all age groups. Outbreaks of noroviruses are likely to occur in close contact communities, such as schools, prisons, hospitals, hotels, or cruise ships [28].

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Norovirus is the leading cause of acute gastroenteritis (AGE) worldwide, and in the United States, norovirus is estimated to cause 19–21 million illnesses, 1.7–1.9 million outpatient visits, 56,000–71,000 hospitalizations, and 570–800 deaths annually [46]. The main mode of transmission is the fecal–oral route which can be through food, water, and environmental contamination. The direct contamination of food during production and contamination of food during preparation have been responsible for foodborne transmission of the virus. Raw fruits, vegetables, and shellfish are main risks of infection. Waterborne transmission of the virus may be due to municipal water systems, commercial ice consumption, water sources at camps, and recreational water exposure. The general population is prone to acquire the viral infection, but the majority of deaths occurs at extreme ages, such as infants and the elderly. Although prevention (good hygiene, washing fruits and vegetables thoroughly, and cooking seafood properly) is the only approach to limit infection, a vaccine is ideal to encourage an immune response in humans. The main obstacles facing vaccine development for noroviruses are the lack of an animal model that resembles the human disease, the viral and human host diversity, and virus evolution [45]. Another major barrier to studying the pathogenesis, virus–host interactions, and effect of control measures to prevent and treat norovirus gastroenteritis has been the lack of a robust and reproducible cell culture system [47]. Vaxart, a US biotechnology company, is developing an oral vaccine platform to protect against norovirus. This tablet vaccine comprises a nonreplicating adenovirus-based vector expressing the VP1 gene from the GI.1 norovirus strain and a double-stranded RNA adjuvant. Sixty-six adult subjects meeting inclusion/exclusion criteria were randomized 2:1 to receive a single vaccine dose or placebo, respectively. This oral norovirus vaccine was well-tolerated and generated substantial immune responses, including systemic and mucosal antibodies as well as memory IgA/IgG. These results are a major step forward for the development of a safe and immunogenic oral norovirus vaccine [48]. Treanor et al. (2020) [49] investigated the safety and immunogenicity of bivalent virus-like particle (VLP) vaccine candidate formulations with and without monophosphoryl lipid A (adjuvant MPL). Adults over 60 years of age displayed no safety concerns and had similar immune responses to the norovirus VLP vaccine candidate as younger adults, unaffected by increasing age, a second dose, or inclusion of MPL.

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Development of Vaccines for Influenza Influenza is an infectious respiratory disease; in humans, it is caused by influenza A (genus influenzavirus A) and influenza B (genus influenzavirus B) viruses. Symptoms associated with influenza virus

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infection vary from a mild respiratory disease confined to the upper respiratory tract and characterized by fever, sore throat, runny nose, cough, headache, muscle pain, and fatigue to severe and in some cases lethal pneumonia owing to influenza virus or to secondary bacterial infection of the lower respiratory tract. Influenza A viruses cause flu pandemics. A pandemic can occur when a new and very different influenza A virus emerges that both infects people and has the ability to spread efficiently between people [50]. The WHO estimates that annual epidemics of influenza result in ~1 billion infections, 3–5 million cases of severe illness and 300,000–500,000 deaths. The severity of pandemic influenza depends on multiple factors, including the virulence of the pandemic virus strain and the level of preexisting immunity. The most severe influenza pandemic, in 1918, resulted in >40 million deaths worldwide. Influenza vaccines are formulated every year to match the circulating strains, as they evolve antigenically owing to antigenic drift. Nevertheless, vaccine efficacy is not optimal and is dramatically low in the case of an antigenic mismatch between the vaccine and the circulating virus strain [50]. Influenza viruses are members of the Orthomyxoviridae family and are enveloped negative-sense single-strand RNA viruses with a segmented genome. Influenza A and influenza B viruses contain eight RNA segments, which encode RNA polymerase subunits, viral glycoproteins (namely, hemagglutinin (HA), with its distinct globular “head” and “stalk” structures, which facilitate viral entry, and neuraminidase (NA), which facilitates viral release), viral nucleoprotein (NP), matrix protein (M1) and membrane protein (M2), the nonstructural protein (NS1), and nuclear export protein (NEP). The HA and NA viral proteins are the most antigenically variable, and in the case of influenza A virus, they are classified into antigenically diverse subtypes. These two viral glycoproteins are located at the surface of the virus particle and are the main targets for protective antibodies induced by influenza virus infection and vaccination [50]. There are 18 different hemagglutinin subtypes and 11 different neuraminidase subtypes (H1 through H18 and N1 through N11, respectively). While there are potentially 198 different influenza A subtype combinations, only 131 subtypes have been detected in nature. Current subtypes of influenza A viruses that routinely circulate in people include A(H1N1) and A(H3N2) [51]. The challenges in developing an influenza vaccine include the dependence on embryonated eggs for vaccine production, the lengthy timeline for vaccine production, the need for annual vaccination, the emergence of antigenically novel viruses, the need for improved immunogenicity in the elderly, and the need for an improved correlate of protection. The ultimate goal of a universal influenza vaccine is to protect against all influenza A viruses, obviating the need for annual revaccination. Influenza vaccines must protect all age groups, particularly those most vulnerable to

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complications of severe influenza. Ideally, new vaccines should increase the breadth of the immune response to include antigenically distinct viruses within the same subtype and viruses of other subtypes, should not be manufactured in eggs, and should require less time to manufacture than currently licensed technologies [52]. Strategies employed to improve vaccine efficacy involve using structure-based design and nanoparticle display to optimize the antigenicity and immunogenicity of target antigens; increasing the antigen dose; using novel adjuvants; stimulating cellular immunity; and targeting other viral proteins, including neuraminidase, matrix protein 2, or nucleoprotein [53].

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Development of Vaccines for Sepsis Sepsis (septicemia) is a medical emergency that describes the body’s systemic immunological response to an infectious process that can lead to end-stage organ dysfunction and death [54]. The global epidemiological burden of sepsis is, however, difficult to ascertain. It is estimated that more than 30 million people are affected by sepsis every year worldwide, resulting in six million deaths annually. Mortality rates from sepsis are approximately 41% in Europe versus approximately 28.3% in the United States [55]. There are currently no effective pharmacological treatments for sepsis, making early recognition, resuscitation, and immediate treatment with appropriate antibiotics the key to reducing the burden of resulting disease. The majority of sepsis, around 70–80%, is community acquired making emergency departments and primary care key targets to improve recognition and early management. Case fatality rates for sepsis are decreasing in many countries with the reduction attributed to national or regional screening and quality improvement programs focused on early identification and immediate treatment. Sepsis, now defined as life-threatening organ dysfunction due to a dysregulated host response to infection, was recently recognized by the World Health Organization as a global health priority. Sepsis causes or contributes to up to half of all in-hospital deaths in the USA [56]. Sepsis may be due to bacterial, viral, mycoplasma, or fungal pathogens. The common species causing sepsis includes Pneumococcus, Staphylococcus, Mycoplasma, Legionella, Escherichia coli, Klebsiella, Enterococcus, Candida, Streptococcus, methicillin-sensitive Staphylococcus aureus and Neisseria sp. [56]. GBS septicemia is caused by the bacterium Streptococcus agalactiae, which is commonly called group B strep, or GBS. GBS causes sepsis in newborns. GBS can also cause serious infections in adults that include bloodstream infections, pneumonia, skin and soft-tissue infections, and bone and joint infections [57].

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Secretion of extracellular vesicles (EVs), a process common to eukaryotes, archaea, and bacteria, represents a secretory pathway that allows cell-free intercellular communication. EVs purified from a S. aureus mutant that is genetically engineered to express detoxified cytolysins are immunogenic in mice, elicit cytolysinneutralizing antibodies, and protect the animals in a lethal sepsis model [58]. Gram-negative bacterial lipopolysaccharide (LPS, endotoxin) is an initiator of sepsis. Vaccines directed against core LPS structures that are widely conserved among Gram-negative bacteria could be used in the prevention of sepsis. Killed whole bacterial vaccines (E. coli O111:B4, J5 [Rc chemotype] mutant and S. minnesota, Re chemotype) protected mice against experimental sepsis [59].

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Development of Vaccines for Tuberculosis Tuberculosis (TB) is a detrimental disease that has affected millions and is the leading cause of death due to an infectious agent. The infectious agent Mycobacterium tuberculosis was discovered in 1882. The only vaccine effective for TB is the Bacillus CalmetteGuerin (BCG) vaccine; however, it is only effective in infants and children [60]. Therefore, there is a dire need to develop a vaccine that is universally effective. It is suggested that a 60% effective vaccine which will provide protection for 10 years could prevent 17 million cases of TB between the years 2040 and 2050. Despite intense efforts, designing a vaccine for TB is difficult. The immunological correlates of protection have not been clarified, the preclinical animal models pose uncertainty, and M. tuberculosis has the ability to evade host immunity. In 2019, many vaccine candidates have been analyzed in clinical trials. The recombinant strains of BCG and the attenuated strains of M. tuberculosis have been tested to replace BCG vaccination in infancy. Protein and adjuvant combinations and recombinant viral vector subunit vaccines have been designed to increase the efficacy of neonatal BCG vaccinations [61]. TB vaccine development has been hindered by the lack of animal models accurately predicting the effects in humans and the lack of knowledge of the M. tuberculosis epitopes. A phase 2b trial of the M72/AS01E tuberculosis vaccine conducted in different countries of Africa. Human immunodeficiency virus (HIV)-negative adults 18–50 years of age with latent M. tuberculosis infection (by interferon-γ release assay) were randomly assigned (in a 1:1 ratio) to receive two doses of either M72/AS01E or placebo intramuscularly 1 month apart. Most participants had previously received the Bacillus Calmette-Gue´rin vaccine. The authors assessed the safety of M72/AS01E and its efficacy against progression to bacteriologically confirmed active pulmonary tuberculosis disease. Clinical suspicion of tuberculosis

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was confirmed with sputum by means of a polymerase chain reaction test, mycobacterial culture, or both. M72/AS01E provided 54.0% protection for M. tuberculosis-infected adults against active pulmonary tuberculosis disease, without evident safety concerns [62]. A further study was conducted among adults infected with M. tuberculosis. Vaccination with M72/AS01E elicited an immune response and provided protection against progression to pulmonary tuberculosis disease for at least 3 years [63]. H4:IC31 (AERAS-404) is a field-reconstituted vaccine with H4 antigen (Sanofi Pasteur) and IC31® proprietary adjuvant (Valneva, formerly Intercell) supplied in different vials. H56:IC31 is a vaccine with the H56 antigen (Statens Serum Institut; SSI) formulated in IC31® adjuvant. H4:IC31 has an acceptable safety profile, immunogenic, and capable of triggering multifunctional CD4+ T cell responses in previously BCG-vaccinated healthy individuals [64]. A trial of BCG revaccination and vaccination with H4:IC31, in South African adolescents, showed efficacy in preventing Mycobacterium tuberculosis infection. BCG revaccination administered as a single-dose ID and both H4:IC31 and H56:IC31 administered as 2 doses IM had acceptable safety profiles in healthy, QFT-negative, previously BCG-vaccinated adolescents [65].

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Development of Vaccines for Tick-Borne Diseases Increases in tick-borne disease prevalence and transmission are important public health issues [66]. Lyme disease, caused by Borrelia burgdorferi, is a major tick-borne disease. Borrelia mayonii are a type of bacteria recently found in North America that can cause Lyme disease [67]. There is currently no vaccine available to prevent Lyme borreliosis in humans. Borrelia outer membrane proteins have been investigated as vaccine candidates but are not successful yet. The other tick-borne diseases include ehrlichiosis (caused by Ehrlichia sp.), babesiosis (caused by the parasite Babesia sp.), Rocky Mountain spotted fever (caused by Rickettsia rickettsia), tularemia (caused by Francisella tularensis), and anaplasmosis (cause by Anaplasma sp.). These diseases are not diagnosed properly and if untreated may be fatal. The diseases caused by these pathogens significantly lower health quality status, may lead to impairment in their ability to work, increased utilization of healthcare services, and greater out of pocket medical costs. Development of vaccines could provide protection against these diseases.

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Development of Vaccines for Flesh-Eating Bacteria Flesh-eating bacteria, also known as necrotizing fasciitis (NF), is a life-threatening infection which infects the subcutaneous tissue, fascia, and muscles. It is responsible for approximately 500,000 deaths annually. Inflammation is the initial sign of NF and then it progresses to skin necrosis that causes deep tissue damage. Escherichia coli, Group A streptococcal infections, methicillin-resistant Staphylococcus aureus, methicillin-sensitive Staphylococcus aureus, and fungal infections are the major bacterial organisms responsible for NF. Those who are infected by flesh-eating bacteria experience systemic toxicity with multiorgan failure, which may result in death. Currently, treatment for flesh-eating bacteria is surgery, antibiotic therapy, skin grafting, reconstructive surgery, negative pressure wound therapy, and even amputation of limbs. The severity of flesheating bacteria on patients is immensely painful and calls for an urgent need to design a vaccine for patients. A vaccine will provide an effective and direct route of elimination without the need for expensive and time-consuming procedures [68].

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Development of Vaccines for Parasites The parasites include ectoparasites, like ticks, mosquitoes, fleas, and itch mite, and endoparasites including Plasmodium, Entamoeba, Leishmania, Trypanosoma, Babesia, Toxoplasma, Wuchereria, Brugia, Giardia, Ascaris, tapeworm, hookworm, pinworm, whipworm, Onchocerca, Fasciola, and Schistosoma. Most of the diseases are classified under neglected tropical diseases and are the major causes of fatality in poverty-stricken regions of the developing world. Though the diseases caused by these parasites affect millions of people in the developing world, in the long term (due to climate change, movement of refugees, etc.) they pose a risk to people all over the world. As yet there are no vaccines against these parasites. Hence there is an urgent need to develop vaccines against these parasites causing misery to millions of people [8].

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Development of Vaccines for Malaria Malaria has affected millions of people over the centuries and has caused 410,000 deaths in 2019 [69]. The protozoan parasite causing malaria stems from the Plasmodium species, which is native to subtropical and tropical regions. Severe and fatal malaria are known to be caused by P. falciparum. An infected female Anopheles mosquito is the causative agent that initiates infection. When the mosquito bites into the human, it injects sporozoite into the skin,

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where it travels through the blood stream to the liver. The sporozoite travels by multiple cells and then resides in a hepatocyte. The sporozoite grows into 40,000 merozoites that invade red blood cells and erythrocytes, which eventually leads to erythrocyte invasion and growth in the blood stream [70]. The epidemiology of transmission varies in different regions, villages, and even personto-person. Therefore, it is challenging to develop a vaccine that caters to the severity of each individual case. Another major challenge facing vaccine development is that many infected people are asymptomatic, who acts as carriers for transmission of malaria parasites. During pregnancy, malaria can cause miscarriages, death of the fetus, decreased birth weight, and premature delivery. Pregnant women are at risk for anemia and sometimes even death. The malaria parasite has an extraordinary ability to evade the immune system, which may explain the failure of malaria vaccines to date [71]. Due to the impact of the disease in the developing world, there is an urgent need to develop vaccines to protect against malaria. The circumsporozoite protein (CSP), the major protein expressed on the surface of the infecting sporozoite, is essential for mediating liver infection. A truncated form of CSP is linked to hepatitis B surface antigen (HBsAg) to produce RTS. RTS is co-expressed in yeast cells with HBsAg to produce RTS,S [72, 73]. Despite the advancements in malaria vaccine development, we do not have an efficient vaccine that protects against young and old adults. RTS,S is the first malaria vaccine to provide protection against malaria in children [74]. The RTS,S/AS01 vaccine against malaria infection completed phase III trials in 2014 and demonstrated efficacy against clinical malaria of approximately 36% over 4 years for a four-dose schedule in children aged 5–17 months [75]. Saponins, particularly those obtained from Quillaja saponaria Molina, are known potent adjuvants and Quillaja saponins (QS) have for long been used in animal vaccines. Saponin-based adjuvants can be formulated in different ways; in free form, with aluminum hydroxide, in ISCOMs (immunostimulating complex) or in ISCOM-Matrix/Matrix structures. The ISCOM, a potent adjuvant formulation, consists of stable complexes composed of saponin, cholesterol, phospholipid, and incorporated antigen(s). The hallmarks of the ISCOM technology are the dose-sparing potential, induction of high and long-lasting antibody titers and potent T cell responses. However, later it was shown that antigen incorporation is not critical for these immune properties. Antigen and empty ISCOMs, i.e., ISCOM-Matrix/Matrix could simply be mixed with sustained vaccine efficacy. A novel adjuvant formulation based on two different Matrix particles made from two separate purified fractions of saponins, yielding Matrix-A™ and MatrixC™. These Matrix particles, approximately 40 nm large, are

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subsequently mixed at defined ratios to get the Matrix-M™ adjuvant [76]. Collins et al. (2017) [77] developed a more immunogenic CSP-based RTS,S-like vaccine called R21. The major improvement is that in contrast to RTS,S, R21 particles are formed from a single CSP-hepatitis B surface antigen (HBsAg) fusion protein, and this leads to a vaccine composed of a much higher proportion of CSP than in RTS,S. In animal studies, R21 is immunogenic at very low doses and when administered with the adjuvants Matrix-M, it elicits sterile protection against transgenic sporozoite challenge. In a phase I study, R21 adjuvanted with Matrix-M adjuvant was considered safe and had good immunogenicity even when administered at a fivefold lower 10 μg dose in UK and African populations [78]. A large-scale malaria vaccine implementation program coordinated by the World Health Organization to investigate RTS,S/AS01 efficacy is now ongoing in Malawi, Ghana, and Kenya. The program aims to vaccinate about 360,000 children per year from 2019 to 2023 and will examine safety, compliance with the booster dose, and reduction in mortality [79]. Serum Institute of India (SII) and the US-based biotechnology company, Novavax, Inc. have announced an agreement for a commercial license for the use of Novavax’s proprietary Matrix-M vaccine adjuvant with SII’s malaria vaccine candidate R21 [80].

11 Development of Vaccines for Cancer, Neurodegenerative Diseases, Substance Abuse, and Autoimmune Diseases Cancer is the leading cause of death in the world. Though there are vaccines for some cancers induced by virus (e.g., human papillomavirus (HPV)–related cervical or oropharyngeal cancer; Merkel cell polyomavirus (MCPyV)–related Merkel cell carcinoma (MCC) and Epstein-Barr virus (EBV)–related head and neck cancers), any epitopes derived from open reading frames (ORFs) in the viral genome could contribute to the potential source of antigens [81]. However, there are no vaccines against the majority of cancers including cancer of the lung, breast, colon, pancreas, skin, brain, blood, etc. Development of cancer vaccines should be a priority as it could reduce the incidence of the disease, thereby reducing emotional and economic hardship to millions of people. Tumor neoantigen, or tumor-specific antigen (TSA), is the repertoire of peptides that displays on the tumor cell surface and could be specifically recognized by neoantigen-specific T cell receptors (TCRs) in the context of major histocompatibility complexes (MHCs) molecules. Neoantigens could play a critical role in tumor-specific T cell-mediated antitumor immune response and successful cancer vaccines [81].

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As people live longer, they are more prone to neurodegenerative diseases including Alzheimer’s and Parkinson’s diseases. As yet there are no cures for these diseases. A vaccine to prevent this disease will decrease the enormous burden on society. The currently available medications for the treatment of drug abuse have had only limited success. Antiaddiction vaccines, aimed at eliciting antibodies that block the pharmacological effects of drugs, have great potential for treating drug abuse [82]. As yet there are no vaccines for arthritis, type I diabetes, allergy, multiple sclerosis, and other autoimmune diseases. A vaccine for these diseases could improve the quality of life of people suffering from these debilitating diseases [8].

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Antibody-Dependent Enhancement Vaccines have been one of the most effective public health initiatives in human history. However, not every attempt at introducing immunity is safe. Paradoxically, antibodies produced in response to a vaccine have actually increased susceptibility to illness from infectious agents and dangerous unforeseen consequences have been reported in response to vaccination. Antibodies neutralize pathogens when they block its attachment and/or entry into a cell, thereby reducing infectivity of the pathogen. Paradoxically, nonneutralizing antibodies can actually increase infectivity of pathogens in a process coined antibodydependent enhancement (ADE) or viral infection [83]. One theory for how this could occur involves antibodies produced in a prior infection or vaccination having affinity for epitopes on a related— but serologically distinct—pathogen and facilitating FcR-mediated endocytosis of this related pathogen into hose immune cells, suppressing antiviral pathways, and ultimately triggering a proinflammatory cytokine storm to exacerbate severity of the infection [83]. This paradox has contributed to futile vaccine development attempts for various flaviviruses, coronaviruses, and immunodeficiency viruses. Dengue virus (DENV) represents perhaps the most infamous example of ADE among flaviviruses. Young infants and children with antibodies for one DENV serotype (produced during a prior, subclinical infection, or passively introduced from their mother) were observed to suffer severe illness upon infection with a different DENV serotype [84, 85]. This phenomenon also showed up in vaccine trials. For example, in the CYD14 phase 3 clinical trial of a DENV vaccine, participants aged 2–5 that received the vaccine were found to be 7.45 times more likely to be hospitalized for virologically confirmed dengue compared to control after 3 years [86].

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Antibodies against related flaviviruses such as dengue virus (DENV) and West Nile virus (WNV) can cross-react with Zika virus (ZIKV) and could thereby increase disease severity. ADE may explain the severe disease manifestations associated with ZIKV outbreaks and highlights the need to exert caution when designing flavivirus vaccines [87]. Bardina et al. (2017) [87] showed that low titers of DENV and WNV antibodies enhanced ZIKV viremia. Similarly, immunodeficiency viruses have demonstrated ADE in multiple species. Antibodies produced in cats in response to FIV vaccine have been shown to enhance infection in subsequent infections [88]. ADE has been observed in macaques infected with SIV as well [89]. HIV-1 has increased infectivity in vitro in the presence of HIV-1 antibodies [90]. Demonstration of ADE in vaccine clinical trials for HIV further complicates this paradox. For example, attenuated recombinant adenovirus-5 (Ad5) expressing HIV genes (gag, pol, and nef) increased HIV-1 infection rate among participants with high Ad5 antibodies prior to the vaccination [91]. It is unclear how these Ad5 antibodies could contribute to HIV-1 infectivity but results like these highlight the complex challenges faced by vaccinologists in ensuring their products are not only effective in stimulating neutralizing antibodies but also safe.

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Future Challenges The COVID-19 pandemic showed the need for developing and manufacturing vaccines with minimum resources and within a short time frame. The pandemic led to the development of mRNA vaccines that were never developed commercially before. The mRNA vaccines developed were highly efficient than the conventional vaccines. The novel vaccines developed showed the importance of investing in basic and translational research as well as the need for collaboration between academia and industries. With climate change and the increase in temperature and the movement of people, there may be pandemics in the future. The scientific and management knowledge of the current pandemic may be helpful in tackling future pandemics. There is a need to develop new adjuvants to induce the immune system and make the vaccine highly potent. There is also a need to develop new classes of vaccines (DNA, RNA, VLPs, AAV vector, structure-based vaccines) that are more efficient in inducing immunity. Use of artificial intelligence (AI), deep learning, and bioinformatic software will accelerate the design of efficient vaccine candidates by predicting better immunogenic regions that may require fewer tests before clinical trials.

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36. Gan´czak M (2016) Zika virus infections from the perspective of the general practitioner. Family Med Primary Care Rev 4(4):487–491 37. Modjarrad K, Lin L, George SL, Stephenson KE, Eckels KH, De La Barrera RA, Jarman RG, Sondergaard E, Tennant J, Ansel JL, Mills K, Koren M, Robb ML, Barrett J, Thompson J, Kosel AE, Dawson P, Hale A, Tan CS, Walsh SR, Meyer KE, Brien J, Crowell TA, Blazevic A, Mosby K, Larocca RA, Abbink P, Boyd M, Bricault CA, Seaman MS, Basil A, Walsh M, Tonwe V, Hoft DF, Thomas SJ, Barouch DH, Michael NL (2018) Preliminary aggregate safety and immunogenicity results from three trials of a purified inactivated Zika virus vaccine candidate: phase 1, randomised, double-blind, placebo-controlled clinical trials. Lancet 391 (10120):563–571 38. Hadinegoro SR, Arredondo-Garcı´a JL, Capeding MR, Deseda C, Chotpitayasunondh T, Dietze R, Muhammad Ismail HI, Reynales H, Limkittikul K, Rivera-Medina DM, Tran HN, Bouckenooghe A, Chansinghakul D, Corte´s M, Fanouillere K, Forrat R, Frago C, Gailhardou S, Jackson N, Noriega F, Plennevaux E, Wartel TA, Zambrano B, Saville M, CYD-TDV Dengue Vaccine Working Group (2015) Efficacy and long-term safety of a dengue vaccine in regions of endemic disease. N Engl J Med 373 (13):1195–1206 39. Halstead SB (2018) Which dengue vaccine approach is the most promising, and should we be concerned about enhanced disease after vaccination? There is only one true winner. Cold Spring Harb Perspect Biol 10(6): a030700 40. Wilder-Smith A, Smith PG, Luo R, KellyCirino C, Curry D, Larson H, Durbin A, Chu M, Tharmaphornpilas P, Ng LC, Sartori ˜ a G, Yoon AMC, Luna EJA, Gubler DJ, Espan IK, Flasche S (2019) Pre-vaccination screening strategies for the use of the CYD-TDV dengue vaccine: a meeting report. Vaccine 37 (36):5137–5146 41. Flasche S, Wilder-Smith A, Hombach J, Smith PG (2019) Estimating the proportion of vaccine-induced hospitalized dengue cases among Dengvaxia vaccinees in the Philippines. Wellcome Open Res 4:165 42. Lo´pez-Medina E, Biswal S, Saez-Llorens X, Borja-Tabora C, Bravo L, Sirivichayakul C, Vargas LM, Alera MT, Vela´squez H, Reynales H, Rivera L, Watanaveeradej V, Rodriguez-Arenales EJ, Yu D, Espinoza F, Dietze R, Fernando L, Wickramasinghe P, Duarte Moreira E Jr, Fernando AD, Gunasekera D, Luz K, da Cunha RV,

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52. Houser K, Subbarao K (2015) Influenza vaccines: challenges and solutions. Cell Host Microbe 17(3):295–300 53. Wei CJ, Crank MC, Shiver J et al (2020) Nextgeneration influenza vaccines: opportunities and challenges. Nat Rev Drug Discov 19:239–252 54. Gyawali B, Ramakrishna K, Dhamoon AS (2019) Sepsis: the evolution in definition, pathophysiology, and management. SAGE Open Med 7:2050312119835043 55. Levy MM, Artigas A, Phillips GS et al (2012) Outcomes of the surviving sepsis campaign in intensive care units in the USA and Europe: a prospective cohort study. Lancet Infect Dis 12 (12):919–924 56. Thompson K, Venkatesh B, Finfer S (2019) Sepsis and septic shock: current approaches to management. Intern Med J 49(2):160–170 57. Furfaro LL, Chang BJ, Payne MS (2018) Perinatal Streptococcus agalactiae epidemiology and surveillance targets. Clin Microbiol Rev 31(4):e00049–e00018 58. Wang X, Thompson CD, Weidenmaier C, Lee JC (2018) Release of Staphylococcus aureus extracellular vesicles and their application as a vaccine platform. Nat Commun 9(1):1379 59. Cross AS (2010) Development of an antiendotoxin vaccine for sepsis. Subcell Biochem 53:285–302 60. Zhu B, Dockrell H, Ottenhoff T, Evans T, Zhang Y (2018) Tuberculosis vaccines: opportunities and challenges. Respirology 23 (4):359–368 61. McShane H (2019) Insights and challenges in tuberculosis vaccine development. Lancet Respir Med 7(9):810–819 62. Van Der Meeren O, Hatherill M, Nduba V, Wilkinson RJ, Muyoyeta M, Van Brakel E, Ayles HM, Henostroza G, Thienemann F, Scriba TJ, Diacon A, Blatner GL, Demoitie´ MA, Tameris M, Malahleha M, Innes JC, Hellstro¨m E, Martinson N, Singh T, Akite EJ, Khatoon Azam A, Bollaerts A, Ginsberg AM, Evans TG, Gillard P, Tait DR (2018) Phase 2b controlled trial of M72/AS01E vaccine to prevent tuberculosis. N Engl J Med 379 (17):1621–1634 63. Tait DR, Hatherill M, Van Der Meeren O, Ginsberg AM, Van Brakel E, Salaun B, Scriba TJ, Akite EJ, Ayles HM, Bollaerts A, Demoitie´ MA, Diacon A, Evans TG, Gillard P, Hellstro¨m E, Innes JC, Lempicki M, Malahleha M, Martinson N, Mesia Vela D, Muyoyeta M, Nduba V, Pascal TG, Tameris M, Thienemann F, Wilkinson RJ, Roman F (2019) Final analysis of a trial of

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Spiessens B, Guinovart C, Espasa M, Bassat Q, Aide P, Ofori-Anyinam O, Navia MM, Corachan S, Ceuppens M, Dubois MC, Demoitie´ MA, Dubovsky F, Mene´ndez C, Tornieporth N, Ballou WR, Thompson R, Cohen J (2004) Efficacy of the RTS,S/AS02A vaccine against Plasmodium falciparum infection and disease in young African children: randomised controlled trial. Lancet 364 (9443):1411–1420 73. Cockburn IA et al (2018) Malaria prevention: from immunological concepts to effective vaccines and protective antibodies. Nat Immunol 19:1199–1211 74. Keating C (2020) The history of the RTS,S/ AS01 malaria vaccine trial. Lancet 395 (10233):1336–1337 75. RTSS Clinical Trials Partnership (2015) Efficacy and safety of RTS,S/AS01 malaria vaccine with or without a booster dose in infants and children in Africa: final results of a phase 3, individually randomised, controlled trial. Lancet 6736:31–45 76. Reimer JM, Karlsson KH, Lo¨vgrenBengtsson K, Magnusson SE, Fuentes A, Stertman L (2012) Matrix-M™ adjuvant induces local recruitment, activation and maturation of central immune cells in absence of antigen. PLoS One 7(7):e41451 77. Collins KA, Snaith R, Cottingham MG, Gilbert SC, Hill AVS (2017) Enhancing protective immunity to malaria with a highly immunogenic virus-like particle vaccine. Sci Rep 7:46621 78. Venkatraman N et al (2019) Phase I assessments of first-in-human administration of a novel malaria anti-sporozoite vaccine candidate, R21 in matrix-M adjuvant, in UK and Burkinabe volunteers. Preprint at Medarxiv 79. Mayer F (2020) The quest for a vaccine against malaria. Nature Portfolio. https://www. nature.com/articles/d42859-020-00021-8 80. Future Medicine (2020) Serum Institute joins hands with Novavax for malaria vaccine. https://futuremedicineindia.com/serum-insti tute-joins-hands-with-novavax-for-malariavaccine/ 81. Jiang T, Shi T, Zhang H et al (2019) Tumor neoantigens: from basic research to clinical applications. J Hematol Oncol 12:93 82. Shen XY, Orson FM, Kosten TR (2012) Vaccines against drug abuse. Clin Pharmacol Ther 91:60–70 83. Takada A, Kawaoka Y (2003) Antibodydependent enhancement of viral infection: molecular mechanisms and in vivo implications. Rev Med Virol 13(6):387–398

Challenges for Vaccinologists 84. Halstead SB, O’Rourke RJ (1977) Dengue viruses and mononuclear phagocytes. I. Infection enhancement by non-neutralizing antibody. J Exp Med 146 (1):201–217 85. Halstead SB, O’Rourke RJ, Allison AC (1977) Dengue viruses and mononuclear phagocytes. II. Identity of blood and tissue leukocytes supporting in vitro infection. J Exp Med 146:218–229 86. Hadinegoro SR, Arredondo-Garcia JL, Capeding MR et al (2015) Efficacy and long-term safety of a dengue vaccine in regions of endemic disease. N Engl J Med 373:1195–1206 87. Bardina SV, Bunduc P, Tripathi S, Duehr J, Frere JJ, Brown JA, Nachbagauer R, Foster GA, Krysztof D, Tortorella D, Stramer SL, Garcı´a-Sastre A, Krammer F, Lim JK (2017) Enhancement of Zika virus pathogenesis by

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Chapter 2 Principles in Immunology for the Design and Development of Vaccines Claudius U. Meyer and Fred Zepp Abstract Vaccinology has come a long way from early, empirically developed vaccines to modern vaccines rationally designed and produced. Vaccines are meant to cooperate with the human immune system, the later largely unknown in the early years of vaccine development. In the recent years, a tremendous depth of knowledge has been accumulated in the field of immunology that has provided an opportunity to understand the mechanisms of action of the vaccine components. In parallel, our knowledge in microbiology, molecular biology, infectiology, epidemiology, and furthermore in bioinformatics has fostered our understanding of the interaction of microorganisms with the human immune system. Strategies engaged by pathogens strongly determine the targets of a vaccine, which should be formulated to stimulate potent and efficiently protective immune responses. The improved knowledge of immune response mechanisms has facilitated the development of new vaccines with the capacity to selectively address the key pathogenic mechanisms. The primary goal of a vaccine design might no longer be to mimic the pathogen but to identify the relevant processes of the pathogenic mechanisms to be effectively interrupted by a highly specific immune response, eventually surpassing natural limitations. Vaccines have become complex sets of components meant to orchestrate the fine-tuning of the immune processes leading to a lasting and specific immune memory. In addition to antigenic materials, which are comprised of the most critical immunogenic epitopes, adjuvant components are frequently added to induce a favorable immunological activation. Furthermore, for reasons of production and product stability preservatives, stabilizers, inactivators, antibiotics, or diluents could be present, but need to be evaluated. While on the one hand vaccine effectiveness is a primary goal, on the other hand side effects need to be excluded due to safety and tolerability. Further challenges in vaccinology include variability of the vaccinees, the variability of the pathogen, the population-based settings of vaccine application, and the process technology in vaccine production. Vaccine design has become more tailored and in turn has opened up the potential of extending its application to hitherto not accessible complex microbial pathogens plus providing new immunotherapies to tackle diseases such as cancer, Alzheimer’s disease, and autoimmune disease. This chapter gives an overview of the key considerations and processes involved in vaccine design and development. It also describes the basic principles of normal immune responses and in their function in defense of infectious agents by vaccination. Key words Vaccine, Vaccination, Immune memory, Pathogen, T cell, B cell, Infectious disease, Adjuvant

Sunil Thomas (ed.), Vaccine Design: Methods and Protocols, Volume 1: Vaccines for Human Diseases, Methods in Molecular Biology, vol. 2410, https://doi.org/10.1007/978-1-0716-1884-4_2, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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Introduction Vaccination has been proven to be one of the most effective medical interventions to reduce morbidity and mortality of infectious diseases. The outstanding theme of vaccination is to induce a protective immune memory using principles identified for the natural interaction of an infectious pathogen (bacteria, viruses, etc.) with the (human) immune system (Fig. 1). In contrast to the natural infectious pathogens, vaccines ideally achieve their protective effects without clinical symptoms of disease or side effects. Vaccine design in principle builds on the structure and biological properties of an infectious agent. It is of utmost importance for vaccine developers to understand the physiology of the pathogen, its underlying genetics, the epidemiology, the pathogenesis, and immunobiology. As pathogens tend to generate variants as a strategy to escape previous immune memory, an ideal vaccine should also have the capacity to induce cross-protective immune responses against those potential variants of the pathogen. The common understanding is that a vaccine should stimulate all those steps naturally initiated by a pathogen contact leading to immune activation and promoting an adequate effector mechanism, involving mediators and cellular responses, which are tailored to address the specific pathogen. At all times in varying degrees, the development of vaccines was started in alignment with observed natural phenomena [1]. Early vaccinologists including Chinese pediatricians and Edward Jenner (in Europe) [2] deducted their concepts from the observation that under certain conditions individuals were spared from highly contagious diseases. Over the last two centuries, vaccination was strongly endorsed by the progress in biological sciences, the emergence of biochemical techniques, and the discoveries in immunology and genetics. The techniques available in the late twentieth century further facilitated the development of new vaccine concepts such as subunit vaccines (purified or recombinant protein or polysaccharide), DNA- or mRNA-vaccines, and the application of reverse vaccinology resulting in rationally designed, genetically engineered antigenic vaccine components [3]. The first two decades of the twenty-first century have seen an ever-growing choice of new vaccine designs, including viral vectors, RNA- or DNA-vaccines, and virus-like protein particles, additionally stimulated by the advent of systems vaccinology, including data-driven approaches based on bioinformatic methodology [4, 5]. The advantages of modern vaccine concepts have often been associated with specific drawbacks, including the fact that highly purified vaccine antigens often provide less potent immunogens. Moreover, there are challenging diseases such as malaria, tuberculosis, or HIV/AIDS that at least to date remain out of reach of

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Fig. 1 After infection with a pathogen a complex interaction between pathogen and the host immune system includes illness and harmful processes. The vaccine offers similar antigens like the pathogen, but the whole part of disease is ommitted, thus directly leading to protection without a need for healing

classical vaccine design. To overcome these impediments, adjuvant formulations had been developed to stimulate innate immune components as natural support for a vaccine-driven immunization [6]. Following the recognition of the important role of the innate immunity for the induction of an adaptive immune response, new adjuvant mechanisms were developed that together with an evergrowing armament from systems biology have generated new insights and options to modulate the type of immune response, specifically increasing the level of immune activity at least to those levels typically seen with original live-attenuated or inactivated vaccines. Thus, the hope is that modern vaccines may have the potential not only to compensate for limitations of natural occurring immune responses but even also to surpass natural limitations. 1.1 A Brief History of Vaccination

Already in the ancient world it was common knowledge that for some infectious diseases an individual was rarely infested twice. This observation led to the practice of inoculation that has been documented in China more than 1000 years before Jenner’s remarkable studies [1]. Even the term “immunity” was used in reference to plague during the fourteenth century. Progress in natural sciences and the development of experimental techniques during the eighteenth century led to the systematic use of inoculation to fight smallpox, one of the most serious threats during that time. In the early eighteenth century variolation, the transmission of small,

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presumably sublethal volumes of liquids from smallpox pustules was introduced to England by Lady Mary Wortley Montagu. Lady Montagu survived infection with smallpox herself. Impressed with the method of variolation, she ordered the embassy surgeon, Charles Maitland, to inoculate her 5-year-old son. After her later return to London in 1721, Lady Montagu introduced the method to the physicians of the royal court. Thereafter variolation became quickly popular among physicians in Europe. However, variolation was not without risks. In average, 2–3% of variolated persons died from the disease but the mortality associated with variolation was 10 times lower than that associated with naturally occurring smallpox. Modern concepts of vaccination date back to 1796 when Edward Jenner based on empirical observation used liquid from pustules of cowpox to induce protective immunity in human individuals. Today, the use of cowpox as a vaccine is considered to be the landmark of modern vaccination concepts. Edward Jenner recognized that milkmaids infected by cowpox, a generally harmless infection for humans, were rendered immune to smallpox. In 1796, Jenner deliberately inoculated people with small doses of cowpox (vaccinia) from pustules and successfully demonstrated that protection against smallpox could be achieved. Jenner termed this preventive measure “vaccination,” and over the following decades, inoculation against smallpox using cowpox became widely accepted in Europe. While Jenner at his time neither understood nor could explain the biological basis of “vaccination,” his concept was successful and provided protection from smallpox apparently due to cross-immunity between cowpox and smallpox. Until the end of the nineteenth century, diseases were believed to be caused by invisible microbes which were “spontaneously generated” in response to “bad air” and other environmental triggers, as well as a belief that imbalance in the body caused what were actually infectious illnesses. Progress in microbiology and virology since the late nineteenth century elucidated the modern concept of communicable diseases. Pasteur and Koch established that microorganisms were the true cause of infectious diseases. These discoveries led to the science of immunology. Hence further advances in vaccinology were gained from an increasing understanding of the etiology of infectious diseases and host–pathogen interactions. Pasteur challenged the spontaneous generation theory of microbes while Koch demonstrated that infectious agents transmit diseases. Koch defined four postulates which established an individual agent as the cause of a disease. In addition, in the late 1870s, Pasteur developed the first attenuation procedure for pathogens. Pasteur’s approach provided microorganisms less pathogenic but still immunogenic. Using animals as a live propagating medium, Pasteur and his team were able to produce attenuated rabies viruses of different strengths of which the weakest could be used to prepare a vaccine.

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In 1885, the first human individual was vaccinated with a live, attenuated rabies vaccine. However, due to technical limitations of vaccine production at that time, fatal cases of rabies in vaccinated individuals occurred. At the end of the nineteenth century, many of the fundamental aspects of vaccinology were established due to the pioneering work of Pasteur and Koch. Probably the most important advance was the insight that the administration of pathogens, either attenuated or killed, resulted in protection against the disease caused by the respective nontreated pathogen. The first inactivated vaccines, developed in the 1890s, were directed against the typhoid and cholera bacilli. Other vaccines consisting of killed whole pathogens, produced in the early twentieth century, were directed against pertussis, influenza, and typhus. These were followed by inactivated vaccines directed against polio (IPV), rabies, Japanese encephalitis, tick-borne encephalitis, and hepatitis A [7]. Although inactivated vaccines exhibit a lower risk of vaccineassociated disease than live vaccines, their efficacy can be reduced by the same factors, i.e., circulating antibodies (maternal antibodies) or concomitant infection. Moreover, multiple doses of inactivated vaccines are generally needed to provide sufficient stimulation of the immune system to induce durable immune responses. To support the mounting of a durable immune response to vaccines, aluminum compounds as vaccine adjuvants have been introduced (from the Latin word adiuvare, meaning “to help or aid”). Still today aluminum salts represent the most frequently used adjuvant system. Further progress in biochemistry facilitated the development of inactivated vaccines based on purified toxins. The first subcellular vaccines made available in the 1920s used diphtheria and tetanus toxoids [1]. As technology improved, it became possible to purify protein or polysaccharide subunits from infectious organisms to develop increasingly specific vaccine antigens. A further important milestone was the development of sophisticated ways to culture and propagate infectious pathogens, like viruses, ex vivo. Based on these new techniques, the development and production of purified attenuated viral pathogens as live vaccines became possible. Typical examples of vaccines that use passage in artificial media or cell culture as means of attenuation include the oral polio virus (OPV) or measles, mumps, rubella, and varicella vaccines [8] as well as the Bacille Calmette-Gue´rin (BCG) tuberculosis vaccine [9]. Recent years have been characterized by an impressive progress in the fields of cell biology and immunology as well as important technical improvements concerning fermentation and purification. Building on the improved knowledge of the principles of host– pathogen interactions, the host’s immune response today can be dissected in order to identify the precise antigenic structures that are most relevant to initiate protective immunity. The appropriate

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antigens are isolated as subcomponents of pathogens and subsequently produced in large quantities either by purification or by using recombinant in vitro technologies. Moreover, innovative adjuvants have been introduced that specifically modify and augment those aspects of the immune response that are most appropriate for protection [10]. These adjuvants have the potential to foster long-lasting immunological memory to maintain protection. Over the last 100 years, vaccine development has evolved from an empirical approach using trial and error, to one of highly rational vaccine design where careful selection of immunogenic components and adjuvants is key to the desired efficacy against challenging pathogens. Modern vaccine design needs to consider target antigen selection in much more detail to improve immunogenicity, while at the same time conserving a favorable reactogenicity and safety profile [11]. With new vaccine technologies currently emerging, it will be possible to custom-design many vaccines for optimal efficacy, low reactogenicity, and excellent safety profiles in the near future, applicable to the whole population, specific groups, or designed as personalized vaccines [12].

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Basic Concepts of Vaccine Immunology The primary goal of vaccination is the induction of protective immunity against disease causing agents, primarily focused on infectious pathogens, i.e., microorganisms like bacteria, viruses, or fungi. During recent years, the scope has been extended to other targets, detrimental physiological components, as far as these are immunologically assessable. To achieve this, vaccines mostly are designed to address natural defense mechanisms and activate the immune system in a manner similar to the natural reaction in case of a challenge. Modern vaccine development, more than ever before, strongly depends on our understanding of the human immune system [13]. An operational view of the human immune system identifies two major cellular networks, the innate and the adaptive immune system (Fig. 2). In this working model, innate and adaptive immunity are sequentially addressed to identify invading pathogens and initiate the most effective defense response. Emerging evidence suggests that these cellular networks set up an intensely interacting and cooperative system with interactions, which are crucial to generate and maintain a protective immune response. A growing number of cell type enable this cooperation, with specialized antigen-presenting cells (APCs) among the most important cell types to orchestrate the effectiveness of the immune system [14, 15].

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Fig. 2 The immune system includes two functional areas, the innate immunity and the adaptive immunity. The innate immunity follows less specific, but vigorous principles, which can be found in the animal kingdom in vertebrates as well as in non-vertebrates, thus representing an evolutionary very old, but successful cellular network of response. The adaptive immunity has exclusively developed in vertebrates, representing only 1% of the total animal species diversity. (Adapted from Abbas et al. Cellular and molecular Immunology, 9th ed.)

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Innate Immunity The innate immune system represents an evolutionary old, first-line surveillance system to detect and defend the host against pathogens that surmount the body’s physical and chemical barriers (e.g., skin, ciliated epithelia, mucous membranes, stomach acids, and defensive enzymes in secretions). Innate defense mechanisms include the production of pro-inflammatory cytokines and chemokines, helping to initiate the recruitment and the activation of inflammatory cells as well as the release of noncellular effector molecules such as complement or lysozyme. The cellular components of the innate immune system are generated in the bone marrow and migrate into blood and different tissues of the body. Tissue-residing (e.g., macrophages and dendritic cells) and circulating phagocytic cells (e.g., neutrophils, eosinophils, and monocytes) as well as natural killer cells and innate lymphoid cells [16] represent major cellular contributors of the innate immunity [14].

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After invasion of a pathogen, the innate immune system is responsible to detect, contain, and ideally eliminate the threat immediately. Pathogens are detected through molecular-sensing surveillance mechanisms, e.g., pattern recognition receptors (PRRs), expressed by the cells of the innate immune system either on the cell surface or in intracellular compartments (i.e., DNA[17], or RNA- [18] sensors). In recent years, the number of identified PRRs has increased considerably, overall covering a broad set of danger signals [19]. Typical examples of PRRs are the transmembrane Toll-like receptors (TLRs) which recognize pathogen-associated molecular patterns (PAMPs) that are shared by several pathogens; for example, lipopolysaccharide expressed by all Gram-negative bacteria. This enables the innate immune system to sense the occurrence of an infectious event. For instance, TLR4 at the cell surface recognizes bacterial, whereas TLR9 is located intracellular and recognizes viral single-stranded RNA. In contrast to the cells of the adaptive system, PRRs lack specificity to single pathogenic species, thus are sensing invaders only when archetypal PAMPs are present. Nevertheless, considerably diverse sets of different receptor systems are in place to sense most of the pathogenic assaults and instigate immediate pro-inflammatory gene expression cascades in order to facilitate host defense capacity [20]. Epithelial cells, fibroblasts, and vascular endothelial cells support innate immune cells when infected, stressed, or damaged by the secretion of chemical messengers like cytokines and chemokines to attract other resident and circulating innate cells to the site of infection. Pathogen elimination or at least reduction may be achieved by innate immune effectors in those cases when innate cells are at site of infection; in case they can be recognized by PRRs and the concentration of the pathogens, e.g., stimulating PAMPs is appropriate [21]. In case of repeated challenging, the innate immune response may change the level of response, depending on the main stimulating molecular structure. Repeated contact to lipopolysaccharide (LPS) results in a dampened, tolerogenic reaction, prestimulation with ß-Glucan sensitizes, and increases the innate reactivity [22]. These details indicate that innate immunity needs to be seen as a considerably more specific system based on an overall complex pattern recognition system, with specific PRR and with even memory development. Such an innate immunological memory may depend mainly on natural killer (NK) cells and macrophages, which together provide protection against reinfection in a manner independent of T or B cells. The memory responses in innate cells involve epigenetic changes based on methylation and acetylation, a functional reprogramming that might induce reactivation in case of a secondary encounter of the pathogen [23].

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Elimination can be accomplished by phagocytosis of the invader and subsequent destruction within intracellular vesicles containing oxygen radicals and digestive enzymes. Monocytes, macrophages, and neutrophils are phagocytes. Additionally, pathogens can be destroyed by soluble chemical factors secreted by innate immune cells or generated in the liver. Complement represents the most important and effective soluble effector system of innate immunity. Complement proteins circulate in the blood in an inactive form. Comparable to the coagulation system, the 25 complement proteins are activated in cascades. When activated, complement components fulfill several effector functions including the recruitment of phagocytes, the opsonization of pathogens to facilitate phagocytosis, and the removal of antibody–antigen complexes. The complement system also strongly promotes the effector function of the adaptive immune response by mediating lysis of antibody-coated pathogens. The innate immune response is enforced by chemotactic stimuli, released by infected epithelial and endothelial cells or other innate immune cells to recruit additional circulating cells from the blood stream to the site of inflammation. While the defense provided by innate immune mechanisms in principle is sufficient to resolve an infection, during evolution many microorganisms have developed escape mechanisms to overcome the effectors of innate immunity. Therefore, in these cases, innate immunity will delay the invasion of pathogens, but intervention of the adaptive immune response is indispensable to overcome and finally clear an infection. Although innate defense mechanisms are fast reacting, for vaccine-related purposes they lack the needed level of specificity and are not equipped to provide a long-term specific immunological memory. In consequence, innate immunity alone is not sufficient for vaccine-related protective immune responses that depend strongly on the induction of lasting immune memory responses [24]. Recent observations termed trained innate immunity might offer therapeutic options in the future, but its application for vaccine development yet needs to be demonstrated [25]. Nevertheless, innate immunity fulfills an indispensable role in the early detection of invading pathogens and subsequent activation of the adaptive immune response. The detection of pathogens and the phagocytosis of antigens by immature dendritic cells (DC) is a prerequisite to initiate adaptive immune responses. After ingestion of antigens, immature DCs transform into activated antigenpresenting cells (APC) that migrate to the draining lymph node. The APC acts as a messenger to precisely define the nature of the perceived danger and convey this information to secondary lymphoid organs, where they activate the relevant adaptive immune response [26, 27]. In the end, vaccines aim to instruct the adaptive immune system, but vaccine antigens must be recognizable by innate immune cells.

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Adaptive Immunity Innate immune response offers some specificity, but the adaptive immune system is facilitated to recognize pathogenic structures with a much higher discriminatory capacity. Not general structural patterns as seen by the innate immune system, but unique molecular details (epitopes) are recognized and memorized. With such a deeply detailed memory, the adaptive immunity assists as a second line of defense in the elimination of a foreign, pathogenic invader and in terminating an infestation. Moreover, in the course of an adaptive immune response, antigen-specific memory cells are generated that will provide a faster and stronger immune response whenever the body is rechallenged by the same pathogen again in the future [15]. The cellular elements of the adaptive immune response are lymphocytes that are able to specifically recognize antigens, i.e., the components of an infectious pathogen “foreign” to the body and potentially dangerous. There are two main subsets of lymphocytes: B cells which initially develop in the bone marrow and T cells which are generated in the thymus. Activated B cells can produce and secrete antigen-specific antibodies, i.e., proteins that will bind to antigens. T cells comprise of different cell types that confer either regulatory or effector functions. T cells with regulatory function preferentially express the cluster of differentiation (CD) 4 cellsurface protein and are referred to as CD4-postive T cells. Effector-T cells are characterized by the expression of the CD8 cell-surface molecule. In contrast to innate immune cells, lymphocytes can express a huge diversity of antigen-specific receptor molecules (theoretically 1015 to 1020; practically at least more than 106 [28, 29]). Antigen receptors are encoded by a set of genes that undergo multiple recombination events, eliciting the random generation of an extensive number of diverse receptor structures. The diversity of the receptor repertoire is further increased by individual changes and random gene insertions. The huge T and B cell repertoires of the human immune system provide the potential to recognize the biggest part of the naturally occurring antigenic structures. During the development of the adaptive immune system within the generated receptor repertoire, lymphocytes expressing receptors that recognize self-antigens will be eliminated by a process named negative selection, while simultaneously cells that recognize nonselfantigens are positively selected. Initially the repertoire is maintained with single or very few cells expressing receptors that will recognize any given antigen. As soon as an individual clone is activated by a specific challenge, the clone expands to a whole population of specific cells. Such a population basically represents the memory for the specific antigens and therefore pathogens.

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T Cells Each T cell expresses a single antigen-specific receptor molecule (TCR). TCRs, however, cannot directly recognize complete pathogenic structures. Instead, the TCR recognizes molecular fragments (small peptides derived from processing of larger protein antigens) that have to be presented in association with major histocompatibility complex (MHC) molecules at the cell surface of antigen-presenting cells (APC). In consequence, activation of T-lymphocytes strongly depends on the interaction with APCs. Professional APCs, derived from specialized phagocytes termed dendritic cells (DCs), ingest pathogen-derived proteins. After phagocytosis, the antigens are broken down and processed, and the resulting peptide fragments are transported to the cell surface where they are embedded into MHC molecules. An individual T cell can only be activated by a peptide antigen for which it expresses the specific receptor. Moreover, besides its antigen specificity, the TCR additionally can only interact with MHC molecules of its own tissue type. This quality is described as self-restriction and ensures that only cells of the same organisms will interact to mount an adaptive immune response. T cells activated by antigen-bearing DCs express the CD4 cellsurface protein and are restricted to recognize antigen in the context of MHC class II molecules. CD4+ T cells fulfill modulatory and effector functions by secreting soluble factors (cytokines) that exert direct antimicrobial properties or affect the activities of other immune cells. In most cases, CD4+ cells will help other immune cells to perform their task and are, therefore, referred to as helper T cells (Th). Based on the instructions they received from the APC while recognizing the antigen, these Th cells secrete appropriate sets of cytokines to assist other subsets of immune cells. Several functional subpopulations of Th cells have been described. Th1 cells secrete mainly interferon-gamma (IFNγ), a cytokine known to limit pathogen survival. IFNγ also promotes the differentiation of cytotoxic lymphocytes (CD8+ cells) that are able to destroy cells infected by intracellular pathogens. Th2 cells produce various cytokines (Interleukins [IL] IL-4, IL-5, IL-13) that preferentially activate innate immune cells (eosinophils, mast cells) especially facilitating the immune response to extracellular parasites (Fig. 3). Another subset, termed follicular T helper cell (Tfh) based on its tissue localization in follicular structures of lymph nodes, is characterized by the secretion of IL-21, a cytokine thought to favor the secretion of antibodies by antigen-specific B cells [30]. An important and diverse subtype are regulatory T cells (Treg cells), belonging to the CD4+ T cells. Treg inhibit immune or inflammatory responses by blocking the activity of effector T cells, helper T cells, and APCs. Treg are crucial to downregulate immune responses

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Tn

IL-1, IL-6, IL-23, TGF-ß

Th1

IFNg, TNF

Defense against Intracellular pathogens

Th2

IL-4, IL-5, IL-13

Acvaon of eosinophils & mast cells

Th9

IL-9

Proinflammatory response

Th17

IL-17, IL-21, IL-22, IL-25, IL-26

Mucosal defense against extra-cellular bacteria & fungi

Th22

IL-22

Defense against Gram-negave Bacteria

Treg

IL-10, TGF-ß

Regulaon of local immune response (gut, lung)

T

IL-21

Promoon of angen secreon from B cells

Fig. 3 Starting with a naive T cell (Tn) a complex functional diversity of T helper cells (Tn) develop to help focussing the adaptive immune response on various specific challenges. (Updated from Moser et al. (2010); doi:10.1016/j.vaccine.2010.07.022)

after an effective protective response, to maintain immunological self-tolerance process, and for the prevention of uncontrolled or chronic inflammatory responses. T cells expressing the CD8 surface molecule represent T effector cells that have the capacity to eliminate cells infested with intracellular pathogens. Antigen recognition by CD8+ T cells depends on the fact that virtually all nucleated cells present fragments of intracellular proteins at their surface MHC molecules as part of a normal surveillance process. While APCs present externally derived antigen fragments in association with MHC class II molecules, nonimmune cells use MHC class I molecules to present peptides derived from intracellular sources. Thus, cells infected by intracellular pathogens will express antigenic fragments of the pathogen in addition to the normal set of self-antigens. CD8+ T cells continuously screen MHC class I molecules to detect nonselfantigens indicative for an intracellular infection. Cells displaying high levels of pathogen-derived peptides, e.g., in the case of a virus infection, subsequently will be killed by CD8+ T cells by

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secretion of cytotoxic factors. In addition, CD8+ T cells can inhibit viral replication without destroying the infected cells by producing cytokines that are able to interfere (Interferon) with pathogen replication. CD8+ cytotoxic cells also can eliminate cells exhibiting abnormal host peptides, such as those presented by tumor cells, and therefore play an important role in the immune control of aberrant cell growth. Although CD8+ T cells can react directly to cells expressing nonself-antigen/MHC class I complexes, their optimal cytotoxic potential is achieved in the presence of cytokines produced by regulatory CD4+ T helper cells. There are many reports on more subtypes of T cells, a field which has seen an ever-growing set of subpopulations. At the same time, ongoing discussion is stimulated by the more or less present plasticity to change or switch subtype. Some reports suggested to leave the basic classification system of rigid T cell subtypes, which relies on few cell-surface proteins and on a small set of secreted cytokines. Data might argue for a broader view on individual T cell qualities, which fall on a continuum of ranges of properties, thus subtypes need to be appreciated as more heterogeneous and complex as has been thought before [31].

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B Cells B cells represent the second effector compartment of the adaptive immune response. Like T cells, each B cell expresses a single antigen receptor (B cell receptor: BCR), which consists of a membranebound copy of the antibody molecule that can be secreted by the B cell after activation. In contrast to T cell receptors, the BCR binds directly to molecular structures of pathogens with no need for previous antigen processing. Antigen binding by the appropriate BCR activates the naı¨ve or the memory B cell and induces proliferation and differentiation into plasma cells. Long-lived plasma cells produce and secrete large amounts of antibodies that are released in the blood and other body fluids. Antigen-specific antibodies are an important effector concept of adaptive immunity. Antibodies can facilitate phagocytosis or complement-mediated killing of pathogens or neutralize toxins by binding to their appropriate antigens (Fig. 4) [32]. Antibody molecules consist of a “constant” fragment (Fc-fragment), a structural feature common to all antibodies of a given isotype, and a “variable” region, which includes the region that defines the antigen specificity (Fab-fragment). The constant part of the molecule exists in five different classes (isotypes) termed immunoglobulin [Ig] A, IgD, IgE, IgG, and IgM. The Ig-isotype determines the ability of an antibody class to localize to particular body sites and to recruit the optimal effector cells. The variable region of the antibody exists in a huge number of randomly

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Claudius U. Meyer and Fred Zepp

Fig. 4 Antibodies are generated with a huge diversity regarding their binding specificity, therfore antbodies are found in various contexts to bind with high specificity to pathogenic invaders

generated different molecular configurations. This BCR repertoire guarantees maximal capability to recognize diverse pathogenic antigen. Activation of naı¨ve B cells after the first encounter with an antigen and subsequent differentiation into long-lived plasma cells usually needs 10–14 days. Initially at least two preliminary cell types are generated, short-lived plasma cells, which will typically produce IgM-type antibodies as a first response level. IgM antibodies are large molecules consisting of five bivalent antibody molecules linked together to exhibit the binding regions. In the further course of the immune response, the newly differentiated second cell type, the germinal center (GC) B cell will undergo extensive proliferation, forming the germinal center wherein the antibody production will switch to the IgG-isotype, which also represents the major isotype of B cell memory responses [32]. Depending on the specific circumstances of B cell activation, antibody production may switch to IgA, which is secreted to mucous membranes or IgE, mainly for the defense of infections by parasites. In most cases, optimal B cell activation and differentiation into antibody-secreting plasma cells will only be achieved when B and T cells are simultaneously activated by elements of the same pathogen

Principles of Vaccination

B

homing receptors

lymph node T

CCR7 CCR4

Dendritic cell

B

T cell zone

CXCR4

41

B

T B

B7 CD28

costimulators

IgG

T cell

CCR4 CCR7

CCR5

T cell

B cell

CCR1

Antigen presentation

T cell help to B cells

Fig. 5 T cells are activated when recognizing antigenic epitopes presented by dendritic cell in the T cell zone of the lymph node. At the border to the B cell follicle activated T cells help B cells stimulate production of antibodies the antigen

(Fig. 5). T cell-independent direct activation of B cells occurs only in response to repetitive antigenic structures, such as carbohydrates found in bacterial cell walls. These T cell-independent immune responses are characterized by the secretion of low-affinity antibodies of the IgM type, lacking the typical memory response upon reexposure to the same antigen. In these instances, activated B cells will recruit the help of T cells to mount an optimal response and to elicit immunological memory. After activation of the B cell by binding to a pathogen antigen, the surface BCR–antigen complex will be internalized, and elements of the antigen are processed and presented to an appropriate CD4+ T helper cell. The interacting CD4+ T cell will differentiate into a follicular T helper cell in order to provide helper signals for the B cell. T cell-dependent B cell responses are characterized by the secretion of high-affinity antibodies and a large spectrum of isotypes (in particular IgG). The quality of antibody response has a bearing on protection, e.g., the antigen binding capability of antibodies (affinity, avidity) and the dynamics of the peak response (priming); long-term protection requires the persistence of antibodies and the generation of immune memory cells capable of rapid and effective reactivation [33].

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Immune Memory As outlined, T helper cells play an important role in the regulation of both, T and B cell responses as well as cytotoxic T-lymphocytes. However, the most important property of adaptive immunity is its capacity to establish an immunological memory, assuring a stronger and faster protective immune response whenever challenged again by the same or related pathogen. While the primary immune response on average takes 10–14 days to build up, immunological memory shortens the immunological reaction time to a couple of days, thereby effectively preventing future reinfection with the same agent (Fig. 6). At the first encounter with an antigen, usually only a small number of naı¨ve lymphocytes expressing a given antigen specificity are available. Upon activation by antigen recognition, T and B lymphocytes will go through rapid proliferation, leading to the accumulation of an increased number of cells expressing receptors for the specific antigen. In a yet not fully understood way, some of these cells will differentiate into effector cells while others will become “memory cells,” able to survive for longer periods of time within the host. Any exposure to an antigen (pathogen or vaccine) therefore leads to a long-term modification of the cellular repertoire, such that the relative frequency of T and B cells specific for an individual antigen is increased in antigen-exposed individuals compared with naı¨ve individuals. Memory T and B cells will develop secondary (recall) responses on reencounter with their specific antigen [34, 35]. The adaptive response on secondary exposure leads to a rapid expansion and differentiation of memory T and B cells into effector cells, and the production of high levels of antibodies. A higher proportion of IgG and other isotypes of antibodies compared with the level of IgM characterizes memory antibody responses. During the process of reactivation, the binding avidity of antibodies can be optimized by somatic hypermutation of the variable antigen binding region. B cell memory has two arms, with B memory cells travelling through the body and with long-lived plasma cells residing in the bone marrow permanently producing antibodies. T memory cells are localized at various locations, circulating in the blood, sampling in the lymphoid tissues, or migrating in most of the organ tissue systems. Local and functional variants of T memory cells have been described; for instance, tissue resident T memory cells (Trm) “trapped” in tissues [36], or stem cell-like T memory cells (Tscm), which might have a prominent role in the long-term persistence of immune responses [37]. The capacity to generate immune memory is the key feature of the adaptive immune system and is crucial for maintenance of longterm protection. This capacity to establish an immunological

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Fig. 6 Dynamics of the adaptive immune response. (Adapted from “Understanding Modern Vaccines: Perspectives in Vaccinology, Volume 1,” 2011 Elsevier, Oberdan, L., Cunningham, A., Stern, P.L.: Chapter 2. Vaccine immunology)

memory response is also the fundamental aim of the biological effects of vaccines. In summary, the initially antigen processing and presentation by dendritic cells (DCs) are key steps that define the environment and the course of an efficient immune response. Therefore, innate immunity sets the stage for the subsequent adaptive response and innate and adaptive immunity have to interact intensely in order to initiate the most effective type of protective immunity.

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How Do Vaccines Mediate Protection? Long-term protection is ensured by the maintenance of antigenspecific effector cells and/or by the induction of immune memory cells that can be rapidly reactivated into immune effectors whenever the organism is challenged with the same pathogen again in the future or encounters some other “nonself”-structure. Vaccineinduced immune effectors are essentially antigen-specific antibodies produced by plasma cells that are capable of binding specifically to a toxin or a pathogen. Other effectors are cytotoxic CD8+ T cells that can limit the spread of infectious microorganisms by killing infected cells or secreting specific antiviral cytokines. The generation and maintenance of both B and CD8+ T cell responses is supported by growth factors and signals provided by CD4+ T helper cells. Most

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antigens and vaccines trigger both B and T cell responses. In addition, CD4+ T cells are required for most antibody responses, while antibodies exert significant influences on T cell responses to intracellular pathogens. With regard to a new, evolving field of vaccinology, the therapeutic vaccination, these principles are also effective, although the targeted structures may not come as an invading pathogen, but may be molecular structures, which are recognized as foreign like neoantigens on cancer cells [38].

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Immune Correlates of Protection Following the idea that a vaccine is constructed as an innocuous replicate of the real pathogen, it might seem obvious to measure the immune response parameters seen in real-live infectious disease. For many decades, antibody titers have been interpreted as main correlates for successful vaccination. Additionally, in the last two decades, T cell-mediated immunity has been measured to complete the understanding of vaccine-induced responses [1]. However, most immune correlates of protection are not well defined. For instance, a considerable number of vaccines target pathogens, which enter the organism via mucosal tissues, challenging the mucosal compartment of the immune system. However, historically, for vaccines against these pathogens, correlates of protection were detected via identification of specific antibodies (or T cells) circulating in the peripheral blood, thus in the systemic compartment [39]. In consequence, these concepts appear to be insufficient or inappropriate for future vaccine development [40]. Conventional immunological assays including ELISA, ELISpot, flow cytometry, and neutralization assays supported the present knowledge, but the developments in modern immunological methodologies, including a range of “OMICs” technologies, envision a much more sophisticated and comprehensive picture of the effects of an immunization series on the immune system. Systems vaccinology techniques like mass detection of single-cell reactions with single-cell RNA sequencing (scRNseq) combined with clonal lymphocyte identity have already been shown to offer the capacity to delineate meaningful data sets [4, 41]. Ideally, a successful immune response is measured by the efficacy to protect from infectious disease. However, this approach usually is difficult to perform regularly on individual basis. Thus, the challenge is to correlate immunogenicity and the protective efficacy of a given vaccine. The goal will be to narrow the information gap between an immunological response characterized with innovative methods on the one hand and the factors conveying effective protection concluded from seminal clinical trials on the other hand. The fast-developing methodologies and the impressive

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gain of knowledge on disease pathology may offer reasonable parameters to comprehensively measure correlates of protection in the near future, but will challenge authorities to interpret and standardize such results from clinical trials.

10

Principles of Vaccine Development Vaccines have been developed using the conventional method basically introduced as variolation before Edward Jenner in the eighteenth century. An isolated pathogen inactivated or at least attenuated was injected after no or minor purification steps. Compared to the preceding incidence rates of smallpox the dropping number of cases was a major achievement, although some of the vaccinees died. Jenner made the next major step by using a basically harmless cowpox virus as vaccine to induce immunity against human smallpox, thereby considerably improving the safety. More than two centuries later, after the elimination of some of the most dreadful natural human infectious diseases, the side effects or even harmful effects of vaccine products receive increasing attention. During a unique evolution for the last 100 years, every new biotechnological innovation has been used to improve effectiveness, safety, and reactogenicity, as well as reliability of production methods of vaccines [42]. Using vaccines on a population level made it a considerable responsibility for health-care authorities to recommend vaccines, thus the requirements for vaccine licensing have continually become more detailed and challenging. Today a growing array of different vaccine concepts has been developed as to raise an appropriate immune response specifically tailored to eliminate the various pathogens and their pathology (Fig. 7) [43]. Symptoms of an infection are either caused directly by the pathogen or, more often, they are consequences of the emerging immune response, representing side effects of our physiological defense mechanisms. Typical complaints such as physical discomfort, malaise, fever, or organ malfunction in most cases are related to inflammatory reactions that occur in course of the immunological defense process. Since vaccines are administered to prevent infections and/or diseases, they are expected to provide protection without the risk of side effects or clinical symptoms of disease. To this end in vaccine development, it is important to understand the life cycle of an infectious agent, how it multiplies and infests the human organism, how the immune system counteracts and overcomes the microbial invasion and finally builds up a protective immunity, i.e., an effective barrier against future challenges by the same agent. Moreover, it is essential to define which elements of the natural immune response are relevant for the elimination of the pathogen and future protection, and which are responsible for

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POLYSACCHARIDE AND CONJUGATED POLYSACCHARIDE Conjugated protein (e.g. toxoid, CRM197, protein D) Polysaccharideconjugate vaccine (e.g. pneumococcal Polysaccharide conjugate vaccines) vaccine Protein triggers T-cell response

Toxic groups

Antigenic determinants

TOXOID ANTIGEN

Infectious pathogen

Treat with heat or chemicals so that it is inactivated but killed/inactivated vaccine still immunogenic (e.g. pertussis whole cell vaccines)

Toxin

Inactivation

Infectious pathogen

Non-toxic toxin

A

A Wild virus is replicated

B C

Antigenic determinants induce antibody production (e.g. tetanus vaccines)

Live attenuated vaccine (e.g. varicella vaccines)

WHOLE PATHOGEN in cell culture B The process is replicated

several times... C ...to produce a

less virulent strain

Surface antigen

SPLIT AND SUBUNIT VACCINES A Purification of subunit vaccine (natural or recombinant proteins) (e.g. acellular pertussis vaccines) B Purification of recombinant antigen (natural assembly into spheres) (virus-like particles; e.g. hepatitis B vaccines)

Insert gene into expression system (e.g. yeast)

Human

purification

Sequence gene encoding antigen

REASSORTANT LIVE

Protein expression

Human/bovine reassortment (e.g. ressortant rotavirus vaccines)

Vectors A B C Antigen may be produced by recombination or purification

Human pathogenic virus Structural vaccinology

C Purification of split vaccine (e.g. influenza vaccines)

Nucleic acid-based vaccines

POTENTIAL FUTURE CONCEPTS

Fig. 7 Different types of vaccines have emerged through the last decades of development. Vaccines can be produced using different processes and may contain live attenuated pathogens (usually viruses), inactivated whole pathogens, toxoids (an inactivated form of the toxin produced by bacteria that causes the disease), or parts of the pathogens (e.g. natural or recombinant proteins, polysaccharides, conjugated polysaccharide or virus-like particles). (Adapted from Vetter et al. (2018) doi.org/10.1080/07853890.2017.1407035)

symptoms of disease and discomfort. Ideally, a vaccine should induce only the elements of the natural immune response that are essential for protection, but simultaneously exclude all negative effects of natural infection. In vaccine development, therefore, not only the elements of the immune response guaranteeing best protection must be considered but also the acceptable tolerability and safety ramifications of the induced inflammatory response [44]. As a consequence, the design of a vaccine has to be based on both structural and biological properties/qualities of an infectious agent and the type and quality of naturally occurring immune responses initiated by the infectious pathogen [45]. While for many decades vaccine development concentrated primarily on targeting components of the adaptive immunity (B cells or immunoglobulins, T cells and cytokines, such as interferon), recent research indicates that innate and adaptive immunity have to interact vigorously to initiate the most potent type of protective immune response [15]. In particular, antigen processing and presentation by DCs are key steps in the development of efficient immune responses.

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Conventional vaccines formulated with whole microbial pathogens usually supply a broad repertoire of different antigens, whereas antigenic epitopes assure sufficient immunostimulatory activity within a heterogeneous B cell and T cell population. In contrast, highly purified antigens consisting only of a limited number of epitopes may pose the risk of insufficient interaction with individuals missing the adequate immune-receptor repertoire. Moreover, genetic heterogeneity of the pathogen may counteract the expected benefit of highly purified vaccine antigens. Keeping this in mind, selection of vaccine antigens has to balance specificity and purity of antigens against sufficient antigenic variety to ensure targeting the immune system of all or at least the majority of individuals in a given population. Today the manufactured vaccine products have evolved into high-tech products including a highly sophisticated mixture of ingredients, which apart from the antigenic molecules may include adjuvants, preservatives, stabilizers, inactivators, antibiotics, diluents, and trace elements from the production and purification process. Most of them fulfill technical purposes, but some are also critical for a successful delivery and function of the vaccine [46]. Knowledge of process technology, e.g., initiatives like “Quality-by-design” (QbD) and Process Analytical Technology (PAT), have become cornerstone technological frameworks to proof identity, stability, and concentration of the vaccine ingredients to the authorities [42]. In summary, vaccine design has made significant advances within more than 100 years. Historically, an empiric “isolation– inactivation–injection” concept has governed most part of the vaccine development. At the same time, this has been coupled with the comprehension that one dose for everyone for a single disease will do. Although we have not left these concepts, the mentioned new, innovative techniques to look deep into the details of the complexity of an infectious disease will enable us to leave the useful, but limited scientific reductionism, and enter into a systems biology-driven conceptual approach for vaccines, which also includes the various needs of different patient groups in a given population, and even heading to find ways to create vaccines fitting the needs of an individual [12].

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Selecting Vaccine Antigens Following the cornerstone definition of the immune system, the active component of a vaccine should display molecules which are clearly recognized as “nonself” by the cellular partners of the immune system. Thus, a prerequisite to start with the design of a vaccine is the identification of appropriate antigenic structures. For example, if a neutralizing antibody response is sufficient to protect

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from infection, usually an antigenic structure from the bacterial/ viral cell surface is selected. The identification of B-cell epitopes, but also Th and cytotoxic T lymphocyte (CTL) epitopes, represents the starting point for developing any successful vaccine candidate. Vaccine immunogens containing the appropriate set of relevant epitopes should induce substantial and protective immune responses [47]. This has been done successfully for the H. influenzae type b, pneumococcal and meningococcal and hepatitis B vaccines, or from secreted toxins, like tetanus or diphtheria. In the course of an antibody response, antigen-specific helper T cells are essential for the evolution of high-affinity antibodies and immune memory. Other antigen-specific T cells, including cytotoxic T cells, accomplish important effector functions, such as the targeted removal of host cells infected by intracellular pathogens. Thus, vaccine antigens have to be selected that enable these T cell effector-mediated responses. Hepatitis B vaccines, for example, induce antibodies as well as hepatitis B-specific T cell responses [48], Pertussis vaccines induce antibodies and stimulate helper T cells to produce interferon [49, 50], and hepatitis A and IPV vaccines probably stimulate both T and B cells. In some instances, the immune response induced by vaccination may even be stronger than the response observed after natural infection as has been shown for Human Papilloma Virus (HPV) vaccines [51]. Progress in biotechnology in recent years has allowed the isolation of subcomponents of pathogen, thereby eliminating unwanted pathogenic components. One successful attempt to separate antigenic structures from unwanted material has been made with split- or subvirion vaccines. These vaccines are prepared by using a solvent (such as ether or a detergent) to dissolve or disrupt the viral lipid envelope, a technology that has been applied most successfully in the inactivated influenza vaccines [52]. Purification steps are also engaged in the production of subunit vaccines, comprising protein or polysaccharide antigens, such as acellular pertussis proteins [49], typhoid Vi-antigen, and pneumococci polysaccharides [53]. With purified antigens, an impaired immunogenicity may also occur that may be unable to address sufficient elements of the immune system needed for the protective response. One such example are polysaccharide antigens, which alone are not able to recruit T helper cells in order to obtain B cell support. This phenomenon is especially significant in young infants and children as well as with the elderly [54]. As a result, immune responses to plain polysaccharide antigens are characterized by the secretion of low-affinity antibodies, mainly immunoglobulin M (IgM) molecules, and display a stereotyped “innate response” behavior. Repetitive encounters with the same antigen fail to induce a secondary, memory-like immune response [55]. This disadvantage was finally surmounted by covalently binding the polysaccharide antigen to a carrier protein. Conjugate vaccines dramatically improve B and T

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cellular immune responses to polysaccharides. The conjugated vaccine is internalized, processed, and the antigen components of the conjugated protein are presented in the context of major histocompatibility complex (MHC) molecules to be recognized by conjugate-protein/peptide-specific T helper cells. Applying this approach, polysaccharide-specific B cells recruit help from conjugate-protein-specific T cells to get all signals needed to promote further activation as well as isotype switching to IgG production and generation of memory B cells. Today, this elegant technique is regularly applied to vaccines containing bacterial polysaccharides for the prevention of invasive diseases caused by encapsulated bacteria. Examples include H. influenzae type b, pneumococcal and meningococcal vaccines. Modern molecular biology has allowed to produce recombinant vaccines that contain only the antigen substructures relevant to elicit protective immunity [55]. The first recombinant vaccine, licensed in 1986, included the gene for hepatitis B surface antigen (HbsAg; [56]). Two recombinant vaccines are available against cervical cancer, both based on human papillomavirus (HPV) virus-like particles assembled from recombinant HPV L1 coat proteins [51]. Even multiple proteins have been produced with recombinant technology for the Meningococcus type B strains. Followed by vaccines against Staphylococcus aureus or Pseudomonas strains, this technology was the first successful example for reverse vaccinology, which opened the field of vaccine development to the possibilities of bioinformatic methodology. Structural vaccinology has evolved to predict the exact location and the design of epitopes relevant for binding B or T cell receptors [57]. Most protective immune responses correspond to polyclonal responses, some vaccines respond only to a few epitopes, but antigenic clusters present in the major epitopic regions of a pathogen are more likely to elicit a protective immune response. Threedimensional structures of antigenic molecules are now deduced from the sequences data leading to epitope-based vaccines covering all possible epitopes helpful for an immune response with a broad and diverse T (or B) cell clonality. High-density surface display of epitopes using nanoparticles or scaffold protein allows an immune cell-targeted vaccine design, further improving immunization efficiency [47].

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Improving Vaccines In spite of the classical, mainly empirical approach to design vaccines, a century of vaccine development has witnessed a tremendous improvement with regard to increased vaccine specificity, activation of appropriate immunological mechanisms, with considerably lower reactogenicity and far better safety profiles. We have

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long left the era of crude inactivated pathogen material as a basic vaccine ingredient. Meanwhile a multitude of vaccine designs try to interfere the highly complex diseases-mediating mechanisms of a still small group of the top frightening pathogens. Today for about 30 human pathogens vaccines have been made available, about the same number of targets are in the pipeline, nevertheless, a limited bunch compared to an estimated number of 1400 known human pathogen specimen, indicating the challenge ahead. Major barriers to improve vaccine design includes [55]: (a) Mechanisms of immune response to vaccines. As stated several times in this text, our current knowledge on how pathogens interact with the immune system and also how vaccines induce immune responses is a preemptive prerequisite for further improving immunizing processes initiated by vaccines. The list is continued with only partially available correlates of protection, limited immunogenicity, suboptimal contribution of the innate immunity, and a lack of appropriate animal models. (b) Variability of the vaccinees. The “one-size-fits-all” concept will not be a choice for future vaccines. Individual genetics but also epigenetic variability need to be further described with genome-wide association studies (GWAS) and have to be tailored to develop individualized vaccine formulation. The tailored vaccine should consider physiological limitations or build on individual capabilities to receive a vaccine. Research on exclusive vaccines will elucidate the reasons why there are nonresponders in some of the vaccine concepts; in addition, it will offer the chance to adapt vaccines to gender, age, and ethnic background and also to the needs of patients with primary immune defects. (c) Variability of the pathogen (or the nonself-structure). We have a wealth of information on the diversity of pathogenic specimen, including antigenic drift and major shift (like an ongoing topic in influenza virus), and the effects of hypervariable regions of viruses, the feedback on a complex biology of the life cycle of a pathogen. In addition, challenges come from latency of infectious/pathogenic processes, often difficult to analyze due to the long and unforeseeable timescales. A major issue regarding the interaction of host with pathogen is the frequently used escape mechanisms, which pathogens use to outsmart the immune system. (d) Safety and reactogenicity of vaccines. The association of adverse reactions with a given immunization needs to be investigated in detail so as to anticipate adverse reactions for each individual vaccinee. Understanding

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adverse reactions to the vaccines and communicating the findings generated will help to tackle vaccine hesitancy, which has lately been recognized as a major obstacle to the necessary high vaccination rates to reach herd immunity at the population level. (e) Frame settings to support successful vaccination. A sufficient immune response to a given vaccine is only possible in a well-functioning organism. Environmental factors like poor nutrition, poor health conditions with coinfections, and intoxications from a polluted living area are counteracting an effective and successful immunological protection of individuals as well as the societies. (f) Process technology in vaccine production. For each new vaccine product, the technological basis needs to be updated. With changing vaccine platform technologies, the challenges to offer technological readiness, handle the technological complexity, provide for consistent up-scaling and appropriate flexibility in adaption of the production processes in case of changing needs, the anticipation of appropriate stability of the finally distributed product, and the overall speed to respond to new demands from epidemiological threats are central challenges to the industry while entering into new concepts of immunization [58]. Still in the interest of each new developmental vaccine project, adjuvant molecules will offer a relevant answer to some of these listed questions, and specifically, how to co-stimulate a helpful innate immune response during a vaccine-induced immunization. Adjuvant research has demonstrated that with the right selection of antigens together with new adjuvants the immune response elicited by vaccines can be adapted to the pathogens and targeted populations. Recognizing that this cannot always be achieved with only one adjuvant type led to the investigation of Adjuvant Systems, which combine classical adjuvants (aluminum salts, o/w emulsion, and liposomes) and immunomodulatory molecules, such as MPL and QS21. This concept has allowed the development of vaccines tailored to the antigen and target population, such as the HPV vaccine with Adjuvant System AS04 and a malaria vaccine with Adjuvant System AS02.

13

Future Prospects From the global perspective, health-care systems urgently await for new or improved vaccines, ready to be applied on population level opposing relevant infectious diseases [59].

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Table 1 List of pipeline vaccine targets (WHO 2020) Campylobacter jejuni

Nipah virus

Chagas disease

Nontyphoidal salmonella disease

Chikungunya

Norovirus

Dengue

Paratyphoid fever

Enterotoxigenic Escherichia coli

Respiratory syncytial virus (RSV)

Enterovirus 71 (EV71)

Schistosomiasis disease

Group B streptococcus (GBS)

Shigella

Herpes simplex virus

Staphylococcus aureus

HIV-1

Streptococcus pneumoniae

Human hookworm disease

Streptococcus pyogenes

Leishmaniasis disease

Tuberculosis

Malaria

Universal influenza vaccine

With an increasing global disease surveillance based on a fast and powerful diagnostic tool set, the pace to identify new and epidemic pathogens is accelerated. With more pathogens causing devastating impact on health, and their geographical spreading between continents can be followed in “real-time,” thus giving reason to extend the set of available vaccines to this increasing number of targets. A growing number of pathogens is listed by the WHO (Table 1), which are in the developmental pipeline. Some have been in this list for years, even decades, indicating the biological complexity research is confronted with, whereas, some others entered the list only recently. Currently, the time between identification of the pathogen and its availability of vaccines is decreasing as the developmental processes benefit from (bio-) technological advances due to available vaccine strategies and as well as powerful and flexible production lines. A critical example of how pressing the need for a vaccine can become is the SARS-CoV2 pandemic. Within less than 12 months after the pathogen had been identified, sequenced, and target epitopes had been identified, the first vaccines were licensed and marketed, after following all necessary regulations for licensure. Some challenges presented by infectious diseases such as malaria, tuberculosis, and HIV/AIDS so far could not be addressed successfully with classical vaccines, including those containing traditional adjuvants. This has led to new approaches including live vectors, DNA vaccines, and new adjuvant formulations as described before. The technology is of particular interest for the development of HIV vaccines and therapeutic vaccines for certain cancers. However, to date, clinical trials performed in the context of HIV have

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been less encouraging, and the potential of such an approach is progressing slowly. DNA and RNA vaccines are composed of genes encoding a key antigenic determinant. They come often inserted into a bacterial plasmid or as stabilized messenger RNA [60]. Administration of the DNA or RNA vaccines leads to the expression of the foreign gene and synthesis of antigens derived from the infectious organism within the host cells. Presentation of the foreign proteins by the host cells can elicit an immune response similar to that induced by natural infection. Depending on the host cells targeted, DNA or RNA vaccines have the potential to stimulate cellular or humoral immune response. Clinical trials of plasmid DNA vaccines for HIV infection, Ebola hemorrhagic fever, West Nile virus infection, avian influenza, and various cancers are currently ongoing. Many more advances in vaccinology can be expected in the future, including therapeutic cancer vaccines that have been tested with promising results with a number of spontaneous tumor animal models, including models of breast, prostate, pancreatic, and colon cancer [61]. These vaccines are designed involving antigen-specific vaccines and DC vaccines formulated with patients’ DCs loaded with tumor-associated antigens. DC vaccines work by isolating and exposing the cancer patient’s DCs ex vivo to compounds that include tumor-associated antigens. After their reintroduction to the patient, these DCs promote a cytotoxic T cell response against the tumor tissue. Another interesting target are allergic diseases, which affect up to 25% of the population in western countries. Novel immunotherapies are currently under development, among them DNA and RNA vaccines [62]. Progress is also expected from vaccines for the treatment of autoimmune diseases like type 1 diabetes, arthritis, Alzheimer’s disease, multiple sclerosis, hypertension, dyslipidemia, substance dependency, as well as vaccine concepts to neonates and to old people. The continuing progress in vaccine technologies and the understanding of the mechanisms underlying immune response are facilitating a more refined approach to vaccine design tailored to the desired effect in combating disease. References 1. Plotkin S (2014) History of vaccination. Proc Natl Acad Sci U S A 111(34):12283–12287. https://doi.org/10.1073/pnas.1400472111 2. Buck C (2003) Smallpox inoculation—should we credit Chinese medicine? Complement Ther Med 11(3):201–202. https://doi.org/ 10.1016/S0965-2299(03)00087-6 3. Rappuoli R, Bottomley MJ, D’Oro U, Finco O, De Gregorio E (2016) Reverse

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Chapter 3 Revisiting the Principles of Designing a Vaccine Shubhranshu Zutshi, Sunil Kumar, Prashant Chauhan, and Bhaskar Saha Abstract Immune principles formulated by Jenner, Pasteur, and early immunologists served as fundamental propositions for vaccine discovery against many dreadful pathogens. However, decisive success in the form of an efficacious vaccine still eludes for diseases such as tuberculosis, leishmaniasis, and trypanosomiasis. Several antileishmanial vaccine trials have been undertaken in past decades incorporating live, attenuated, killed, or subunit vaccination, but the goal remains unmet. In light of the above facts, we have to reassess the principles of vaccination by dissecting factors associated with the hosts’ immune response. This chapter discusses the pathogen-associated perturbations at various junctures during the generation of the immune response which inhibits antigenic processing, presentation, or remodels memory T cell repertoire. This can lead to ineffective priming or inappropriate activation of memory T cells during challenge infection. Thus, despite a protective primary response, vaccine failure can occur due to altered immune environments in the presence of pathogens. Key words Leishmania, Vaccine, Subcellular localization, Antigen priming, Epitope crypticity, Immune synapse, T cell plasticity, Reverse vaccinology, Proteomics

1

Introduction The principle of vaccination is the induction of antigen-specific memory immune response that is fast reactivated upon second exposure to the pathogen expressing the antigen (Fig. 1). The goal is to provide life-long prophylactic immunity to that pathogen. Since the time of Jenner, vaccination-induced host protection is achieved by priming hosts with live, killed, attenuated, or subunitbased vaccines. The antigen priming establishing a pool of hostprotective memory T cells that are reinvoked upon exposure to a pathogen expressing those priming antigens protecting the host [1, 2]. This dogma helped us to develop successful vaccines against many diseases such as typhoid, cholera, and preparations such as

Shubhranshu Zutshi and Sunil Kumar contributed equally to this work. Sunil Thomas (ed.), Vaccine Design: Methods and Protocols, Volume 1: Vaccines for Human Diseases, Methods in Molecular Biology, vol. 2410, https://doi.org/10.1007/978-1-0716-1884-4_3, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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Fig. 1 Characteristics of the primary and secondary immune response. Qualitative characteristic of primary immune response is dependent on host, antigen, and vaccine formulation-based factors. The secondary immune response is characterized by a quick and efficacious response against the pathogen

MMR and BCG. However, this principle has not been able to achieve the long-standing goal of vaccination against diseases such as leishmaniasis, Trypanosomiasis, malaria, HIV, and tuberculosis—all dreadful infectious diseases killing millions across the globe. Therefore, in this introduction to the whole practical nuances of vaccination, we will revisit the principles of vaccination and propose how vaccines may be failing so that during the next edition of protocols, these points are considered and vaccines are reformulated. Leishmaniasis is a neglected tropical disease for which vaccination is the need of the hour. Many vaccine candidates have been tried against leishmaniasis in preclinical and clinical trials, but they were unable to achieve the goal of serving as an efficacious antileishmanial vaccine [3]. Antigen priming generates memory T cells in an environment free of pathogen’s inhibitory or deviating influences. During the challenge infection, pathogen-driven perturbation of the host immune system may fail to recall memory T cells. This may occur at the level of (1) antigen processing and presentation, (2) APC–T cell interface, and (3) Th phenotype plasticity. These factors contribute to a weakened or nonrobust memory recall response during the infection, resulting in an ineffective generation of protective immunity. In this introductory chapter, we thus critically reassess these factors in detail and propose that vaccinology needs a significant reconsideration of the principles of vaccinology to translate into working vaccines. The following sections describe different pathogen-induced effects on the host immune system significantly affecting the generation of a protective and robust antileishmanial immune response.

Fig. 2 Leishmania-induced alteration in antigen processing and presentation inside APC leads to ineffective priming. (a) Receptor-specific endocytic mechanism leads to different fates of degradation and presentation pathway. (b) Leishmania inhibits phagolysosome biogenesis by gp63-mediated inhibition of NOX2 and

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Disruption in Antigenic Priming Affects Vaccine Efficacy

2.1 Stage-Specific Vaccine Candidate Imparts Specificity

The principle of vaccination is to induce a host-protective immune response against any form of the pathogen. During an encounter with a pathogen, our immune cells respond quickly against the pathogenic intruder by generating memory cells. Vaccination protocols employing killed, live-attenuated, or xenogenic microorganisms for protection studies have shown promise against many pathogens, Leishmania remains a threatening exception. Based on earlier vaccination modules, we hypothesize to select the infective stage gene for a vaccine candidate. Leishmania has two stages viz. promastigote (noninfective, extracellular) and amastigote (infective, intracellular). The proteome of promastigotes and amastigotes revealed differences as stage-specific expression and differential expression of multiple proteins were observed [4, 5]. As an amastigote is observed in humans and its proteome is available for degradation inside APCs, it becomes imperative to focus on the amastigote-specific vaccine candidates. Although promastigote is noninfective, people have tried promastigote-specific antigen for vaccination such as gp63, gp46, and LACK. They met with success in mice but sterile protection is still far away [6–9]. As amastigotes propagate in humans and are infective, the best way is to select amastigote-specific proteins. Hence, numerous gene-knockout parasites defective in A2, HASPB1, p27, and Ldcen have shown promise against both homologous and heterologous species of Leishmania by generating Th1 response but ultimately failed in subsequent trials [10–13].

2.2 Relative Antigen Abundance During Processing Inside APCs

Leishmania harbors inside the antigen-presenting cell (APC) such as macrophage whereas Th cells execute antileishmanial immunity. Thus, unavailability or less availability of an antigen during the degradation mechanism may deter antigen-specific epitopes from presenting onto the surface of APC during the MHC presentation pathway. Depending on their binding kinetics with MHC molecules, readily available epitopes are efficiently presented on the surface of APC [14]. Increased epitope presentation correlates with higher expression of their source proteins, although their preferential presentation could not be established [15]. However, a study involving p17and p24 of HIV showed that abundant epitopes are presented readily via MHC molecules. The epitope selection process is dependent upon the propensity for proteasomal digestion, an affinity for

ä Fig. 2 (continued) preventing antigen degradation. Leishmania can also (c) reduce expression, (d) sequester, and (e) decrease the extracellular release of immunogenic protein to decrease their chances of degradation and MHC-presentation. (f) Parasite-induced alteration in lysosomal protease activity can result in the increased presentation of cryptic epitopes as against immunodominant ones

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TAP, and ER-aminopeptidases rather than MHC affinity [16]. A recent theoretical model predicted that higher antigenic abundance may help a low-affinity peptide to be efficiently presented by APCs and vice-versa [17]. Thus, downregulation of the primed antigen during Leishmania infection may lead to the suboptimal presentation of epitopes and may not induce memory T cell activation. When pathogens enter a host, the immune system recognizes the antigen and presents it in the context of MHC molecules. The main function of class I MHC molecules is to present endogenous foreign peptides. After degradation by protease, the nascent generated peptides are loaded onto class I MHC which presents the antigen to T cells. Class-II MHC molecules present peptides generated from exogenously acquired antigens. In a pathogenic organism, the abundance of protein varies from a few molecules to million molecules per cell. The highly expressed antigens are in very low amount but easily bind with MHC-II molecules forming MHC-II–peptide complex for recognition by the immune system facilitating protective response. [18]. However, a small amount of antigen unable to form the MHC-II complex facilitate a low zone tolerance providing no protection. Approximately 24,000 HLA class I peptides were identified by mass spectrometry which indicated that gene expression and MHC–peptide affinity share a multiplicative relationship where a tenfold increase in expression was found to compensate for a 90% decrease in binding affinity [19]. MARIA, a tool designed to predict HLA peptides, was based on antigen expression and peptide affinity data to generate epitopes most likely to induce strong CD4+ T cell responses [20]. Another tool HLAthena predicts epitopes based on expression and processing data. It was experimentally validated to be of high predictive power, as it correctly identified >75% HLA-peptides across 11 tumor-derived cell lines [21]. Pulsechase experiments revealed abnormalities in cancer cells by turnover kinetics of MHC peptides and cellular proteins. Besides cellular expression, the rate of protein degradation influences MHC presentation [22]. In a separate study, six best epitopes of HLA-I restricted alleles of L. tropica were selected based on immunoinformatics and molecular dynamics simulation approaches. By this approach, a stronger binding affinity with HLA-I alleles was selected as probable vaccine candidates [23]. 2.3 Subcellular Localization and Availability of Antigens for Processing

Leishmania cleverly engineers the macrophage for their survival inside the lysosome. Infected macrophages form phagolysosome and fragmented peptide reach to the cell surface via MHC, but the availability of antigen for presentation and processing depends on subcellular localization in Leishmania. Leishmania-infected macrophages prevent antigens from processing and antigen presentation mechanisms by sequestering in endocytic compartments [24]. Availability of antigen is imperative for epitope presentation

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and reactivation of memory T cells generated during priming. Infection-induced antigen sequestration may inefficiently induce protective immunity during postpriming challenge infection. Antigen processing and peptide presentation occur through either endocytic or cytosolic route. After challenging the host with the parasite, immune cells recognize the infected peptide to present them at the cell surface through the engagement of the MHC complex. Ag localization is thus considered to be an important factor for antigen recognition, as in vitro studies against intracellular pathogen Leishmania, Mycobacteria, and Salmonella have shown [25]. L. mexicana cysteine proteases are localized within the parasite’s lysosomal compartment for killing the amastigotes in phagosomes. When presented in the context of MHC, cysteine protease-specific T cells recognize the antigens. However, membrane-bound acid phosphatase is located in the intracellular vesicles of the amastigote stage of this parasite preventing it from recognition by T cells. So, the localization of antigen has a crucial impact on antigen recognition. Recognition of antigen can be corrected by overexpression or engineering in such a way to express the soluble protein [26, 27]. Leishmania Alba (acetylation lowers binding affinity)-domain proteins help parasite develop through differential subcellular localization. Alba-domain proteins co-localize predominantly in the cytoplasm of both stages of Leishmania promastigote and amastigote and translocate to nucleolus and flagellum upon differentiation to amastigotes. Moreover, RNA-binding proteins also have a role in gene expression through developmental stages in Leishmania, as an alba-domain protein shuttle between cytoplasm to the nucleolus [28]. Bone marrowderived macrophages infected with T. gondii parasites were able to present secretory OVA but not the parasites with intracellular OVA. A strong correlation was observed between antigen secretion and presentation of antigen, class I MHC-peptide activates T cell response in vitro through ova-expressing transgenic T. gondii parasitic infection [29]. In vivo immunization with DNA constructs encoding OVA belonging to various subcellular sites leads to differential activation of OVA-specific CD4+ and CD8+ T cells. DNA vectors encoding cell-associated OVA antigen resulted in greater CD8+ T cell division than soluble forms of OVA antigen. Exogenous and membrane-associated antigens have better CD4+ division than cytosolic forms of OVA antigen [30]. Dendritic cells were able to present membrane-bound OVA to OVA-specific T cells but not a cytosolic form of OVA antigen. Dendritic cells infected with L. major secrete NT-OVA and could prime OT-I CD8+ T cells for IFN-γ secretion. However, DC infected with L. major nonsecreted NT-OVA could not activate or diminished T cell response [31]. Now, one can speculate antigenic epitope competition for

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the availability of MHC-II molecules in APCs. Thus, less abundant or poorly processed antigens can be outnumbered by more abundant or highly processed antigens for MHC-based presentation on the surface of APC.

3

Leishmania-Associated Inhibitions in Antigenic Processing/Presentation

3.1 Endocytic Mechanisms for Leishmania Uptake

Leishmania, a protozoan parasite, alternates between both promastigote and amastigote forms. Mammalian macrophages internalize both forms of parasite at the different stages during the entrance or infection establishment process. To establish an infection, parasites have to cross the host cell plasma membrane layer by phagocytosis [32]. Hence, the internalization process requires numerous receptors to recognize cognate ligands on the parasite such as complement receptors CR1, CR3, fibronectin receptor, mannose-fucose receptor, and receptor for advanced glycosylation end products [33]. The pathway utilized by APCs for parasitic uptake has implications in antigenic processing. DC-SIGN, DCIR2, or SR-mediated uptake directs the antigen/parasite towards the MHC-II–mediated presentation pathway, while MR and various other receptors favor the formation of early endosomes which do not fuse or acidify and remains stable for extended periods [34]. These are used for cross-presentation, but they can also give adequate time for the phagocytosed parasite to adapt to the intracellular niche, thus favoring parasite growth and inhibiting antigen presentation. The uptake of the Leishmania parasite in the host cells initiates phagocytosis. L. infantum chagasi metacyclic promastigotes use CR3 (complement receptor) not MR (mannose-fucose receptor) to enter macrophages. However, avirulent promastigotes use both CR3 and MR for entry into macrophages. MR activates the inflammatory profile whereas the CR3 receptor does not activate NADPH activation, hence respiratory burst at the phagosome membrane. GP63, LPG, and proteophosphoglycans (PPG) are present on the surface of L. major promastigote target the complement component C3, mannose-binding protein, and galectin to initiate phagocytosis for its survival [35]. Macrophage internalized phagocytic receptors such as CR and FCγ receptors promote promastigote and amastigote phagocytosis by suppressing the production of IL-12 whereas FCγ–ligand interaction induces IL-10 production. Therefore, the nature of the initial ligand–receptor interaction that precedes phagocytosis has a role in the successful infection of the macrophages or resistance of the host [36]. Macrophage normally exists as naı¨ve macrophages (M0), and a change in milieu activates a signal to M1 or M2 subtype. Activation of M1 macrophages by Th1 lymphocytes that produce IFN-γ, TNF-α helps in killing the intracellular pathogen by a process

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known as an oxidative burst. However, host cells increase the production of ROS, superoxide, hydrogen peroxide, hydroxyl radicals, and nitric oxide that have high microbicidal activity. In contrast, Th2 lymphocytes activate the M2 phenotype by producing IL-4, and IL-13, thereby activating the arginase enzyme that favors Leishmania to survive inside the macrophage. The balance between M1 and M2 subtypes dictates the phenotypic plasticity of Leishmania infection. Rapamycin, a mammalian target of the mTOR pathway, helps Leishmania to survive inside M2-type macrophage. Also, L. donovani infected host activates the mTOR pathway by decreasing the expression of M1 markers such as ROS, NO, iNOS, NOX-1, IL-12, IL-1β, and TNF-α and increasing the expression of M2 markers like arg-1, IL-10, TGF-β, CD206, and CD163 [37]. 3.2 Inhibition of Phagolysosome Formation

Phagolysosome biogenesis is a complex process regulated by soluble N-ethylmaleimide-sensitive factor attachment receptor (SNARE), sequential fusion events between phagosomes and intracellular organelles, including early endosomes, late endosomes, and lysosomes. Phagolysosome formation marks the beginning of parasitic adaption inside the unfavorable environment of macrophages by transforming into amastigotes. Phagolysosome biogenesis alters the antigen processing and presentation in Leishmania infection by many ways: First, during the phagosome maturation process, parasitic proteins are proteolytically cleaved hence free peptides were available for direct loading on MHC class I molecules in phagosomes or translocated to the cytoplasm for further processing by the proteasome [38]. Second, the LPG molecule present on the surface of promastigotes inhibits the phagolysosome biogenesis by altering the intracellular trafficking (by impairing NADPH oxidase assembly or exclusion of vesicular proton-ATPase from phagosomes) [39]. Third, GP63 also inhibits the assembly of NOX2 complex on phagosomes, by impairing the ability of infected cells [40]. Thus, Leishmania inhibits this key process for its survival inside macrophages which leads to a subsequent decrease in antigen presentation. As a consequence, reduced leishmanicidal activity leads to depleted epitopes for presentation to memory T cells. To prevent the lysosome from fusion intracellularly, Leishmania arrests the fusion process for its survival in the host. The process of fusion of recycling endosome with cell surface is governed by R-SNARE, VAMP3 (vesicle-associated membrane protein), and controlled by Syt V, a major regulator of phagosome biogenesis [41]. The APCs expressed cathepsins B and D help in antigen processing and facilitate degradation of the invariant chain (Ii) molecule. Cathepsin S helps in the removal of invariant chain from B cells and dendritic cells, and cathepsin L plays an essential role in the removal of invariant chain from thymic epithelial cells. L. major-infected BALB/c mice treated with CA074 (cathepsin B inhibitor) alters the endo-lysosomal protease activity in APC by

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inducing TH1 response. In contrast, L. major-infected BALB/c mice treated with CLIK148 (cathepsin L) modulates the antigenprocessing protease activity by inducing TH2-type response. This differential immune response in the TH cell repertoire may be due to: first, in the antigen processing compartment various proteases were present that changes the digestion pattern. Second, disruption of protease-specific recognition sites on the target proteins may either abolish or create new epitopes altering the active TH profile following challenge infection [42]. Syt XI, a negative regulator of phagocytosis and cytokine secretion, is expressed in murine macrophages and localizes to recycling endosomes following which lysosome recruits nascent phagosomes [43]. Late endosome and lysosome aid in phagocytosis through VAMP7 and Syt-VII through Ca2+-dependent exocytosis [44]. Recently, a strain-specific difference in phagosome maturation was found among L. Braziliensis strains, where ML-associated strain was found to be most potent in inhibiting phagolysosome formation upon infection [45]. These studies suggest that Leishmania inhibits phagolysosome formation for its survival, and it has implications in the process of antigen presentation and induction of protective immunity. 3.3 Dysregulation of Protease Activity by Leishmania

Lysosomal cysteine proteases known as cathepsins play an important role in antigen processing and presentation mechanisms. In macrophages, cathepsins L and S are two predominant subtypes that are present in their active form. These enzymes are now unanimously known as site-specific proteases [46, 47]. Thus, any alteration in their peptidase activity would thereby alter the generation of predicted epitope repertoire. An immunodominant epitope requiring cysteine-specific cleavage for MHC presentation may be rendered subdominant due to cathepsin inhibition. The transcription factor STAT1 alpha and IFN regulatory factor 1 (IRF-1) controlled the expression of iNOS and MHC-II gene. As the interaction between STAT1α and importin-alpha5 was blocked in L. donovani-infected macrophages, henceforth it hampered IFN-γ–induced STAT1α nuclear translocation or downregulated IFN-γ–induced gene regulation without affecting STAT1α [48]. Lysosomal cysteine proteases such as cathepsins S and L assist APC for efficient binding of foreign peptides to MHC-II molecule [49]. Moreover, cathepsins B and L selectively induced the secretion of cytokines like TNF-α, PDGF, β-FGF, and IFN-γ in synovial fibroblast-like cells from rheumatoid arthritis patients [50]. Also, cathepsins L and K were observed to induce inflammatory pathways by inducing IL-6 in macrophages and help in eliminating S. aureus [51]. Thus, it is plausible that Leishmania may tweak the cytokine milieu, which plays an essential role in TH subset differentiation, towards its survival by cathepsin-dependent mechanisms.

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During Leishmania infection, cathepsin B and cathepsin L have been associated with Th-type immune response. Cathepsin B (Ctsb) was shown to downregulate host protective Th1 response. Th1 response was observed to be potentiated by treatment with either cathepsin B inhibitor or in Ctsb / mice [52, 53]. On the other hand, cathepsin L was found to work antagonistically during leishmaniasis, whereby it was observed to be important for the induction of Th1 response during Leishmania infection and inhibition was found to strengthen anti-inflammatory Th2 response [42, 54]. Therefore, lysosomal proteases like cathepsins play a role in Th polarization. An anti-inflammatory environment induces macrophages towards the M2 phenotype, and thus it implicates the role of Leishmania in disturbing antigen processing mechanisms by cathepsin-dependent mechanisms. This again may be responsible for the inefficient generation of epitopes under pathogen challenge condition, thereby hampering the activation of vaccine-induced, antigen-specific memory T cells. 3.4 Destruction of Antigen Presentation Machinery inside Macrophages

Fragmented antigen was loaded to MHC class II molecules in late endosome compartments. The invariant chain helps MHC class II molecules to reach the endocytic compartment in ER and prevents premature peptide loading as well as guides MHC class II transport to MHC class II-like molecules via cytoplasmic tail. Thereafter, the invariant chain gets degraded by hydrolysis and the fragment generated during hydrolysis is called CLIP (class II-associated invariant chain peptide), and it is controlled by chaperone HLA-DM (a negative regulator of HLA-DO). 9–15mer peptides bind to MHC class II following lysosomal proteolysis; hence MHC class II molecules can exit from the endocytic compartment to reach the cell surface for T cell activation. [55]. This entire process is the key to successful activation of the host’s T cells. Adequate MHC– peptide–TCR interaction is essential for the generation of memory T cells during priming as well as for recall response during challenge infection. Any blockade in antigen presentation can lead to ineffective immune activation, even in vaccine recipients harboring protective memory T cells. Since APCs are a key niche for intracellular parasites like Leishmania, it is quite evident that the parasite has to evolve strategies to overcome intracellular degradation. Impaired antigen presentation is a characteristic feature of progressive Leishmania infection [56]. H-2 M is a cofactor responsible for the release of CLIP peptide from the binding cleft of class II molecules and facilitates the binding of antigenic peptides. However, MHC-II and H-2 M molecules were internalized by Leishmania amastigotes within parasitophorous vacuoles (PV). This leads to degradation of MHC-II molecule causing inefficient presentation of the antigenic peptide to Th cells and failure or ineffective activation of T cell functions [57]. L. amazonensis infection downregulates the surface

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expression of MHC-II alongside costimulatory molecules such as CD40 postinfection. Moreover, BMDCs infected with L. braziliensis and L. major showed decreased expression of MHC-II, CD86, and decreased potential for induction of T cell proliferation [58, 59]. Thus, challenge infection impairs antigen presentation leading to ineffective activation of protective memory T cells generated during vaccination. 3.5 Immunodominance and Epitope Crypticity Affects Vaccination

Although a polypeptide antigen can generate multiple epitopes, more often than not the immune response is developed against a restricted few known as immunodominant epitopes. The peptide portions which are not presented on MHC are termed as cryptic epitopes. However, the epitopes that do not have dominant potential to bind with MHC molecule are called as subdominant epitopes, and this phenomenon is termed as epitope crypticity. Many constraints are responsible for epitope crypticity such as inefficient processing, the poor unfolding of antigen, inter-epitope competition for binding with MHC molecule, excessive or indolent proteolysis, unbiased action of HLA-DM, delayed interaction with MHC or TCR, flanking determinants on the antigen might exclude binding of the index determinant, and competition between adjacent determinant on the same antigen may lead to preemption of binding of the desired determinant [60]. The phenomenon of immunodominance has huge implications in vaccine development against T cell activating pathogens as pathogen-induced factors affecting the dominant epitope repertoire could lead to failed immune response [61]. Studies showed that APCs have a key role to play in immunodominance and antigen folding, peptide-MHC affinity, and epitope editing affects the generation of immunodominant repertoire [62]. In the case of Leishmania infection, selection of preferred epitope out of many is a tedious task. Immunodominance is the phenomenon by which immune responses are mounted against only a few selected epitopes from the pool of antigenic peptides and present them to the immune system for antipathogenic immune response. The selection of immunodominant epitopes from preferred antigenic pool relies on various parameters such as relative abundance, structural topographies of antigens, the affinity of epitope towards MHC molecule, T cell precursor frequency, receptor affinity between peptide and T cell, and differential sensitivity to cathepsins or resistance to HLA-DM mediated dissociation [63]. L. major LACK antigen with an immunodominant epitope 156–173 amino acids failed to protect from L. major infection [64]. In another study, the C-terminal region of GRA-6 immunodominant antigen elicits a protective response against Toxoplasma gondii parasite by inducing a CD8 T cell response [65]. ASP-2 gene with different mutations for making cryptic or subdominant epitope confers a protective response against experimental Trypanosoma cruzi infection by inducing

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CD8+ T cell response [66]. This phenomenon could be possible by alteration in the parasite genome to reduce the breadth and magnitude of the immune response to favor parasitism, hence artificially broadening the T cell repertoire to favor the host. Thus, the shifting of the dominant antigen in Leishmania infection affects the host-protective T cell response in vaccine outcome. 3.6 Dysfunctional Epitope Loading to MHC Molecule

Leishmania-infected bone marrow-derived macrophages inefficiently present OVA, beta-galactosidase, and L. major-derived antigen to specific T cell hybrids. Interference in antigen presentation could be due to diminished antigen processing, and unavailability of MHC class II molecules on the surface of infected cells finally hampers the antigen loading to MHC-II molecules [67]. Within the endocytic compartment, Leishmania amastigotes reside in an organelle known as parasitophorous vacuoles (PV) which is supposed to have the highest amount of parasite antigens in the host cell. There are specific distribution and polarization patterns for both MHC class I and II molecules as MHC-I is not loading compartments for MHC class I molecules; however, MHC class II compartments can load with MHC class II molecules [68]. Factors responsible for antigen loading to MHC class I molecule are endogenous antigen, synthesis of the new small size of protein in cytosol, degradation of newly synthesized protein in the cytosolic proteasome, TAP molecules in the endoplasmic reticulum, and pH-independent. However, factors responsible for antigen loading to class II MHC molecule are exogenous antigen, phagocytosis by professional APC, antigen have to reach endosome, antigen has to reach to MHC molecule, TAP independent, PH dependent, proteolytic trimming of CLIP, and proper expression of HLA-DM [69]. The interactions between MHC class I or class II molecules and ligands are regulated by chaperones, which support the peptideaccessible conformation of the molecule. However, ligand-free MHC proteins usually do not reach the cell surface, because of proteasomal degradation in the cytosol. MHC molecules reach the cell surface either by reloading their ligand-binding site with peptides or by internalization in intact form into peptide-rich endosomes. MHC class I molecules require 8–10 residue peptides in length for binding, whereas MHC class II molecules can adopt much longer peptides than MHC-I. MHC class I molecules have H-2Ld protein, associated with weak interaction between peptides and Beta2-microglobulin, hence low-level cell surface expression. In contrast, MHC-II has strong interaction with peptide residues, has HLA-DM/H-2 M chaperone molecule associated with the invariant chain, and functions as a chaperone to target them to endosomal compartments and retards them from the cell surface in an inactive state. Removal of the invariant chain is done by class II-associated invariant chain peptide (CLIP), which bind to various

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MHC class II molecules [70]. These variations affect the T cell repertoire due to perturbed antigen presentation with Leishmaniainfected macrophages. 3.7 MHC-II Affinity Determines the Immunodominant Nature of Generated Epitopes

MHC-II–peptide interaction affinity is central to the phenomenon of immunodominance which is affected by both peptide-intrinsic and extrinsic factors. DM acts as a positive regulator for immunodominant peptide binding but as a negative regulator for cryptic epitope binding to MHC-II. This DM-dependency is also linked to MHC–peptide affinity, as DM-facilitated, immunodominant epitopes are kinetically more stable than the DM-inhibited, cryptic epitopes [71, 72]. Kinetic stability and immunodominance of a peptide can be changed by single amino acid substitutions at binding sites. For instance, low-stability 1166A variant of an immunodominant LACK epitope from L. major reduces its kinetic stability towards the I-Ad molecule [71]. These findings indicate that Leishmania-induced changes to these factors such as DM-degradation or gene mutation shift the antigen repertoire. The strength of the signal generated by TCR relies on antigen concentration and antigen affinity for optimal T cell response. Although the TCR binding affinity to a peptide is strong enough to activate T cell response, few exceptions such as long half-lives result in an impaired T cell response [73]. To predict the CD4 T cell epitope, one needs to look at the pattern of antigen cleavage, MHC-II presentation, and TCR recognition. First, the prediction of antigen cleavage sites depends on cleavage by cathepsin S, B, and H. Second, each 12-mer peptide antigen sequence was selected based on the scores of modeled peptide–MHC-II complexes that will bind with TCR molecule. Third, top-scoring peptide–MHC-II complexes were selected based on the scores of modeled ternary peptide–MHC-II–TCR complexes and the distribution of predicted cleavage sites that can bind to a given TCR [74]. The affinity of an epitope towards MHC binding determines the immunogenicity, and stronger interaction of the peptide with MHC leads to higher immune response. This is not the exclusive reason for determining immunogenicity, as many factors—antigen or protein abundance, immunodominance, processing of antigen, efficient T cell repertoire, and optimal binding of TCR—are responsible. In an estimate, around 30,000 dengue virus peptides were derived, only 0.3% of those were predicted to bind HLA-A, but did not find any direct correlation with HLA-B [75]. By site-directed mutagenesis, dibasic sites have created a target of intracellular proteases adjoining one of the three cryptic epitopes of ML-M providentially mutated lysozyme proteins that were shown to be immunogenic but not unmutated ML-M. These alterations in lysozyme proteins may not always favor the cryptic determinant rather activation of epitopespecific T cells [76]. All the process of selection of best-fitted epitopes into a particular MHC groove is a complex event in

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which chaperones and accessory molecules participate. For selection and binding of immunodominant epitopes to MHC-II molecule, multiple factors such as the favorable denaturing environment in the endocytic compartment, proteolytic enzyme for trimming of antigenic determinant, simultaneous processing of multiple proteins, the involvement of multiple alleles of MHC-II that can bind the same antigen, and competition among best-fitted epitopes on a single protein antigen are involved [77]. Endogenously derived LACK-specific T cells stimulate macrophages at 6 h post-L. donovani infection by inducing the production of IL-12 but at 48 h, infected macrophages failed to stimulate MHC-II restricted T cell may be due to T cell-stimulating capacity get compromised late after infection [78]. L. donovani-infected splenic macrophages decrease the membrane cholesterol by altering the kinetic parameters of peptide–MHC complex formation and impair T cell stimulating capacity. But it can be corrected by liposomal delivery of cholesterol. Reduced membrane cholesterol leads to faster dissociation of peptides lessening the probability of optimal T cell activation [79]. These findings can help in epitope prediction and optimal T cell response (Fig. 2a–f).

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Impairment of Immune Synapse at the APC–T Cell Junction An immunological synapse comprises three sets of receptors: TCR, adhesion, and costimulatory. While adhesion receptors/molecules keep APC-T cell together for sustained interaction and proper T cell activation, TCR, and costimulatory molecules determine the specificity and functional outcome of the interaction (Fig. 3). Hence, the interaction between TCR–MHC-II during Leishmania infection is an important factor that determines the further course of cytokine milieu [80]. The earliest attempts for single-cell analysis of MHC–TCR interaction showed that Leishmania infection severely limits the capability of macrophages to activate effector T cells to varying degrees, depending on respective TCR signaling thresholds [81]. Infected and bystander DCs were found to be different in their Th-inducing capability. While Leishmania-infected MHC-IIhi DCs convert naive CD4+ T cells to a Tbet+ IFNγ+ IL-10+ nonprotective phenotype, bystander DC’s promote protection [82]. APC–T cell interaction through TCR-associated Tyrosine Kinases LCK and Fyn co-cluster into a central supramolecular activation cluster (cSMAC), whereas LFA and talin (cytoskeletal protein) co-cluster into a peripheral SMAC (pSMAC) meet at cell junction interface termed as segregated supramolecular activation clusters (SMACs) [83]. At the cell junction interface, a central cluster of T cell-based antigen receptors and a ring of three concentric membrane proteins are present. Here, the inner circle is covered mostly by TCR and CD28 associated with

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Fig. 3 APC–T cell junction leads to various effector and activator functions. MHC–TCR interaction leads to the activation of the immune system activating signaling cascades with the help of accessory costimulatory molecules such as CD80, CD86, CD28, CD40, and CD40L. Naı¨ve/memory T cell activation leads to Th differentiation and cytokine production such as IL-12, IL-10, IL-4, IFNγ, and IL-17. T-cell activation is also responsible for activating humoral immunity and secretion of leishmania-specific IgGs

the central supramolecular activation cluster (cSMAC), and the peripheral is covered by integrins (pSMAC). Outer or distal SMAC (dSMAC) is surrounded by large ectodomains proteins CD43 and CD45 [84]. These clusters together comprise immune synapse. Interaction of T cell with anti-CD3 antibodies formed a small TCR cluster by recruiting ZAP70 (zeta chain-associated protein kinase 70), adaptor molecules Grb2, LAT (linker for activation of T cells), and SLP76 (SH2 domain-containing leukocyte protein of 76) [85]. LFA-1, whose ligand is ICAM-1, (intercellular adhesion molecule) binds with the ICAM family proteins. LFA-1 (leukocyte function-associated antigen-1) helps in the transmigration of naı¨ve lymphocytes into lymph nodes and recruits neutrophils to an invading tissue site. ICAM-1 is expressed on endothelial cells, and leukocytes transmigrate by following the same route of LFA-1 ligand [86]. Signal transduction is a series of cascade events that require an assembly of multiple receptors, phosphorylated protein kinases, and adapter molecules such as LAT, grb2, SLP-76, PLC-γ isoforms to initiate T cell activation response in glycolipid-enriched membrane domains [87]. Moreover, TRAIL co-stimulation also

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induced phosphorylation of LCK and ZAP70 to activate a transcription factor NF-KB that helps in the recruitment of lipid raft assembly to induce T cell activation response [88]. L. major affects CD40 signalosome function in lipid raft by depleting membrane cholesterol and forms a stable IS [89]. L. donovani alters membrane lipid content in lipid raft and thereby interferes with the MHC–peptide complex’s interaction with TCR [78]. Supplementation with exogenous cholesterol rectifies both CD40 signaling [89] and MHC-II molecules’ conformation in the L. donovani-infected macrophages [90]. Likewise, treatment with a liposomal formulation of cholesterol molecule restored the functioning of normal immune synapse and antileishmanial T cell immunity during L. Donovani infection [91]. This could be related to the biomarkers—resolvin D1 (RvD1), leukotriene B4 (LTB4), prostaglandin F2α (PGF2α), IL-1β, IL-6, IL-8, IL-10, IL-12p70, TNF-α, and TGF-β1—identified in the serum of L. donovaniinfected host, as compared to control groups [92]. Within 10 min of activation, NK cells readily form cSMAC and pSMAC-like compartments that could lead to cytotoxic or inhibitory immune synapses as a ratio of activating tyrosine kinases Syk, ZAP70, and Lck to the tyrosine phosphatase SHP-1 in the cSMAC; the ratio is high in cytotoxic synapses whereas low in inhibitory synapses [93]. For efficient priming and Th-mediated host-protective response against Leishmania infection, a balance between cytotoxic and inhibitory synapse is crucially important. The immune synapse generated during interaction between DCs and T cells plays a decisive role in CD154/CD40-dependent IL-12 production [94]. CD28, a costimulatory molecule, forms a complex with TCR called TCR-CD28 micro-cluster and associates with PKC-θ to induce the activation of T cell [95]. CD80 and CD86 are costimulatory molecules on antigen-presenting cells that engage CD28 and CTLA-4 to balance between activation and inhibition of T cells. CTLA-4 is constitutively expressed on Tregs and also on effector T cells post-activation. The binding strength between CTLA-4 and costimulatory molecules such as B7 (CD80 and CD86) have higher affinity, and the recognition by B7 leads to a shift of PKC-θ and CD28 away from the immune synapse [96]. CTLA-4 induces TGF-β production rendering host susceptible to Leishmania infection. TGF-β blocks the NO production and thereby deactivates the antileishmanial functions of macrophages. Therefore, interference with the immunological synapse by parasites could impair the efficient activation of memory T cell repertoire. This can lead to suboptimal or nonprotective Th responses upon challenge infection.

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T Cell Associated Events Affecting Vaccination Outcome In general, during acute viral infection, T cell response can be multivariate: clonal expansion, differentiation into effector T cells, death of activated T cell population, generation of long-lived memory T cell as well as phenotypic alteration during infection [97]. Antigen-primed T cells have higher antigen responsiveness than that of un-primed or naı¨ve T cells, as elucidated by a process known as functional avidity maturation [98]. The strength of antigen responsiveness of T cells can be regulated by DC numbers, antigen concentration, TCR avidity, and finally differentiation and proliferation of T cell into a non-effector to effector stage [99]. The responsiveness of T cells can be manifested through plasticity, anergy, exhaustion, or apoptosis. In the next section, we describe each parameter in detail.

5.1

T-Cell Plasticity

CD4 T cells play a vital role in host defense against microbial pathogens. Naı¨ve CD4-T cells first differentiate into Th1 and Th2 cells based on signature cytokine IFN-γ and IL-4. In Leishmania infection, it is well established that Th1 and Th2 paradigms are associated with parasite clearance or parasite persistence. The T cell population can be redefined based on the transcription factor or cell surface markers such as Th1, Th2, Th9, Th17, Tfh (T follicular helper cells), Th22, and Treg. Signature cytokine or master transcription factor differentiates T cell from their subtypes such as t-bet and STAT4 signals through Th1, GATA-3, and STAT6 signals through Th2, PU.1, and TGF-β signals through Th9, RORγt, TGF-β, SMAD proteins, and STAT3 signals through Th17, STAT3, and BCL-6 signals through Tfh, TNF-α signals through Th22, FOXP3, STAT5, and TGF-β signals through Treg [100, 101]. All these T cell subsets are associated with some intrinsic phenotype that governs the T cell fate. TGF-β plays a decisive role in cell phenotypes either for switching into Th17 or Treg. At low concentrations, TGF-β1 favors the Th17 phenotype; however, at high concentrations, TGF-β favors induced FoxP3+ regulatory cells (iTregs) by blocking IL-23R expression [102]. Moreover, TGF-β is also required for Th9 cell development by stimulating transcription factor PU.1 and IRF-4 by inhibiting t-bet expression. IL-2-driven STAT5 activation also favors Th9 development by inhibiting the Th17 phenotype [103]. Priming with DNA or protein activates, proliferates, and expands antigen-specific T cells. This expansion phase has a higher population of effector cells in which mostly die during the contraction phase. Primed cells have longterm memory because of successive exposure to antigen. Primed antigen-specific memory T cells compete with infection T cell memory in the finite space of the immune system for a variety of immunodominant epitopes. Plasticity in the phenotype of memory

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T cell population following priming and challenge with pathogen has many reasons: competition between different epitopes, change in TCR stimulus, cytokine, or inflammatory environment [104, 105]. 5.2

T-Cell Anergy

Following an antigenic encounter, lymphocytes are unable to perform an intrinsic function but remain in a dormant state for a certain time interval based on the tolerance mechanism. This leads to a state known as T cell anergy. T cell anergy can be clonal (growth arrest stage), or adaptive (unable to proliferate and perform effector function). T cell anergy can be seen in Leishmania amazonensis infection due to parasite–antigen-specific apoptosis of the cells, hence leading to defective antigen presentation [106, 107]. T cell anergy can be due to dendritic cells expressing enzyme indoleamine-2,3-dioxygenase (IDO) responsible for tryptophan catabolism [108]. Moreover, anergy can be induced due to FOXP3-expressing Treg population or coinhibitory molecule CTLA-4 [109, 110]. T cell anergy can be due to T memory cells are less dependent on second signal or costimulatory signal. In one study, both in vivo and in vitro naı¨ve T cells had stimulated by an anti-CD3 monoclonal antibody (mAb) not responded to superantigen staphylococcal enterotoxin B nor soluble forms of anti-CD3 mAbs and APC hence not induce anergy. Strong signal 1 delivered by anti-CD3 mAbs without costimulatory molecules does not cause T cell anergy rather induces naive T cells to mitogen responsiveness memory T cells [111]. T cell anergy can be induced in Leishmania amazonensis due to defects in antigen-presentation machinery. It was not due to decreased production of growth factors not due to anti-CD3– induced T cell activation nor due to necrosis rather due to apoptosis as it was confirmed through exogenous supplements of growth factors and DNA fragmentation in lymph node cells [107]. In a separate study, anergy can be seen in CD8 T cell in human visceral leishmaniasis as gene and surface marker expression get upregulated in splenic tissue samples at the transcription level. This study indicates splenic CD8 cells get exhausted or display anergic phenotype; however, CD8 cells from PBMC have features of both anergic cells and CTLs [112]. The reasons for vaccine failure may comprise reduced B and T cell proliferation, alteration in antigenpresentation machinery, fragmentation of immunodominant antigen, and persistent exposure of T cell to high-level antigens.

5.3

T Cell Exhaustion

T-cell exhaustion is a state of T cell dysfunction in which physical deletion of antigen-specific both CD4+ and CD8+ T cells during chronic infection and cancer. T cell exhaustion is characterized by the production of IL-2, loss of ex vivo killing capacity, and production of IFN-γ, TNF-α, or beta chemokines in a very low amount with high viral load capacity [113, 114]. Defective and anergic

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CD8 T cell response was generated against chronic L. donovani infection thereafter die from T cell exhaustion. As CTLA-4 and PD-1 negatively influence T cells, exhaustion of T cells in chronic infection is observed [112]. The blockade of PD-L1 signaling generates both CD4+ and CD8+ T cell protective response and promotes clearance of the L. donovani parasite by inhibiting autophagy [115]. 5.4 Programmed Cell Death or Apoptosis

Cell death is a natural process characterized by loss of integrity in the plasma membrane, changes in cell shape and morphology, shrinkage of the cell, retraction of pseudopods, condensation of chromatin, membrane blebbing, changes in the cytoplasmic organelle, fragmentation of cell as well as the nucleus, phagocytosis of fragmented cell, and formation of apoptotic bodies. Apoptosis-like phenotype can also be seen in Leishmania-infected macrophages such as the production of reactive oxygen species through NO and H2O2, the effect of anticancerous and antiinfectious molecules such as amphotericin B, miltefosine and endonuclease G, metacaspase, aquaporin Li-BH3AQP, calpains, cathepsin B-like protein, LmjHYD36, and Lmj.22.0600, respectively [116, 117]. Posttranslational modification in Leishmania is associated with deglutamylation and polyglutamylation, and overexpression of polyglutamylase promotes leishmanial apoptosis, while overexpression of deglutamylase inhibits Leishmania-regulated cell death [117]. Antigen-specific T cells were generated against foreign invaders after infection clearance; thereafter, pathogen-specific CTL dies by apoptosis although some exception like memory T cells has antigen-independent long-term survival capacity [118]. To generate high-level memory T cells against pathogens, the implementation of a prime-boost approach is required. Generally priming is done by DNA and boosting is done by protein. So, the classical approach, i.e., homologous prime-boost, was used for neutralizing the primary antibodies but unable to protect against chronic infection because it only works against immunodominant epitope. To overcome the problem, the heterologous prime-boost strategy is applied because it is not only covering the broad range of epitope but also enhancing the magnitude and functionality of T cell and antibodies response [119]. As booster is needed to increase the CD8 T cell memory population, but memory cell formation depends on various parameters such as several antigens, quality, and nature of pathogen encounter, immunization time interval, inflammatory response, nature of antigen used for immunization, and the immune response against the specific antigen [120]. Priming with antigen initiates the inflammatory response at the site of injection; thereafter, effector and memory T cells are generated to clear the infection. Later, clearance of infection is followed by withdrawal of inflamed cytokines and differentiation of effector cells into long-lived memory T cell through epigenetic modification [121] (Fig. 4).

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Fig. 4 Differentiation of T cell to various subsets. Pictorial representation of various fates of T cell after differentiation into their T cell subtypes based on their surface marker and secretory cytokines molecule associated with a signal strength that decides the T cell phenotype

6

Reverse Vaccinology: An Extension of Immunoinformatics From its conception, vaccine development was largely based on Pasteur’s principles of isolating a pathogen, inactivating, and injecting into the hosts’ body to induce immunity. This had been the central guiding principle of vaccine development for almost a century. Until recently, antigen identification from a pathogenic organism relied heavily on biochemical and immunological techniques involving in vitro culture of organisms. This technique for antigen search was time-consuming, laborious, and difficult to achieve with uncultivable microbes [122, 123]. Also, we already discussed the probable roadblocks in antileishmanial vaccine development associated with immune system activation. In the next few sections, we try to describe how omics-generated information in the twenty-first century could help us with our goal of a successful vaccine against this neglected parasitic disease. Immunoinformatics gained momentum in the 1980s when DeLisi and Berzofsky designed the first tool for predicting the T cell epitopes by modeling the interactions between epitopes and TCR with utmost accuracy, thereby identifying probable T cell epitopes [124]. This requires scanning of proteome and genome of pathogenic organisms for identifying potential epitopes. It,

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therefore, broadened the scope of vaccine design by allowing researchers to make in silico prediction of previously unknown virulence factors and immunogenic sequences which would have been to determine by conventional methods [125]. As a result, immunoinformatics helped in digging out many novel and potential immunogenic proteins from the M. tuberculosis genome and revealed the similarity between the epitopes of Francisella tularensis and Homo sapiens, i.e., self-epitopes [126, 127]. Thereafter, the advent and popularization of automatic sequencing in the 1990s led to a “big leap forward” in the wholegenome sequencing of various organisms. It enabled a huge amount of in silico data at our disposal to identify multiple vaccine candidates by in silico analysis of a pathogen’s genome [128]. Also, the genome sequence of multiple strains of a microorganism gave rise to the field of comparative genomics and proteomics. This started an alternate revolution in the field of immunoinformatics which was termed as “reverse vaccinology.” It was so-called as it represented a sensitive, in silico “top-down” approach starting from the genome of the pathogenic organism and ending in antigen identification, in complete contrast to the traditional method of antigen identification [129]. One of the earliest approaches utilizing reverse vaccinology is to find the surface-expressed proteins in a bacterium using subcellular localization tools as they would be easily accessible by B cell receptors and antibodies during the immunization and subsequent infection, respectively. Around 25 vaccine candidates against Meningococcus-type B were identified in just 18 months, as against a dozen of them identified through conventional methods in four decades [130]. It gave new hope to the field of vaccine development and accelerated the process of antigen identification to a great extent. At present, this approach is being utilized as an important tool in the search for an effective vaccine against various bacteria, viruses, and parasitic microorganisms. Reverse vaccinology encompasses many techniques such as determination of putative epitopes, subcellular localization of the protein, MHC-haplotype–specific peptides, as well as comparative genomics and proteomics by in silico applications.

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Reverse Vaccinology in T Cell-Based Vaccine Design

7.1 Epitope Prediction and Mapping

B cell epitope prediction is quite complicated than the corresponding analysis in T cells due to the recognition of conformational epitopes by the B cell receptor. On the other hand, T cell immunity is based on the recognition of linear peptide fragments bound to MHC-I or MHC-II by CD8+ and CD4+ T cells, respectively. T cell-based immunity is largely important in viral infections and other intracellular pathogens such as M. tuberculosis and

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Leishmania sp. Thus, information about linear peptide fragments/ epitopes can be easily generated from the side chain of amino acids in an epitope which interacts with MHC grooves. Interaction between the MHC molecule and epitope is a balance between affinity and restrictions with main and side chains of peptides [131]. Moreover, different MHC allomorphs have specific requirements for peptide sequences that can fit in the pockets which depend on the size and charge properties of peptides. It thus results in the selective nature of peptide–MHC interaction [132]. In silico proteome scanning for prediction of nonredundant and strong-affinity MHC-I and MHC-II epitopes belonging to conserved surface proteins of Leishmania identified 10 epitopes. Consequently, 5/10 peptides were able to induce PBMC proliferation from recovered individuals, thus indicating immunogenicity. Later, these peptides were observed to induce differential immune responses in subclinical, active, and posttreatment patient groups, although a Th1 response was a predominant characteristic among all peptides [133, 134]. Also, Leishmania LeIF and sandfly protein PpSp15 were used as model antigens to design an immunogenic, Th1 inducing and broadly conserved multiepitope vaccine against L. major, indicating the utility of in silico analytics in designing a potent vaccine by high-throughput immunoinformatics [135]. In recent years, high-throughput techniques have greatly increased the HLA-specific peptidome data for both HLA-I and HLA-II ligands. These MS-derived datasets have been incorporated in epitope prediction tools to increase their accuracy. Immune epitope database (IEDB) is the most widely used HLA-I allele-specific predictor, while NetMHCPan is a pan-allelic MHC-I ligand predictor. Ligand predictors for MHC-II alleles include NetMHCII, MHCPred, TEPITOPE, while NetMHCIIPan is a pan-allelic predictor for MHC-II molecule [136]. Predivac 2.0 was designed to tackle the problem of HLA polymorphism, and this software predicts the best immunodominant CD4+ T cell epitopes which bind to a wide variety of allelic variants of MHC found in geographically distinct human populations [137]. This is of prime importance when we are designing vaccines against diseases that spread across distinct continents. More recently, Dhandha et al. designed IFNEPITOPE which predicts IFNγ-inducing epitopes and regions in a protein sequence based on machine learning, motif search, and hybrid approach. A multiepitope chimeric vaccine peptide possessing higher antigenicity was designed for onchocerciasis using IFNEPITOPE and other in silico tools [138, 139]. After high-throughput sequencing techniques revolutionized the availability of whole-genome data, it became evident that interstrain variability is a common feature among all microbes’ viz. bacteria, viruses, parasites. Comparative genomics allows us to

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7.2 Utility of Comparative Genomics and Pangenome Analysis

mine the strain-wide genomic data of a pathogenic species to look for “conserved” and “unique” genes. It gave rise to the term “pangenome” which was first formally written down in bacterial context [125]. Comparative study of Streptococcus agalactiae (Group B streptococcus or GBS) isolates revealed that there are at least 30 unique genes present in each of the sequenced genome. Thus, pangenome was divided into three groups: (1) core genome, which is shared by all strains, (2) dispensable genes which are found in some of the strains, and (3) strain-specific genes which are a unique characteristic of a specific strain. Also, species are grouped as “open” or “closed” pangenome depending on the higher and lower diversity of strain-specific genes, respectively [125, 140]. Pangenome analysis becomes essential in vaccinology as it may lead to the identification of common genes and epitopes which can serve as a vaccine candidate for different pathogenic strains (ex. Staphylococcus aureus) or closely related species in a genus (Leishmania sp.). Indeed, there has been a plethora of evidence to prove that intra-species diversity exists at all levels of classification. Due to this fact, polyvalent vaccines have been designed for a broad level of protection against bacteria such as Streptococcus and Staphylococcus sp. The pangenome vaccinology approach may help us to develop vaccines that can be effective against all pathogenic serovars of a pathogen [141]. PanRV is one such multiapproach-based tool that identifies core, accessory, and unique genes by analysis of multiple genomes of a species. It identified seven genes that can be potential vaccine candidates by analysis of 301 genome sequences of Staphylococcus [142].

7.3 Genome Annotation, Subcellular Localization, and Antigenicity Prediction

The raw material for starting a reverse vaccinology experiment is the whole-genome sequence of the pathogen. Hence, the first step in a workflow utilizing a reverse vaccinology approach is to identify the putative coding regions in a genome which bookmarks the genecoding segments. It is based on the unique characteristics of start (ATG, GTG, and TTG) and stop codons (TAA, TAG, and TGA). So, any long genomic fragment within the boundary of these codons is assumed as a coding DNA sequence (CDS) [125]. Genome annotation helps us to predict novel hypothetical proteins with no functional annotation, which can be aligned with known protein sequences for predicting their putative functions via motif search tools. One such user-friendly tool is NCBI Conserved Domain Database (CDD) which aligns your query sequence with its database and predicts functionality based on protein domain matching. Prokaryotic genome annotation tools include GLIMMER based on Hidden Markov Model, while newer ones include DFAST. Eukaryotic annotation tools include web-based server tools such as EGPred [143–145].

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Subcellular localization prediction tools were primarily designed for bacterium where B cell immunity plays a major role in pathogen clearance. The main idea behind this was to identify the membrane/surface-exposed proteins which can directly interact with B cell receptor and elicit protective immunity. For intracellularly harboring pathogens such as viruses, some bacteria, and parasites, surface localization is not a prerequisite for a potential vaccine candidate, as the immunity is T cell based. Here, every protein has an equal chance of being presented via MHC molecules. Still, immunodominant and cryptic epitopes are shown to present in these conditions as well; however, factors are multivaried and have been reviewed elsewhere [146]. In short, subcellular localization can anyway provide us with the preliminary data needed for antigen identification or functional annotation, especially in the case of hypothetical proteins. These tools are generally based on the detection of N-terminal signal peptide sequences such as leader sequence, transmembrane helices, and certain motifs [125]. The presence of these signatures in a protein sequence determines its fate after translation in the ER. PSROTb is the most widely used tool for analyzing prokaryotic, archaeal genomes while SCLPred is used exclusively for eukaryotic genomes [147, 148]. Antigenicity of a protein/peptide sequence can also be predicted using in silico prediction tools. This is particularly useful for validating our designed vaccine peptide as a confirmatory step. There are many web-based tools for predicting the antigenic properties of protein/peptide sequences. Gupta et al. developed a machine learning-based tool, ProInflam, which groups the peptides into “inflammatory” and “noninflammatory” subtypes depending on their composition and motifs [149]. Context-Free Encoding Scheme (CfreeEns) is a machine learning-based approach for predicting the antigenicity and pathogenicity among different serotypes of a virus such as influenza [150]. Other popular tools include VaxiJen 2.0 and ANTIGENPro, which give an antigenic score for the input sequence [151, 152].

8 Reverse Vaccinology Against Leishmaniasis: Redefining Vaccine Candidate Selection In post-genomic era, reverse vaccinology gained momentum and became popular for the rapid identification of protective antigens against many pathogens. Since a vaccine against leishmaniasis has been a lingering subject despite years of investigations, scientists looked towards reverse vaccinology in search of new avenues. Since antileishmanial immunity is majorly T cell-based which is dependent on elicitation of host-protective CD4+, and recently, CD8+ as well. So, the strategy of in silico-based vaccine design focused on

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Fig. 5 Future of vaccination. Refinements in the delivery systems using nano vaccines, genetic engineeringguided approaches for improving vaccines include the use of synthetic antigens and gene circuits. Humanized mouse and primate models aim to closely mimic the host response, while in silico-based approaches such as reverse vaccinology based on machine learning improves the vaccine candidate selection

determining high-affinity, broad coverage epitopes to MHC alleles for generating an efficient multiepitope vaccine (Fig. 5). While Dikhit et al. predicted 9 high-affinity MHC-I epitopes that have binding capability and promiscuity to 40 HLA-I supertypes and are conserved among Leishmania sp., Kashyap et al. identified two peptides from LPG-3 and one peptide from nucleoside hydrolase (NH) of L. donovani which showed similar features. Moreover, Dikhit et al. observed that a cocktail of selected peptides was able to induce IFNγ and NF-κB–mediated antileishmanial immune response in BALB/c mice upon vaccination [153, 154]. A more comprehensive approach involved analyzing more than 8000 proteins common proteins among L. major and L. donovani, thus selecting 526 surface-localized proteins with nonsimilarity to the host proteome. Ultimately, 19 epitopes were selected which efficiently binds with predominant alleles of MHC-I and MHC-II molecules which are more likely to be selected for TCR recognition [155]. Secretory and membrane-exposed proteins of Leishmania are evaluated using reverse vaccinology-based approaches to generate efficacious vaccine candidates. A multiepitope vaccine consisting of high-affinity epitopes from five secretory proteins and 13 membrane proteins of L. donovani was constructed along with ribosomal 50S L7/L12 as an adjuvant. It was further refined for structural parameters and tested for physicochemical and immunogenic parameters by in silico tools, thus designing a potent and

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IFNγ-inducing vaccine candidate [156, 157]. In another study, Singh et al. designed a multiepitope containing immunogenic, nontoxic, nonallergenic, and host-protective immunity generating vaccine candidates by reverse vaccinology [158]. On a similar note, in silico-verified epitopes from 26 amastigote-expressing secretory proteins were used to design a multiepitope vaccine containing a TLR4-inducing property as well. A simulated vaccination study predicted the candidate to generate a skewed Th1 response [159]. A sequential subtractive approach was employed to screen membrane proteins expressed in all life stages of the parasite. Immunogenicity, nonallergenicity, and nonsimilarity to human proteome were also applied to sequentially filter the noncomplying candidates, which ultimately resulted in 16 potential vaccine candidates [160]. Interestingly, when an in silico designed multiepitope vaccine candidate was tested in vivo, vaccinated mice were able to generate a host-protective immune response consisting of higher IFN-γ, NO, and IgG2a levels and protected from a chronic VL infection. This suggests the potential of in silico deterministic tools in designing a better and efficacious vaccine against leishmaniasis [161]. Recently, 11 immunogenic epitopes were selected from promastigote-surface antigen (PSA), L. major large RAB GTPase (LmlRAB), and H2B (histone subunit) in a study. 3/25 peptide pools were able to induce IFNγ production from PBMC of recovered individuals and able to induce multifunctional Th1-specific effector and central memory T cells [162]. Thus, reverse vaccinology is advancing steadily in the field of antileishmanial vaccine development. It is bound to become more precise and accurate in its predictive ability due to an exponential increase in omics-based experimental data. Hence, in the future, we can see it actively complementing in vivo studies by a highthroughput selection of best antigenic candidates for vaccine trials (Fig. 6).

9 Proteomics-Based Approaches for High-Throughput Vaccine Discovery against Leishmaniasis With the development and refinement in cellular fractionation protocols, two-dimensional electrophoresis (2-DE), chromatographic, and spectroscopic techniques such as liquid chromatography and mass spectrometry, proteomics research have grown leaps and bounds. It very well complemented the genomics-based approaches to vaccine development [163]. While reverse vaccinology and comparative genomics allowed us to select the candidate antigens based on the sequence-inferred properties, proteomics

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Fig. 6 Timeline of vaccine discovery. Evidence-based (wet lab or in silico) selection of candidate antigen is followed by formulation design and preclinical trials in animal models to determine the best vaccine candidates which are taken up for clinical trials. Phase I, II, and III trials are performed to determine the safety and efficacy of vaccine candidates in the host population on a large scale. Postmarketing (Phase IV) trials are performed after commercialization to determine any unaccounted side effects

helped us in rapidly identifying vaccine candidates, characterizing a vaccine and deducing host response to vaccination. The proteomics approach has been successful in developing efficacious and superior vaccines against a variety of bacterial, viral, and parasitic diseases. Although the parameters for intended vaccine characteristics require to be different for each of these pathogens due to the varying nature of targeted immune profile, the crux of the approach remains largely similar. For vaccines targeting bacterial species, proteomic studies have allowed segregating the intracellular protein population into surfome (surface exposed) and secretome (extracellularly secreted) which are most likely candidates for antibacterial vaccine due to their extracellular nature. Using proteomics data, a multicomponent, surface proteinbased, wide-coverage vaccine preparation Trumenba® was developed successfully [164]. A combinatorial approach involving MS and in silico validation, Hornburg et al. identified jhp_0775 as a potential candidate that was inducing protective immunity in mice and humans as well [165]. Proteomic profiling of membrane and secretory proteins of P. falciparum by MS, followed by in silico and in vitro validation revealed novel candidate antigens which can serve as potential vaccine candidates [166]. Similarly, the exoproteome of Leishmania is long believed to be rich in immunomodulatory biomolecules [167] and has been established as a major player in the early as well as the late phase of infection [168, 169]. Thus, exoproteome analyses can be done in various Leishmania sp. for identification and validation of distinct as well as pan-species proteins which can be potent antigens in the context of vaccination. Low- and high-passage strains of

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L. amazonensis exhibited distinct proteome profiles, where 37 proteins (including 6 hypothetical proteins) were identified to be significantly downregulated in high-passage strains and proposed as virulent factors and potential vaccine candidates [170]. Recently, LC-MS–based proteomic analysis was performed to characterize the differences associated with a virulent and avirulent strain of L. major. It was observed that there are significant differences if proteome profile between the strains, indicating pathogenic attenuation. Consequently, an avirulent strain vaccine based on this study was found to induce significant protection against leishmaniasis [171]. 2-DE–based profiling of L. amazonensis and L. major revealed distinct interspecies protein expression variability as well; hence proteomic studies like these can complement the “pan-genome” approach and help us in achieving the goal of a pan-species antileishmanial vaccine [172]. Moreover, 97–68 kDa and 68–43 kDa fractions of splenic L. donovani amastigotes analyzed by proteomic approach identified 47 proteins (25 were hypothetical) which were able to induce host-protective Th1 response in mice and can serve as potential vaccine candidates [173]. Hence, as elucidated above, proteomics approaches have helped us in unraveling the protein compartmentalization, interspecies, and interstrain proteome variability and helped us in identifying novel vaccine candidates in our quest for an efficacious vaccine. In the coming years, proteomics would surely help us to decipher the molecular signatures associated with disease-specific protection postvaccination [174]. Also, it may turn into our main arsenal in the identification of the molecular nature of vaccines in the context of MHC–peptide interaction as well as predicting the nature of hosts’ response to subunit vaccines [175]. References 1. Zepp F (2016) Principles of vaccination. Methods Mol Biol 1403:57–84 2. Moser M, Leo O (2010) Key concepts in immunology. Vaccine 28(Suppl 3):C2–C13 3. Iborra S, Solana JC, Requena JM, Soto M (2018) Vaccine candidates against Leishmania under current research. Expert Rev Vaccines 17:323–334 4. McNicoll F, Drummelsmith J, Mu¨ller M et al (2006) A combined proteomic and transcriptomic approach to the study of stage differentiation in Leishmania infantum. Proteomics 6:3567–3581 5. Biyani N, Madhubala R (2012) Quantitative proteomic profiling of the promastigotes and the intracellular amastigotes of Leishmania donovani isolates identifies novel proteins

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Chapter 4 Status of COVID-19 Pandemic Before the Administration of Vaccine Sunil Thomas Abstract Severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) is responsible for the disease COVID-19 that has decimated the health and economy of our planet. The virus causes the disease not only in people but also in companion and wild animals. As yet we do not know why the virus is highly successful in causing the pandemic within 3 months of its first report. Lack of a voice on how to handle the pandemic impacted the management of the disease globally. Publication of the importance of masks and social distancing in preprint servers reduced the spread of the disease and deaths associated with it. Very few countries have invested in science and research and development and that has impacted the development of therapies for the pandemic. Though vaccination against COVID-19 started in December 2020, slower rate of immunizations has resulted in rapid spread of the mutant strains of SARS-CoV-2. Lack of transparency and accountability coupled with anergic leadership was responsible for the high incidence of disease and death associated with the COVID-19 pandemic. Key words COVID-19, SARS-CoV-2, Pandemic, Virus structure, Anergic leadership, Social distancing, Vaccine, Disease management

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Introduction “The saddest aspect of life right now is that science gathers knowledge faster than society gathers wisdom.”—Isaac Asimov. The coronavirus disease 2019 (COVID-19) is currently responsible for the pandemic that has decimated the health and economy of every country. COVID-19 is regarded as a respiratory disease that manifests with fever, cough, shortness of breath or difficulty breathing, chills, muscle pain, headache, sore throat, and loss of taste and smell. Other symptoms include diarrhea, nausea, and vomiting [1, 2]. Many patients with the COVID-19 are asymptomatic but

The original version of the chapter has been revised. A correction to this chapter can be found at https://doi.org/ 10.1007/978-1-0716-1884-4_38 Sunil Thomas (ed.), Vaccine Design: Methods and Protocols, Volume 1: Vaccines for Human Diseases, Methods in Molecular Biology, vol. 2410, https://doi.org/10.1007/978-1-0716-1884-4_4, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022, corrected publication 2022

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are able to transmit the virus to others [3, 4]. The prolonged pandemic has resulted in social distancing, travel restrictions, decreased trade, high unemployment, commodity price decline, closure of educational institutes, and financial stress that has impacted the global economy. COVID-19 disease is caused by the severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2), a member of the betacoronavirus genus. The disease has resulted in a mortality of 0.5–8.0%. The death rate has varied with time. As yet, there are no effective drugs available for the treatment of the disease [5]. Vaccinations against COVID-19 started in December 2020. As of writing this chapter, it is not known how long it takes to vaccinate the global population, at least to people who are at risk of the disease.

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Origin and Transmission In December 2019, a group of patients with pneumonia of unknown cause were confirmed to be infected with a novel coronavirus, known as 2019-nCoV, in Wuhan, Hubei province, China, which had previously not been detected in humans or animals. Epidemiological evidence suggested that most of these patients had visited the Huanan seafood market in Wuhan and that the gene sequence of the virus obtained from these patients was highly similar to that identified in bats. The virus was subsequently renamed SARS-CoV-2 as it was similar to the coronavirus responsible for severe acute respiratory syndrome (SARS-CoV) [6]. On Dec 30, 2019, Li Wenliang, an ophthalmologist at Wuhan Central Hospital, China, sent a message to a group of fellow doctors warning them about a possible outbreak of an illness that resembled severe acute respiratory syndrome (SARS) in Wuhan, Hubei province, China, where he worked. Li raised the alarm after he saw seven patients with SARS-like symptoms. Li reported the suspected outbreak to his colleagues in a closed group on the WeChat social media platform after learning that patients were being quarantined. Meant to be a private message, he encouraged them to protect themselves from infection. Days later, he was summoned to the Public Security Bureau in Wuhan and made to sign a statement in which he was accused of making false statements that disturbed the public order. Nevertheless, Li decided to speak out about his experience because, “I think a healthy society should not have just one voice,” as he told Caixin Media. Li returned to work after signing the statement and contracted severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2), apparently from a patient. Dr. Li died after becoming infected with SARS-CoV-2 in Wuhan, China, on Feb 7, 2020, aged 33 years [7]. The World Health Organization (WHO) on March 11, 2020, declared the novel coronavirus (COVID-19) outbreak a global pandemic. Bioinformatic analyses revealed that SARS-CoV-2 has characteristics typical of coronavirus family. It belongs to the betacoronavirus

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2B lineage [8]. Data indicated the closest relationship of SARS-CoV2 with the bat SARS-like coronavirus strain BatCov RaTG13, with an identity of 96%. It is hypothesized that SARS-CoV-2 could be of bat origin, and SARS-CoV-2 might be naturally evolved from bat coronavirus RaTG13 [9, 10]. Betacoronavirus isolated from pangolins has a sequence similarity of up to 99% with the currently infected human strain [11]. Another study indicates that SARS-CoV-2 and the coronavirus from a pangolin in Malaysia has high genetic similarity. The gene similarity between these two viruses in terms of E, M, N, and S genes is 100, 98.6, 97.8, and 90.7%, respectively, suggesting the potential for pangolins to be the intermediate host [12]. SARS-CoV-2 replicates efficiently in cats, tiger, lion, minks, ferrets, and golden hamsters [13–16]. SARS-CoV-2 is transmitted via fomites and droplets during close unprotected contact between the infected and uninfected. Symptomatic and asymptomatic patients are the main source of infection. The virus can also spread through indirect contact transmission. Virus-containing droplets contaminate hands which upon contact with the mucous membranes of the mouth, nose, and eyes cause infection. Several studies have demonstrated the aerosol transmission of SARS-CoV-2 [17]. In some pediatric SARS-CoV-2 infection cases, although children’s nasopharyngeal swabs are negative, rectal swabs are consistently positive, indicating the possibility of fecal-oral transmission [18]. Recent studies demonstrate that SARS-CoV-2 could replicate effectively in human intestinal organoids and intestinal epithelium. The study demonstrated that SARSCoV-2 has the potential to spread through intestinal tract [19].

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Symptoms Caused by the SARS-CoV-2 Virus Some people with SARS-CoV-2 infection remain asymptomatic, while in others the infection can cause mild to moderate COVID19 disease and COVID-19 pneumonia, leading some patients to require intensive care support. In some cases, the patients die due to the disease; older adults are more prone to death. Symptoms such as fever or cough, and signs such as oxygen saturation or lung auscultation findings, are the first and most readily available diagnostic information. Such information could be used to either rule out COVID-19 disease or select patients for further diagnostic testing. People with mild COVID-19 might experience cough, sore throat, high temperature, diarrhea, headache, muscle or joint pain, fatigue, and loss of sense of smell and taste. Symptoms of COVID-19 pneumonia include breathlessness, loss of appetite, confusion, pain or pressure in the chest, and high temperature [20]. Diabetes mellitus is consistently one of the most common comorbidities found in patients with COVID-19. A large proportion of patients with COVID-19 requiring hospitalization and/or

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succumbing to the disease have had diabetes and other chronic conditions as underlying risk factors [21, 22]. Patients with diabetes and severe COVID-19 have higher pro-inflammatory cytokines and a worse survival rate than nondiabetics [23, 24]. Individuals with diabetes are more susceptible to COVID-19 than those without diabetes. SARS-CoV-2 infection could induce hyperglycemia. Angiotensin-converting enzyme 2 (ACE2) may be the key regulator that involved in the association between COVID19 and hyperglycemia. The main role of ACE2 is to incise angiotensin II to generate angiotensin 1–7 and thereby mediates the protective effects of vasodilation, anti-inflammatory, and antiproliferation. In addition, ACE2 is identified as a receptor that facilitates coronavirus entry into cells [25]. A recent research noted that ACE2 expression was substantially increased in patients with diabetes mellitus than those without diabetes [26]. The enhanced susceptibility to COVID-19 infection in patients with diabetes may be attributed to overexpression of ACE2. The expression of ACE2 was higher in the pancreas than in the lungs, indicating that SARS-CoV-2 may bind to ACE2 in the pancreas and cause pancreatic injury [27]. COVID-19 infection is also known to induce an increase in blood glucose levels even for those not diagnosed with diabetes before admission [25, 28, 29]. Hypoxia is a leading cause of multiple organ injuries and death in COVID19 patients. Aggressive oxygen therapy is known to cure these patients [30]. COVD-19 could also impact eyesight for a cohort of patients. Acute viral retinitis is manifested in some patients with COVID-19, and there are also reports of severe blindness due to viral infection [31]. Mucormycosis (black fungus) is reported in COVID-19 patients in India. The disease is caused by a group of molds called mucormycetes. Mucormycosis may be caused by Rhizopus species, Mucor sp., Rhizomucor sp., Syncephalastrum sp., Cunninghamella bertholletiae, Apophysomyces sp., and Lichtheimia sp. The mucor mold is commonly found in soil, plants, manure, and decaying fruits and vegetables [32]. Drop in immunity due to medications may be the reason for mucormycosis [33]. In addition to mucormycosis, COVID-19 patients are susceptible to invasive candidiasis (white fungus) caused by Candida auris [34, 35].

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Structure of SARS-CoV-2 Coronaviruses belongs to the subfamily Coronavirinae in the family of Coronaviridae and the subfamily contains four genera: Alphacoronavirus, Betacoronavirus, Gammacoronavirus, and Deltacoronavirus. The genome of CoVs (27–32 kb) is a single-stranded positive-sense RNA (+ssRNA) which is larger than any other

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RNA viruses [36]. The size of the SARS-CoV-2 is around 120 nm in diameter. SARS-CoV-2 is an enveloped virus with a singlestranded RNA genome of 29.8 kb. More than two-thirds of the genome comprises Orf1ab encoding 16 nonstructural proteins (nsps) followed by mRNAs encoding structural proteins, spike (S), envelop (E), membrane (M), and nucleocapsid (N). These genes are interspaced with several accessory genes (open reading frames [Orf] 3a, 3b, 6, 7a, 7b, 8, 9b, 9c, and 10). The functions of these proteins are of particular interest for understanding the pathogenesis of SARS-CoV-2. Several of the nsps (nsp3, nsp4, nsp6) and Orf3a are transmembrane proteins involved in regulating the host immunity, modifying host cell organelles for viral replication and escape and hence considered drug targets [37]. The major structural proteins of SARS-CoV-2 are spike (S), membrane (M), envelop (E), and the nucleocapsid (N) proteins [5]. The trimeric spike glycoprotein of SARS-CoV-2 uses the host angiotensin-converting enzyme 2 (ACE2) as the entry receptor [38]. The virus binds to host cells through its trimeric spike glycoprotein, making this protein a key target for potential therapies and diagnostics. Amraie et al. [39] recently reported that the C-type lectin receptors CD209L/L-SIGN and CD209/DSIGN serve as alternative receptors for SARS-CoV-2 entry into human cells. The C-type lectin domain could function as a calcium-dependent glycan-recognition domain. The most abundant structural protein of coronaviruses is the M glycoprotein; it spans the membrane bilayer, leaving a short NH2-terminal domain outside the virus and a long COOH terminus (cytoplasmic domain) inside the virion. The M protein can bind to all other structural proteins. Binding with M protein helps to stabilize N proteins and promotes completion of viral assembly by stabilizing the N protein–RNA complex, inside the internal virion. As the M protein cooperates with the S protein, mutations may influence host cell attachment and entry of the viruses. The S protein of the virus is glycosylated, and this modification may aid in immune evasion. However, it is not known how the S protein is glycosylated. The M glycoprotein is the most abundant envelope protein of SARS-CoV-2. In silico analyses of the M protein of SARS-CoV2 using Protter demonstrated that it has a triple-helix bundle and forms a single three-transmembrane domain (Fig. 1). In addition, the M glycoprotein has a short amino terminal domain outside the viral envelope and a long carboxy-terminal domain inside the viral envelope. The M protein of SARS-CoV-2 resembles the sugar transporter, Sugars Will Eventually be Exported Transporters (SWEET). Upon analysis, it was observed that other coronaviruses including SARS-CoV, bat SARS-CoV, pangolin SARS-CoV, and MERS-CoV have M proteins homologous to the sugar transporter SWEET. Further analysis by residue-based structure demonstrated

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a

b SARS-COV-2 membrane protein (M) H2N-

PTMs variants disulfide bonds signal peptide N-term: UniProt TMRs: UniProt 10

Intravirion

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Fig. 1 The membrane (M) protein of SARS-CoV-2. (a) Predicted M protein structure of SARS-CoV-2 (ribbon diagram) using the software I-TASSER. (b) Membrane topology of proteins (snake diagrams) determined using Protter. The membrane (M) glycoprotein of SARS-CoV-2 has a triple helix bundle and formed a single threetransmembrane domain

that the protein has the characteristic structure of SemiSWEET, the sugar transporter of prokaryotes [5]. It is still not understood the functions of the M proteins. However, it is known that viral infection triggers metabolic reprogramming in host cells to facilitate optimal virus production. It could be hypothesized that the SemiSWEET sugar transporter-like structure of the M protein may be involved in multiple functions that may aid in the rapid proliferation, replication, and immune evasion of the SARS-CoV2 virus [5]. In silico analysis demonstrated that M protein of SARS-CoV2 is 98.6% similar to the M protein of bat SARS-CoV and maintains 98.2% homology with pangolin SARS-CoV, and 90% homology with the M protein of SARS-CoV; whereas, the similarity is only

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38% with the M protein of MERS-CoV. Thus, the M protein of SARS-CoV-2 resembles the M protein of bat and pangolin SARSCoV to a greater extent than MERS-CoV [5]. To evade detection by host innate immune sensors, viruses that replicate in the cytoplasm compartmentalize their genome transcription in organelle-like structures, thereby protecting the virus against host cell defenses and increasing the replication efficiency [40]. The transmembrane nsp3, nsp4, and nsp6 are known to rearrange endoplasmic reticulum (ER) membranes thereby inducing curvature of the ER membrane, essential for virus replication. Lack of antiviral therapies is the paucity of knowledge regarding the β-coronavirus–host cell interface [41]. We recently mapped the transmembrane and lumen domains of the nsps of SARS-CoV2 [37].

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Variants of SARS-CoV-2 Viruses generally acquire mutations over time, giving rise to new variants. Several variants of the SARS-COV-2 are reported recently. Lineage B.1.1.7 is also known as the UK variant. B.1.1.7 variant may be associated with an increased risk of death compared with other variants. This variant has a mutation in the receptor-binding domain (RBD) of the spike protein at position 501, where the amino acid asparagine (N) has been replaced with tyrosine (Y). This variant is associated with increased transmissibility (i.e., more efficient and rapid transmission). The 501Y.V2 variant (lineage B.1.351) is the South African strain. This variant shares some mutations with B.1.1.7. This variant has multiple mutations in the spike protein, including K417N, E484K, N501Y. Some of the vaccines may not be protected by this variant. P.1 is a close relative of the B.1.351 lineage and is the Brazil variant. This variant has 17 unique mutations, including three in the receptor-binding domain of the spike protein. This variant was first reported by the National Institute of Infectious Diseases (NIID) in Japan in four travelers from Brazil. The P.1 lineage contains three mutations in the spike protein receptor-binding domain: K417T, E484K, and N501Y [42]. Lineage B.1.429 is the California variant (CAL.20C). The S protein L452R mutation is within a known receptor-binding domain that has been found to be resistant to certain S protein monoclonal antibodies [43]. The new variant (from India) is the “double mutant” as it contains two key mutations, called E484Q and L452R, that have been found separately in other variants but not together in a single strain.

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The Indian SARS-CoV-2 variant is B.1.617. It is one of the most contagious variants known [44] (https://www.who.int/en/ activities/tracking-SARS-CoV-2-variants/). Some of the potential consequences of emerging variants include the ability to transmit rapidly, ability to cause milder or severe disease, ability to evade detection by PCR, ability to decrease susceptibility to monoclonal antibodies, and ability to evade vaccine induced immunity [42].

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Immediate Scenario After Partial or Complete Lockdown Due to COVID-19 The essential commodities including napkins, tissue rolls, disinfectant, and bleach were not available in stores or were rationed for a couple of weeks. In some countries, public transportation services were halted without notification and people were stranded without food or water [45]. The Olympic Games that was to be held in July 2020 in Japan was canceled due to fear of the rapid spread of the disease. Movie theaters, universities, schools, churches, offices, and games were closed since the start of the partial or complete lockdown. Public transports including air travel were reduced due to fear of the disease. Countries that had enforced strict lockdowns could control the disease; whereas countries including USA and Sweden, where economy took precedence than health of the population, had a high increase in COVID-19. The death rate in the USA was the highest compared to other countries primarily due to lack of enforcement of lockdown during the pandemic. There was a shortage of masks, including N95 mask for hospital personnel. Most countries including United States had limited stock of masks, gloves, PPEs, and ventilators for patients. Hospitals were reusing masks during the initial pandemic months. By February 23, 2021, more than 500,000 people died due to COVID-19 in the United States alone since the start of the pandemic. The global death was around 2.5 million, with 112.5 million infected with the disease. The unofficial figures (death and infection) in most countries may be several times higher than the official figures.

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Drugs Used in the Treatment of COVID-19 As yet there are no effective drugs for the treatment of COVID-19. During the COVID-19 pandemic, several people in key positions touted scrupulous drugs including Lysol, a disinfectant for treatment. Gullible people resort to anything to ward off infection. Several drugs including hydroxychloroquine and remdesivir, though not tested, were tried on patients. None of the drugs lowered the death in people affected by COVID-19. In developing

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countries where people pay for treatment (out of pocket, not through medical insurance), the hospitals extorted people who came down with mild COVID-19. The treatment drugs consisted of a cocktail of drugs including zinc, vitamin C, and some antivirals and hospital stay for a week! All these drugs are not effective for the treatment of COVID-19. In May 2021, emergency use approval was granted in India for anti COVID-19 therapeutic application of the drug 2-deoxy-Dglucose (2DG). 2-DG is a repurposed cancer drug. Cancer cells require significantly more glucose than normal cells to grow. Inhibition of glycolysis could lead to shrinkage of some cancers. In patients with COVID-19, 2DG works by inhibiting glycolysis as well as inhibition of glycolytic enzymes 6-phospho-fructo-2kinase/fructose-2,6-bisphosphatase-3, a positive regulator of phosphofructokinase-1 as well as lactate dehydrogenase A that inhibits viral replication [46].

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Behavioral Pattern that Slowed the Virus Preprint servers are online repositories containing scholarly papers that are not yet peer reviewed or accepted by traditional academic journals. Authors can submit revised versions of their papers to the preprint server at any time. The pandemic saw exponential increase in publication in preprint servers. The information on the use of mask and social distancing was initially published in the preprint servers. Public health policies could be formulated and updated based on the papers published in the preprint servers. In the absence of effective and safe vaccines or antivirals to control the disease, strategies for mitigating the burden of the pandemic are focused on nonpharmaceutical interventions, such as social distancing, hand washing, contact tracing, quarantine, isolation, and the use of face masks in public [47]. Handwashing has received considerable attention during the COVID-19 pandemic. Handwashing with soap and water for at least 20 s or the use of alcohol-based hand sanitizers when soap and water are not available is the first line of defense in stopping the spread of infection. It has been shown that wearing a mask reduces the contact transmissibility by reducing transmission of infected droplets in both laboratory and clinical contexts. Public mask-wearing is most effective at reducing the spread of the virus when compliance is high (Fig. 2). The decreased transmissibility could substantially reduce the death toll and economic impact while the cost of the intervention is low [48]. The community-wide benefits are likely to be greatest when face masks are used in conjunction with other nonpharmaceutical practices such as social distancing, and when adoption is nearly universal (nation-wide) and compliance is high. Chu et al. [49] support physical distancing of 1 m or more and

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Fig. 2 A Google doodle image encouraging the use of face mask and social distancing during the COVID-19 pandemic

hypothesized that contact tracing could reduce the disease transmission. Use of face mask and social distancing has saved millions of lives.

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Poor Leadership Impacted the Spread of the Virus Globally Leaders are not super humans to understand and to react to every incident or needs around them. However, they have experienced advisers giving suggestions and advice on important matters. It is up to the leader to execute or implement decisions in a timely manner based on suggestion of the advisers, his/her gut instincts, education, experience, and knowledge. Anergy is a term in immunology that describes a lack of reaction by the body’s immune system to fight foreign substances. Unresponsive or anergic leaders delay decisions to the extent that their slow actions have a negative impact on the well-being of the society they serve. Anergic leaders ignore or are not aware of the need of the society they serve. Anergic leaders have less compassion and empathy to the needs of the society. Anergic leaders also populate leadership positions with unqualified personnel that pay homage and only listen to him. Some of these leaders see everything in the prism of economics. A delayed decision has ramification that extends to several generations. In many countries, the pandemic showed how anergic leadership negatively impacted the society. They downplayed the importance of the pandemic, and this led to the flaring of the disease and the deaths associated with it. Lockdowns and cease of economic activities would have drastically cut deaths due to COVID-19.

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COVID-19 in India Most of the countries had adequate time to prepare against COVID-19. However, none of the countries (developed or developing) were prepared to face COVID-19. India’s healthcare budget is just 1.3% with respect to its GDP, which is woefully inadequate. There is a lack of functioning healthcare facilities in most rural areas. Though India is home to some of the largest vaccine companies, the government did not place orders for COVID-19 vaccines (same as the Trump administration in the US). In most Indian states, there was no social distancing or mask requirements. The government also encouraged mass religious gatherings and elections in different states that had large crowds. By April 2021, the second COVID-19 wave had set in. The new variant (B.1.617) was deadlier than any other variant. Millions of people were infected, and thousands (or millions) perished. Hospitals were overflowing; there were no beds and people died due to lack of oxygen. Crematoriums were working 24  7. In the states of Uttar Pradesh (UP) and Bihar, bodies were found floating in rivers due to lack of cremation. It was stated in local media that more than 1600 teachers lost their lives due to elections (where the teachers had to work as election officials). Even after the incidence of death spiraled there was no immediate lockdown. Early lockdown would have decreased the spread of the disease. The federal government was not keen to order vaccines but spend huge amounts of money building new Parliament, offices, and residences for the governing class. The full scale of disease and deaths due to the second wave of COVID-19 will only be known after a year.

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Distribution of Vaccines The COVID-19 vaccines have been distributed since December 2020. The current vaccines are based on mRNA, adenovirus vectors, or attenuated viruses (Fig. 3). The developed countries have more access to vaccines compared to developing countries. The rate of vaccination is slower than anticipated. As of February 15, 2021, only 2.5 doses were administered per 100 people. Israel had the best immunization programs. By February 2021, more than 84% of Israel’s population 70 and older had received both shots of Pfizer’s Covid-19 vaccine. Mass vaccinations have led to a drop in severe cases among the 70-and-older population [50]. COVAX is one of three pillars of the Access to COVID-19 Tools (ACT) Accelerator, which was launched in April 2020 by the World Health Organization (WHO), the European Commission, and France in response to this pandemic. Bringing together governments, global health organizations, manufacturers,

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Fig. 3 Pfizer BioNTech COVID-19 vaccine

scientists, private sector, civil society, and philanthropy, with the aim of providing innovative and equitable access to COVID-19 diagnostics, treatments, and vaccines for developing countries. The COVAX pillar is focused on the vaccines. It is the only truly global solution to this pandemic because it is the only effort to ensure that people in all corners of the world have access to COVID-19 vaccines once they are available, regardless of their wealth [51]. COVAX hopes to deliver more than two billion vaccine doses to people in 190 countries in less than a year. The organization is interested to ensure 92 poorer countries will have access to vaccines at the same time as 98 wealthier countries.

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Wastage of Vaccines Pfizer’s and Moderna’s vaccine vials contained more than the number of doses displayed on their labels. It is common for vaccine manufacturers to overfill their vials to account for regular waste as doses are prepared and administered. The Pfizer vial originally intended for five 0.3-milliliter (ml) doses, or 1.5 ml total, actually contains 2.25 ml of vaccine once the liquid is diluted for injection. Moderna vials intended for 10, 0.5-ml doses, 5 ml total, often have 6 ml. But getting those extra doses out requires the right syringe. The standard vaccine syringe holds 3 ml, but a thinner, 1-ml syringe is often used for vaccines with small doses, like Pfizer’s and Moderna’s. Because it is narrower, the smaller syringe traps less liquid between the plunger and the tip of the needle after a dose is expelled. And specially designed “low dead-volume” syringes,

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which come in both sizes, have plungers that slide all the way down to the needle to eliminate most of this trapped liquid. As the vaccine distribution centers were not supplied by the right syringe, there was a wastage of vaccine. Given that the United States has bought 300 million Moderna doses so far, it stands to waste some 30 million doses by failing to enable vaccinators to draw out an 11th dose [51].

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Impact of COVID-19 on Climate Change Wearing of mask, social distancing, work-from-home during the COVID-19 pandemic led people to confine to their homes. There were very few incidences of influenza in the communities during the pandemic [52]. As the economic activity has slowed considerably, there are fewer vehicles on the road and few airplanes in the sky. During COVID-19 pandemic, employees had the option to work from home. Overall, the slow economic activity has resulted in cleaner air in most regions of the planet. The Philadelphia region (where this author is based) has more snowy days in early 2021 than in the previous 5 years. As people have seen and experienced the change in climate due to decreased pollution during COVID-19 pandemic, it may lead to new policies and environmental regulation in the future.

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Preparing for a Pandemic in the Future The COVID-19 pandemic demonstrated that one-fourth of the population of a country should have access to hospitals in case of a pandemic. There should be enough PPE and other protective clothing for 2 years. There should be access to ventilators and oxygen for one-fiftieth of the population. Food should be available, free of cost for people in middle- and low-income countries. Economic activities should cease, and mandatory lockdown initiated based on the incidence of the disease. People should be encouraged to work online, educational institutes and places of worship closed during the pandemic. Elections should be postponed if possible. If vaccines are available, order should be placed for all the population and vaccine administration should be completed within 6 months. The quicker the vaccines are administered, chances of highly contagious variants arising will be lesser.

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Part II Trends in Vaccinology

Chapter 5 mRNA Vaccines to Protect Against Diseases Sunil Thomas and Ann Abraham Abstract Infectious diseases are a leading cause of death worldwide, and vaccines are the cheapest and efficient approach to preventing diseases. Use of conventional vaccination strategies such as live, attenuated, and subunit has limitations as it does not fully provide protection against many infectious diseases. Hence, there was a need for the development of a new vaccination strategy. Use of nucleic acids—DNA and RNA—has emerged as promising alternative to conventional vaccine approaches. Knowledge of mRNA biology, chemistry, and delivery systems in recent years have enabled mRNA to become a promising vaccine candidate. One of the advantages of a mRNA vaccine is that clinical batches can be generated after the availability of a sequence encoding the immunogen. The process is cell-free and scalable. mRNA is a noninfectious, nonintegrating molecule and there is no potential risk of infection or mutagenesis. mRNA is degraded by normal cellular processes, and its in vivo half-life can be regulated by different modifications and delivery methods. The efficacy can be increased by modifications of the nucleosides that can make mRNA more stable and highly translatable. Efficient in vivo delivery can be achieved by formulating mRNA into carrier molecules, allowing rapid uptake and expression in the cytoplasm. The severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) emerged in late 2019 and spread globally, prompting an international effort to accelerate development of a vaccine. The spike (S) glycoprotein mediates host cell attachment and is required for viral entry; it is the primary vaccine target for many candidate SARS-CoV2 vaccines. Development of a lipid nanoparticle encapsulated mRNA vaccine that encodes the SARSCoV-2 S glycoprotein stabilized in its prefusion conformation conferred 95% protection against Covid-19. Key words mRNA vaccine, DNA vaccine, Disease, Infection

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Introduction Vaccines have proven to be the most effective and efficient approach to preventing diseases and controlling outbreaks. Conventional vaccine approaches, such as live-attenuated and inactivated pathogens and subunit vaccines, provide durable protection against several diseases. Despite this success, there remain major hurdles to vaccine development against a variety of infectious pathogens, especially those better able to evade the adaptive immune response or those that do not induce any cellular response. For most

Sunil Thomas (ed.), Vaccine Design: Methods and Protocols, Volume 1: Vaccines for Human Diseases, Methods in Molecular Biology, vol. 2410, https://doi.org/10.1007/978-1-0716-1884-4_5, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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emerging virus vaccines, the main obstacle is not the effectiveness of conventional approaches but the need for more rapid development and large-scale deployment. Conventional vaccine approaches may not be applicable to noninfectious diseases, such as cancer. Hence, there is a need for the development of more potent and versatile vaccine platforms [1]. Over the years, researchers have been on the search for novel and advanced approaches to vaccine design and development. Therapeutics using nucleic acid have emerged as promising alternatives to conventional vaccine approaches. Wolff et al. (1990) for the first time reported injecting RNA of chloramphenicol acetyltransferase, luciferase, and beta-galactosidase separately into mouse skeletal muscle in vivo. Protein expression was readily detected in all cases, and no special delivery system was required for these effects [2]. However, these early promising results did not lead to substantial investment in developing mRNA therapeutics, largely owing to concerns associated with mRNA instability, high innate immunogenicity, and inefficient in vivo delivery [1]. Major technological innovations over the past several years have enabled mRNA to become a promising therapeutic tool in the fields of vaccine development. The use of mRNA has several beneficial features over subunit, killed, and live-attenuated virus, as well as DNA-based vaccines. The most important being safety; mRNA is a noninfectious, nonintegrating molecule and there is no potential risk of infection or mutagenesis. mRNA is degraded by normal cellular processes, and its in vivo half-life can be regulated by different modifications and delivery methods. The efficacy can be increased by modifications of the nucleosides that can make mRNA more stable and highly translatable. Efficient in vivo delivery can be achieved by formulating mRNA into carrier molecules, allowing rapid uptake and expression in the cytoplasm. mRNA is not combined with other vectors; hence, there is no concern for anti-vector immunity, and mRNA vaccines can be administered repeatedly. mRNA vaccines have the potential for rapid, inexpensive, and scalable manufacturing, primarily due to the high yields of in vitro transcription reactions [1]. The severe acute respiratory syndrome coronavirus 2 (SARSCoV-2) emerged in the late 2019 and spread globally, prompting an international effort to accelerate development of a vaccine. The S glycoprotein mediates host cell attachment and is required for viral entry; it is the primary vaccine target for many candidate SARS-CoV-2 vaccines. Multiple mRNA vaccines from different companies were tested for their efficacy. The candidate vaccine mRNA1273 (developed by Moderna) was a lipid nanoparticle– encapsulated, nucleoside-modified messenger RNA (mRNA)– based vaccine that encodes the SARS-CoV-2 spike (S) glycoprotein stabilized in its prefusion conformation. The mRNA-1273 vaccine induced anti–SARS-CoV-2 immune responses in all participants,

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and no trial-limiting safety concerns were identified (Jackson et al. 2020). The BNT162b2 mRNA vaccine (developed by Pfizer/ BioNTech) candidate conferred 95% protection against Covid-19 in persons 16 years of age or older. Safety over a median of 2 months was similar to that of other viral vaccines [3, 4]. One of the advantages of an mRNA vaccine is that within weeks of design and development, clinical batches can be generated after the availability of a sequence encoding the immunogen. The process is cell-free and scalable. The greatest advantage is that a facility dedicated to mRNA production should be able to rapidly manufacture vaccines against multiple targets, with minimal adaptation to processes and formulation. In addition, new targets requiring multi-antigen approaches will benefit from the speed in which mRNA can render multiple constructs [5].

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Nucleic Acid Vaccines Protect Against Infection Compared to conventional protein/peptide-based vaccines intended to induce antigen-specific adaptive immune responses, DNA vaccines are more stable, cost-efficient, easy to manufacture, and safe in handling. DNA vaccines are considered as a thirdgeneration vaccination approach in which antigenic materials are encoded as DNA plasmids for direct in vivo production to elicit adaptive immunity. As compared to other platforms, DNA vaccination is considered to have a strong safety profile, as DNA plasmids neither replicate nor elicit vector-directed immune responses in hosts [6]. Successful in vivo transfection of mammalian cells following injection of purified DNA was first reported by Atanasiu et al. (1962) [7]. However, its potential went largely unrealized until Wolff et al. (1990) reported that proteins could principally be expressed either by direct injection of DNA or messenger RNA (mRNA) into mouse skeletal muscles [2]. Tang et al. (1992) delivered DNA of human growth hormone into the skin of mice using a “gene gun” [8]. They demonstrated that the technique was an easy method to generate antibody in the host and could be a method for the development of a vaccine. Ulmer et al. (1993) generated a viral antigen for presentation to the immune system without the limitations of direct peptide delivery or viral vectors [9]. They injected plasmid DNA encoding influenza A nucleoprotein into the quadriceps of BALB/c mice. This resulted in the generation of nucleoprotein-specific CTLs and protection from a subsequent challenge with a heterologous strain of influenza A virus. DNA vaccine concept has been tested and applied against various pathogens and tumor antigens. In theory, the DNA vaccines are safe, as non-live vaccine approach is a unique and technically simple means to induce immune responses. DNA vaccines

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affect not only humoral immunity but also cellular immunity. The optimized gene sequence of interest is delivered to the skin (intradermally), subcutaneously or muscle by one of several delivery methods. Using the host cellular machinery, the plasmid enters the nucleus of transfected cells, including resident antigen presenting cells (APCs). Here, expression of plasmid-encoded genes is followed by generation of foreign antigens as proteins that have been converted to peptide strings. These host-synthesized antigens can become the subject of immune surveillance in the context of both major histocompatibility complex (MHC) class I and class II molecules of APCs in the vaccinated host. Antigen-loaded APCs travel to the draining lymph nodes where they “present” antigenic peptide–MHC complexes in combination with signaling by co-stimulatory molecules to naive T cells. This interaction provides the necessary secondary signals to initiate an immune response and to activate and expand T cells or, alternatively, to activate B cell and antibody production cascades. Together, both humoral and cellular immune responses are engendered [10]. In a DNA vaccine, the genetic material must first enter the host cell’s nucleus. In the nucleus, messenger RNA is created, which travels out of the nucleus into the cytoplasm, where protein is formed from it. However, genetic information can only enter the nucleus when the cell is dividing, making the process inefficient. Hence, DNA vaccines are not very popular due to the low efficiency in inducing antigenic protein production in the cells. The ability to make new DNA vaccines without the need to handle a virulent pathogen or to adapt the pathogen for manufacturing purposes demonstrates the potential value of this vaccine technology for use against emerging and epidemic pathogens [11]. Currently, there are no approved DNA vaccines for use in humans. Nevertheless, some DNA-based vaccines were approved by the FDA and the USDA for veterinary use, including a vaccine against West Nile virus in horses and canine melanoma. One of the first human clinical trials with DNA vaccines evaluated the therapeutic and prophylactic effects against HIV in which no significant immune response but a potential immunogenicity was detected. Another clinical trial which targeted the hepatitis B virus showed the induction of a humoral response in patients not responding to conventional vaccination. The overall safety of DNA vaccines has been thoroughly proven in several clinical trials, underlined by the fact that no antibody response against prokaryotic parts of the DNA vaccine itself has been observed and that adverse effects are limited to mild local reactivity at the injection site [12]. Issues have been raised with regard to the safety of DNA vaccines, these include the potential to integrate into cellular DNA, the development of autoimmunity, and the possibility of antibiotic resistance. DNA vaccines that are currently being tested do not show relevant levels of integration into host cellular DNA.

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However, vectors that are modified or adjuvanted with the goal of increasing immunogenicity could increase the chances of integration. A further concern is that an integrated vaccine might cause insertional mutagenesis through the activation of oncogenes or the inactivation of tumor suppressor genes. In addition, an integrated plasmid DNA vaccine could, in theory, result in chromosomal instability through the induction of chromosomal breaks or rearrangements. However, none of these concerns have been witnessed in the preclinical or clinical evaluation of DNA products [10]. The principal issue regarding the future of DNA vaccines concerns improving their immunogenicity in larger animals and in humans. The DNA vaccine platform has driven significantly weaker immune responses in nonhuman primates and in humans compared with mice. It seems to be less immunogenic compared with recombinant viral vectors such as adenoviral vectors or recombinant protein for induction of antibody responses [10].

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Development of mRNA as a Vaccine The feasibility of an mRNA vaccine was first demonstrated by Martinon et al. [13]. The authors induced anti-influenza cytotoxic T lymphocytes (CTL) in vivo by immunizing mice with liposomes containing messenger RNA (mRNA) encoding the influenza virus nucleoprotein (NP). Furthermore, with the same mRNA-liposome preparation, virus-specific CTL responses could be also elicited in mice of three different haplotypes each of them known to present a distinct NP peptide in an MHC-restricted fashion. mRNA vaccines were not popular initially due to the instability of mRNA, high innate immunogenicity, inefficient in vivo delivery difficulties to ensure long-term storage, the high cost of manufacturing GMP grade material, and inefficient in vivo delivery [1, 14]. The development of mRNA as a vaccine benefitted due to FPLC purification, and the modification of the nucleosides. RNA activates cells of the innate immune system by stimulating Toll-like receptors (TLRs), specifically TLR3, TLR7, and TLR8. However, when naturally occurring modified nucleosides, for example, pseudouridine (Ψ), 5-methylcytidine (m5C), N6-methyladenosine (m6A), 5-methyluridine (m5U), or 2-thiouridine (s2U), were incorporated into the transcript, most of the TLRs were no longer activated. Dendritic cells (DCs) exposed to such modified RNA express significantly less cytokines and activation markers than those treated with unmodified RNA. DCs and TLR-expressing cells are potently activated by bacterial and mitochondrial RNA, but not by mammalian total RNA, which is abundant in modified nucleosides. The modified nucleosides abrogated 50 -triphosphate RNA-mediated activation of another RNA-responsive immune sensor, retinoic acid-inducible protein I (RIG-I). Thus, in vitro

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transcripts containing nucleoside modifications would remain translatable and also avoid immune activation in vivo, such an mRNA is preferred as a therapeutic tool for vaccination [15]. mRNAs containing pseudouridines have a higher translational capacity than unmodified mRNAs when tested in mammalian cells and or administered intravenously into mice. The delivered mRNA and the encoded protein could be detected in the spleen 24 h after the injection, where both products were at significantly higher levels when pseudouridine-containing mRNA was administered. Even at higher doses, only the unmodified mRNA was immunogenic, inducing high serum levels of interferon-α (IFN-α). Incorporating pseudouridine, a naturally occurring modified nucleoside, into mRNA not only suppresses RNA-mediated immune activation in vitro and in vivo but also enhances the translational capacity of the RNA. These characteristics and the ease of generating such an RNA by in vitro transcription make Ψ-containing mRNA a unique tool for expression of any protein in vitro and in vivo [16]. There are several advantages of in vitro transcribed mRNA that contribute to its vaccine potential. The development process of an mRNA vaccine is faster than conventional protein vaccines. mRNA vaccine enables the synthesis of antigen proteins in situ, eliminating the need for protein purification and long-term stabilization which are challenging for some antigens. Advanced industrial setup can manufacture mRNA up to kilogram scales. mRNA vaccine enables the synthesis of antigen proteins in situ, eliminating the need for protein purification and long-term stabilization which are challenging for some antigens. Due to these advantages, mRNA vaccines have great potential to be manufactured and deployed in a timely manner in response to rapid infectious disease outbreaks [17].

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Types of mRNA Vaccines There are two types of mRNA vaccines: conventional mRNA vaccines and self-amplifying mRNA vaccines. Conventional mRNAbased vaccines encode the antigen of interest and contain 50 and 30 untranslated regions (UTRs) and a terminal poly(A) tail, whereas self-amplifying RNAs encode not only the antigen but also the viral replication machinery that enables intracellular RNA amplification and abundant protein expression. Self-amplifying mRNA can be derived from the engineered genomes of Sindbis virus, Semliki Forest virus, Kunjin virus, among others. After the purified RNA replicon is delivered into host cells, either as viral particles or as synthetically formulated RNA, it is translated extensively and amplified by its encoding RNA-dependent RNA polymerase. Compared with the rapid expression of conventional mRNAs, published results have shown that vaccination with self-amplifying mRNA vaccines results in higher antigen expression levels, although delayed in time, which

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persist for several days in vivo. Equivalent protection is conferred but at a much lower RNA dose. Due to the lack of viral structural proteins, the replicon does not produce infectious viral particles. Conventional and self-amplifying mRNA vaccines cannot integrate into the host and degrade naturally during antigen expression. These factors indicate the viability and safety of mRNA vaccines compared to traditional approaches to vaccine design [18].

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Formulation and Delivery of mRNA Vaccines Despite mRNA’s appealing features and advances in the field, in vivo delivery of mRNA remains challenging. The first challenge is the instability of mRNA mostly due to enzymatic degradation by RNases. RNases are present ubiquitously throughout the body to degrade exogenous RNAs. Naked mRNA is quickly degraded by extracellular RNases and is not internalized efficiently. mRNA, consisting of hundreds to thousands of nucleotides, has to reach the cytosol in full length for active translation. Hence, protection against RNases is critical for most in vivo delivery strategies. Efficient intracellular delivery of mRNA is a challenge owing to the negative charge and large size of mRNA molecules. The negative charge prevents most mRNA from translocating across the negatively charged cell membrane. The large size makes efficient encapsulation and delivery more challenging than other molecules. Various delivery strategies have been investigated to address these obstacles with different delivery materials, formulation methods, and routes of administrations [17]. Several transfection reagents have been developed that facilitate cellular uptake of mRNA and protect it from degradation. Once the mRNA transits to the cytosol, the cellular translation machinery produces protein that undergoes posttranslational modifications, resulting in a properly folded, fully functional protein. This feature of mRNA pharmacology is particularly advantageous for vaccines and protein replacement therapies that require cytosolic or transmembrane proteins to be delivered to the correct cellular compartments for proper presentation or function. mRNA is finally degraded by normal physiological processes, thereby reducing the risk of metabolite toxicity [1]. Efficient delivery of mRNA into target cells in vivo is a major challenge. A variety of formulations have been developed to protect the nucleic acid from RNases and facilitate its uptake into cells. Positively charged lipids, cationic polypeptides, polymers, micelles, or dendrimers have been used for in vivo RNA delivery. Lipid nanoparticles (LNPs) are appropriate carriers for mRNA in vivo and are valuable tools for delivering mRNA encoding therapeutic proteins [19]. Lipids, lipid-like compounds, and lipid derivatives are widely used to formulate lipid and LNPs for in vivo delivery of mRNA

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vaccines. LNPs are generally defined as nano-sized particulate systems that are composed of synthetic or physiological lipid materials. LNPs can encapsulate RNA molecules, protecting RNA from enzymatic degradation. LNPs effectively deliver mRNA molecules into the cytosol of cells by endocytosis mechanisms. The endocytosis process transports mRNA-loaded LNPs into cell membrane-bound vesicles, including endosomes and lysosomes [17]. The final step of RNA release from LNPs into the cytosol might involve the membrane disruption of endosomes. The cationic or ionizable lipid materials, such as 1,2-di-O-octadecenyl-3-trimethylammonium propane (DOTMA), N,N-Dimethyl-2,3-bis[(9Z,12Z)-octadeca9,12-dienyloxy]propan-1-amine (DLinDMA), and N1,N3,N5tris(3-(didodecylamino)propyl)benzene-1,3,5-tricarboxamide (TT3), usually contain one or multiple amino groups. These lipid materials can be positively charged at a certain pH to encapsulate the negatively charged RNA molecules via electrostatic interactions and help interact with the cell membrane on target cells [17]. Delivery routes can affect the in vivo distribution pattern and expression kinetics of encapsulated mRNA vaccines. Immunization by intramuscular (IM), intradermal (ID), subcutaneous (SC) administrations deliver LNPs mRNA vaccine to resident/ infiltrating APCs and related immune cells, stimulating strong and prolonged local expression [17]. Polymeric materials, including polyamines, dendrimers, and copolymers, are functional materials capable of delivering mRNA vaccines. Similar to functional lipid-based carriers, polymers can also protect RNA from RNase-mediated degradation and facilitate intracellular delivery. Cationic polymers, such as polyethylenimine (PEI), polyamidoamine (PAMAM) dendrimer, and polysaccharide, condensed and delivered negatively charged RNA molecules. PEI is a widely used polymeric material for mRNA vaccine delivery. PEI formulations are often prepared by direct mixing PEI solution with RNA solution. Besides cationic polymer materials, anionic polymers, such as PLGA, are also used to deliver mRNA vaccines. As an anionic polymer is not able to efficiently encapsulate the negatively charged mRNA molecules, cationic lipid materials are added to create lipid–polymer hybrid formulations [17]. The polymeric materials used in combination with mRNA are only tested in animals. Other agents used in the delivery of mRNA include peptides, or the use of virus-like replicon particle (VRP). As yet, the only mRNA vaccine successfully developed and available commercially is the SARS-COV-2 vaccine.

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mRNA Vaccines Against SARS-CoV-2 SARS-CoV-2 is responsible for the disease COVID-19 that has decimated the health and economy of our planet. The virus causes the disease not only in people but also in companion and wild

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animals. The first reported case turned up in Wuhan, China, and transmission of the disease has quickly plagued the world. Transmission of the disease is primarily due to direct contact with someone who is infected or by respiratory droplets that are in close proximity (~6 feet) and coming in contact on a time-dependent manner (~15 min or longer) [20]. By the end of March 2021, 125 million people were infected with the virus, leading to around three million deaths due to the virus. However, it has been recognized that reported numbers may be underestimates of actual infection cases since individuals may be asymptomatic and carriers of the virus, without showing signs of any symptoms of the virus. Common symptoms of the virus have been identified as fever, dry cough, fatigue, and dyspnea, while severe symptoms include systemic infection and pneumonia. Age, health, and behavior of the population impacted the death rate due to COVID-19. Old people, people with underlying diseases such as diabetes, lung diseases (due to smoking), liver disease, cardiovascular disease, and obesity are more prone to death due to COVID-19 [20, 21]. Taxonomically, SARS-CoV-2 is classified as a member of the species SARS-related coronavirus (SARSr-CoV) in the genus Betacoronavirus of the family Coronaviridae. Zhou et al. (2020) identified a closely related SARSr-CoV genome sequence, RaTG13, which shared a 96% whole-genome sequence identity with SARSCoV-2, indicating a probable bat origin of SARS-CoV-2 [22]. Though many non-vaccination treatment options have been explored (antiviral drugs, cell-based therapy, convalescent plasma therapy), a vaccine is a proven and most efficient method necessary to ensure individual protection and the development of herd immunity. Across the world, more than 120 vaccine candidates were in various preclinical and phase 1 to 3 clinical trials that included inactivated, live-attenuated, viral-vectored replicating and nonreplicating, protein- and peptide-based, and nucleic acid approaches [23]. Due to the ease of manufacture, safety, and efficiency, developing an mRNA-based vaccine against SARS-COV2 was found to be more feasible [23]. Data from preclinical studies, animal models, and other data have indicated the S protein is the major target of neutralizing antibodies, and many of these antibodies target the RBD of the S protein, neutralizing antibodies produced by vaccination are protective in animal models, and most of the vaccine candidates induced cellular immunity [24]. In March 2020, Moderna and the Vaccine Research Center (VRC) of the National Institute of Allergy and Infectious Diseases (NIAID) conducted the first phase I clinical trial of a novel lipid nanoparticle (LNP)-encapsulated mRNA-based vaccine, which encoded the spike protein (S protein) [2]. Since SARS-CoV-2 is a novel and emerging virus, there is very limited knowledge on the immune response to SARS-CoV-2. However, with existing vaccine candidates, data and animal models from SARS and MERS; the early

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publication of the full-length genome sequence of SARS-CoV-2; the similar sequence in the S protein between SARS-CoV-1 and SARS-CoV-2; and the use of DNA and RNA vaccine methods, researchers and biotech companies were able to fashion vaccine candidates for the virus. The less-restrictive regulatory guidelines for vaccine development were also instrumental in fast-tracking the process of vaccine development. The outcomes of phase III clinical studies of mRNA vaccines proved efficacy in preventing SARS-CoV-2 infection by 95% and 94.5%. Pfizer and BioNTech COVID vaccine, BNT162b2, became the first mRNA-based vaccine to be authorized for emergency use in the UK and the USA [25]. With much vigor and persistence, Pfizer and Moderna designed effective vaccines against SARS-CoV2 using mRNA technology that was released to the public in December 2020. Emergency Use Authorization of Pfizer and BioNTech’s mRNA vaccines have provided hope for the eradication of exponential spread of the COVID-19 virus. However, longterm immunity still needs to be monitored, in the chance a booster shot might be required for continued immunity against the virus. With optimism, the SARS-COV-2 mRNA vaccines will diminish the spread of the virus. Immune response to a pathogen is often heterogeneous and varies between individuals according to age, underlying health conditions, and the environment. In order to achieve herd immunity, at least 70% of the total population would need to be vaccinated [25]. Another drawback to the COVID vaccines is long-term storage as the vaccine must be stored at 70  C, which poses challenges in distributing. Nevertheless, these vaccines were used in the immunization programs of North America. The SARS-CoV-2 mRNA vaccines are the only mRNA vaccines available commercially.

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mRNA Vaccines Targeting Influenza Avian influenza A viruses (H5N1, H10N8, H7N9, and H1N1) have caused severe respiratory diseases in humans. Hemagglutinin (HA) and neuraminidase (NA) are crucial glycoproteins for infection and both are expressed on the surface of the influenza A virus. HA is the gatekeeper that allows viral entry into the host cells. Existing antiviral therapeutics have been proven ineffective; therefore, vaccination is the most effective method of protection against influenza. Conventional influenza vaccines use the HA protein to generate neutralizing antibodies against the globular head domain and are produced in embryonated chicken eggs or cell substrates. The process is lengthy and demanding, as sufficient supplies are needed to grow the virus [26]. The demands of the epidemic spread exceed vaccine production; therefore, quicker methods of

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vaccine production are necessary to fight the perpetual influenza epidemic. The influenza virus has been the most extensively studied pathogen due to the availability of tools, the ease of testing efficacy in small-animal models, and the beneficial aspects of an mRNA-based vaccine for influenza. While conventional vaccines have been approved, production of vaccine takes about 6 months. In comparison, self-amplifying mRNA vaccines can be generated within a shorter period of time. In 2013, during an outbreak of a deadly strain of H7N9 influenza in China, the HA gene was cloned in a self-amplifying mRNA vaccine pDNA template, and the selfamplifying mRNA vaccine was produced within 8 days after publication of the HA gene sequence [27]. Petsch et al. (2012) demonstrated that an injection with unmodified conventional mRNA encoding various influenza virus antigens combined with a protamine-complexed RNA adjuvant was immunogenic in mice, ferrets, and domestic pigs [28]. Several studies afterwards showed that mRNA-based influenza vaccines formulated in LNP or CNE-induced T and B cell immune responses. It has also been shown that chitosan, polyethylenimine (PEI), and dendrimer-based formulations of self-amplifying mRNA are effective. Lutz et al. [29] used a single IM injection of LNP-formulated HA-encoding mRNA to induce hemagglutination inhibition (HAI) titers in NHPs [29]. Whereas, Lindgren et al. [30] demonstrated the protective levels of antibodies to hemagglutinin were induced after two immunizations of modified nonreplicating mRNA, which encodes influenza H10 in lipid nanoparticles (LNP) in nonhuman animals [30]. A “universal” influenza vaccine capable of inducing strong immune responses has been introduced as a viable approach. Using an LNP-formulated mRNA vaccine, Pardi et al. [31] demonstrated that one immunization induced antibodies against the immune subdominant HA stalk in mice, ferrets, and rabbits [31]. Recently, the first human trial of an mRNA-based influenza vaccine was reported. The vaccine was composed of an LNP-formulated, nucleoside-modified conventional mRNA encoding of an H10N8 HA antigen. The vaccine was immunogenic in all subjects with 100% and 87% of vaccinated subjects achieving HAI titers 40 and micro-neutralization titers 20 [32].

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mRNA Vaccines Targeting Rabies Rabies is a neurological disease that infects mammals, including humans. The virus is a single-stranded RNA virus of the Lyssavirus family. Humans are primarily infected by way of dog bites, but the virus can be transmitted through coyotes, skunks, and bats. Rabies is estimated to cause 59,000 human deaths annually [33]. Rabies is most prevalent in low- and middle-income countries where the

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contact with canines is high [34]. Development of a rabies vaccine has met many roadblocks due to production capacities, required administration schedules, storage requirements, and cost. Bacterial and viral vaccine approaches were pursued, but the efforts were met with variable success. A promising approach is encoding the rabies virus antigen, RABV-G in mRNA. CureVac applied mRNA technology to develop a rabies vaccine candidate, CV7201 [34]. Alberer et al. demonstrated the first-in-human clinical trial in healthy adults of a prophylactic mRNA-based vaccine encoding the rabies virus glycoprotein [35]. Volunteers received three doses of CV7201, mRNA rabies vaccine candidate, intradermally or intramuscularly by needle-syringe or needle-free devices. The study showed that mRNA vaccine candidates are capable of inducing functional antibodies against a viral antigen. The trial determined that vaccination with CV7201 is safe regardless of the route of administration; however, the vaccine was immunogenic only when administered using needle-free devices. Preclinical studies have shown B-cell and T-cell immune responses in mice and pigs when injected intradermally and intramuscularly. CV7201-induced neutralizing antibodies in mice that protected against intracranial challenge with the rabies virus. In guinea pigs, needle-free injection of CV7201 resulted in increased immunogenicity [35]. Unfortunately, immune responses in subjects declined after 1 year. It was determined that immune responses from the mRNA rabies vaccine could be increased by improving the delivery system. Recently, a CV7202 mRNA vaccine in phase I clinical trial was developed. The mRNA that carries the information for the formation of the rabies virus protein (RABV-G) is injected into muscle. Two 1 μg or 2 μg doses of CV7202 were well tolerated and elicited rabies neutralizing antibody responses that met WHO criteria in all recipients [36].

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mRNA Vaccines Against Zika Virus The Zika virus, primarily transmitted by mosquitoes, was first found in a rhesus macaque and in mosquitoes in the Zika forest (Uganda) in 1947 and 1948. In 1952, the first human case arose [37]. The Zika virus caught the world’s attention when it caused an epidemic on Yap Island, Micronesia, French Polynesia, and other regions of the South Pacific, and later spread to the Americas infecting >700,000 people [38]. The virus has infected approximately 1.5 million people. ZIKV stems from two genetic lineages (African and Asian) with MatchMaker. Select the reference model (closest PDB structure) and the structure to match (the I-TASSER choose model) > Ok (Fig. 2). Use the coordinates of lipid bilayer to select only surface-exposed regions on the outer membrane. 4. Set up MicroPulser™ electroporator (Bio-Rad, USA) for bacterial transformation following manufacturer’s instructions. Add 1–2 μL of each ligation reaction to 40 μL of cell suspension, mix gently with pipette tip, and incubate on ice for 1 min. Transfer the cells to the chilled electroporation cuvette. Electroporate the cells as recommended by manufacturer, using the preprogrammed setting Ec2 (2.5 kV and time constant of  5 ms). Immediately, add 500 μL room temperature LB medium. Transfer the solution to a 1.5 mL microtube and shake (180 rpm) for at least 1 h at 37  C. Centrifuge the tubes for 1 min at 8000
g, suspend the pellet in 100 μL of

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Fig. 2 Parameters for align of PDB reference closest structure with I-TASSER predict 3D model for target protein

LB, and spread it on a prewarmed LB plate containing ampicillin (100 μg/mL). Incubate the plates for 16–18 h at 37  C. Select colonies and lyse the cells for plasmid isolation (see Note 5). 5. Select and identify colonies grown on selective LB medium. Add 15 μL of phenol–chloroform solution and 15 μL of lysis buffer to a 1.5 mL microtube. Add one identified colony to this tube, mix well, and centrifuge at 14,000
g for 4 min. Apply the upper phase (approximately 15 μL) in agarose gel 1%. Check for recombinant clones using as control the pAE vector without insert. 6. Besides Buffers B and D containing urea, a buffer containing N-lauroylsarcosine (0.2 g for each 100 mL) can also be evaluated to protein solubilization. In this case, after sonication, allow protein solubilization for 72 h under mild agitation (70 rpm), then follow the protocol as described above. 7. If basal expression is observed in the preinduction sample, a better control can be achieved by addition of glucose on expression medium. For glucose 1 M solution, dissolve 180.156 g of glucose (dextrose) into 1 L of distilled water. Sterilize through a 0.22 μm filter. During inoculum

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preparation, add glucose solution to a final concentration of 1% per mL of LB medium. 8. For an initial volume of 1 L of Buffer B (initial concentration of 8 M of urea), discard 250 mL and replace it for 250 mL of PBS every 1 h. N-lauroylsarcosine (0.2%) could be used to replace 8 M urea for solubilization of insoluble proteins that reaggregated during dialysis. In this case, the inclusion bodies are solubilized in Buffer B, containing 0.2% N-lauroylsarcosine instead of 8 M urea, incubated at 4  C for 72 h for complete solubilization. Purification and dialysis are performed as described above. 9. General anesthesia protocols, like ketamine/xylazine, can also be used. For more information about general anesthesia and other ways to collect hamster blood by gingival vein puncture, consult Rodriguez et al. 2017 [46]. References ˜ ones MDC et al 1. Martı´nez R, Pe´rez A, Quin (2004) Efficacy and safety of a vaccine against human leptospirosis in Cuba. Rev Panam Salud Publica 15(4):249–255. https://doi.org/10. 1590/S1020-49892004000400005 2. Verma R, Khanna P, Chawla S (2013) Wholecell inactivated leptospirosis vaccine: future prospects. Hum Vaccin Immunother 9:763–765. https://doi.org/10.4161/hv. 23059 3. Pereira MM, Schneider MC, Munoz-Zanzi C et al (2017) A road map for leptospirosis research and health policies based on country needs in Latin America. Rev Panam Salud Pu´blica 41:e131. https://doi.org/10.26633/ RPSP.2017.131 4. Xu Y, Ye Q (2018) Human leptospirosis vaccines in China. Hum Vaccin Immunother 14:984–993. https://doi.org/10.1080/ 21645515.2017.1405884 5. Garba B, Bahaman AR (2018) Advances & challenges in leptospiral vaccine development. Indian J Med Res 147:15–22. https://doi. org/10.4103/ijmr.IJMR_1022_16 6. Adler B, de la Moctezuma A (2010) Leptospira and leptospirosis. Vet Microbiol 140:287–296. https://doi.org/10.1016/j.vetmic.2009.03. 012 7. Vernel-Pauillac F, Werts C (2018) Recent findings related to immune responses against leptospirosis and novel strategies to prevent infection. Microbes Infect 20:578–588. https://doi.org/10.1016/j.micinf.2018.02. 001

8. Bulach D, Adler B (2017) Leptospiral genomics and pathogenesis. In: Adler B (ed) Spirochete biology: the post genomic era, 1st edn. Springer International, Berlin, p 295 9. Dellagostin OA, Grassmann AA, Rizzi C et al (2017) Reverse vaccinology : an approach for identifying Leptospiral vaccine candidates. Int J Mol Sci 18:158. https://doi.org/10.3390/ ijms18010158 10. Grassmann AA, Kremer FS, dos Santos JC et al (2017) Discovery of novel leptospirosis vaccine candidates using reverse and structural vaccinology. Front Immunol 8:463. https://doi. org/10.3389/fimmu.2017.00463 11. Zeng LB, Wang D, Hu NY et al (2017) A novel pan-genome reverse vaccinology approach employing a negative-selection strategy for screening surface-exposed antigens against leptospirosis. Front Microbiol 8:396. https://doi. org/10.3389/fmicb.2017.00396 12. Conrad NL, McBride FWDC, Souza JD et al (2017) LigB subunit vaccine confers sterile immunity against challenge in the hamster model of leptospirosis. PLoS Negl Trop Dis 11:e0005441 13. Coutinho ML, Choy HA, Kelley MM et al (2011) A ligA three-domain region protects hamsters from lethal infection by Leptospira interrogans. PLoS Negl Trop Dis 5:1–10. https://doi.org/10.1371/journal.pntd. 0001422 14. Raja V, Sobana S, Mercy CSA et al (2018) Heterologous DNA prime-protein boost immunization with RecA and FliD offers

Human Recombinant Leptospirosis Vaccines cross-clade protection against leptospiral infection. Sci Rep 8:1–9. https://doi.org/10. 1038/s41598-018-24674-8 15. Oliveira TL, Rizzi C, da Cunha CEP et al (2019) Recombinant BCG strains expressing chimeric proteins derived from Leptospira protect hamsters against leptospirosis. Vaccine 37:776–782. https://doi.org/10.1016/j.vac cine.2018.12.050 16. Oliveira TL, Schuch RA, Inda GR et al (2018) LemA and Erp Y-like recombinant proteins from Leptospira interrogans protect hamsters from challenge using AddaVax™ as adjuvant. Vaccine 36:2574–2580. https://doi.org/10. 1016/j.vaccine.2018.03.078 17. Monaris D, Sbrogio-Almeida ME, Dib CC et al (2015) Protective immunity and reduced renal colonization induced by vaccines containing recombinant Leptospira interrogans outer membrane proteins and flagellin adjuvant. Clin Vaccine Immunol 22:965–973. https:// doi.org/10.1128/CVI.00285-15 18. Teixeira AF, Cavenague MF, Kochi LT et al (2020) Immunoprotective activity induced by Leptospiral outer membrane proteins in Hamster model of acute leptospirosis. Front Cell Infect Microbiol 11:568694. https://doi. org/10.3389/fimmu.2020.568694 19. Berven FS, Flikka K, Jensen HB, Eidhammer I (2004) BOMP: a program to predict integral β-barrel outer membrane proteins encoded within genomes of gram-negative bacteria. Nucleic Acids Res 32:W394. https://doi.org/ 10.1093/nar/gkh351 20. Madan Babu M, Sankaran K (2002) DOLOPdatabase of bacterial lipoproteins. Bioinformatics 18(4):641–643 21. Setubal JC, Reis M, Matsunaga J et al (2006) Lipoprotein computational prediction in spirochaetal genomes. Microbiology 152 (Pt 1):113–121. https://doi.org/10.1099/ mic.0.28317-0 22. Yu NY, Wagner JR, Laird MR et al (2010) PSORTb 3.0: improved protein subcellular localization prediction with refined localization subcategories and predictive capabilities for all prokaryotes. Bioinformatics 26:1608–1615. https://doi.org/10.1093/bioinformatics/ btq249 23. Almagro Armenteros JJ, Tsirigos KD, Sønderby CK et al (2019) SignalP 5.0 improves signal peptide predictions using deep neural networks. Nat Biotechnol 37:420–423. https://doi.org/10.1038/s41587-019-0036z 24. Krogh A, Larsson B, von Heijne G, Sonnhammer ELL (2001) Predicting transmembrane

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Chapter 17 Induction of T Cell Responses by Vaccination of a Streptococcus pneumoniae Whole-Cell Vaccine Emily M. Roy, Fan Zhang, Richard Malley, and Ying-Jie Lu Abstract The induction of T cell responses by vaccination is important for protection against infection. We have previously shown that immunization with a killed Streptococcus pneumoniae whole-cell vaccine (SPWCV) by either intranasal immunization or subcutaneous immunization induced T cell responses to SPWCV. Protection against colonization by S. pneumoniae is dependent on CD4+ IL-17A production induced by immunization. Here, we present detailed protocols for preparation of SPWCV, immunization of mice, and assay for T cell responses in blood and splenocytes in immunized mice. Key words T cell, Vaccine, Immunization, Streptococcus pneumoniae

1

Introduction Vaccination is designed to train the immune system how to battle potential future infections. Vaccines are available for prevention of infection by many bacteria and virus and are in urgent need for the COVID-19 pandemic, which has caused many illnesses and deaths worldwide due to lack of preventative medicine [1, 2]. In order for a vaccine to be fully effective, it needs to activate the innate immune system and both arms of the adaptive immune system, the humoral immunity and the cell-mediated immunity. Humoral immunity protects the host from extracellular pathogens and toxins, while cell-mediated immunity protects against intracellular pathogens [3]. B cells and T cells are the major cellular components of the adaptive immune response. B cells are primarily responsible for humoral immunity, whereas T cells are involved in cell-mediated immunity. With help from T cells, B cells can produce high-affinity antibody against proteins, polysaccharides, and other surface markers on pathogen cells. B-cells can also recognize pathogenic antigens in their native form without the help of antigen-presenting

Sunil Thomas (ed.), Vaccine Design: Methods and Protocols, Volume 1: Vaccines for Human Diseases, Methods in Molecular Biology, vol. 2410, https://doi.org/10.1007/978-1-0716-1884-4_17, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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cells and T-helper cells. These antigens are typically large repeating molecules such as capsular polysaccharide, dextran, and bacterial polymeric flagellin [3]. 1.1 Subtypes and Functions of T Cells

T cells are identified by CD3, a surface cluster of differentiation (CD) molecule, and are comprised of two major groups: the CD4 and CD8 cells. The CD4 cells provide helper activities on other populations of cells and are subdivided into at least Th1, Th2, Th9, Th17, and T regulatory (Treg) cells. The CD8 T cytotoxic population is the second major group of T cells that functions in killing target cells. CD8 T cells are comprised of Tc1 and Tc2 subpopulations with similar cytokine profiles as Th1 and Th2 cells. The function of T cells is mainly the regulation of all functions of the adaptive immune system and can be specified into these four areas: promotion of inflammation by cytokine production (Th1 and Th17 cells); helping B cells (Th2 cells); regulating immunosuppressive responses (T regulatory cells); and killing of unwanted target cells (CTL). Naı¨ve T cells must be activated and differentiated in order to induce the appropriate immune response to a pathogen. T cell differentiation is, in part, dictated by the stage of development the T cell is in when it receives the secondary co-stimulatory signal along with the primary interaction between the T cell receptor and the antigen-bound major histocompatibility complex (MHC). CD4 cells recognize the MHC II complex found on all immune cells. CD8 cells recognize MHC I protein which is a marker of body cells and is found on all nucleated body cells except for mature erythrocytes.

1.2 T Cell Induction by Infection of Bacteria and Virus

T cells can be activated during infection by bacteria or viruses, and this activation of T cells may form a memory response that protects the body from future infections by the same pathogen. Mycobacterium tuberculosis, Klebsiella pneumoniae, and Salmonella enterica induce activation of Th17 cells to protect against extracellular bacterial infections at mucosal sites [4]. Similarly, infection by Streptococcus pneumoniae colonization or Staphylococcus aureus also induces protective Th17 cells against future infections [5, 6]. Virus infection will lead to the induction of CD8 CTL cells, and memory CD8 T cells are capable of protecting against secondary infections [7]. Components of viruses and fungi have been shown to induce differentiation of Th17 cells [4]. When the cornea is infected with Herpes simplex virus, IL-23 and IL-17A production are induced, indicating that T cells are activated [8]. IL-17A is also induced when cells are infected with influenza [8].

1.3 T Cell Induction by Vaccination

As Th1 and Th17 pathways have been shown to be important during infection with certain organisms, it stands to reason that it

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would be essential to activate these components of the immune system with vaccines. Historically, vaccines have been known to induce antibody responses to the infection. More recently, studies have shown that inducing T cell activation is also important. Vaccine-induced T cell response persists much longer and contains phenotypically and functionally diverse populations of cells comparing to vaccine-induced antibody response [9]. In general, the injection of proteins without adjuvant or with aluminum adjuvant does not lead to detectable Th1 or Th17 responses [10–12]. Induction of Th1 or Th17 responses to protein antigens requires novel adjuvants such as TLR ligands or curdlan [13]. Such a requirement may create significant hurdles for clinical trials, particularly in children in whom novel adjuvants are challenging to test. In part to circumvent this problem, multiple antigen presenting system (MAPS), a novel technology that enables the creation of macromolecular complexes consisting of polysaccharides that are affinity-bound to proteins, was developed [12, 14]. When adjuvanted with aluminum hydroxide and injected subcutaneously or intramuscularly in mice, these complexes induce robust Th1/Th17 responses to any incorporated protein. The MAPS technology has been evaluated for the preclinical development of vaccines against several pathogens, including S. pneumoniae, Mycobacterium tuberculosis, Salmonella typhi, and Staphylococcus aureus, and showed T cell induction for the protein antigens [12, 15]. 1.4 T Cells Induced by Whole-Cell Vaccine

There are many different types of vaccines, each with their benefits and drawbacks. Live-attenuated vaccines produce long-term immunity and a robust immune response. Examples of this group of vaccines include smallpox, yellow fever, measles, mumps, rubella, chickenpox, and typhoid fever. Inactivated vaccines are more stable but produce a weaker immune response. These vaccines require booster shots to maintain immunity [3]. An example of an inactivated vaccine is the whole-cell pertussis vaccine. The whole-cell pertussis vaccine induces Th1 and Th17 immunity and provides better protection against pertussis lung colonization than the acellular pertussis vaccine in animal models [16]. Similarly, a killed Streptococcus pneumoniae whole-cell vaccine (SPWCV) induces Th17 responses and protected pneumococcal colonization in the nasal pharyngeal in mice models [10, 17–19]. Mice can be immunized either by intranasal route or by other mucosal routes such as buccal or sublingual to generate T cell responses against S. pneumoniae whole-cell antigen (SPWCA); however, any of these routes require a mucosal adjuvant, for which enterotoxins such as cholera or nontoxic mutated derivatives have been used [19]. Later, we showed that T cell generation could also be induced by immunizing mice with SPWCA mixed with aluminum adjuvants, which are routinely and safely used in human vaccination

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[18]. These mice similarly displayed accelerated nasopharyngeal clearance, which correlated with their priming for IL-17A responses. In addition to T cell responses against SPWCA, parenteral immunization also induced much higher antibody responses than that from intranasal immunization and protected mice against invasive disease infections in mice models. Thus, parenteral immunization of mice with SPWCA revealed a bifunctional immunity; with a nonlethal intranasal challenge, clearance is CD4-dependent and IL-17A–mediated; in a lethal aspiration pneumonia model, survival is CD4-independent and dependent on plasma antibodies [20]. Recently, we demonstrated that parenteral immunization of SPWCA generated tissue resident memory T cell populations, in addition to the circulating memory T cells, which play an important role in protecting mice from nasal colonization of S. pneumoniae [21].

2

Materials Induction of T cell response by whole-cell vaccine includes the following steps: preparing whole cell vaccine, vaccine formulation and immunization, and assay for T cell responses in immunized mice. In the following sections, we will describe the necessary process to generate a S. pneumoniae whole-cell vaccine and measure immune responses by immunization of SPWCV in mice. Strain Selection

For induction of T cell response, an unencapsulated bacterium is preferred since the presence of capsule on the surface might affect the internalization of the bacteria and thus interfere with antigen presentation after immunization. Another consideration for strain selection is the detoxification of toxins. For example, the Streptococcus pneumoniae toxin, pneumolysin, can cause hemolysis of red blood cells. Mutations need to be introduced to remove the hemolysis activity of the toxin, but still keep the correct conformation to induce neutralization antibody when being used as an immunogen. One example is the introduction of PdT (triple mutations: Asp-385 to Asn, Cys-428 to Gly, and Trp-433 to Phe substitutions) in the pneumococcal whole-cell strain. The third consideration for strain selection is to remove the autolysis property. S. pneumoniae undergoes autolysis at stationary phase so it is very important to knock out lytA gene from the whole-cell strain. As a result, the strain used for our pneumococcal whole-cell vaccine strain was a Rx1 (no capsule) strain with delta lytA and PdT mutations.

2.2 Culture Medium Preparation

For convenience, make the medium as two parts: BROTH, which is autoclaved, and SUPPLEMENTS, which are filter-sterilized as a 20
concentrate.

2.1

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1. Broth: 1000 mL 2% soy (see Note 1). 5
950 mL

950 mL ®

20 g

Difco Bacto Soytone enzymatic digest of soybean 100 g

20 g

Difco Bacto® yeast extract

100 g

5.0 g

K2HPO4

25 g

0.63 g

L-glutamine

3.15 g

0.10 g

L-asparagine

0.5 g

Autoclave and cool before adding other components 50 mL

20
supplements concentrate

2. Supplements. 500 mL

20
concentrate

200 g

Dextrose

10 g

NaHCO3

1g

Thioglycolic acid

0.1 g

Choline

2 mL

10
salt solution 

Filter sterilize and store at 4 C

3. 10
salt solution. 50 mL

10
concentrate

2.5 g

Magnesium sulfate

0.18 g

Manganese sulfate

5 mL

HCl

Filter sterilize and store at 4  C

2.3 Prepare Bacteria Culture

1. 2% soy medium. 2. Glass bottles and tubes. 3. Blood agar plates. 4. 50% glycerol.

2.4 Inactivation of Pneumococcal Whole Cell

1. Lactate ringer (LR, 102 mM NaCl, 28 mM NaC3H5O3, 1.5 mM CaCl2, and 4 mM KCl) stored at 4  C. 2. LR with 10% sucrose. 3. Chloroform.

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4. Beta-propiolactone. 5. Autoclaved centrifuge bottles. 6. Autoclaved cylinders. 7. Stir bars. 2.5 Immunize Animals by Intranasal Route

1. Cholera toxin (CT). 2. SPWCA. 3. 20 μL pipet and tips. 4. 4–6 weeks old C57/BL6 mice.

2.6 Immunize Animals by Subcutaneous Route

1. SPWCA. 2. Aluminum hydroxide (Alhydrogel from Brenntag). 3. 1 mL syringes. 4. 27 G Needle.

2.7 Detection of T Cell Responses

1. DMEM/F-12 with L-glutamine, 10% FCS, 2-mercaptoethanol, and 10 μg/mL ciprofloxacin.

50

μM

50

μM

2. SPWCA. 3. 96-well plate. 2.8 In Vitro Stimulation of Splenocytes

1. DMEM/F-12 with L-glutamine, 10% FCS, 2-mercaptoethanol, and 10 μg/mL ciprofloxacin. 2. SPWCA. 3. 96-well plate.

3

Methods

3.1 Prepare Bacteria Culture

1. Day 1: Streak bacteria strain on blood agar plate. 2. Leave the plate in a 37  C incubator with 5% CO2 for 16–18 h (see Note 2). 3. Prepare two bottles of 1 L and 1 bottle of 200 mL of 2% soy medium as described above. 4. Keep a tube of 40 mL 2% soy medium in 37  C incubator. 5. Day 2: Transfer all the colonies on the blood agar plates to the 40 mL liquid medium. 6. Keep the liquid culture at 37  C incubator with 5% CO2 without shaking. Measure OD600 until it reaches 0.8. 7. Spin the culture down at 4000
g for 10 min at 4  C. 8. Remove 30 mL of supernatant from the tube and resuspend the pellet in the remaining 10 mL of medium.

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9. Add 10 mL of 50% glycerol in the resuspension and freeze the tube at 80  C as starter. 10. Keep the 2 L soy medium in the 37  C incubator overnight. 11. Day 3: Take the frozen starter out of 80  C freezer and thaw it in a water bath at room temperature. 12. Inoculate 10 mL of starter into each of the 1 L prewarmed soy medium, monitor OD600. 13. Take culture out of incubator when OD600 reaches 1. 3.2 Inactivation of Pneumococcal Whole Cell

SPWCV can be inactivated by addition of 70% ethanol, or 1/40 (v/v) of chloroform, or 1/40 (v/v) trichloroethylene, or 1/4000 (v/v) of beta-propiolactone (BPL). For this protocol, we will use chloroform and BPL as examples for inactivation. 1. Transfer pneumococcal culture to 1 L centrifuge bottle. 2. Harvest cells at 8000
g for 10 min at 4  C. 3. Remove supernatant and resuspend cell pellet in 160 mL of lactate ringer (LR). 4. Repeat steps 2 and 3 twice. 5. Remove supernatant and resuspend cell pellet in 20 mL of LR with 10% sucrose. 6. Measure total volume (Vt). 7. Dilute samples to 1:50 and 1:100 and measure OD600 of diluted samples. Calculate OD of undiluted sample by multiplying measured OD to its dilution factor (see Note 3). 8. Calculate final volume V ¼ Vt*OD600/32 and add additional volume (V  Vt) of LR/10% sucrose. 9. Transfer all the cell suspension to a glass bottle with stir bar. 10. Stir the suspension at 4  C. 11. Inactivation process: (a) Inactivation by chloroform (see Note 4). l

Adding a volume of V/40 chloroform into cell suspension.

l

Continue to stir for 2 h.

l

l

Aliquot to 400 μL each tube and freeze at 80  C for at least 1 h. Lyophilize.

(b) Inactivation by beta-propiolactone. l

Adding a volume of V/4000 beta-propiolactone into cell suspension.

l

Continue to stir for 16–18 h.

l

Move the cell suspension to 37  C and stir for 2 h.

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Aliquot to 400 μL each tube and freeze at 80  C.

l

Lyophilize (optional).

12. Dilute killed whole-cell 1:10, 1:100 in saline. 13. Plate 100 μL of whole-cell vaccine neat, 1:10 and 1:100 on blood agar plates. Incubate at 37  C overnight and check colony growth. A successful killed vaccine should have no colony growing. 14. Determine protein concentration by Lowry method [22]. 3.3 Immunize Animals by Intranasal Route

1. Prepare CT by dissolving the powder in sterile water to make 1 mg/mL solution. 2. Prepare SPWCA by adding 400 μL of sterile water into a lyophilized vaccine. Vortex for 20 s to ensure all powder is in solution. 3. Mix 100 μL of prepared SPWCA with 11 μL of CT for every 10 mice. 4. Mice are held gently without anesthesia, and 10 μL of vaccine is instilled into the nostrils of mice with a pipet. 5. Mice are immunized twice every week. 6. Collect blood or splenocyte samples 2–3 weeks after the last immunization to perform T cell analysis.

3.4 Immunize Animals by Subcutaneous Route

1. Prepare SPWCA as described in Subheading 3.2. 2. Mix SPWCA (final concentration 500 μg/mL) with aluminum hydroxide (final concentration 1.2 mg/mL) overnight on a rotating platform at 4  C. 3. Subcutaneous immunize group of 10 C57/BL6 mice every 2 weeks three times, in a volume of 200 μL for each time. 4. Collect blood or splenocyte samples 2–3 weeks after the last immunization to perform T cell analysis.

3.5 Detection of T Cell Responses

1. Dilute SPWCA to a final concentration of 11 μg/mL in medium. 2. Add 225 μL of stimulate containing medium into each well. 3. In each well, add 25 μL of heparinized blood to 225 μL of stimulate containing medium. 4. Incubate at 37  C CO2 with cover for 6 days. 5. Centrifuge at 2000
g for 2–3 min. 6. Determine cytokine concentration with IFN-γ and IL-17A ELISA kits.

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1. Harvest spleens from immunized mice and process it according to spleen processing protocol. 2. Resuspend splenocytes in stimulation medium at a concentration of 1
107 cells/mL. 3. Dilute SPWCA to a final concentration of 100 μg/mL in medium. 4. Add 225 μL of splenocytes into each well. 5. Add 25 μL of diluted SPWCA in medium. 6. Incubate at 37  C for 3 days. 7. Collect supernatant by centrifuging at 2000
g for 2–3 min. 8. Determine cytokine concentration with IFN-γ and IL-17A ELISA kits. Figure 1 shows an example of IL-17A response after immunization of SPWCA intranasally with CT or subcutaneously with alum.

Fig. 1 Measurement of IL-17A production from blood stimulation of SPWCA immunized mice. Mice were immunized with CT or CT with 100 μg of SPWCA intranasally twice every week and blood was taken 3 weeks after the second immunization. Another two groups of mice were immunized with alum or alum with 100 μg of SPWCA twice every 2 weeks. Blood was taken 2 weeks after the second immunization. Blood was stimulated for 6 days with 10 μg of SPWCA in 96-well plates, and IL-17A was measured with a mouse IL-17A duoset kit from R&D

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Notes 1. Soy concentration can be from 0.5 to 2%. 2. S. pneumoniae undergoes autolysis so plates cannot be in the incubator for too long. 3. When calculating OD from diluted samples, dilution with OD600 in the range of 0.2–1 should be used. An average of two calculated ODs will be used if both dilutions fall into this range. Sometimes a lower dilution is required if none of the dilutions is in this range. 4. Chloroform is heavier than water and will need continuous mixing during inactivation and aliquot steps.

References 1. Gea-Mallorqui E, Compeer EB (2020) SARSCoV-2 vaccine—think globally, act locally. Nat Rev Immunol 20:590 2. Dagotto G, Yu J, Barouch DH (2020) Approaches and challenges in SARS-CoV2 vaccine development. Cell Host Microbe 28 (3):364–370 3. Clem AS (2011) Fundamentals of vaccine immunology. J Glob Infect Dis 3(1):73–78 4. Khader SA, Gaffen SL, Kolls JK (2009) Th17 cells at the crossroads of innate and adaptive immunity against infectious diseases at the mucosa. Mucosal Immunol 2(5):403–411 5. Zhang Z, Clarke TB, Weiser JN (2009) Cellular effectors mediating Th17-dependent clearance of pneumococcal colonization in mice. J Clin Invest 119(7):1899–1909 6. Montgomery CP et al (2014) Protective immunity against recurrent Staphylococcus aureus skin infection requires antibody and interleukin-17A. Infect Immun 82 (5):2125–2134 7. Schmidt ME, Varga SM (2018) The CD8 T cell response to respiratory virus infections. Front Immunol 9:678 8. Kolls JK, Khader SA (2010) The role of Th17 cytokines in primary mucosal immunity. Cytokine Growth Factor Rev 21(6):443–448 9. Pulendran B, Ahmed R (2011) Immunological mechanisms of vaccination. Nat Immunol 12 (6):509–517 10. Lu YJ et al (2009) Protection against pneumococcal colonization and fatal pneumonia by a trivalent conjugate of a fusion protein with the cell wall polysaccharide. Infect Immun 77 (5):2076–2083

11. Lu YJ et al (2012) A bivalent vaccine to protect against Streptococcus pneumoniae and Salmonella typhi. Vaccine 30(23):3405–3412 12. Zhang F, Lu YJ, Malley R (2013) Multiple antigen-presenting system (MAPS) to induce comprehensive B- and T-cell immunity. Proc Natl Acad Sci U S A 110(33):13564–13569 13. LeibundGut-Landmann S et al (2007) Sykand CARD9-dependent coupling of innate immunity to the induction of T helper cells that produce interleukin 17. Nat Immunol 8 (6):630–638 14. van Sorge NM et al (2014) The classical lancefield antigen of group a Streptococcus is a virulence determinant with implications for vaccine design. Cell Host Microbe 15(6):729–740 15. Zhang F et al (2017) Antibody-mediated protection against Staphylococcus aureus dermonecrosis and sepsis by a whole cell vaccine. Vaccine 35(31):3834–3843 16. Ross PJ et al (2013) Relative contribution of Th1 and Th17 cells in adaptive immunity to Bordetella pertussis: towards the rational design of an improved acellular pertussis vaccine. PLoS Pathog 9(4):e1003264 17. Lu YJ et al (2008) Interleukin-17A mediates acquired immunity to pneumococcal colonization. PLoS Pathog 4(9):e1000159 18. Lu YJ et al (2010) GMP-grade pneumococcal whole-cell vaccine injected subcutaneously protects mice from nasopharyngeal colonization and fatal aspiration-sepsis. Vaccine 28 (47):7468–7475 19. Lu YJ et al (2010) Options for inactivation, adjuvant, and route of topical administration of a killed, unencapsulated pneumococcal

Vaccine Induced T Cell Responses whole-cell vaccine. Clin Vaccine Immunol 17 (6):1005–1012 20. Malley R, Anderson PW (2012) Serotypeindependent pneumococcal experimental vaccines that induce cellular as well as humoral immunity. Proc Natl Acad Sci U S A 109 (10):3623–3627 21. O’Hara JM et al (2020) Generation of protective pneumococcal-specific nasal resident

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memory CD4(+) T cells via parenteral immunization. Mucosal Immunol 13(1):172–182 22. McDonald CE, Chen LL (1965) The Lowry modification of the Folin reagent for determination of proteinase activity. Anal Biochem 10:175–177

Chapter 18 Development of a Bacterial Nanoparticle Vaccine Against Escherichia coli Melibea Berzosa, Yadira Pastor, Carlos Gamazo, and Juan Manuel Irache Abstract Currently, different subunit-based vaccine strategies against enterobacteria are being investigated. Among those, bacterial outer membrane vesicles (OMV) are promising candidates because of their immunogenic properties and safety. In order to develop an effective vaccine against this kind of pathogens, it is important to induce both systemic and mucosal immunity. For that reason, the oral route of administration would be an adequate option; although it still represents a challenge due to the particular and harsh conditions of the gut. To overcome these inconveniences, different strategies have been proposed, including the use of polymeric nanoparticles based on the copolymer between methyl vinyl ether and maleic anhydride (Gantrez AN). In the present work, a simple procedure for the preparation of heat-induced OMV (named as HT) obtained from Enterotoxigenic Escherichia coli (ETEC) loaded into these poly(anhydride) nanoparticles is described. Key words Outer membrane vesicles, Nanoparticles, Gantrez® AN, Escherichia coli, ETEC, Acellular vaccines

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Introduction Subcellular or acellular vaccines are currently popular for being the safest option against infectious diseases [1]. Among them, outer membrane vesicles (OMV) have demonstrated their efficacy as subunit-based vaccine candidates, since they contain multiple immunodominant antigens from the outer membrane (OM) of bacteria, including lipopolysaccharide (LPS) and OM proteins [2]. These vesicles have already shown successful immunogenicity and safety profiles in vivo against many different pathogens such as Shigella, Salmonella, Escherichia coli, Brucella, Pseudomonas, or Neisseria meningitidis serogroup B, against which the first licensed vaccine has recently been approved by regulatory agencies [3]. Since the use of OMV increases, recent studies have described different methods to augment the yield of the process, such as the

Sunil Thomas (ed.), Vaccine Design: Methods and Protocols, Volume 1: Vaccines for Human Diseases, Methods in Molecular Biology, vol. 2410, https://doi.org/10.1007/978-1-0716-1884-4_18, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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heat treatment (HT)-based strategy that makes the process safer without any significant alteration of the immunogenicity of these antigenic compounds [4, 5]. However, despite the promising results obtained in vaccination studies with OMV against different enterobacteria like Enterotoxigenic E. coli (ETEC) [6], the route of administration is critical for targeting mucosa and induce the right systemic and local immune response, and some nonliving vaccines may require the use of adjuvants or delivery systems to reach a protective immunity [7]. Among the different mucosal routes, the oral route of administration is preferred for being an easy, painless, and safe way of administration, avoiding the risk of transmitting blood-borne infections through needles and, importantly, inducing both mucosal and systemic immunity [8]. However, this route may be challenging because of the important physical, chemical, and enzymatic hurdles of gastrointestinal tract. Consequently, polymeric nanoparticulate delivery systems (NPs) have become a useful tool which can overcome these problems as well as improve the immune response of the antigens [9–11], being an alternative for the classical adjuvants [12]. For ETEC, different nano-vaccines have been studied for their administration through the oral route, and these NPs are mainly based on biodegradable polymers (i.e., chitosans, poly(lactic-co-glycolic acid) (PLGA), or poly(anhydrides) as poly(methyl vinyl ether-co-maleic anhydride) commercialized as Gantrez® AN), where the antigenic material obtained from ETEC is encapsulated [13–18]. Among those, some studies have recently described the adjuvant properties of Gantrez AN, since it is able to induce Th1 and CD8+ T cell immune responses mediated by a TLR2, TLR4, and TLR5 dependent manner [19, 20], which makes this polymer attractive for vaccination purposes. In the present work, we describe a simple and safe procedure for the extraction of heat-induced OMV (named as HT) from ETEC as antigen model loaded into poly(anhydride) nanoparticles based on Gantrez AN.

2

Materials All solutions are prepared using ultrapure water and analytical grade reagents. Reagents are stored at room temperature (unless indicated otherwise). Final products are lyophilized and stored at room temperature.

2.1 Bacterial Growth and Antigen Extraction

1. Enterotoxigenic E. coli (ETEC) growth: Trypticase-soy broth (Biome´rieux, SA, Marcy l’Etoile, France) on a rotatory shaker at 37  C. 2. Saline isotonic solution: 150 mM NaCl in water.

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3. Membrane-based tangential flow filtration (TTF): 300-kDa size-pore tangential filtration concentration unit (EMD Millipore, USA). 2.2 Protein and Lipopolysaccharide Content Determination

1. Protein content determination by Lowry method: (a) Dilution buffer: 0.4% CuSO4 5  H2O, 0.4% NaOH, 2% Na2CO3, 0.16% sodium tartrate and 1% sodium dodecyl sulfate (SDS). (b) Folin-Phenol reactive: 50% reactive in water. (c) Standard curve: Bovine serum albumin. 2. Lipopolysaccharide (LPS) content by determination of 2-keto3-deoxyoctonate (KDO) content: (a) Oxidation buffer: 0.042 N Periodic acid in 1.25 N H2SO4. (b) Stop reactive: 2% Sodium arsenite in 0.5 N HCl. (c) KDO detection buffer: 0.3% Thiobarbituric acid in water and dimethyl sulfoxide (DMSO). (d) Standard curves: Pure KDO (50 μg/mL) and D-deoxyribose (25 μg/mL).

2.3 SDS Polyacrylamide Gel Components

1. 15% acrylamide-Bis-acrylamide (37.5:1) in 125 mM Tris-HCl, pH 6.8 adjusted with HCl. 2. Electrode buffer: 30 mM Tris-HCl pH 8.3, adjusted with HCl, 192 mM glycine, and 0.1% SDS. 3. SDS-PAGE running buffer: 30 mM Tris-HCl, pH 8.3, 192 mM glycine, 0.1% SDS. 4. SDS lysis buffer: 62.5 mM Tris-HCl pH 6.8, 10% glycerol, 2% SDS, 5% β-mercaptoethanol, and 0.002% bromophenol blue. Store the aliquots at 20  C. 5. Periodate-silver staining buffer: (a) Fixation buffers: 50% methanol and 10% acetic acid in water, 7.5% methanol, and 5% acetic acid. (b) Oxidation buffer: For protein staining protocol, 10% glutaraldehyde in water. For the LPS staining protocol the samples are pre-treated with 0.7% paraperiodic acid, 7.5% methanol, and 5% acetic acid in water. (c) Staining buffer: 4% AgNO3, 0.75% NaOH, and 1.4% NH3 in water. 6. Coomassie blue staining: (a) Incubation buffer: 3% Trichloroacetic acid in water. (b) Staining buffer: 0.25% Coomassie blue in 50% methanol and 10% acetic acid in water.

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7. SDS-PAGE staining molecular mass standard: Rainbow RPN756 (GE Healthcare Bio-Science, USA) containing myosin (220 kDa), phosphorylase B (97 kDa), bovine serum albumin (66 kDa), ovalbumin (45 kDa), carbonic anhydrase (30 kDa), trypsin inhibitor (20.1 kDa), and lysozyme (14.3 kDa). 2.4 Immunoblotting Components

1. Transfer buffer: 0.2 M glycine, 24 mM Tris-HCl 10% methanol (pH 8.3) in water. Store at 4  C. 2. Nitrocellulose membrane. 3. Semidry electroblotter. 4. Blocking buffer: 5% skimmed milk in 10 mM phosphatebuffered saline (pH 7.4). 5. Antibody dilution buffer: 1% skimmed milk with 0.15% Tween-20 in 10 mM phosphate-buffered saline (pH 7.4). 6. Incubation solution: H2O2, 4-chloro, 1-naphtol.

2.5 Nanoparticle Formulation Preparation

1. Polymer: Poly(methyl vinyl ether-co-maleic anhydride) (Gantrez AN-119), (M.W. 200,000, Ashland Inc. Covington, KY, USA). 2. Acetone. 3. Sonicator Microson TM (Misonix Inc., USA). 4. Cryoprotectant: 5% sucrose in water. € 5. Spry-drier (BUCHI Labortechnik AG, Switzerland). 6. Lyophilizer.

2.6 Nanoparticle Characterization

1. Determination of particle size and polydispersity index (PDI): (a) Deionized water. (b) Zetasizer analyzer Malvern, UK).

system

(Malvern®

Instruments,

2. Determination of zeta-potential: (a) 0.1 mM KCl solution in water adjusted to pH 7.4 with HCl. (b) Zetasizer analyzer Malvern, UK).

system

(Malvern®

Instruments,

3. Determination of the morphology of the particles by cryoelectron microscopy (cryoEM): (a) Glow discharged holey carbon grids. (b) Liquid ethane. (c) FEI Vitrobot™.

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2.7 Nanoparticle Loading Capacity

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1. Deionized water. 2. Sonicator. 3. Rotary evaporator.

2.8 Determination of Antigen Integrity

3

See Subheading 2.4.

Methods

3.1 Bacterial Strain and Growth Conditions

Outer membrane vesicles (HT antigenic complex released by heat treatment, see below) are obtained from Enterotoxigenic E. coli grown in trypticase-soy broth on a rotary shaker at 37  C, 140 rpm for 24 h (other conditions may be applied).

3.2 Antigenic Complex (HT Membrane Vesicles)

The extraction process of the antigenic complex HT is shown in Fig. 1. 1. Grow bacteria in trypticase-soy broth on a rotary shaker at 37  C, 140 rpm for 24 h. 2. Heat the cultures at 100  C in flowing steam for 15 min. 3. Harvest cell by centrifugation (6000  g, 20 min). 4. The supernatant of cultured bacteria containing the HT extract is filtrated by 0.22 μm pore-filter. 5. The filtrated supernatant is purified, dialyzed, and concentrated by diafiltration (300-kDa tangential filtration concentration unit). The final product is recovered in the retentate fraction. 6. The retentate fraction is collected by centrifugation at 40,000  g, for 75 min (see Note 1). 7. The resulting pellet is finally resuspended in deionized water, lyophilized and stored at room temperature.

3.3 Characterization of the Antigenic Extracts

1. Total protein content determination: Total protein content may be quantified by the method of Lowry [21], with bovine serum albumin as standard. 2. Lipopolysaccharide (LPS) content determination: LPS content may be quantified by the determination of 2-keto-3-

Fig. 1 Schematic extraction process of the antigenic complex HT from Enterotoxigenic Escherichia coli

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deoxyoctonate (KDO) content, performed by the method of Warren [22] as modified by Osborn [23]. 3. Dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE): Protein and LPS profiles may be determined by SDS-PAGE by the method of Laemmli [24] followed by staining with Coomassie blue [25] or with the alkaline silverglutaraldehyde method for proteins [26], or for LPS [27]. 4. Immunoblotting: The antigenicity of HT components may be analyzed by immunoblotting, carried out as described by Towbin [28] using an appropriate specific antiserum (see Note 2), with the following modifications. 5. After SDS-PAGE, transfer the gel in a transfer buffer (0.2 M glycine; 24 mM Tris; 10% methanol, pH 8.3) to nitrocellulose membrane by using a semidry electroblotter (200 mA; 5 V; 30 min). 6. Place the membrane in blocking buffer (5% skimmed milk in 10 mM phosphate-buffered saline, pH 7.4) overnight at room temperature. 7. Incubate for 4 h at room temperature with specific antibodies (e.g., hyperimmunized or naturally infected animals) diluted 1:100 (or as convenient) in primary buffer: 1% skimmed milk with 0.15% Tween-20 in 10 mM phosphate-buffered saline, pH 7.4. 8. After 4 h, wash the membrane 5 times in 0.15% Tween-20 in 10 mM phosphate-buffered saline, pH 7.4. 9. Incubate the membrane for 1 h at room temperature with the appropriate immunoconjugate: Peroxidase-conjugated diluted 1:1000 in the antibody dilution buffer. 10. Repeat step 8. 11. Membrane is developed by incubation in a solution containing H2O2 and 4-chloro, 1-naphtol for 20 min in the dark. 3.4 Preparation of HT-Loaded Nanoparticles

The preparation of nanoparticles was carried out by a solvent displacement method [29] (Fig. 2). 1. Dissolve 100 mg of the copolymer of methyl vinyl ether and maleic anhydride (Gantrez AN) in 5 mL acetone under magnetic stirring at room temperature. 2. In parallel, 5 mg HT are dispersed by ultrasonication with a probe Microson ™ in 10 mL water for 1 min. 3. In order to obtain the nanoparticles, add the water phase containing HT to the solution of the polymer in acetone. 4. Maintain the just formed nanoparticles under magnetic agitation during 15 min (see Note 3).

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Fig. 2 Schema of the preparative process of HT-loaded nanoparticles

5. Eliminate the organic solvent under reduced pressure (Bu¨chi R-144, Switzerland). 6. The nanoparticles are collected by centrifugation 27,000  g, 20 min, 4  C. Wash with water twice.

at

7. Finally, nanoparticles are freeze-dried using sucrose (5%) as cryoprotectant. 3.5 Nanoparticle Characterization

1. Determination of particle size and polydispersity index (PDI): Disperse the nanoparticles in deionized water and measure the particle size and PDI by photon correlation spectroscopy (PCS) at 25  C with a scattering angle of 90 using a Zetasizer analyzer system. 2. Determination of zeta-potential: Dilute 200 μL of the HT-loaded nanoparticles in 2 mL of a 0.1 mM KCl solution adjusted to pH 7.4. Measure the zeta-potential by electrophoretic laser Doppler anemometry using a Zetasizer analyzer system. 3. Determination of the morphology of the particles by cryoelectron microscopy (cryoEM): Apply 3 μL of reconstituted nanoparticles onto glow discharged holey carbon grids (Quantifoild®) and subsequently plunge-frozen in liquid ethane using the FEI Vitrobot™.

3.6 Nanoparticle Loading Capacity

1. Degradation of loaded nanoparticles. (a) Disperse 15 mg of HT-loaded NPs in deionized water by vortexing 1 min. (b) Centrifuge the dispersed NPs at 27,000  g, 15 min. (c) Resuspend the pellet in NaOH 0.1 M. (d) Sonicate the suspension and incubate it for 1 h. 2. Determination of the amount of HT released from the nanoparticles.

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The amount of HT released from the nanoparticles could be determined using microbicin choninic aicd (microBCA) protein assay described by Pierce, Rockford, CA, USA. 3.7 Determination of Antigen Integrity

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The antigenicity integrity of the HT could be determined by immunoblotting under the conditions described above in Subheading 3.3.

Notes 1. Previous to the centrifugation, the supernatant can be frozen and thawed in order to induce vesicles fusion and thus facilitate subsequent harvesting by centrifugation. 2. This step is performed as a quality control of the antigenicity conservation after bacterial antigenic extraction. 3. The nanoparticles suspension should have an homogeneous aspect like a milky white bluish opalescent fluid.

Funding Information This work was financially supported by “Instituto de Salud Carlos III,” the European Regional Development Fund (ERDF) (PI19/ 00146). References 1. Cozzi R, Scarselli M, Ferlenghi I (2013) Structural vaccinology: a three-dimensional view for vaccine development. Curr Top Med Chem 13 (20):2629–2637 2. Schwechheimer C, Kuehn MJ (2015) Outermembrane vesicles from gram-negative bacteria: biogenesis and functions. Nat Rev Microbiol 13:605–619 3. Van der Pol L, Stork M, van der Ley P (2015) Outer membrane vesicles as platform vaccine technology. Biotechnol J 10(11):1689–1706 4. Pastor Y, Camacho A, Gil AG, Ramos R, Lo´pez ˜ uelas I et al (2017) Effective De Cera´in A, Pen protection of mice against Shigella flexneri with a new self-adjuvant multicomponent vaccine. J Med Microbiol 66(7):946–958 5. Ochoa J, Irache JM, Tamayo I, Walz A, DelVecchio VG, Gamazo C (2007) Protective immunity of biodegradable nanoparticlebased vaccine against an experimental challenge with Salmonella enteritidis in mice. Vaccine 25(22):4410–4419

6. Roy K, Hamilton DJ, Munson GP, Fleckenstein JM (2011) Outer membrane vesicles induce immune responses to virulence proteins and protect against colonization by enterotoxigenic Escherichia coli. Clin Vaccine Immunol 18(11):1803–1808 7. Aoshi T (2017) Modes of action for mucosal vaccine adjuvants. Viral Immunol 30 (6):463–470 8. Lycke N (2012) Recent progress in mucosal vaccine development: potential and limitations. Nat Rev Immunol 12(8):592–605 9. Irache JM, Esparza I, Gamazo C et al (2011) Nanomedicine: novel approaches in human and veterinary therapeutics. Vet Parasitol 180 (1–2):47–71 10. Jin Z, Gao S, Cui X et al (2019) Adjuvants and delivery systems based on polymeric nanoparticles for mucosal vaccines. Int J Pharm 15:572–118731 11. Vela Ramirez JE, Sharpe LA, Peppas NA (2017) Current state and challenges in

Nanoparticle Vaccine Against Escherichia coli developing oral vaccines. Adv Drug Deliv Rev 114:116–131 12. Pati R, Shevtsov M, Sonawane A (2018) Nanoparticle vaccines against infectious diseases. Front Immunol 9:2224 13. Nazarian S, Gargari SLM, Rasooli I et al (2014) A PLGA-encapsulated chimeric protein protects against adherence and toxicity of enterotoxigenic Escherichia coli. Microbiol Res 169(2–3):205–212 14. Doavi T, Mousavi SL, Kamali M et al (2016) Chitosan-based intranasal vaccine against Escherichia coli O157:H7. Iran Biomed J 20 (2):97 15. Wu KY, Wu M, Fu ML et al (2006) A novel chitosan CpG nanoparticle regulates cellular and humoral immunity of mice. Biomed Environ Sci 19(2):87–95 16. Khan MS, Vishakante GD (2013) Development and evaluation of porous chitosan nanoparticles for treatment of enterotoxigenic Escherichia coli infection. J Biomed Nanotechnol 9(1):107–114 17. Noroozi N, Mousavi Gargari SL, Nazarian S et al (2018) Immunogenicity of enterotoxigenic Escherichia coli outer membrane vesicles encapsulated in chitosan nanoparticles. Iran J Basic Med Sci 21(3):284–291 18. Vandamme K, Vesna M, Eric C et al (2011) Adjuvant effect of Gantrez®AN nanoparticles during oral vaccination of piglets against F4+enterotoxigenic Escherichia coli. Vet Immunol Immunopathol 139(2–4):148–155 19. Camacho AI, Da Costa MR, Tamayo I, de Souza J, Lasarte JJ, Mansilla C et al (2011) Poly(methyl vinyl ether-co-maleic anhydride) nanoparticles as innate immune system activators. Vaccine 29:7130–7135 20. Tamayo I, Irache JM, Mansilla C, OchoaRepa´raz J, Lasarte JJ, Gamazo C (2010) Poly

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(anhydride) nanoparticles act as active Th1 adjuvants through toll-like receptor exploitation. Clin Vaccine Immunol 17(9):1356–1362 21. Lowry OH, Rosebrough NJ, Farr AL et al (1951) Protein measurement with the Folin phenol reagent. J Biol Chem 193(1):265–275 22. Warren L (1963) Thiobarbituric acid assay of sialic acids. Methods Enzymol 6:535–538 23. Osborn MJ (1963) Studies on the gramnegative cell wall, i. evidence for the role of 2-keto-3-deoxyoctonate in the lipopolysaccharide of Salmonella typhimurium. Proc Natl Acad Sci 50(3):499–506 24. Laemmli UK (1970) Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227 (5259):680–685 25. Fairbanks G, Steck TL, Wallach DFH (1971) Electrophoretic analysis of the major polypeptides of the human erythrocyte membrane. Biochemistry 10(13):2606–2617 26. Merril CR, Switzer RC, Van Keuren ML (1979) Trace polypeptides in cellular extracts and human body fluids detected by two-dimensional electrophoresis and a highly sensitive silver stain. Proc Natl Acad Sci U S A 76(9):4335–4339 27. Tsai CM, Frasch CE (1982) A sensitive silver stain for detecting lipopolysaccharides in polyacrylamide gels. Anal Biochem 119 (1):115–119 28. Towbin H, Staehelin T, Gordon J (1979) Electrophoretic transfer of proteins from polyacrylamide gels to nitrocellulose sheets: procedure and some applications. Proc Natl Acad Sci U S A 76(9):4350–4354 29. Arbo´s P, Wirth M, Arangoa MA et al. (2002) Gantrez AN as a new polymer for the preparation of ligand-nanoparticle conjugates. J Control Release 83:321–330

Chapter 19 Construction of Novel Live Genetically Modified BCG Vaccine Candidates Using Recombineering Tools Mario Alberto Flores-Valdez and Michel de Jesu´s Aceves-Sa´nchez Abstract One of the strategies for the construction of live vaccine candidates is through the generation of genetically defined isogenic strains, containing single or multiple mutations in target-specific genes generated by allelic exchange. This approach allows to produce rational attenuation of or, alternatively, sequence-specific modifications to produce variants of antigenic molecules or change their expression levels. Genetic tools amenable for their use in mycobacterial strains have allowed the identification and validation of potential targets for the diagnosis, prevention, and treatment of tuberculosis. However, the genetic manipulation of Mycobacterium tuberculosis and other slow-growing strains such as Mycobacterium bovis BCG has been delayed by various factors related to their physiology and cell wall characteristics. Notwithstanding the foregoing, the high frequency of illegitimate recombination and the availability of few antibiotic selection markers limit the feasibility of genetic manipulation of mycobacterial strains. This chapter describes a protocol for the generation of defined mutants using recombination tools in an inducible recombination system driven by mycobacterial Che9c phage RecET proteins, originally developed in Dr. Graham Hatfull’s group, combined with linearized recombination substrates containing flanking sequences of a locus of interest and an antibiotic resistance gene. These recombination substrates contain sites for removal of antibiotics selection markers. This system allows to make marked and unmarked mutations by homologous recombination in a single step as a result of a double crossover between the homologous regions on the genome and the allelic exchange substrate. In addition, this genetic tool used for engineering mycobacterial genomes performs with lower rates of illegitimate recombination and take on average less time to create knock-out (KO) mutant compared with other techniques. Key words Recombineering, Gene replacement, Homologous recombination, Mutation, Vaccine

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Introduction The construction of genetically defined isogenic strains containing single or multiple mutations in target-specific genes generated by allelic exchange has been a common strategy for gene inactivation in bacteria. Targeted inactivation offers complete control over the nature of the introduced mutation, although it is a relatively demanding work. Knock-out strains offer the possibility of testing

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specific pathways, identifying functions that might be important for growth and pathogenicity, and it has been very useful in demonstrating the essentiality of a gene, validating drug targets, for rational attenuation in engineering improved live-attenuated vaccine strains, and is a potential strategy for mutation of specific amino acid residues by site-directed mutagenesis. Furthermore, the construction of site-directed mutants has the advantage of improving the intrinsic machinery of the cell for homologous recombination [1]. However, genetic manipulation of slow-growing mycobacteria such as M. tuberculosis and Mycobacterium bovis BCG has been hampered by various factors related to the physiology and pathogenicity of this microorganism. First of all, the extremely slow growth rate of this bacterium reduces the speed to carry out any experiment [2]. Second, this bacterium requires specific biosafety level 2 (BCG) or level 3 (M. tuberculosis) conditions, substantial training before handling, and presents with an implicit risk of accidental exposure and infection [3]. Additionally, other factors also reduce the rate of success in manipulating these mycobacteria, such as the characteristics of its cell wall and the tendency to form aggregates that limit the efficient entry of DNA, even with the use of electroporation [4]; in addition to the lack of systems to carry out gene transfer in slow-growing mycobacteria, make genetic manipulation of this microorganism difficult [2]. In addition to these challenges, it is necessary to consider the limited repertoire of available antibiotics that can be applied for the selection of recombinant strains in this bacterial genus due to the natural resistance of mycobacteria to many antibiotics, including the production of β-lactamases. Further to this, a limited permeability of hydrophilic compounds because of the inherent characteristics of its cell wall also affects this capacity of selective pressure. Furthermore, the need to use antibiotics with prolonged stability, required for slow-growing species, as well as the requirement to use antibiotics with a low frequency of spontaneous resistance reduces the alternatives for a relatively easy selection of mutants where an antibiotic marker has been introduced [5, 6]. However, despite the difficulties described above, the main obstacle to genetic manipulation in slow-growing mycobacteria is the relatively high level of illegitimate recombination compared to the homologous recombination observed in fast growing mycobacteria such as M. smegmatis due to its inefficient endogenous recombination system [7–12]. In order to overcome these difficulties, specific systems and protocols have been developed for mycobacteria, which allow efficient recovery of mutants by allelic exchange. The identification of critical points in the mutagenesis process has led to the development of new protocols and modifications to existing ones. Together, these new approaches improve the efficiency of

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competent cells to allow DNA entry for allelic exchange and promote legitimate recombination events with the bacterial chromosome. The first requirement for all genetic manipulation is an easy and efficient method to introduce foreign DNA into mycobacterial cells. The most successful method in mycobacteria has been electroporation considering various parameters in the conditions to carry out this process [5]. Other methods used to efficiently introduce foreign DNA into mycobacteria rely on the use of mycobacteriophages [13] and a novel method to transform mycobacteria, namely shockwave-assisted bacterial transformation, which uses a number of consecutive shockwaves to introduce DNA into mycobacteria [14]. It has been observed that to obtain an efficient transformation in slow-growing mycobacteria using electroporation, it is essential to use temperatures between 25
C and 37
C during the incubation of the cells before electroporation. In contrast to the low temperatures of 4
C that are preferred for M. smegmatis [15]. Furthermore, compared to cells that are stored at 80
C before use, the suspension of fresh cells tends to produce higher transformation efficiencies [16]. Another common practice to moderately improve transformation efficiency has been the addition of sublethal amounts of chemical agents to the culture medium that affect the integrity of the cell wall such as glycine or ethionamide [17]. Conversely, in other protocols, detergent agents such as 0.025% tyloxapol are incorporated in the electroporation solution in order to avoid cell aggregates and/or the suspension is prepared in 10% glycerol to promote the solubility of the cells in the aqueous medium [5]. Another important strategy employed along with mycobacterial genetic tools has been to modify the allelic exchange substrate to make it more susceptible to recombining with the genome of the receiving cell. Here, strategies to stimulate homologous recombination include the use of long and linear DNA substrates, DNA substrates short and linear, UV-irradiated or alkaline denaturationtreated plasmids, and single-stranded DNA. Other strategies aimed to improve selection include suicide plasmids with counter-selective markers, incompatible plasmids, heat-sensitive counter-selective replicable plasmids, and heat-sensitive mycobacteriophages [18]. On the other hand, tools focused on improving the recombination machinery have also been described, thus successfully increasing the levels of homologous recombination and the recovery of mutants by allelic replacement with the use of proteins that stimulate homologous recombination, such as recombineering, CRISPR/Cas9, and ORBIT (oligonucleotide-mediated recombineering followed by Bxb1 integrase targeting) [19–22]. “Recombineering” or Recombination-mediated Genetic Engineering is based on the utilization of the bacteriophage λ “Red” functions to facilitate recombination events, an independent system

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to that of endogenous recombination based on RecA activity [23– 25]. The λ red system encodes the Exo (Redα), Beta (Redβ), and Gam (Redγ) proteins [26]. In addition, the functional analogues of Exo and Beta have been described in the RecE and RecT proteins, respectively, encoded by the E. coli Rac prophage [27]. Exo λ and RecE are exonucleases that degrade dsDNA-dependent linear DNA in the 50 -30 direction, with little or no activity in dsDNA holes or nicks [28]. Beta, λ or RecT are cDNA binding proteins that promote complementary DNA strand alignment and chain exchange but are unable to invade double-stranded DNA as RecA does [26], although it has been proposed that the λ recombination system can also invade dsDNA in E. coli [28, 29]. Gam λ functions inactivate the RecBCD system by binding to the RecB subunit and prevent DNA degradation [28]. These systems are useful because they promote recombination at levels higher than the host’s own mechanisms [26, 30], it can use short regions of DNA homology (~50 bp) [31] and can recombine with cDNA substrates without the need of using Exo (RecE) or Gam [28, 32]. The mycobacterial recombineering system represents an attractive strategy to overcome limitations in mycobacterial recombination. This strategy was proposed by van Kessel and Hatfull [28], who reported a recombination system in Mycobacterium tuberculosis and Mycobacterium smegmatis based on the activity of two recombination proteins encoded by the gp60 and gp61 genes of the mycobacteriophage Che9c, and which are RecE and RecT homologs, respectively. This system uses the recombination plasmid pJV53 that allows the inducible expression of these genes regulated by an acetamidase inducible promoter. The high expression of these proteins, as in other bacterial systems, appears to be toxic to Mycobacterium, with lethal effects observed or affecting its growth; therefore, a regulated expression system must be used [19, 26, 28]. Furthermore, these proteins are more effective in promoting homologous recombination than endogenous mycobacterial systems [6]. To apply this tool, once induction of the proteins for recombination has been carried out, the strain carrying the recombineering plasmid must be electroporated with a linear DNA fragment containing the mutated allele and subsequently the transformants must be plated on a selective medium. Allelic exchange substrates in mycobacteria typically contain regions of homology upstream and downstream of a target gene that flank a hygromycin resistance cassette. The ability to generate mutants with 100 to 300 ng of substrate containing ~50 bp to 500 bp homologous regions on both sides of the antibiotic resistance marker shows a frequency of ~1  104. Furthermore, the tool was reported to achieve over 90% efficiency for recombinants obtained being product of homologous recombination [28].

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In addition to improving recombination efficiency, it is also possible to remove antibiotic resistance markers, which is an advantage when subsequent genetic manipulations are required in the mutants obtained or as a requirement of vaccine candidates intended for evaluation in advanced clinical studies [1]. The elimination of these markers can be achieved by using site-specific recombination-mediated marker gene removal systems, included in pYUB854 and pUC-Hyg cloning plasmids. pYUB854 is a plasmid that employs a γδ/res system and that places resolvase sites flanking the hygromycin resistance cassette and uses the removal machinery mediated by a bacteriophage to remove the antibiotic marker gene [33]. The pUC-Hyg plasmid has two dif RecAindependent recombination, sequence-specific sites, at the ends of the hygromycin resistance cassette and its antibiotic marker gene removal is based on a Xer/dif system that requires successive passages without antibiotics to eliminate the antibiotic resistance cassette from the chromosome [34].

2

Materials

2.1 Chromosomal DNA Preparation

1. Mycobacterium strain cell pellet (preferably obtained from mid-log phase liquid cultures). 2. Lysis buffer (SET solution): 25% sucrose, 50 mM EDTA, 50 mM Tris–HCl pH 8. 3. Lysozyme solution (20 mg/mL in water). 4. RNAse A solution (10 mg/mL in water). 5. Proteinase K solution: 400 μg/mL proteinase K, 100 mM Tris–HCl pH 8, 0.5% w/v SDS. 6. Phenol–chloroform–isoamyl alcohol (25:24:1). 7. Chloroform–isoamyl alcohol (24:1). 8. 3 M sodium acetate solution, pH 5.2. 9. Isopropanol. 10. 70% (v/v) ethanol. 11. Molecular biology grade water. 12. Agarose. 13. 50 TAE buffer. 1 L: 242 g Tris base, 57 mL glacial acetic acid, 100 mL 0.5 M EDTA, pH 8.0. 14. Electrophoresis chamber. 15. Electrophoresis power supply. 16. Nanodrop spectrophotometer.

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Fig. 1 Maps of cloning vectors 2.2 Plasmid Construction for Recombination Substrate

1. pYUB854: Plasmid vector for cloning regions flanking the target gene and to create antibiotic-marked mutants. The flanking cassette includes multiple restriction enzyme sites for the cloning of homologous regions upstream and downstream of the hyg cassette and γδres sites; it confers resistance to hygromycin to the resulting mutants after homologous recombination [33]. A graphic map can be seen in Fig. 1. 2. pUC-Hyg (GenBank KU306403): Plasmid vector for cloning regions flanking the target gene and create antibiotic-less mutants. The flanking cassette includes multiple restriction enzyme sites for the cloning of homologous regions upstream and downstream of the dif-hyg-dif cassette; it confers hygromycin resistance for initial selection, which can be eliminated after further growth and selection [34]. A graphic map can be seen in Fig. 1. 3. Luria Broth (LB) and LB plates with hygromycin 150 μg/mL. 4. High-fidelity polymerase. 5. Restriction nzymes. 6. T4 DNA ligase. 7. Kits for purification of PCR products and plasmids. 8. Electrophoresis chamber. 9. Electrophoresis power supply. 10. Thermocycler.

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11. Nanodrop spectrophotometer. 12. Electroporation cuvettes (1 mm gap). 13. Competent Escherichia coli DH5α cells (other strains can also be used depending on the nature of the DNA to be cloned). 14. Hygromycin supplied as a 50 mg/mL stock solution. For selection use at 50–100 (μg/mL). 15. PCR primers. 16. 50 TAE buffer. 17. Electrophoresis chamber. 18. Electrophoresis power supply. 19. Taq DNA polymerase. 20. TAE buffer. 21. Gel documentation system. 2.3 Preparation of Recombineering Substrates

1. LB broth. 2. Hygromycin solution 50 mg/mL. 3. Miniprep plasmid purification kit. 4. Restriction enzymes. 5. Thermocycler. 6. Agarose gel 1.5%. 7. DNA gel purification kit/PCR purification kit. 8. Molecular biology grade water. 9. Nanodrop or similar spectrophotometer.

2.4 Preparation of Recombinogenic/ Electrocompetent BCG

1. pJV53: Expresses the Che9c bacteriophage genes gp60 and gp61 under the control of the Pacet promoter [16]. 2. 20% (w/v) succinate solution: Dissolve 2 g of disodium succinate hexahydrate in 10 mL of distilled water and filter sterilize through 0.2 μM membrane filter. Store at 4
C for 2 months. 3. 20% (w/v) acetamide solution: Dissolve 2 g of acetamide in 10 mL of distilled water and filter sterilize through 0.2 μM membrane filter. Store at 4
C for 2 months. 4. 20% (v/v) Tween 80: Mix 20 mL Tween 80 with 80 mL of distilled water, stir the solution at 50
C to dissolve the Tween 80, cool to room temperature (RT) and filter sterilize through 0.2 μM membrane filter. Store at 4
C for 2 months. 5. 7H9-OADC-Tween 80 media: Dissolve 4.7 g of Middlebrook 7H9 powder and 2 mL glycerol in 900 mL deionized water and autoclave at 121
C, 15 psi. Add 100 mL of Middlebrook OADC enrichment supplement and 2.5 mL of sterilized 20% Tween 80. Store at 4
C for 2 months.

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6. Induction media: Dissolve 4.7 g of Middlebrook 7H9 powder and 2 mL glycerol in 1 L deionized water and autoclave. Add 2.5 mL of sterilized 20% Tween 80 and 1 mL 20% succinate solution. Store at 4
C for 2 months. 7. 7H10 agar plates: Add 19 g of Middlebrook 7H10 powder and 5 mL glycerol to 900 mL of deionized water. Autoclave, cool to 45
C, and add 100 mL of OADC. Mix and pour (20 mL per plate). 8. Cell culture flasks (175 cm2). 9. 2 M glycine: Dissolve 150.14 g in 1 L distilled water; filter sterilize through 0.2 μM membrane filter. Store at room temperature for 6 months. 10. 10% (v/v) glycerol: Mix 100 mL glycerol with 900 mL distilled water and autoclave. 2.5 Electroporation of Recombineering Substrates

1. Electroporation cuvettes (4 mm gap). 2. 7H9-OADC-Tween 80 media and 7H10 plates with hygromycin 100 μg/mL. 3. Small sterile disposable transfer pipettes (1 mL) individually packaged consisting of a small bulb and extended fine tip. 4. Electroporation apparatus: Capable of delivering a pulse of 2.5 kV, 1000 Ω, 25 μF setting. 5. Fifteen milliliters tubes (15 mL). 6. Aluminum foil.

2.6 Growth, Verification of Allelic Replacement Mutants, and Recombineering Plasmid Curation

1. Cell culture flasks (25 cm2). 2. 7H9-OADC-Tween 80 and 7H10 plates. 3. Kanamycin: Dissolve in deionized water to a concentration of 20 mg/mL and filter sterilize. Store at 20
C for 6 months. 4. Hygromycin B supplied as a 50 mg/mL stock solution in PBS; store at 4
C in the dark. 5. Taq DNA polymerase. 6. Cassette and target-specific primers. 7. Inoculation needles. 8. PBS-Tween 80: Add 2.5 mL of 20% Tween 80 (w/v) to 1 L PBS. Filter sterilize.

3

Methods

3.1 Chromosomal DNA Preparation

1. Grow 20 mL of a BCG culture in 7H9 OADC broth at 37
C with 100 rpm shaking until it reaches an OD600 of 0.8–1.0. 2. Pellet the bacteria by centrifugation at 5000 rpm (4500  g) for 10 min at room temperature. Discard the supernatant.

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3. Resuspend the bacterial pellet in 250 μL SET solution, boil at 100
C for 15 min, and add 50 μL of lysozyme. 4. Incubate the mixture overnight at 37
C. 5. Add 10 μL of RNase A and incubate at 37
C for 30 min. 6. Add 250 μL of proteinase K solution and incubate at 55
C for 2 h. 7. Add 500 μL of phenol–chloroform–isoamyl alcohol to the sample, mix vigorously for 2 min, and let stand at room temperature for 5 min. 8. Centrifuge at 12,000 rpm (15000  g) for 10 min and recover the top aqueous layer in a new tube. 9. Add 0.1 volume of 3 M sodium acetate (pH 5.2), followed by 0.7 volume of isopropanol. 10. Invert the sample until well mixed and incubate at 20
C for at least 1 h. 11. Centrifuge the sample at 12,000 rpm (15000  g), 4
C for 30 min. 12. Remove supernatant, add 1 mL of 70% ethanol, and centrifuge the sample at 12,000 rpm (15000  g), 4
C for 5 min. 13. Repeat step 12. 14. Remove carefully supernatant with a micropipette and let dry at room temperature or with a vacuum concentrator. 15. Resuspend DNA in molecular biology grade water and store it at 4
C. 16. Quantify and check purity in Nanodrop and check integrity in an agarose gel electrophoresis (see Note 1). 3.2 Plasmid Construction to Produce the Recombination Substrate

Amplify the two PCR products corresponding to the upstream and downstream regions of the target gene. Each amplicon should contain at least ~500 bp sequence that flanks the target gene (see Note 2). The oligos used for PCR must contain unique restriction sites, introduced artificially at the 50 -end of each primer if they are absent from the flanking sequences intended to be cloned. Alternatively, you can take advantage of native sites that are also present in the vector flaking the hygR gene, making sure these sites are absent from the other flaking region, in order to be able to conduct the sequential cloning of the flanking regions. These restriction sites must be present in the vector intended to produce the recombinant plasmid as substrate for homologous recombination. Furthermore, the location of these restriction sites must be the one that, after cloning of the PCR products, conserves the orientation that these sequences have with respect to the target sequence as present in the genome of reference. Purify PCR products. Perform their cloning into pYUB854 or pUC-Hyg (Fig. 1) in two steps. Each step

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consists of digestion and ligation reactions, followed by transformation into E. coli. As first step, digest, with suitable restriction enzymes, the empty cloning vector and either PCR product. Purify both DNA sequences, quantify them, and perform the ligation reaction. Later, transform competent E. coli with different amounts of the ligation reaction and select clones on LB plates with hygromycin (150 μg/ mL). Verify the successful insertion of the PCR fragment (PCR1) either by PCR or restriction digestion. Grow the E. coli colonies that harbor the pYUB854/pUC-Hyg plasmids with the PCR1 inserted into them, to purify said plasmids. For inserting the second fragment, digest the previously obtained recombinant plasmid (we will call it “vector + PCR1”) and the second PCR product (PCR2) with the corresponding restriction enzymes. No sequence that can be cleaved by these enzymes must be present in PCR1. Purify both DNAs (vector1 + PCR1, and PCR2), quantify them, and perform the ligation reaction. Next, transform competent E. coli with different amounts of the ligation reaction and select clones on LB plates with hygromycin (150 μg/mL). Verify the successful insertion of PCR2 either by PCR or restriction digestion. To make sure you have a substrate for homologous recombination, confirm that the recombinant plasmids contain both PCR fragments, either by PCR or restriction digestion, followed by DNA sequencing to verify identity and fidelity of the sequences being used for recombination. Whenever possible, proceed straight to DNA sequencing. After obtaining the DNA sequence of the recombination substrate, perform a BLAST comparison between the recombination substrate (target sequence plus its flanking regions) and the genome of reference. The generation of the linear dsDNA recombineering fragment from the substrate for homologous recombination (target sequences plus upstream and downstream flanking regions) can be performed by endonuclease restriction of the recombinant plasmid constructed as described above or by PCR amplification using the same recombinant plasmid as template. To proceed via endonuclease restriction, digest the plasmid generated in Subheading 3.2 with the enzymes for which unique restriction sites were introduced during PCR to produce PCR1 and PCR2. These enzymes must be those capable of cleaving the most 50 -upstream and 30 -downstream sites flanking the target sequence. This is to keep up to ~500 bp the length of the sequences flanking the target gene. It is convenient to eliminate sequences remaining from the backbone plasmid, particularly because of the presence of oriE, also contained in pJV53, given that this may drive recombination between these vectors, therefore reducing recombination efficacy between the substrate for homologous recombination and genomic DNA.

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Separate the substrate for homologous recombination from vector backbone sequences by electrophoresis in an agarose gel. Purify the band corresponding to the substrate for homologous recombination by using a gel purification kit. Alternatively, purify the restriction enzyme digest directly using a PCR purification kit (see Note 3). In both cases, include an additional washing step that is recommended in the manufacturer’s instructions, to remove residual salts; elute in molecular biology grade water. The substrate for homologous recombination can also be produced by PCR using the recombinant plasmid as template and both the most 50 -upstream and 30 -downstream primers annealing to each one of the flanking sequences (P1 and P4 primers, Fig. 2), which were used to produce PCR1 and PCR2, as described in Subheading 3.2 (see Note 4). Store recombineering substrates at 20
C prior to electroporation (see Note 5). 3.3 Preparation of Recombinogenic/ Electrocompetent Slow Growing Mycobacteria

1. Grow the BCG strain harboring pJV53 in 20 mL of 7H9 OADC-Tween 80 broth containing 20 μg/mL kanamycin in static conditions, at 37
C and 5% CO2, until it reaches an OD600 of 0.8–1.0 (see Note 6).

Fig. 2 Primers for cloning and deletion characterization

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2. Transfer 5 mL of the culture to 15 mL tube and centrifuge at 5000 rpm (4500  g) for 10 min. 3. Discard the supernatant and resuspend the pellet in 10 mL of induction medium and centrifuge at 5000 rpm (4500  g) for 10 min. 4. Discard the supernatant and resuspend the pellet in 1 mL of induction medium. 5. Transfer the saturated culture to 20 mL of induction medium containing 20 μg/mL kanamycin (see Note 7), 6. Grow the culture in static conditions, at 37
C and 5% CO2 until it reaches late-log phase (OD600 ~ 1.0). 7. Dilute the culture up to an OD600 0.05 in a 175 cm2 tissue culture flask containing 100 mL of induction medium. 8. Transfer the tissue culture flask to an incubator and grow at 37
C and 100 rpm. 9. When the culture reaches the mid-log phase (OD600 of 0.4–0.5), induce the expression of the RecET proteins by adding 1 mL of acetamide solution (see Note 8). 10. Incubate with shaking at 37
C for ~8 h and then add 10 mL of 2 M glycine to the culture and continue shaking at 37
C and 100 rpm for 16 h. 11. Centrifuge at 5000 rpm (4500  g) for 10 min at RT in two 50 mL tubes. 12. Add 1 mL of 10% glycerol to each pellet and gently resuspend the cells by pipetting back and forth with a 1 mL pipet tip (see Note 9). 13. Transfer each suspension to 1.5 mL tubes. 14. Centrifuge at 5000 rpm (4500  g) for 1 min. 15. Remove supernatant with micropipette. 16. Add 1 mL of 10% glycerol to each pellet and resuspend the cells by pipetting back and forth with a 1 mL pipet tip. 17. Repeat points 10–12 three more times (see Note 10). 18. Transfer each last suspension to a 15 mL tube and mix with 3 mL of 10% glycerol. 19. These cells are now electrocompetent and recombinogenic (see Note 11). 3.4 Electroporation of the Substrates for Homologous Recombination

1. Add 400 μL of the competent cells to a 4 mm cuvette. 2. Add 100–1000 ng of the substrate for homologous recombination to the cells and mix gently with a micropipette. Include one electroporation without DNA as control (see Note 12).

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3. Place cuvette into the electroporation chamber and apply a pulse at 2.5 kV, 1000 Ω, 25 μF setting. 4. Remove the cuvette from the chamber, recover the cells with a sterile 1 mL disposable transfer pipet, and transfer the cells to a 15 mL tube containing 5 mL of 7H9-OADCTween 80 (see Note 13). 5. Incubate the cultures at 37
C for 24–72 h. 6. Collect the cells by centrifugation at 5000 rpm (4500  g) for 10 min, and resuspend in ~100–300 mL of 7H9-OADCTween 80. 7. Plate the entire suspension on 7H10 plates containing hygromycin 100 μg/mL by using 3 different volumes of the transformation and spread onto agar (e.g., 50, 200, 500 μL). As a control, also plate the culture electroporated with no DNA (control in step 1) (see Note 14). 8. Extend the inoculum with 3 mm glass beads and allow the plates to dry at RT within a biosecurity safety cabin. Then, introduce the plates within plastic bags and incubate with aluminum foil at 37
C for 3 weeks (see Note 15). 9. Examine the plates for colonies indicative of gene replacement. Plates inoculated with control aliquots (BCG electroporated with no DNA) should have no (or just a few) colonies (see Note 16). 3.5 Growth, Verification of Double Homologous Recombination Events, and Curation of the Recombineering Plasmid

1. Select 20 colonies for verification of the presence of the intended mutation by colony PCR. 2. With a sterile toothpick or a sterile micropipette tip, pick up a small portion of each one of the 20 candidate colonies and place each one of them into individual PCR tubes (PCR strips or individual wells from PCR multiwell plates) containing 30 μL of molecular biology grade water. Boil the samples for 15 min. 3. Using 2 μL of each one of the boiled colony samples, perform PCR reactions with primers annealing to upstream and downstream regions flanking the gene presumably exchanged for the hygromycin resistance gene. These primers could be the most 50 -upstream and 30 -downstream primers annealing to each one of the flanking sequences, which were used to produce PCR1 and PCR2, as described in Subheading 3.2, or, if desired, you can design and check with new primers, as long as these anneal further upstream and downstream with respect to those used to produce PCR1 and PCR2 (P5 and P8 primers, Fig. 2). Should the expected PCR product be too long, or the gene replacement was produced using a pYUB854-derived recombination substrate, it may be best to amplify each one of the upstream

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and downstream flanking regions separately. For this, use primers that anneal within the hygromycin resistance gene and that will be extended outward from each one of the 50 - and 30 ends, respectively. Combine each one of these reactions with a primer that anneals to each one of the regions flanking the target gene, which will be extended inward (P5/P6 and P7/P8 primer combinations, Fig. 2). 4. Select 5–10 colonies that in step 3 gave a PCR result indicating they contain the desired gene knock-out and pick them up with a sterile plastic loop to inoculate each one of them into 25 cm2 cell culture flasks containing 20 mL of 7H9-OADC-Tween 80 (see Note 17). 5. Incubate at 37
C and 100 rpm. 6. When the OD600 reaches 0.8–1.0, transfer 200 μL of each culture into new 25 cm2 tissue culture flasks containing fresh 20 mL 7H9-OADC-Tween 80 (see Note 18). 7. Again, when the OD600 reaches 0.8–1.0, transfer 20 μL into a new 25 cm2 tissue culture flasks containing fresh 7H9-OADCTween 80 and incubate at 37
C and 100 rpm. 8. When cultures reach late-log phase (OD 600 ~ 1.0), dilute cells 1:20 in 1 mL PBS-Tween 80 and prepare serial dilutions to reach 1:104. 9. Plate 100 μL of the 1:104 dilution onto 7H10 OADC agar plates and incubate at 37
C and 5% CO2 for 3 weeks to get isolated single colonies. 10. Randomly, select 50 colonies and screen these BCG knock-out candidates for kanamycin sensitivity (indicative of pJV53 plasmid loss) and hygromycin sensitivity (unmarked mutants) or resistance (marked mutants) by streaking each one of these candidates onto 7H10 OADC agar plates containing 20 μg/ mL kanamycin, 100 μg/mL hygromycin, and 7H10 plates without antibiotics (see Note 19). 11. Select and grow the kanamycin sensitive candidates up to latelog phase (OD 600 ~ 1.0) to produce stocks of the successfully constructed live genetically modified BCG strains in vials with glycerol to a final 25% and store them at 80
C and/or lyophilized. A schematic representation of the possible events after double homologous recombination is shown in Fig. 2.

4

Notes 1. The A260/280 ratio of the nucleic acid must be 1.8–2.0. Intact gDNA is large and therefore migrates very slowly forming a band at the top of the gel. Any bands/smears seen in the

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middle of the gel could be an indication of fragmented gDNA or the presence of RNA. Should protein/solvent/salts contamination be suspected (A260/280 ratio < 1.2), perform a second phenol–chloroform–isoamyl alcohol gDNA extraction, precipitation, and wash steps. For RNA contamination, perform a second RNase treatment (add 10 μL RNAse A) followed by steps aforementioned. 2. The target gene cloned in each vector should include enough flanking region on either side of the gene so that homologous recombination can occur. The minimum length of homologous region using pJV53 recombineering tool is 500 bp. However, longer fragments on either side of the target gene than 1 kb improve homologous recombination efficiency [17]. 3. The linear substrate for homologous recombination generated by restriction digestion can be purified from gel, or not. The main consideration is that contaminating, undigested or partially digested plasmid present in the whole restriction digest reaction, might recombine with pJV53 because of their sharing oriE sequences, which could lead to generate drug-resistant transformants (false positives). These false positives can be discarded after PCR because these will produce an amplicon of the same size as wild type BCG will do. The simultaneous presence of amplicons with sizes expected from wild type and gene knock-out can suggest a single homologous (merodiploid) event. The choice of purifying or not the substrate for recombination is up to the user. The main recommendation here is getting 0.1–1 μg of dsDNA recombineering substrates for each electroporation. 4. To obtain the substrate for homologous recombination by PCR requires a high-fidelity DNA polymerase. Amplification can be performed using a program with no more than 25 cycles and 0.1–1 ng of plasmid in a 50 μL PCR reaction as template to reduce incorporation of errors in sequence and plasmid contamination after amplicon purification. The PCR product should be purified using a PCR purification kit, washed twice, and eluted in molecular biology grade water. 5. PCR product purified is stable for 1 or 2 months kept in 4
C and for longer for a year stored at 20
C. However, when transforming a digestion product as substrate for homologous recombination, it is recommendable using recently digest from plasmid cloning vector. 6. All BCG culture and genetic manipulation must be carried out in appropriate containment facilities inside a class II safety cabinet and BSL2 laboratory. There is a risk of creating aerosols during the electroporation if arcing is violent dispersion of the cell suspension; it could also lead to lid blown off the cuvette. It

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is extremely important to maintain a good aseptic technique. Culture contamination can be evidenced by duplication time and media coloration in liquid cultures and in solid medium by the colony presence in less than 10 days of culture and a non-characteristic colony morphology. Also, purity can be checked using acid-fast staining. Fungal and bacterial contamination can be prevented by adding cycloheximide (100 μg/ mL) or amphotericin B (50 μg/mL) and carbenicillin (50 μg/ mL) to agar plates. 7. The acetamidase promoter is controlled by a form of catabolite repression and it has been observed in minimal medium. Eliminating the use of the dextrose-containing OADC supplement and including 0.2% glycerol and 0.2% succinate as carbon sources has been observed to increase colony numbers compared with the richer (glucose-containing) medium [28, 35]. It is recommended to avoid using OADC for induction and washing cells in induction medium before starting the culture for transformation. Mid-saturated pre-culture let cells adapt a medium without OADC. 8. Prolongated time of induction of recombinases drives cell toxicity. It is therefore highly recommended to check that induction time does not exceed 24 h [17]. 9. The electroporation solution for preparing cell suspension has effect on the cell solubility and the time constant. 10% glycerol provides a better solubility than water and a high-resistance medium, allowing longer time constants to be achieved. Solubility can improve adding 0.025% tyloxapol to avoid clumping. 10. Each washing step helps removing residual salts from culture media that can interfere with the efficiency of electroporation. A time constant of >20 ms should be expected for salt-free cell suspension using 4 mm cuvettes. A time constant of 90% of TB recurrence occurs in patients within 12 months of treatment completion [28]. The POR efficacy study design is smaller and shorter, but with a complex study population, compared to the POD studies [23, 27]. An effective POR vaccine may have a more significant impact in effectively controlling recurrence in countries where multi-drug resistant TB (MDR-TB) and extreme-drug resistant TB (XDR-TB) cases are prevalent and treatment for which are more protracted, expensive, and complicated [23, 25]. (d) Therapeutic Vaccines: Therapeutic vaccines are administered as an adjunct to anti-TB drug treatment, to increase the effectiveness and/or shorten therapy duration [24]. Current TB treatment regimens involve combinations of multiple antibiotics administered for 6 months for drug-susceptible TB and typically 9–20 months for rifampicin-resistant-TB, MDR- and XDR-TB. With the existing treatment regimen, the global success rate of the bacteriological cure is 85% for drugsusceptible TB, 56% for MDR-TB, and 39% for XDR-TB [1]. The therapeutic vaccines are useful in this scenario, mainly targeting MDR- and/or XDR-TB [22]. A therapeutic vaccine should ideally reduce disease severity and treatment failures and significantly improve the personal, logistical, and financial burden of TB management [24].

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5

Radha Gopalaswamy and Selvakumar Subbian

Candidate TB Vaccines in Clinical Trials A total of 16 vaccine candidates are in different phases of a clinical trial to fulfill the goals mentioned above. These vaccine candidates are divided into four major categories: live attenuated whole-cell vaccine, inactivated whole-cell vaccine, adjuvanted protein subunit vaccine, and viral-vectored vaccine. In general, these vaccines carry various mycobacterial antigens and induce divergent levels of hostimmune response.

5.1 Live Attenuated Whole-Cell Vaccine

Recently developed, live attenuated mycobacterial vaccine candidates aimed to replace the current BCG vaccine include recombinant BCG (rBCG) strains, such as VPM1002 and recombinant Mtb mutants that have a deletion in one or many genes and MTBVAC, respectively.

5.1.1 VPM1002

VPM1002 is an rBCG Prague (ΔureC hly + rBCG) strain that expresses the listeriolysin from Listeria monocytogenes and lacks the urease C gene developed by Max Planck Institute, Germany [29, 30]. Animal studies were done with rBCG containing hly+, which showed better protection than BCG when challenged with Mtb. However, the listeriolysin gene was known to form transmembrane β-barrel pores in the phagosomal membrane and works best at an acidic pH of 5.5. This acidic pH was created in the improvised strain by deleting the ureC gene, and the strain provided antigenic translocation into the cytoplasm as well as apoptosis of host macrophages [29]. The rBCG has been tested for safety and efficacy in mice, guinea pigs, rabbits, and nonhuman primate models with or without Mtb challenge and compared to BCG [31]. The Vakzine Projekt Management (VPM) of Germany facilitated further development of this vaccine candidate and sponsored clinical testing of this novel rBCG vaccine. In collaboration with the nonprofit product development organization, Aeras [32], phase I trials conducted in Germany and South Africa showed that VPM1002 was safe and immunogenic for B-cell and T-cell responses. This was followed by a phase IIa randomized clinical trial in healthy South African newborns, which showed that VPM1002 was safe, well-tolerated, and immunogenic in newborn infants. A phase IIb study in HIV exposed and unexposed newborns has been completed and requires unblinding [22, 33, 34]. VPM1002 is currently investigated for its efficacy against TB relapse among adolescents and adults treated for active TB, in a phase III trial in India [22, 35].

5.1.2 MTBVAC

MTBVAC is a live attenuated vaccine derived from a clinical isolate of Mtb, MT103, by deleting the phoP and fadD26 genes, made by the University of Zaragoza, Biofabri, Spain, and Tuberculosis

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Vaccine Initiative (TBVI) [36]. PhoP is a critical transcriptional regulator, which controls 2% of Mtb coding genes, and Fad26 is essential for the synthesis of phthiocerol dimycocerosates (DIM), a family of surface lipids involved in Mtb virulence [37]. MTBVAC was found safe in all preclinical studies where mice and guinea pig models were used and conferred superior protection in mice, compared to BCG [37]. MTBVAC, the first live attenuated vaccine based on a clinical Mtb isolate, entered the first-in-human clinical evaluation in January 2013. In this study, MTBVAC showed a similar safety profile as BCG [38]. Revaccination of MTBVAC seemed to improve the efficacy of BCG, as indicated in the guinea pig model [39]. In May 2018, phase Ib clinical trials were completed for MTBVAC in adults and infants. Two phases (IIb/IIa) of clinical trials and epidemiological studies have been carried out in South Africa by Aeras and Biofabri with adults and/or infants. MTBVAC immunogenicity and its effects on IGRA conversion and reversion were also investigated and demonstrated a promising induction of mycobacterium-specific CD4+ T-cells [40]. Two additional phase II trials are ongoing in South Africa [17, 24]. 5.1.3 AERAS 422

The AERAS 422 is a live attenuated vaccine, a novel recombinant BCG (derived from Danish 1331 strain) expressing perfringolysinO, associated with lysis of the endosome compartment and the over-expression of vital immunodominant antigens—Ag85A, Ag85B, and TB10.4. This prime strategy vaccine developed by Aeras showed improved immune responses in mice and guinea pig models compared to parental BCG and acceptable safety in preclinical studies [41]. However, in phase I clinical trial, it was shown to reactivate varicella-zoster virus infection in 2 of 8 healthy adults, and further studies with the vaccine were discontinued [42]. Safety is a significant concern while using a live attenuated vaccine since they can persist longer in vivo and possibly exhibit unacceptable adverse events, particularly among immunecompromised hosts, despite their ability to elicit a robust immune response.

5.2 Killed Mycobacterial Vaccines

The second type of vaccine candidates uses killed mycobacterial cells or cell extracts as a source of antigens to invoke a protective immune response. Killed mycobacterial vaccine preparations currently in clinical trials include RUT-1, DAR-901, Mycobacterium vaccae, and Mycobacterium indicus pranii (MIP).

5.2.1 RUT-1

RUT-1 is made of detoxified and fragmented Mtb and delivered through liposomes developed by Archival Pharma. This vaccine reduced the duration of treatment for LTBI and induced a more robust immune response that contains bacillary load and pulmonary pathology in preclinical studies with mice and guinea pig models of infection [43, 44]. Besides, these vaccines safety and

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efficacy have been validated in numerous experiments conducted in mice, guinea pigs, goats, and minipigs [45]. Phase I clinical trials of RUT-1 were conducted to evaluate the safety profile and T-cell immune responses following subcutaneous injection of 4 different dose regimens over 6 months. These studies established the safety profile, prophylactic, and immunotherapeutic capacity of RUT-1 against TB [45, 46]. Phase II studies demonstrated the safety, efficacy, and immunogenicity of the vaccine for LTBI among HIV-infected and uninfected cases. It showed reasonable tolerability and a polyantigenic response with a single dose at the highest tested concentration proving efficacious [47]. An additional trial in individuals with multidrug-resistant TB is ongoing and expected to be completed in July 2020 [24]. 5.2.2 Dar-901

DAR-901 is a whole-cell, heat-killed vaccine prepared from M. obuense, based on a previously tested vaccine, SRL172, using a large-scale broth culture method. This vaccine was developed by Dartmouth University, along with Aeras (Rockville, MD, USA). Although the mycobacterial strain used in DAR-901 was initially identified as M. vaccae, by phenotypic methods, detailed 16S rRNA sequencing determined the strain as M. obuense [48]. SRL-172, an inactivated whole-cell vaccine, was safe, well-tolerated, and immunogenic in humans and became the first vaccine after BCG to enter the phase III clinical trial. However, the results showed a lack of efficacy in preventing confirmed TB cases, and the vaccine study was stopped after 7 years [49, 50]. In preclinical studies using a mice model, DAR-1 showed safety and protective efficacy. DAR-901 induced cellular and humoral immunity and boosts protection against the aerosol challenge of Mtb compared to a homologous BCG boost [51]. Phase I clinical trial was conducted with IGRA-negative healthy volunteers and found that DAR-901 had an acceptable safety profile, well-tolerated with no serious adverse events and induced cellular and humoral immune responses to mycobacterial antigens [48]. A randomized, placebo-controlled, double-blind phase IIb clinical trial has been initiated with adolescents who had previously received BCG in the United Republic of Tanzania [24, 48].

5.2.3 M. Vaccae

Heat-killed M. vaccae, a nontuberculous mycobacteria based vaccine, is manufactured by Anhui Zhifei Longcom Biologic Pharmacy Co., Ltd. (Anhui, China). The National Institutes for Food and Drug Control, China, and the 309th Hospital of the People’s Liberation Army (Beijing, China) jointly developed this vaccine [17]. M. vaccae has been shown to potentiate the Th1 response to manifest a more protective antibacterial immunity and suppress the Th2 component [52]. Studies using a mouse model with low-dose immunization of M. vaccae induced Th1 response and

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was more protective and displayed lower bacterial load when challenged with Mtb [53]. Five doses of inactivated whole-cell M. vaccae were shown to be safe and immunogenic in phase I studies done with HIV-infected adults and children in Zambia [54]. Phase II clinical trials in HIV-infected and BCG-vaccinated adults in Finland indicated an exact host-protective role for M. vaccae that prevented HIV-associated TB [55]. A phase III trial in Tanzania suggested that the administration of a multipledose series of M. vaccae to HIV-infected adults with childhood BCG immunization is safe and is associated with significant protection against HIV-TB [50]. M. vaccae is already licensed as an adjunctive therapeutic vaccine in TB patients in China [24]. 5.2.4 MIP

Mycobacterium indicus Pranii (MIP) was a killed vaccine, which was developed at the Talwar Research foundation in India and Cadila Pharmaceuticals for use as a vaccine for leprosy. However, testing it in a guinea pig model showed that the vaccine could protect against Mtb infection. The same was confirmed in various mouse models of Mtb infection, in comparison to BCG, and proven to offer protection in live and killed form, unlike BCG, which works in only the attenuated live form [56]. In mouse and guinea pig models, MIP was given as a booster to BCG, Th1 and Th17 immune responses were induced, and the vaccine displayed protective efficacy [57]. The immunotherapeutic potential of MIP, as an adjunct therapy to anti-TB treatment, was tested in category II patients; in this study, MIP vaccination of patients showed improvement in treatment and reduction in the conversion to MDR [58]. MIP is phase III completed vaccine for pulmonary TB [22].

5.3 Adjuvant Protein Subunit Vaccines

TB antigens, when administered alone, fail to elicit a robust immune response. Subunit vaccines require another component called the adjuvant to stimulate and activate the immune response to the vaccine. The choice of vaccine is critical, and the TB vaccine requires an adjuvant to stimulate the Th1 and Th17 responses. Potential TB vaccines currently use TLR agonists, liposomal formulation, or a combination along with different delivery vehicles [59].

5.3.1 H1: IC31

The H1:IC31 is a subunit adjuvant vaccine developed by Statens Serum Institut, Denmark, Valneva, France, and TBVI. The H1 is a hybrid of TB antigens ESAT-6 and Ag85B combined with IC31, which utilizes a cationic polypeptide as a delivery vehicle [60]. H1: IC31 showed safety and protection in mouse and guinea pig models of TB [61]. Besides, the vaccine-elicited adult-like protective T-cell responses in a neonatal murine immunization model proved its use in neonates [62]. This vaccine has passed three different phase I trials. When tested on naı¨ve human volunteers, it showed

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safety and immune response to both antigens, ESAT-6 and Ag85B and induced substantial memory response as followed up for 2.5 years [63]. In another trial, safety and immune response were compared to healthy BCG-vaccinated individuals and prior or latently TB-infected individuals with the administration of vaccine twice. It showed persistent memory response when followed up for 32 weeks and rare local or systemic effects due to vaccination indicating good safety [64]. In the third study, H1: IC31 was tested in individuals with or without TB in a highly TB endemic area. Except for two serious adverse events, in which the patients recovered without sequelae in 72 h, the rest of the study population showed a strong immune response, and the vaccine was welltolerated [65]. Further, in phase II studies, H1: IC31 showed safety and immunogenicity in the HIV-infected population [66]. 5.3.2 H56:IC31

The H56:IC31 is an adjuvanted protein vaccine developed by Statens Serum Institute, Denmark, in collaboration with Valneva, France, and Aeras. H56 is a hybrid antigen formed by the fusion of Ag85B, ESAT6, and Rv2660c added to adjuvant IC31 [60]. Rv2660c is a well-known latency-associated protein, and H56 has shown better protection against reactivation of bacterial loads in mouse models of latent TB [67]. In the nonhuman primate model of low- and high-dose aerosol challenge, H56:I31 showed reasonable control of late-stage infection and latent TB [68]. The safety and tolerance of H56:IC31 and its immune response were studied in a phase I trial. Both uninfected and Mtb-infected individuals were included and examined using different doses of the vaccine. The study indicated both antigenic IgG response and CD4-specific T-cell induction. Besides, a lower dose prompted a good immune response and reiterated the monitoring of bradycardia in future clinical trials [69]. In another phase I trial, dose optimization and schedule for H56:IC31 were studied, and two to three vaccination at low dose was found ideal with acceptable safety and tolerated well [70]. Phase II trial for reducing recurrence of TB in HIV-negative people who are treated with anti-TB drugs is ongoing in South Africa and in the United Republic of Tanzania [17].

5.3.3 H4: IC31

H4: IC31 is another adjuvant vaccine developed by Sanofi Pasteur and Staten’s Serum Institute, Denmark, and Valneva, France. The H4 is a fusion peptide of Ag85B and TB10.4 of Mtb, co-formulated with the adjuvant IC31 [60]. In mouse and guinea pig models, the safety and immunogenicity of the vaccine, as well as its optimal dose, were established [71]. Phase I trial was carried out in BCG immunized individuals, with different quantities of the antigen and adjuvant. Although this study was conducted in Sweden and Finland, countries with a low TB incidence, the vaccine was

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safe, efficacious, and induced a T-cell response under the tested conditions [72]. In phase II trial, the vaccine was given to previously neonatal BCG immunized individuals, who were negative for Mtb infection, as estimated by QuantiFERON conversion [73]. The H4:IC31 appears to be a promising vaccine candidate, similar to H1:IC31 and H56:IC31. All the three vaccine candidates are currently pursued by Aeras, after phase II trial sponsored by them. 5.3.4 ID93: GLA/SE

The ID93:GLA/SE is an adjuvant vaccine developed by the Infectious Diseases Research Institute and has a fusion protein of four mycobacterial antigens—Rv2608, Rv3619, Rv3620, and Rv1813 administered with GLA-SE as an adjuvant [23]. Preclinical safety and immunogenicity of ID93:GLA/SE were studied in mice, guinea pigs, and nonhuman primates. The protein subunit vaccine ID93/GLA-SE showed protection against TB and MDR-TB in animals and is a candidate for boosting the protective efficacy of the childhood BCG vaccine [74, 75]. Besides, ID93:GLA/SE showed adequate therapeutic immunization against Mtb when combined with first-line drugs, rifampicin, and isoniazid, in mouse and nonhuman primate models [76]. Phase I clinical trials were conducted in the USA and South Africa. In Phase I trial conducted in the USA, different doses of the vaccine-adjuvant were given as three injections 28 days apart in BCG-naı¨ve, uninfected (QuantiFERON negative) healthy individuals. The ID93: GLA/SE showed acceptable safety in all doses and elicited antibody response. In combination with GLA/SE, ID93 showed good humoral and helper T-cell responses [77]. Another phase I trial was conducted among BCG-vaccinated healthy individuals in South Africa, a high incidence TB endemic setting. This study showed a safety profile for the different doses administered without any adverse events. Further, ID93:GLA/SE elicited long-lasting antigen-specific IgG and Th1 responses [78]. A phase IIa clinical trial of the ID93:GLA-SE vaccine was carried out in South Africa to assess the safety and immunogenicity of TB-HIV coinfected adults following treatment completion. This trial showed encouraging CD4+ T-cell and antibody responses to vaccination. Phase IIb clinical trials are currently underway in the same population intended to prevent TB recurrence [17, 79].

5.3.5 M72/AS01E

The M72/AS01E is an adjuvant vaccine developed by GlaxoSmithKline in association with Aeras consisting of a fusion protein of two mycobacterial antigens, Rv1196 and Rv0125 [22]. Preclinical trials were done using M72/AS01B formulation in mouse and guinea pig model wherein the vaccine showed safety and protection against a low-dose aerosol Mtb challenge [80]. The M72/AS01B elicited a robust and comprehensive CD4+, CD8+ T cell-mediated

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immune response in the mouse model. Immunization of guinea pigs with the vaccine also resulted in prolonged survival (>1 year) after aerosol challenge with virulent Mtb, comparable to BCG immunization [80]. Besides, the protective effect of BCG was increased upon coadministration of M72/AS01A in a guinea pig model [81]. M72/AS01A was evaluated in a nonhuman primate model and found immunogenic with no adverse effect. When used to boost BCG, it showed long term protection [82]. After phase I and II trials with AS01 formulations, AS01E was finally progressed [80, 83, 84]. The vaccine conferred protection to adults who received or receiving TB treatment in a phase II trial. Still, there was a local reaction observed in vaccinated individuals, and the trial was suspended [85]. In another phase II trial, the safety, reactogenicity, and immunogenicity of M72/AS01E were evaluated in HIV-negative adolescents in a TB endemic region. In this study, no adverse events were noted, and both CD4 and CD8 T-cell responses were elicited by this vaccine [86]. In a phase IIb trial in Kenya, South Africa, and Zambia, M72/AS01E showed 54% protection among Mtb-infected adults against active pulmonary TB, without much safety concerns, giving promise that M72/AS01E may aid in controlling TB transmission [87]. In another phase II trial conducted in India, the vaccine was shown to elicit cellular and humoral responses, which persisted for nearly 3 years in HIV-negative and HIV-positive individuals [88]. 5.3.6 GamTBVac

GamTBVac was manufactured by Gamaleya Federal Research Centre, Russia. This vaccine consisted of a fusion protein of TB antigens, ESAT6, CFP10, and Ag85A combined with new adjuvant composition dextran/CpG. GamTBVac was proven safe and immunogenic in mice and guinea pig models. It showed a protective effect against Mtb infection in an aerosol and intravenous challenge. It was also influential in boosting the impact of prior BCG vaccination [89]. Phase I trial of GamTBVac was done in BCG-vaccinated, uninfected, healthy individuals with different doses of the vaccine and found to be immunogenic and safe in tested conditions [90]. Currently, phase IIa trial is ongoing in Russia in healthy BCG-vaccinated adults [23, 24].

5.4 Viral-Vectored Vaccines

Live, attenuated, and non-replicating viruses can be engineered to deliver genes encoding the antigens of interest into host cells. Viral vector vaccines do not require adjuvants, and the viral backbone presents the antigen in vivo. These groups of vaccines can induce robust and sustained immune responses in the lungs. This technology also helps to clone multiple immune-dominant antigens into the viral vectors, which can be easily scaled up in high titers, allowing easy vaccine production. However, the vector must be safe enough to prevent the occurrence of vector-based pathogenesis and/or the induction of vector-specific immunity that can interfere

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with subsequent booster vaccinations, particularly in immunosuppressed individuals [36, 79, 91]. 5.4.1 MVA85A

The MVA85A is a subunit viral-vectored vaccine in which a recombinant, replication-deficient strain of vaccinia virus Ankara is used as a vector to carry Mtb antigen 85A. It was developed by Oxford University along with Aeras and hence named as AERAS-485. Modified vaccinia virus Ankara (MVA) was attenuated through multiple passages, but it retains the ability to express Mtb proteins [23, 91, 92]. A preclinical evaluation has been done in mice, guinea pigs, and nonhuman primates for MVA85A. In mice, MVA85A administration showed Ag85A-specific CD4 and CD8 T-cell response. The vaccine boosted a higher level of protection and reduced bacterial load following the Mtb challenge, compared to BCG alone [93]. In guinea pig and nonhuman primates, MVA85A boosted BCG and adequate protection against Mtb aerosol challenge [94–96]. The safety and efficacy of MVA85A have been shown in several phase I studies, which involved different doses, administration routes, and population groups. In the first phase I study in the UK, MVA85A induced a high level of IFNγ secreting T-cells, which increased many-fold after a booster dose when tested in BCG-naı¨ve BCG-vaccinated individuals promising to be useful in TB endemic setting [97]. The safety and immunogenicity of MVA85A were assessed in mycobacteria-exposed but uninfected healthy adults in TB endemic South Africa as well as in latently infected individuals in the UK; both studies show significant efficiency for this vaccine [98, 99]. Further analyses identified the ideal dose in BCG-vaccinated infants, HIV-infected adults, and BCG-vaccinated healthy adults; all these studies confirmed good safety and immunogenicity with no adverse events [100–103]. In another phase I trial, the delivery route was compared between the intramuscular and intradermal route in BCG-vaccinated healthy adults, where the intramuscular proved safer and more immunogenic [104]. Besides, the aerosol way of administration was compared to the intradermal route in a phase I trial. Ag85A-specific CD4 T-cells were detected in bronchoalveolar lavage cells from both groups; however, the immune response was robust in the aerosol group than in the intradermal group. While MVA-specific cellular responses were detected in both groups, serum antibodies to MVA were detected only after intradermal administration of the vaccine [105]. In phase IIa trial, MVA85A was found to be safe and immunogenic when tested in HIV and/or Mtb-infected individuals in TB endemic settings [106]. However, in a Phase IIb trial, MVA85A showed no efficacy in protecting infants previously vaccinated with BCG against TB. Hence, a later trial planned with adults in South Africa was suspended [23, 24, 105]. Several reasons were outlined for the lack of efficacy, including use of a single antigen, hypo-immune responsiveness in infants, immunological

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interference by EPI vaccines, boosting at the peak of the BCG response, immunosuppression in HIV-infected adults, decreased Ag85A expression after Mtb infection, and reduced Ag85A availability in the lungs during chronic infection [24]. Currently, MVA85A is tested for its effect as a booster vaccine combined with other viral-vectored vaccines in phase I studies [107]. 5.4.2 ChAdOx1.85A/ MVA85A

Several unique features of Adenovirus with the lung as its target, ability to tolerate multiple antigens and safety, and adjuvant effect made it a favored choice. The immune interference observed with human Adenovirus prompted using related ones like attenuated chimpanzee adenoviral vector (ChAd) as vaccine backbone for use in humans [108]. ChAdOx1.85A is a simian adenoviral vector carrying Mtb Ag85A developed by Oxford University. In preclinical studies, ChAdOx1.85A was tested alone or in combination with MVA85A for its safety and efficacy against Mtb challenge. When administered to naı¨ve and BCG-primed mice, the combination of ChAdOx1.85A/MVA85A administered either by mucosal or systemic route improved the BCG vaccine [109]. Phase I studies in BCG-vaccinated healthy adults in the UK showed that ChAdOx1.85A prime—MVA85A boost vaccination was safe and efficacious with no serious adverse events [110].

5.4.3 Ad5Ag85A

AdAg85A is a replication-deficient serotype 5 adenoviral vector containing the natural Mtb antigen 85A. This vaccine was developed by McMaster University and CanSino [92]. In a murine model, Ad5Ag85A showed better protection compared to BCG when administered intranasally, compared to the intramuscular route. However, a systemic administration of Ad5Ag85A followed by a mucosal booster with the same vaccine showed better T-cell responses than BCG [111]. Further studies indicated that intranasal immunization elicited a more robust IFN-γ positive CD4 and CD8 T-cell response in the mice‘s airway lumen either when given as a primary vaccine or a boost to BCG-primed animals [112, 113]. In a guinea pig model, intranasal and intramuscular administration of Ad5Ag85A was tested in a prime or prime-boost to BCG. The study showed that prime boosting by the intranasal mucosal route elicits long term survival of BCG-primed guinea pigs following the Mtb challenge [112]. In a phase I study, the vaccine was well-tolerated and immunogenic in BCG-naive and BCG-vaccinated healthy adults with more potently boosted CD4 and CD8 T-cells in BCG-vaccinated individuals. Concerns have been raised about using Ad5 as a vaccine vector due to the high prevalence of preexisting serotype 5 adenovirus-neutralizing antibodies. Still, the study showed little effect in the vaccines potency against TB and needed further validation [114].

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5.4.4 Crucell Ad35

Adenoviral vectors based on adenovirus 35 show low natural immunity and strong T and B-cell response. Stable Ad35 adenoviral vector carries Mtb antigens Ag85A, Ag85B, and TB10.4 as a fusion protein was generated by Crucell in collaboration with Aeras, USA, and named Crucell Ad35 or AERAS-402. In a mouse model, the vaccine-elicited CD4 and CD8 response to Mtb antigens [115]. Further, the vaccine conferred better protection than BCG against Mtb by intranasal or intramuscular route with a varied T-cell response in two different mouse models [116]. The AERAS-402 has been shown to elicit an immune response similar to BCG using an aerosol route in the nonhuman primate model. However, it failed to show protection against high-dose Mtb challenge, indicating that model needs other Mtb antigens or rAd serotypes or different challenge doses [117]. In Phase I trial done in low TB endemic area, the AERAS-402 showed a good T-cell response when used to boost BCG-primed volunteers, healthy BCG-vaccinated infants as well as HIV-infected adults [118– 120]. In phase IIb clinical trial using healthy BCG-vaccinated infants, the vaccine was tolerated except in one case, the T-cell response was lower than that observed in healthy adults. The addition of another dose failed to improve the response [121]. However, the trial was stopped based on the above results and did not move on to the efficacy phase.

5.4.5 TB/FLU-04L

The TB/FLU-04L vaccine consists of a live attenuated influenza A/Puerto Rico/8/34 H1N1 virus vector that expresses Mtb antigens Ag85A and ESAT-6. This is delivered through the intranasal route and found to be safe and immunogenic [24]. This was developed by the Research Institute of Influenza, Saint Petersburg, Russia. Preclinical studies have shown the protective efficacy of this vaccine in mice, guinea pig, and nonhuman primate model [23]. A phase I study conducted on previously BCG-vaccinated, IGRAnegative, healthy volunteers identified the vaccine to be safe and immunogenic with no serious adverse effects, as well as no influenza infection was reported [23, 122]. A phase IIa is planned with QFT-positive adults for TB/FLU-04L vaccine [24, 60]. The vaccines VPM1002 and MTBVAC were developed as prime vaccine, while DAR-901, H1:IC31, H56:IC31, H4: IC31, ID93: GLA/SE, M72/AS01E, GamTBVac, ChAdOx1.85A, MVA85A and Crucell Ad35 were all developed as a prime booster vaccine. The vaccines RUT-1, M. vaccae, and MIP were developed as an immunotherapeutic vaccine [17, 60].

6

Conclusion Vaccination is vital in reducing the incidence of any infectious disease. The END-TB strategy aims at eliminating TB deaths and reduce the prevalence by the year 2035. However, BCG is the only

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widely used vaccine for TB control, despite the number of TB cases did not significantly decline over the years. This warrants the need for a new TB vaccine to be used in global TB management. To this end, a wide variety of vaccines have been developed—one required for prime, boost, or prime-boost, prevention of infection, disease, or recurrence/reactivation of latent Mtb infection. Different approaches have been employed to generate these types of vaccines, including whole live attenuated vaccine, killed vaccine, adjuvant vaccine, and viral-vectored vaccine. All of the candidate vaccines described here have been tested in multiple preclinical models, such as mice, guinea pigs, and/or nonhuman primates, and showed a good T-cell response. These vaccines showed adequate safety and efficacy in human clinical trials in various populations in TB endemic and non-endemic countries. These comprehensive vaccine development studies show 16 candidate vaccines in the pipeline at multiple phases—I, II, and III of clinical development. However, only a few showed a superior protective effect against TB compared to BCG. Interestingly, many of the vaccines carry mostly the same Mtb antigens, and the search for novel and more effective/efficient antigens is still a promising field. Apart from being used to prevent TB disease, therapeutic vaccines can offer better treatment outcomes while being used along with chemotherapy, particularly in TB endemic countries with a rising number of MDR- and XDR-TB cases. Another potential requirement towards developing an improved TB vaccine is to thoroughly analyze the consolidated outcome of various clinical trials conducted with different candidate vaccines, rather than focusing on the individual vaccine performance report. Such an analysis would reveal the immune correlations of the protective response needed to combat TB. Nonetheless, the knowledge gained from vaccine biology, preclinical findings, and clinical trials helps investigators worldwide devise the most effective strategy to prevent Mtb infection and/or protect against active disease. References 1. WHO (2019) Global tuberculosis report 2019. https://www.who.int/tb/ publications/global_report/en/ 2. WHO (2015) End TB strategy. The World Health Organization, Geneva, Switzerland. https://www.who.int/teams/globaltuberculosis-programme/the-end-tbstrategy# 3. Colditz GA, Brewer TF, Berkey CS, Wilson ME, Burdick E, Fineberg HV et al (1994) Efficacy of BCG vaccine in the prevention of tuberculosis. Meta-analysis of the published literature. JAMA 271(9):698–702

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Chapter 21 Structure-Based Design of Diagnostics and Vaccines for Lyme Disease Sunil Thomas Abstract Changes in climate have increased the geographical range of insect vectors responsible for the transmission of several diseases. Lyme disease, caused by the bacterial pathogen Borrelia burgdorferi, has become recognized as the most prevalent arthropod-borne infection in the USA. It is transmitted to humans through the bite of infected blacklegged ticks. As yet, there are no commercial vaccines available that effectively provide protection against Lyme disease. Vaccination strategies involving use of subunit vaccines developed in many laboratories have been found to be less efficient in protecting against the disease. Hence, there is a need to develop powerful vaccines that provide robust protection against Borrelia. Recently, using the principle of structure-based design, we designed and developed novel diagnostics and vaccine candidates that protected against Lyme disease in animal models. This chapter describes design and development of peptides based on the principle of structure-based design for use in diagnostics and vaccines to protect against Lyme disease in an animal model. Key words Lyme disease, Lyme borreliosis, Borrelia burgdorferi, Structure-based vaccines, Diagnostics, Animal model

1

Introduction Lyme borreliosis or Lyme disease is a tick-borne disease that predominantly occurs in temperate regions of the northern hemisphere and is primarily caused by the bacterium Borrelia burgdorferi in North America and Borrelia afzelii or Borrelia garinii in Europe and Asia [1]. The genus Borrelia is a member of the family Spirochaetaceae (spirochetes). The incidence of Lyme disease in the USA has been increasing since national surveillance with the use of a standardized case definition was instituted in 1991. There is no way of knowing exactly how many people get Lyme disease. A recently released estimate based on insurance records suggests that each year approximately 476,000 Americans are diagnosed and treated for Lyme disease. This number is likely an overestimate of actual infections because patients are sometimes

Sunil Thomas (ed.), Vaccine Design: Methods and Protocols, Volume 1: Vaccines for Human Diseases, Methods in Molecular Biology, vol. 2410, https://doi.org/10.1007/978-1-0716-1884-4_21, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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treated presumptively in medical practice. Regardless, this number indicates a large burden on the health care system and the need for more effective prevention measures [2]. The great majority of cases occur in Northeastern United States, with additional foci in northern midwestern states (Wisconsin and Minnesota). Lyme disease also occurs in the Pacific coastal regions of Oregon and Northern California. Although the geographic range of Lyme disease remains limited, it has been expanding, primarily due to changes in climate. The incidence of Lyme disease is highest among children 5 to 14 years of age and middleaged adults (40 to 50 years of age), and it is slightly more common among males than among females [3]. The major natural reservoirs for B. burgdorferi are mice, chipmunks, and other small mammals, as well as birds. Deer are not competent hosts for B. burgdorferi but are important in sustaining the life cycle of the vector ticks [3]. B. burgdorferi spirochetes are transmitted through the bite of tick species belonging to the genus Ixodes, which are largely confined to temperate climate zones of the northern hemisphere. In North America, the Ixodes species that transmits B. burgdorferi is primarily Ixodes scapularis, whereas Ixodes pacificus may act as a vector in the western coastal regions. In Europe, Ixodes ricinus is the tick species primarily responsible for transmitting B. burgdorferi, whereas Ixodes persulcatus is predominant in large parts of Russia and Asia [4]. Transmission is most likely during the nymphal stage, since nymphs are abundant in the spring and early summer and are small and difficult to detect [3]. The most common sign of Lyme disease is erythema migrans. Erythema migrans usually begins as a small erythematous papule or macule that appears at the site of the tick bite 1 to 2 weeks later and subsequently enlarges. The skin lesion is frequently accompanied by influenza-like symptoms, such as malaise and fatigue, headache, arthralgias, myalgias, fever, or regional lymphadenopathy, and these symptoms may be the presenting manifestation of the illness [1, 5]. The spread of B. burgdorferi within the nervous system has been demonstrated in nonhuman primates [6]. In up to 5% of untreated patients, B. burgdorferi may cause chronic neuroborreliosis, sometimes after long periods of latent infection [7]. 1.1 History of Lyme Disease

In the early 1970s a cohort of children and adults in Lyme, Connecticut, and the surrounding areas were suffering from an idiopathic and debilitating disease. Their symptoms included swollen knees, paralysis, skin rashes, headaches, and severe chronic fatigue. Visits to doctors and hospital stays had become all too common. All of them were bitten by ticks of that region. The condition was given the name Lyme disease in 1977 [8]. Willy Burgdorfer and his associates in 1982 isolated the infectious agent that causes Lyme disease that now bears his name:

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Borrelia burgdorferi [9]. Burgdorfer and colleagues collected and dissected Ixodes ticks from Shelter Island, New York (a region with a high prevalence of Lyme disease) and found that they contained spirochetes, specifically in the midgut region. Indirect immunofluorescence revealed that antibodies from sera of Lyme diseaseinfected patients reacted positively with the spirochete, whereas sera from healthy subjects did not, thereby confirming the link between the tick-derived spirochete and Lyme disease [10]. 1.2 B. burgdorferi Structure

B. burgdorferi is a Gram-negative spiral bacterium bound by an inner cytoplasmic membrane and an outer membrane. The outer membrane lacks lipopolysaccharide and consists of a lipid bilayer that is composed of phospholipids and glycolipids. Cholesterol glycolipids in the outer membrane form lipid-raft-like microdomains that change in order and size in response to temperature, which is an important environmental cue for B. burgdorferi during transmission between the tick vector and the mammalian host. The B. burgdorferi outer membrane also contains surface lipoproteins, which can change depending on the environment. The outersurface proteins of B. burgdorferi include OspA, OspB, OspC, OspD, OspE, OspF, DbpA, DbpB, CspA, VlsE, BptA that are lipid-modified outer-surface proteins anchored to the outer leaflet of the outer membrane through their lipid moieties [11]. P13, P66, BesC, BamA, Lmp1, and BB0405 are outer-surface proteins that have one or more transmembrane domains that anchor them into the outer membrane [11]. B. burgdorferi bacteria live in the midgut of ticks. In ticks, B. burgdorferi predominantly express the outer-surface protein A (OspA) before the blood meal, when the bacteria is in the midgut. OspA mediates the attachment of B. burgdorferi to the tick midgut by binding the midgut receptor TROSPA (tick receptor for OspA). When infected ticks feed blood meal, the spirochetes multiply within the gut, express high levels of OspC, migrate to the tick’s salivary glands, and infect the vertebrate host. OspC has been shown to bind a tick salivary protein, Salp15, in vitro and in vivo, indicating a possible role for OspC in transmission and/or survival early during host colonization [1]. Thus, OspA is required for colonizing the vector, whereas OspC is required for infecting the host. OspF is identified as a potential adhesin and binds to organs of the human host. B. burgdorferi is able to persist in patients for extended periods and establish chronic infection in host tissues. VlsE is an important virulence determinant of B. burgdorferi [11]. The periplasmic flagella of B. burgdorferi is responsible for the flat-wave morphology of the bacteria. The flagella are attached to each cell pole and wind around the cell cylinder in the periplasmic space between the peptidoglycan layer and the outer membrane. Flagellar motors are located at the cell poles and are situated next to the methyl-accepting chemotaxis proteins that direct movement of

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the bacteria toward chemoattractants (including nutrients) and away from repellants including organic solvents [1]. The genome of the bacterium Borrelia burgdorferi contains a linear chromosome of 910,725 base pairs and at least 17 linear and circular plasmids. The chromosome contains 853 genes encoding a basic set of proteins for DNA replication, transcription, translation, solute transport, and energy metabolism [12]. B. burgdorferi has a very limited metabolic capacity and is highly dependent on its tick vector and vertebrate host for most essential factors. B. burgdorferi lacks genes encoding proteins that have a role in the tricarboxylic acid cycle and oxidative phosphorylation and relies exclusively on glycolysis for energy production. For this purpose, B. burgdorferi uses several host or vector-derived carbohydrates, including glucose, glycerol, maltose, N-acetylglucosamine, trehalose, and chitobiose. The B. burgdorferi genome also lacks genes that are required for the synthesis of amino acids, lipids, nucleotides, and cofactors; to obtain these factors, the B. burgdorferi genome encodes 16 distinct membrane transporters, many of which have broad substrate specificity. Owing to the inability of B. burgdorferi to synthesize fatty acids, its lipid composition reflects that of the host tissues; B. burgdorferi exchanges lipids with the plasma membrane of eukaryotic cells, either through direct contact or via outer membrane vesicles [1]. Arthritis is a prominent manifestation in patients with Lyme disease. The majority of individuals with Lyme disease have the HLA-DRB1*0401 or HLA-DRB1*0101 allele; these alleles also occur more frequently in patients with rheumatoid arthritis [13]. HLA-DRB1 belongs to MHC-II and is recognized by T-cell. In this study we selected the epitope regions of the antigenic proteins that bind to HLA-DRB1.

2 2.1

Materials Bioinformatics

1. NCBI protein database (https://www.ncbi.nlm.nih.gov/pro tein/). 2. DNAStar Lasergene. 3. Immune epitope database and analysis resource (IEDB) (iedb. org). 1. Peptides conjugated to KLH (Table 1).

2.2 Peptides and Reagents

2. Phosphate-buffered saline (PBS, pH 7.4).

2.3

1. Peptides (Table 1).

ELISA

2. Lyme disease patient sera. 3. ELISA plate (Nunc MaxiSorp).

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Table 1 Amino acid sequence of the peptides synthesized OspA: kvtskdks steekfnekg evs OspC: nsgkdgntsa nsadesvkgp OspE: filigac kihtsydeqs sgesk VlsE: ffvfin cksqvadkdd ptnkfy p100: ykgpydstn tyeqivgige flar

4. Phosphate-buffered saline (PBS; pH 7.4). 5. Blocking buffer: 5% fetal calf serum (FBS) in PBS/0.1% Tween20. 6. Wash buffer: 1
PBS. 7. Goat anti-mouse-HRP. 8. 2,20 -Azinobis [3-ethylbenzothiazoline-6-sulfonic diammonium salt (ABTS) substrate.

acid]-

9. Hydrogen peroxide (H2O2). 10. ELISA plate reader. 2.4 Use of Peptides as a Vaccine

1. Mice (C3H/HeN) (5 weeks old). 2. AddaVax. 3. Syringes and needle. 4. Borrelia burgdorferi cells. 5. Quantitative PCR (qPCR) (thermal cycler). 6. Primers to detect flaB. 7. Sonicator.

2.5

3

Software

1. GraphPad Prism (GraphPad Software, CA).

Methods

3.1 Identification of Antigenic Epitopes and Peptide Synthesis

1. The outer-surface proteins of B. burgdorferi are downloaded from NCBI protein database. The proteins include OspA (Accession No. CAA32579), OspC (Accession No. AFA36430), OspE (Accession No. AAA22959), p100 (Accession No. CAA53022), and VlsE (Accession No. CAH61549). 2. Using DNAStar Lasergene software, we determined the B cell epitopes of the antigenic proteins of Borrelia OspA, OspC, OspE, VlsE, and p100 (Fig. 1) (see Note 1).

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OspA (ACCESSION CAA32579) 1 mkkyllgigl ilaliackqn vssldeknsv svdlpgemkv lvskeknkdg kydliatvdk 61 lelkgtsdkn ngsgvlegvk adkskvklti sddlgqttle vfkedgktlv skkvtskdks 121 steekfnekg evsekiitra dgtrleytgi ksdgsgkake vlkgyvlegt ltaekttlvv 181 kegtvtlskn isksgevsve lndtdssaat kktaawnsgt stltitvnsk ktkdlvftke 241 ntitvqqyds ngtklegsav eitkldeikn alk OspC (ACCESSION AFA36430) 1 mkkntlsail mtlflfiscn nsgkdgntsa nsadesvkgp nlteiskkit esnavvlavk 61 evetllasid evakkaignl iaqnglnaga nqngsllaga yvistliaek ldglknseel 121 kekiedakkc nkaftdklks shaelgiang aatdanakaa ilktngtkdk gaqeleklfe 181 svknlskaaq etlnnsvkel tspvvaespk kp OspE (ACCESSION AAA22959) 1 mnkkmkmfiv yavfiligac kihtsydeqs sgeskvkkie fskftvkikn kdksgnwtdl 61 gdlvvrkeen gidtglnagg hsatffslee evvnnfvkvm teggsfktsl yygykeeqsv 121 ingiqnkeii tkiekidgte yitfsgdkik nsgdkvaeya isleelkknl k p100 (ACCESSION CAA53022) 1 mkeldkeklr dfvnmdlefv nykgpydstn tyeqivgige flarplinsn snsiyygkyf 61 inrfiddqdk kasvdvfsig srsqldsiln lrriltgyli ksfdyerssa eliakvitih 121 navyrgdlny ykevyieaal ksltkenagl srvysqwagk tqifiplkkn ilsgkvesdi 181 didslvtdkv vaallsenea gvnfarditd iqgethkadq dkidieldnv hksdsnitet 241 ienlrdqlek atdeehrkei esqvdakkkq keeldkkaid ldkaqqklds sednldiqrd 301 tvrekiqedi deinkeknlp kpgdvsspkv dkqlqikesl edlqeqlket sdenqkreie 361 kqieikksde ellkskdpka ldlngdlnsk vsskekikgk egeivkeesk asladlnnde 421 nlmrpedqkl sedkkldskk nlkpvseier vneisksnnn eisessplyk psysdmdske 481 gidnkdvnlq etksqtksqp tslnqdlttm sidssnpvfl evidpitnlg tlqlidlntg 541 vrlkestqqg iqrygiyere kdlvvikmds gkaklqilnk lenlkvises nfeinknssl 601 yvdskmilvv vkdsgnvwrl akfspknlne filsenkilp ftsfsvrknf iylqdefksl 661 itldlntlkk vk VLSE (ACCESSION CAH61549) 1 mntkkissai llttffvfin cksqvadkdd ptnkfyqsvi qlgngfldvf tsfgglvaea 61 fgfksdpkks dvktyfttva aklektktdl nslpkeksdi ssttgkpdst gsvgtavega 121 ikevselldk lvkavktaeg assgtaaige vvdnaaaaka adkdsvtgia kgikeiveaa 181 ggskklkaaa akgennkgag klfgkagdaa hgdseaaska agavsavsge qilsaivkaa 241 aagdqegkkp geaknpiaaa igegdgdaef nqdgmkkddq iaaaialrgm akdgkfavkn 301 dekgkaegai kgaaesavrk vlgaitglig davssglrkv gdsvkaaske tppalnk

Fig. 1 The Borrelia burgdorferi proteins and its epitopes for the development of diagnostics and structurebased vaccines. The B cell epitopes of the antigenic proteins of Borrelia OspA, OspC, OspE, VlsE, and p100 are shaded in yellow. The most promiscuous and high affinity binders to MHC-II allele HLA-DRB1 are shown underlined in black. The high affinity MHC-II binders are the most probable T-cell epitopes. The regions are selected based on ranking

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3. To determine the T-cell epitope, we chose alleles (HLA-DRB1) that bind to MHC-II (Fig. 1) (see Note 2). 4. The epitopes are ranked and peptides synthesized based on ranking. The peptides are conjugated to keyhole limpet hemocyanin (KLH) (see Note 3). 5. The peptides could be used in diagnostic applications as well as to determine the immunogenicity. 3.2 Isolation of Serum from Patients

1. Collect blood from patients with Lyme disease and healthy subjects. 2. After collection of the whole blood, allow the blood to clot by leaving it undisturbed at room temperature. This usually takes around 1 h. Remove the clot by centrifuging at 2000
g for 10 min in a refrigerated centrifuge. 3. The pale-yellow supernatant liquid is the serum. Collect the serum using a Pasteur pipette or micropipette. Store at 20  C (for long-term storage) or refrigerate for short-term storage. Thaw to room temperature before use.

3.3 Peptides for Diagnostic Applications

1. To determine whether the peptides could be used in diagnostic applications, coat ELISA plate (MaxiSorp) with peptides (in Table 1). The peptides are diluted in PBS (2 μg/mL) and incubated in the ELISA plates for 2 days (4  C). The working volume of each well is 100 μL. 2. The coated ELISA plate is left at room temperature (RT) for 10 min, washed in PBS, and blocked in blocking buffer for 30 min at room temperature (see Note 4). 3. Remove blocking buffer. The sera (of patients and healthy subjects) is diluted in blocking buffer (see Note 5) and incubated in the ELISA plate for 1 h at RT. 4. Wash with washing buffer (3
) (see Note 6). 5. Incubate with Goat antihuman HRP for 1 h at RT (dilute with blocking buffer). 6. Wash with washing buffer (3
). 7. Prepare ABTS substrate by adding 1 μL 30% H2O2 to 1 mL ABTS substrate. Use immediately. Perform the step under diffused lighting. Cover the plate immediately. 8. Read the ELISA plate at 410 nm (see Note 7).

3.4 Use of Peptides as a Vaccine to Protect Against B. burgdorferi

1. Immunize C3H/HeN mice with 50 μg (each) of the peptides in the presence of the adjuvant (AddaVax) (0.2 mL) (subcutaneous immunization). 2. The B. burgdorferi bacteria are subjected to sonication at 40 kHz, 1 min (repeat 5 times) (SONIX vaccine) (see Note 8).

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3. The mice should be immunized two doses of the peptides 2 weeks apart. 4. Two weeks after the second immunization the mice are challenged with 105 B. burgdorferi cells per mouse. 5. Two weeks postinfection blood is collected to determine antibody titers (see Note 9). 6. The mice are euthanized and tissues (heart and joints) are collected to evaluate bacterial burden. 7. The bacterial burden is quantitated by qPCR by determining the flagellin gene flaB [14, 15] (see Note 10).

4

Notes 1. To determine a protein sequence for potential antigenic epitopes, sequences that are hydrophilic, surface-oriented, and flexible are selected. Most naturally occurring proteins in aqueous solutions have their hydrophilic residues on the protein surface and hydrophobic residues buried in the interior. We selected protein sequences that had good hydrophilicity as predicted by the Lasergene software (DNAStar, WI, USA). 2. Lyme arthritis patients in whom the arthritis resolves within 3 months of the infection show an increased frequency of the HLA–DRB1. 3. The KLH is conjugated to the peptides through cysteine in C terminal. Peptides per se does not induce antibodies; hence, they are conjugated to high molecular KLH (from a marine mollusk). 4. Cover ELISA plate with aluminum foil or plastic wrap to prevent vaporization of reagents. 5. The blocking buffer is used in the dilution of sera or other unconjugated or conjugated antibodies. 6. For washing buffer double distilled water, PBS, or PBS/0.01% Tween 20 can be used. If using PBS/0.01% Tween 20, make sure no air bubbles are present in the wells of the ELISA plate after washing. 7. The peptides bound to the antibodies from Lyme disease patient sera, not in healthy sera controls (Fig. 2). The graphs are plotted, and statistics analyzed by GraphPad Prism. 8. Sonicate on ice to prevent heating of samples. There should be an interval of 2 min on ice between sonication. Sonicate in 5 mL. tubes to prevent aerosol and splashing of samples. A sonication for 5 min is sufficient to lyse all the bacteria. The sonicate is different from heat attenuated vaccines. Increase in

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Fig. 2 Detection of Borrelia specific antibodies in blood samples (positive: known patients) (negative: healthy controls). **p < 0.01 as determined by Student t test

temperature during heat inactivation can degrade some proteins and nucleic acids. Sonication does not destroy proteins and nucleic acids including DNA or RNA if performed in a short time. The DNA and RNA are fragmented into small sizes on sonication. 9. To determine the titer of the peptides, follow Subheading 3.3. Briefly, the peptides are coated in the ELISA plates for 2 days. The excess peptides are discarded, washed, and blocked with blocking buffer. The plates are incubated with the mouse antibodies (diluted in blocking buffer) for 1 h at RT. After washing, they are further incubated with goat anti-mouse HRP for 1 h at RT. Finally, they are treated with the ABTS substrate and the results read in an ELISA plate reader at 410 nm (Fig. 3). 10. The B. burgdorferi bacterial load is determined by qPCR based on the copy number of flaB gene. The naı¨ve mice (unvaccinated) had the highest bacterial load in the heart and joint tissues. Mice immunized with OspC and VLSE had a significant reduction in the bacterial load in the skin, heart, and joint demonstrating that the peptides provided protection against Borrelia. Mice immunized with OspA and OspE had significant lower levels of bacterial burden in the joint compared to the heart. However, the peptides of p100 provided no significant protection against Borrelia (Figs. 4 and 5). The SONIX vaccine also provided protection against the pathogen. The study demonstrated that structure-based vaccine provides significant protection against the pathogenic Borrelia and may be considered vaccine candidates for Lyme disease.

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Fig. 3 The antigenic peptides of Borrelia reacted with peptide-specific antibody before and after challenge with the pathogen

Fig. 4 C3H/HeN mice were immunized at day 0 and 14 and were infected at 4 weeks post-priming with 105 Borrelia cells per mouse. Two weeks postinfection mice were euthanized and tissues were collected to evaluate bacterial burden. Bacterial load (Borrelia) in the skin of mice vaccinated with the antigenic epitopes followed by challenge with Borrelia burgdorferi. *p < 0.05, **p < 0.001 as determined by the Student t test

This is a preliminary study to identify the epitopes of the B. burgdorferi antigenic proteins to provide protection against the pathogen. Additional studies are required to determine the efficacy of the antigenic proteins in providing protection. Alternate vaccination strategies to provide protect against B. burgdorferi include use of adenovirus vector-based vaccines or mRNA-based vaccine.

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Fig. 5 C3H/HeN mice were immunized at day 0 and 14 and were infected at 4 weeks post-priming with 105 Borrelia cells per mouse. Two weeks postinfection mice were euthanized and tissues (heart and joint) were collected to evaluate bacterial burden. *p < 0.05 as determined by the Student t test

Acknowledgements Sincere thanks to Maria Esteve-Gassent, Texas A&M University for technical support. References 1. Steere AC, Strle F, Wormser GP, Hu LT, Branda JA, Hovius JW, Li X, Mead PS (2016) Lyme borreliosis. Nat Rev Dis Primers 2:16090 2. Centers for Disease Control and Prevention (2021) Lyme disease. https://www.cdc.gov/ lyme/stats/humancases.html 3. Shapiro ED (2014) Clinical practice. Lyme disease. N Engl J Med 370:1724–1731 4. Kullberg BJ, Vrijmoeth HD, van de Schoor F, Hovius JW (2020) Lyme borreliosis: diagnosis and management. BMJ 369:m1041 5. Steere AC (2001) Lyme disease. N Engl J Med 345:115–125 6. Roberts ED, Bohm RP Jr, Lowrie RC Jr, Habicht G, Katona L, Piesman J, Philipp MT (1998) Pathogenesis of Lyme neuroborreliosis in the rhesus monkey: the early disseminated and chronic phases of disease in the peripheral nervous system. J Infect Dis 178:722–732 7. Logigian EL, Kaplan RF, Steere AC (1990) Chronic neurologic manifestations of Lyme disease. N Engl J Med 323:1438–1444

8. Bratton RL, Whiteside JW, Hovan MJ, Engle RL, Edwards FD (2008) Diagnosis and treatment of Lyme disease. Mayo Clin Proc 83:566–571 9. Burgdorfer W, Barbour AG, Hayes SF, Benach JL, Grunwaldt E, Davis JP (1982) Lyme disease—a tick-borne spirochetosis? Science 216:1317–1319 10. Elbaum-Garfinkle S (2011) Close to home: a history of Yale and Lyme disease. Yale J Biol Med 84:103–108 11. Kenedy MR, Lenhart TR, Akins DR (2012) The role of Borrelia burgdorferi outer surface proteins. FEMS Immunol Med Microbiol 66:1–19 12. Fraser C, Casjens S, Huang W et al (1997) Genomic sequence of a Lyme disease spirochaete, Borrelia burgdorferi. Nature 390:580–586 13. Iliopoulou BP, Guerau-de-Arellano M, Huber BT (2009) HLA-DR alleles determine responsiveness to Borrelia burgdorferi antigens in a mouse model of self-perpetuating arthritis. Arthritis Rheum 60:3831–3840

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14. Barbour AG, Maupin GO, Teltow GJ, Carter CJ, Piesman J (1996) Identification of an uncultivable Borrelia species in the hard tick Amblyomma americanum: possible agent of a Lyme disease-like illness. J Infect Dis 173:403–409

˜oz15. Sa´nchez RST, Santodomingo AM, Mun Leal S, Silva-de la Fuente MC, Llanos-Soto S, ˜ a D (2020) Rodents Salas LM, Gonza´lez-Acun as potential reservoirs for Borrelia spp. in northern Chile. Rev Bras Parasitol Vet 29(2): e000120

Chapter 22 Development of a SONIX Vaccine to Protect Against Ehrlichiosis Sunil Thomas Abstract The obligately Gram-negative intracellular bacterium Ehrlichia that resides in mononuclear phagocytes is the etiologic agent of human monocytotropic ehrlichiosis (HME). HME is an emerging and often lifethreatening, tick-transmitted infectious disease in the USA. Currently, three different Ehrlichia species can cause ehrlichiosis in humans in the USA—Ehrlichia chaffeensis, Ehrlichia ewingii, and Ehrlichia muris subspecies eauclairensis. Ehrlichia also causes diseases in companion animals and domesticated ruminants. Ehrlichia are vector-borne diseases and transmitted by tick bites. As yet there are no commercially available vaccines to protect against these pathogens. Previously we developed structure-based vaccines and subunit vaccines to protect against ehrlichiosis in animal models. Though the vaccines are efficient in inducing protection, there is a delay in clearing the pathogens in challenge studies. In this chapter we demonstrate the development of a SONIX vaccine that is more potent than conventional vaccines. The vaccination strategy may be useful in Emergency Use Authorization (EUA) scenarios during public health emergencies. Key words Ehrlichia, SONIX vaccine, Bacteria, Ehrlichiosis

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Introduction The Gram-negative, obligate intracellular bacterium Ehrlichia that resides in mononuclear phagocytes is the etiologic agent of human monocytotropic ehrlichiosis (HME). HME is an emerging and often life-threatening zoonotic, tick-transmitted infectious disease in the USA. Lack of early diagnosis and treatment of HME are the main factors that lead to severe and fatal disease. Currently, three different Ehrlichia species can cause ehrlichiosis in humans in the USA—Ehrlichia chaffeensis, Ehrlichia ewingii, and Ehrlichia muris subspecies eauclairensis. Ehrlichia also causes diseases in companion animals and domesticated ruminants. E. chaffeensis and Ehrlichia canis cause canine ehrlichioses in dogs, whereas Ehrlichia ruminantium causes heartwater in cattle, sheep, and goats. As yet

Sunil Thomas (ed.), Vaccine Design: Methods and Protocols, Volume 1: Vaccines for Human Diseases, Methods in Molecular Biology, vol. 2410, https://doi.org/10.1007/978-1-0716-1884-4_22, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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there are no commercially available vaccines to protect against these pathogens [1, 2]. Ehrlichia are vector-borne diseases and transmitted by tick bites. The major factors associated with the emergence or re-emergence of vector-borne diseases include climate change, including global warming and shorter winter season that impact the development and activity of ticks, outdoor activities, global traveling, urbanization, changes in land use, deforestation, habitat fragmentation, natural environment encroachment, which together predispose to a higher contact among wildlife, humans, and domestic animals; the employment and easier access to molecular tools, favoring the diagnosis and identification of vector-borne agents; and the increase of awareness of tick-borne agents by veterinarians, physicians, scientists, and public health authorities [2, 3]. Amblyomma americanum (Lone Star tick) is the most frequently encountered tick species, likely responsible for a majority of tick bites. It is the primary vector and amplifying reservoir of Ehrlichia species [4]. The laboratory strain, Ehrlichia muris was originally isolated from a mouse in Japan in 1983. In 2009, an E. muris-like agent (EMLA) (currently named Ehrlichia muris subspecies eauclairensis) was identified as a causative agent of human ehrlichiosis. A retroanalysis of 760 Ixodes scapularis ticks collected from 1992 through 1997 in Wisconsin revealed an EMLA infection rate of 0.94%, indicating presence of this pathogen in the upper midwestern region since at least the mid-1990s. E. muris is thought to be transmitted by Haemaphysalis flava ticks in Japan, by I. persulcatus ticks in Eastern Europe, and by I. ricinus ticks in Western Europe [5]. Canine monocytic ehrlichiosis (CME) is a cosmopolitan disease of dogs that is caused by Ehrlichia canis. Rhipicephalus sanguineus, the “brown dog tick,” is considered the primary vector of E. canis [6]. Ehrlichia ruminantium causes serious economic losses to cattle and goat farmers in Africa and the Caribbean and is recognized as an agricultural biothreat. Amblyomma variegatum is the tick vector of this pathogen [7]. We previously developed structure-based vaccines [1] and subunit vaccines [8] to protect against ehrlichiosis in animal models. Though the vaccines are efficient in inducing protection, there is a delay in clearing the pathogens in challenge studies. Vaccine development is a long, ardent, and complex process involving several regulatory agencies and often takes around 10–15 years to reach the clinic. An Emergency Use Authorization (EUA) is a mechanism to facilitate the availability and use of medical countermeasures, including vaccines, during public health emergencies, such as the current COVID-19 pandemic. Under an EUA, FDA may allow the use of unapproved medical products, or unapproved uses of approved medical products in an emergency to diagnose, treat, or prevent serious or life-threatening diseases or conditions when

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certain statutory criteria have been met, including that there are no adequate, approved, and available alternatives. Taking into consideration input from the FDA, manufacturers decide whether and when to submit a EUA request to FDA [9]. We have developed a SONIX vaccine to protect against Ehrlichia infection [10]. We tested the vaccine in an animal model of ehrlichiosis. This paper provides a protocol for the development of SONIX vaccine. The SONIX vaccine is more potent than conventional vaccines.

2 2.1

Materials Cell Culture

1. DH82 cells. 2. Dulbecco’s Modified Eagle medium (DMEM). 3. Fetal bovine serum. 4. Ehrlichia muris. 5. T 75 flask. 6. Diff-Quik stain. 7. Light microscope. 8. Glass slides. 9. Sterile plastic loop. 10. Sonicator.

2.2 Animals and Consumables

1. C57BL/6 mice (male and female) (5 weeks old).

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1. Real-time PCR detection system (Bio-Rad, CA).

Quantitative PCR

2. Syringe (3 mL) and needle (preferably 27G).

2. Real-time PCR reagents (Bio-Rad, CA). 3. Primers for disulfide bond formation protein gene (dsb). 2.4

ELISA

1. Peptides (Ehrlichia Hsp60, P28–19). 2. Ehrlichia muris specific antibodies. 3. ELISA plate (Nunc MaxiSorp). 4. Phosphate-buffered saline (PBS; pH 7.4). 5. Blocking buffer: 5% fetal calf serum (FBS) in PBS/0.1% Tween20. 6. Wash buffer: 1 PBS. 7. Goat anti-mouse-AP. 8. Blue Phos™ phosphatase substrate (Kirkegaard and Perry Laboratories, MD). 9. ELISA plate reader.

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Software

1. GraphPad Prism (GraphPad Software, CA).

Methods 1. Culture DH82 cells in T 75 flask. The cells are grown in DMEM. Incubate cells at 37  C and 5% CO2. When the cells are 50% confluent, add E. muris infected DH82 stock (see Note 1). 2. Observe the infected cells by placing a loopful of cells on a glass slide 1 week after culture. The percentage of infectivity is determined by Diff-Quik staining. Generally, 90% of the cells are infected within a week after culture. 3. Once the infection is 90%, the DH82 cells infected with E. muris are collected and centrifuged at 200  g for 10 min. The supernatant is removed and washed twice with PBS. The cells are suspended in 5.0 mL buffer. 4. The DH82 cells infected with E. muris are subjected to sonication at 40 kHz, 1 min (repeat 5 times) (see Note 2). 5. After sonication of the bacteria the resulting product, SONIX vaccine could be directly available for use as a vaccine. No further additions or purification of the resulting material is necessary. The sonicated vaccine can be injected as is, or for convenience of administration can be added to a pharmaceutically acceptable adjuvant (eg. AddaVax). 6. Measure protein concentration (Bradford or Lowry methods). Inject 50 μg per mouse (subcutaneous or intramuscular), two doses, 2 weeks apart. 7. Two weeks after the second dose. Challenge with E. muris intraperitoneally (i.p.) with a high dose of E. muris (1  104 bacterial genomes) and observe daily (see Note 3). Controls included unchallenged naı¨ve mice as well as unvaccinated mice injected with E. muris alone. 8. Mice are sacrificed on days 7 and 14 after ehrlichial challenge, and spleen and liver harvested and sera collected. 9. The ehrlichial load in spleen and liver is determined by quantitative RT-PCR [1] (see Note 4). 10. ELISA is performed to measure the concentration of E. muris specific antibodies. The ELISA plates are coated with 50 μL of peptide (Ehrlichia Hsp60 43–63) or recombinant P28–19 protein at a concentration of 2 μg/mL in PBS. Serum samples are diluted 1∶100, and 100 μL of each sample is added to peptide-coated wells and incubated at 25  C for 1 h. Alkaline phosphatase-conjugated goat anti-mouse antibodies are added at a dilution of 1∶300, and color is developed using

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BluePhos™ phosphatase. Optical densities are measured using an ELISA plate reader at 650 nm after 30 min. Incubation at room temperature. All assays are performed in triplicate wells, and the average values are calculated.

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Notes 1. Our purpose is to culture the Gram-negative intracellular bacteria E. muris. Do not add antibiotics to the medium. 2. Sonicate on ice to prevent heating of samples. There should be an interval of 2 min on ice between sonication. Sonicate in 5 mL. tubes to prevent aerosol and splashing of samples. A sonication for 5 min is sufficient to lyse all the bacteria. The sonicate is different from heat attenuated vaccines. Increase in temperature during heat inactivation can degrade some proteins and nucleic acids. Sonication does not destroy proteins and nucleic acids including DNA or RNA if performed in a short time. The DNA and RNA are fragmented into small sizes on sonication. 3. Controls included unchallenged naı¨ve mice as well as unvaccinated mice injected with E. muris alone. 4. Mice immunized with the sonicated lysate had no bacteria in the spleen and liver after E. muris challenge as determined by quantitative real-time PCR (Fig. 1). Prior experience with

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Fig. 1 SONIX vaccines provide significant protection against E. muris challenge. (a) The SONIX vaccine reduced E. muris on day 7 after bacterial challenge. (b) The SONIX vaccine reduced E. muris on day 14 after bacterial challenge. *P < 0.05, **P < 0.01 as determined by students T-test

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Ehrlichia vaccines had demonstrated that vaccinated mice produced high levels of E. muris specific antibodies on day 14 after challenge. To determine whether antibody is responsible for clearance of Ehrlichia we performed a sandwich ELISA to probe for Ehrlichia specific antibody. Mice immunized with sonicated E. muris and later challenged with the pathogen had low antibody response, whereas mice vaccinated with recombinant p28–19 or infected with E. muris 2 months prior to challenge produced high levels of antibody (Fig. 2). References 1. Thomas S, Thirumalapura NR, CrocquetValdes PA, Luxon BA, Walker DH (2011) Structure-based vaccines provide protection in a mouse model of Ehrlichiosis. PLoS One 6 (11):e27981 2. Thomas S (2016) Development of vaccines for Ehrlichiosis. In: Thomas S (ed) Rickettsiales. Biology, molecular biology, epidemiology, and vaccine development, pp 177–196 3. Andre´ MR (2018) Diversity of Anaplasma and Ehrlichia/Neoehrlichia agents in terrestrial wild carnivores worldwide: implications for human and domestic animal health and wildlife conservation. Front Vet Sci 5:293

4. Anderson BE, Sims KG, Olson JG, Childs JE, Piesman JF, Happ CM, et al (1993) Amblyomma americanum: a potential vector of human ehrlichiosis. Am J Trop Med Hyg 49:239–244 5. Xu G, Pearson P, Rich SM (2018) Ehrlichia muris in Ixodes cookei ticks, northeastern United States, 2016–2017. Emerg Infect Dis 24:1143–1144 6. Schaefer JJ, Needham GR, Bremer WG, Rikihisa Y, Ewing SA, Stich RW (2007) Tick acquisition of Ehrlichia canis from dogs treated with doxycycline hyclate. Antimicrob Agents Chemother 51:3394–3396

SONIX Vaccine to Protect Against Ehrlichiosis 7. Esemu SN, Besong WO, Ndip RN, Ndip LM (2013) Prevalence of Ehrlichia ruminantium in adult Amblyomma variegatum collected from cattle in Cameroon. Exp Appl Acarol 59:377–387 8. Crocquet-Valdes PA, Thirumalapura NR, Ismail N, Yu X, Saito TB, Stevenson HL, Pietzsch CA, Thomas S, Walker DH (2011) Immunization with Ehrlichia P28 outer membrane proteins confers protection in a mouse

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model of ehrlichiosis. Clin Vaccine Immunol 18(12):2018–2025 9. US Food and Drug Administration (2020) Emergency use authorization for vaccines explained. https://www.fda.gov/vaccinesblood-biologics/vaccines/emergency-useauthorization-vaccines-explained 10. Thomas S, Walker DH (2019). Vaccine to protect against Ehrlichia infection (US 10,335,476 B2) (US patent)

Part V Vaccines for Human Parasitic Diseases

Chapter 23 Development of the Antileishmanial Vaccine Sunil Kumar, Shubhranshu Zutshi, Mukesh Kumar Jha, Prashant Chauhan, and Bhaskar Saha Abstract Search for an efficacious antileishmanial vaccine has led to clinical trials of numerous vaccine candidates in the past few decades. As no promising candidate has emerged from these studies, novel vaccine modalities and vaccine assessment techniques are still emerging for antileishmanial vaccine development. Briefly, this chapter discusses: (a) history and timeline of antileishmanial vaccine development; (b) techniques utilized for developing whole-parasite and subunit-based antileishmanial vaccine formulations, and (c) immunogenicity and post-challenge protective efficacy assessment of vaccine candidates. Key words Antileishmanial vaccine, Electroporation, Ni-NTA affinity chromatography, Mass spectrometry, Immunogenicity, Parasite load, T-helper response, Flow cytometry

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Introduction Leishmaniasis is a complex spectrum of vector-borne protozoan disease transmitted through the bite of the female sandfly, viz. Phlebotomus sp. and Lutzomyia sp. in the old and new world, respectively. Leishmania lives as round, nonmotile amastigotes (3–7 μm in diameter) inside macrophages, whereas it survives in a flagellated form known as promastigotes in the sandfly vector [1, 2]. It is considered a neglected tropical disease with an estimated 12 million cases existing worldwide. About 1.5 to two million new cases are occurring annually, putting 350 million people at high risk across five continents [3]. Chemotherapeutic treatment options are limited; drugs are associated with a high risk of side effects, toxicity, and emerging resistance remains a major con-

Sunil Kumar and Shubhranshu Zutshi have contributed equally to this work. Sunil Thomas (ed.), Vaccine Design: Methods and Protocols, Volume 1: Vaccines for Human Diseases, Methods in Molecular Biology, vol. 2410, https://doi.org/10.1007/978-1-0716-1884-4_23, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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Fig. 1 A schematic flow chart representing the characterization of virulent Leishmania antigen from a pool of antigen by numerous techniques to narrow down the most potential immunogenic vaccine candidates for immunization study in mouse model

cern in endemic areas. A prophylactic preventive measure such as a vaccine could be cost-effective, safe, and protective for a long duration. Herein, we briefly describe a novel vaccination protocol as shown in Fig. 1.

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1.1 Timeline of Antileishmanial Vaccine Development

The basic principle of vaccination is to induce the immune system by mimicking the natural course of the disease, albeit noninfectious, thereby generating immunological memory against subsequent encounters. However, preclinical and clinical trials in the hope of generating an effective antileishmanial vaccine have met with little or no success. The next section briefly describes various vaccination modalities employed for antileishmanial vaccine development.

1.1.1 First-Generation Vaccine

It was observed that individuals recovered from leishmaniasis became resistant to subsequent infection from the same pathogen. This observation raised hope for successful antileishmanial vaccination. Predating antiquity, tribal communities in the Middle-East exposed their babies buttock to be bitten by sandflies to remove scars. This technique, known as leishmanization (LZ), was 100% efficient in the former Soviet Union, Middle-East, and Israel [4, 5]. After the preliminary success of LZ, it was discontinued due to safety and difficulty in the standardization of the inoculum, individuals developing nonhealing lesion, and immunosuppression. Drawbacks associated with the use of virulent parasites had shifted focus toward the use of killed Leishmania strains utilizing autoclaving and irradiation. Although it was proved to be effective in field studies, a decline in potency over longer periods was observed [6, 7]. Killed parasite vaccine preparations were tested in different endemic foci containing population susceptible to different Leishmania species and seemed to be more successful as an accessory to chemotherapeutic treatment than as a prophylactic vaccine alone. Although economical and easy to prepare, difficulty in standardizing Leishmania cultures and inconsistent results with respect to efficacy have made this an unpopular vaccination strategy. Failure of killed parasites in eliciting a protective immune response has led to the employment of live but attenuated parasites. Single geneattenuated parasites were designed and few promising results were obtained in mouse trials, but drawbacks associated with the risk of regaining virulence turned this strategy non-viable [8–11].

1.1.2 Second Generation Vaccine

With the advent of biotechnology and an increase in knowledge about the genomics and proteomics of Leishmania, various antigenic proteins, expressed in either promastigotes or amastigotes, were identified. Thus, many antigenic proteins of Leishmania, either in native or recombinant form, were tried as vaccine candidates against leishmaniasis. Fucose-Mannose Ligand (FML) was administered in the native form with different adjuvants, and it is still used as a licensed vaccine against canine leishmaniasis [12– 14]. Many proteins such as gp63, dp72, GP’s, LD1, A2 were tested in animal models, but the majority of them did not yield satisfactory results [15–19]. Leish-111f, a fusion protein-based vaccine, adjuvanted with MPL has been tested against L. major and L. infantum

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infection [20, 21]. After initial success in mice and hamster models, it successfully completed phase I and phase II trials. Of late, the addition of sandfly salivary components, such as maxadilan and LJM11, in vaccine preparation showed promise in animal models of leishmaniasis [22–24]. 1.1.3 Third Generation Vaccine

Third generation vaccine consists of nucleic acid in which a plasmid DNA vector encoding the gene of interest under the control of an eukaryotic promoter is injected directly into the muscle. Although the uptake efficiency and translation of the encoded protein by the myocytes are quite low, it is adequate to evoke cellular immunity against the antigen in various animal models [25]. An added advantage of DNA vaccination is the induction of cytolytic T-cell (CD8+ T-cell) responses possibly due to the cross-presentation of antigen by professional APC which may engulf apoptotic cells, represent free peptides or peptides complexed with heat shock proteins. This is particularly important in the case of Leishmania as CD8+ T-cells have been shown to play a protective role in antileishmanial immunity [26]. LACK (Leishmania homolog of receptors for activated C kinase) DNA vaccine showed protection against CL, but not VL [27, 28]. Candidates such as ORFF, KMP-11 exhibited partial protective response in different preclinical studies [29–32]. The satisfactory result was achieved with multivalent DNA vaccine preparation incorporating NH36 and was found to elicit an immunoprotective profile [33]. A multigenic DNA vaccine known as LEIS HDNAVAX gave encouraging results in initial studies [34]. Moreover, a recent technique employing DNA vaccination is a heterologous prime-boost in which plasmid DNA and purified protein are used for priming and boosting, respectively. LACK and cysteine proteases have been used in this form against experimental visceral leishmaniasis in mice as well in dogs by inducing TH1 type cytokines associated with reduced clinical pathology in visceral organs [35–37]. Despite various preclinical studies, a successful vaccine is yet to emerge victorious from clinical trials in humans.

1.2 Importance of Vaccine Candidate and Strategy Selection

Leishmania sp. contains approximately 9000 genes that encode for proteins. Rationale-based selection of a pathogens’ protein as a vaccine candidate is an important factor in designing an effective vaccine. However, more often than not, it is overlooked in many vaccination studies. Immune system milieus are evidently different during antigenic priming and natural infection. Hence, it may affect the antigenic processing and presentation compartment inside the APCs (mainly macrophages). Also, it may tweak the immunological synapse toward generating pro-parasitic immune signaling [38]. Hence, these factors need to be accounted for during antileishmanial vaccine development as it may significantly affect the outcome of vaccination.

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1.3 Determinants of an Effective Antileishmanial Vaccine

Immune responses, whether host-protective or disease progressing, are guided mainly by the secretion of cytokines by T-cells and APCs. IFN-γ is a major cytokine involved in the killing of intracellular Leishmania. It activates macrophages to produce free radicals and inflammatory cytokines like TNFα and IL-6. It also inhibits the production of IL-10, a major anti-inflammatory cytokine. IL-12, another antileishmanial cytokine, is mainly secreted by macrophages and induces IFN-γ production in T-cells by binding to IL-12R [39, 40]. Major anti-inflammatory or pro-leishmanial cytokine includes IL-10, chiefly produced by CD4+CD25+ Tregs, which alongside TGFβ is mainly responsible for dampening of Th1 and other Leishmania-killing host responses involving IFN-γ, IL-6, and TNFα. Also, IL-4 is a Th2 cytokine that is also responsible for inhibiting IFN-γ mediated host-protective response. It switches the macrophages to the alternative (M2) activation state that promotes antiinflammatory environment and parasite persistence. Moreover, Th17 exhibits dual nature in leishmaniasis progression. On the one hand, it increases hosts’ susceptibility to CL (L. major); on the other hand, it promotes Th1 response in visceral leishmaniasis (Labrus donovani) [39, 41]. Hence, an ideal vaccine should increase the antigen-specific T-cell repertoire and skew the T-cell response toward the host-protective Th1 subtype while inhibiting the expansion of anti-inflammatory Th2, Treg, Th17 subsets (in CL).

1.4 Major Techniques Used in Antileishmanial Vaccine Development

The use of high voltage charge to introduce DNA inside cells, termed as electroporation, was first employed in the 1980s. A linear DNA is introduced into cells by application of electric shocks, which then undergoes homologous recombination to integrate with the cellular genome. This technique is widely used in genetic manipulation studies to generate “knock-in” and “knock-outs” [42]. Genetically modified Leishmania sp. for immunization is also generated using this technique [43].

1.4.1 Electroporation

1.4.2 Ni-NTA Affinity Chromatography

Ni-NTA affinity chromatography is a type of immobilized metalion affinity chromatography (IMAC) that utilizes the capability of histidine amino acid to interact and form coordination bonds with immobilized metal-ion matrices (Ni2+, Co2+, Cu2+, etc.). This technique is widely used for purification of proteins to be utilized in various downstream applications. The target protein is tagged with an array of histidine residues and bound protein is then purified by passing imidazole, which replaces the protein-Ni2+ interaction owing to its high affinity with metal ions. In vaccination studies, this technique is used for purification of the recombinant form of antigenic proteins [44].

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1.4.3 PeritoneumDerived Primary Macrophages

Peritoneal macrophages (PM) are being used as experimental macrophages in mice for a long time. PM express higher levels of inducible nitric oxide synthase, IL-12, and lysosomal contents than macrophages from other sources such as bone marrow and spleen. Moreover, these cells express higher levels of MHC-II and co-stimulatory molecules, for example, CD86, indicating their immunostimulatory potential [45, 46]. Due to these reasons, they are widely used in leishmaniasis studies for mimicking Leishmania infection in vitro.

1.4.4 Mass Spectrometry

High-throughput proteome analysis has become a major tool in cellular analysis. SELDI-TOF is a widely used method in mass spectroscopy. It is quite similar to MALDI-TOF; however, it gives better reproducibility and sensitivity. Here, proteins to be analyzed are crystallized and vaporized by a beam of UV-based laser and ionized. m/z ratios of different proteins are derived from the velocity of charged protein ions in an electric field. Recently, Orbitrap mass spectrometers have been designed for better accuracy and a higher resolution of protein mass data [47, 48]. Proteome profiling of virulent and avirulent strains of pathogens is widely used to identify potential virulence factors [49].

1.4.5 Indirect and Sandwich ELISA

Enzyme-linked immunosorbent assay (ELISA) replaced RIA in the 1970s to become the popular method for measuring the concentration of antigens, as it uses nonradioactive methods. In sandwich ELISA, the antigen is trapped between the surface-coated capture and detection antibodies to the antigen, which is then detected by a chemiluminescent reaction. It is a widely used method for detecting antigen or cytokine concentration in cell culture supernatants or biological fluids [50]. Another variant, indirect ELISA, is used for measuring antigen-specific Ig levels in blood sera. An antibody present in blood serum is captured by a coated antigen, which is again detected by a peroxidase enzyme-based chemiluminescent reaction [49, 51]. ELISAs are the standard methods for analyzing the antigen-specific immune response in vaccination studies, accurately and specifically.

1.4.6 Macrophage T-Cell Co-Culture System

To measure the killing activity or specifically measure the responses of T-cells in the presence of infected host cells, co-culture systems are considered best in vitro models. In antileishmanial vaccine studies, they can explicitly give information about the type of response elicited by antigen-primed T-cells upon encounter by parasitized macrophages. Secretory levels of cytokines and NO can be measured and intracellular parasite load can be quantified simultaneously by this culture system [52, 53].

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1.4.7 In Vivo Parasitic Load

As vaccination is aimed at reducing pathogenic burden, parasite load in the draining lymph node (CL) or spleen and liver (VL) assesses disease severity. Draining lymph node is isolated and the supernatant obtained by crushing the organ is added to the culture medium for the transformation of intracellular amastigotes to promastigotes, which are then enumerated after 5 days.

1.4.8 Flow Cytometry

Flow cytometry is the most popular method for single-cell based immunophenotyping of Th cell subsets. As the antileishmanial immune response is T-cell based, Th subset profiling and functional characterization become essential. Moreover, memory T-cell subsets can be identified by this technique which helps in assessing the robustness of protective immune response elicited by vaccination. Antigen-specific population and cytokine secretion can be read by stimulation using purified antigens or CSA, ionomycin, and Golgi transport inhibitors [54].

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Materials

2.1 Maintenance and In Vitro Passaging of L. major and L. donovani Strain

1. RPMI-1640 medium. 2. Penicillin-streptomycin. 3. β-mercaptoethanol. 4. Fetal bovine serum, USA origin. 5. Hanks’ Balanced Salt Solution.

2.2 Generation of Dominant-Negative Mutant Parasites

1. pIRSAT vector. 2. Escherichia coli DH5-α competent cells. 3. Genomic DNA extraction kit. 4. Gel extraction kit. 5. PCR purification kit. 6. Micropulser electroporation apparatus.

2.3 PCR-Based Molecular Cloning and Sequence Confirmation

1. pcDNA6/HisA (Invitrogen, Carlsbad, CA, USA). 2. pet28a + (Novagen, Merck & Co., Kenilworth, NJ, USA). 3. Restriction endonucleases. 4. Genelute™ PCR purification kit (MilliporeSigma, St. Louis, MO, USA). 5. Genelute™ Gel extraction kit (MilliporeSigma, St. Louis MO, USA). 6. T4 DNA ligase. 7. E. coli DH5-α competent cells. 8. Single-pass DNA sequencing service (SS1001, first Base Laboratories, Sdn. bhd.).

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2.4 Recombinant Protein Purification

1. IPTG. 2. Clontech™ His60 superflow resin (Takara Bio, Mountain View, CA, USA). 3. Sodium phosphate: Mono- and dibasic. 4. Glycerol. 5. Sodium chloride. 6. β-mercaptoethanol. 7. Triton-X 100. 8. Novagen™ Benzonase™ Nuclease (Merck Millipore, Burlington, MA, USA). 9. Lysozyme. 10. Imidazole.

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SELDI

1. Tris-base. 2. NaCl. 3. Triton-X 100. 4. EDTA. 5. Protease inhibitor cocktail tablets. 6. Pierce™ BCA protein assay kit (Pierce Biotechnology, Rockford, IL, USA). 7. Pierce™ BSA protein standard ampoules (Pierce Biotechnology, Rockford, IL, USA). 8. ProteinChip Q10 array (Bio-Rad, Hercules, CA, USA). 9. Sinapinic acid mixture. 10. PCS4000 SELDI-TOF instrument (Bio-Rad, Hercules, CA, USA).

2.6 Mass Spectrometry

1. TCEP. 2. Iodoacetamide. 3. Trypsin. 4. Dionex™ SolEx™ C18 silica-based SPE cartridge (Thermo Fisher Scientific, Waltham, MA, USA). 5. Acetonitrile. 6. Formic acid. 7. EASY-nLC 1000 system (Thermo Fisher Scientific, Waltham, MA, USA). 8. Q Exactive Orbitrap mass spectrometer (Thermo Fisher Scientific, Waltham, MA, USA). 9. PicoFrit™ columns (Thermo Fisher Scientific, Waltham, MA, USA).

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1. Fluid thioglycollate medium. 2. NaCl. 3. KCl. 4. Na2HPO4.7H2O. 5. KH2PO4. 6. Gibco® RPMI-1640 medium. 7. L. major/L. donovani strain. 8. RPMI-1640 medium. 9. Penicillin-streptomycin. 10. Sodium pyruvate. 11. L-Glutamine. 12. β-mercaptoethanol. 13. Fetal bovine serum. 14. Hanks’ Balanced Salt Solution. 15. CO2 incubator.

2.8 Giemsa Staining for Amastigote Count

1. Methanol. 2. Giemsa stain. 3. Nikon™ Eclipse E600 with U-III Film Camera System (Nikon Corporation, Tokyo, Japan).

2.9 Cytokine/ Indirect ELISA

1. 8-well strips, 96-well format, Corning® (Corning Inc., NY, USA). 2. Sodium phosphate dibasic, heptahydrate. 3. NaCl. 4. Potassium orthophosphate. 5. KCl. 6. Bovine serum albumin Fraction V. 7. Tween-20. 8. Sulfuric acid. 9. Purified Rat anti-mouse IL-4/IL-10/IFNγ/IL-12/TNFα. 10. Biotin Rat anti-mouse IL-4/IL-10/IFNγ/IL-12/ TNFα. 11. Streptavidin-POD conjugate. 12. BD OptEIA™ TMB substrate reagent set. 13. SpectraMax® M5 microplate reader (Molecular Devices, LLC, San Jose, CA, USA).

2.10 NO Release Assay

1. Fluid thioglycolate medium. 2. RPMI-1640 medium. 3. L. major/L. donovani strain.

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4. RPMI-1640 medium. 5. Penicillin-streptomycin. 6. Sodium pyruvate. 7. L-Glutamine. 8. β-mercaptoethanol. 9. Fetal bovine serum. 10. Hanks’ Balanced Salt Solution. 11. CO2 incubator. 12. Nunc® Labtek® Chamber Slide ™ system, 8 well Permanox® (Merck). 13. Griess reagent. 14. SpectraMax® M5 microplate reader. 2.11 Macrophage TCell Co-Culture Assay

1. Nunc® Labtek® Chamber Slide ™ system, 8 well Permanox®.

2.12 Antibody/ Sandwich ELISA

1. CSA from Leishmania. 2. Blood serum samples. 3. Sulfuric acid. 4. Bovine serum albumin fraction V. 5. Tween-20. 6. Biotin rat anti-mouse IgG2a/IgG1/IgM. 7. Streptavidin-POD conjugate. 8. BD OptEIA™ TMB substrate reagent set. 9. SpectraMax® M5 microplate reader.

2.13 Western Blotting

1. Acrylamide. 2. N, N0 methylene bis(acrylamide). 3. Sodium dodecyl sulfate. 4. Ammonium persulfate. 5. Tetramethylethylenediamine (TEMED). 6. 1.5 mm glass and spacer plates. 7. Basic power supply. 8. Immobilon-P 0.22 μM PVDF membrane. 9. Tris-base. 10. NaCl. 11. Primary antibody—blood monoclonal IgG1.

serum/anti-his

tag

12. Goat Anti-Mouse IgG (H + L)-HRP Conjugate.

antibody,

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13. ClarityMax™ western ECL substrate. 14. X-ray films. 2.14 In Vivo Challenge Infection or Priming with Leishmania

1. Stationary phase culture of L. major/L. donovani. 2. Hanks’ Balanced Salt Solution. 3. Inverted 40 microscope. 4. Neubauer’s chamber. 5. 32G needle syringe.

2.15 Footpad Measurement for CL Progression

1. Screw gauge.

2.16 In Vivo Parasite Load Assay

1. RPMI-1640 medium. 2. Penicillin-streptomycin. 3. β-mercaptoethanol. 4. Sodium pyruvate. 5. Falcon® 25 cm2 non-vented flask. 6. Frosted-end slides. 7. Bench-top centrifuge.

2.17 Ldu for VL Severity

1. Spleen from different study groups of mice. 2. Microscopic slides. 3. Methanol. 4. Giemsa stain.

2.18

qPCR

1. Hanks’ Balanced Salt Solution. 2. Gene-specific primers. 3. SYBR® premix Ex Taq containing Tli RNaseH (Takara Bio Inc., Kusatsu, Shiga, Japan). 4. 8-well strips/96-well plate. 5. StepOnePlus™ Real-time PCR system (Applied Biosystems Inc., Foster City, CA, USA).

2.19

Flow Cytometry

1. Cell strainer, 60 μM. 2. Fetal Bovine serum. 3. NaCl. 4. Sodium phosphate, dibasic. 5. KCl. 6. Potassium orthophosphate. 7. HEPES, free acid.

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8. Cytofix/Cytoperm-Plus kit with Golgi Plug. 9. (a) FITC Rat anti-mouse-CD3, PB Rat anti-mouse-CD4, PerCPCy5.5 Rat anti-mouse CD44, APCCy7 Rat antimouse-CD62L, APCCy7 Rat anti-mouse-CD25, APC Rat anti-mouse-FOXP3, PE Rat anti-mouse-IFNγ, PE Rat anti mouse-IL4, and PE Rat anti-mouse-IL10. (b) PECy7 Rat anti-mouse/human CD44, PerCPCy5.5 Rat anti-mouse-CD62L, APC Rat anti-mouse-Tbet, APC Rat anti-mouse-IL17, (c) APC Rat anti-mouse-GATA3, PE Rat anti-mouse-RORγT. 10. Isotype controls—FITC mouse IgG1, κ isotype control, PE Rat IgG2a, κ isotype control, PerCP-Cy5.5 mouse IgG1, κ isotype control, PE-Cy7 Hamster IgG1, κ isotype control, APC Rat IgG2b, κ isotype control, APC-Cy7 Rat IgG1, λ isotype control, Pacific Blue Rat IgG2a, κ isotype control.

3

Methods

3.1 Preparation of Vaccine Formulation

1. Isolate draining lymph node (dLN) from infected BALB/cJ mice under sterile conditions.

3.1.1 Generation of In Vitro Culture-Based Avirulent Strain

2. Crush lymph node in RPMI-1640 (supplemented with 100 μg/mL penicillin-streptomycin, 50 μM 2-ME, 1 μg/mL sodium pyruvate). 3. Centrifuge cell suspension at 200  g for 8 min at 25  C. 4. Meanwhile, keep a T25-flask ready with fresh RPMI-1640 and decant the supernatant obtained from step 3 into the flask and screw the cap tightly. 5. Keep the flask at 25  C for 6–8 days for allowing the transformation of parasites (see Note 1). 6. Once transformed into promastigotes, passage culture each time at the stationary phase into a fresh flask at 1:20 ratio of inoculum to the parasite (see Note 2).

3.1.2 Generation of Overexpression Based Avirulent Strain (See Note 3)

1. PCR amplifies the N-terminal segment of LdeK1 (1–840 bp) using primers with BgIII restriction site. 2. Digest pIRSAT1 vector by BgIII and clone the BgIII-digested fragment of LdeK1. 3. Select the positive clones by transforming ligated pIRSAT1 in E. coli DH5α and confirm fragment insertion by restriction digestion. 4. Digest cloned plasmid with SwaI and gel-purify. 5. Meanwhile, pellet mid-log phase Leishmania at 1300 g for 10 min and wash in cytomix electroporation buffer (120 mM

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KCl2, 0.15 mM CaCl2, 10 mM K2HPO4, 25 mM HEPES, 2 mM EDTA, and MgCl2; pH 7.6) in half of original volume. 6. Pellet cells again and resuspend in cytomix buffer at 2  108 cells/mL. 7. Take 10 μg of linear-plasmid from step 4 in a 4-mm cuvette and add 500 μL of cell suspension. 8. Electroporate at 25 μF, 1500 V (3.75 kV/cm) for 2 pulses with a 10-s gap in between. 9. Transfer electroporated cells to appropriate media. 10. Add nourseothricin (50 μg/mL) to culture 24 h postelectroporation and screen single-cell clones by limiting dilution method. 11. Screen positive clones by analyzing the expression of the N-terminal segment of LdeK1through RT-PCR and western blotting. 3.1.3 Preparation of E. coli DH5α and BL21 Competent Cells

1. Streak E. coli DH5α onto a fresh LB agar plate and keep overnight at 37  C. 2. Pick a well-differentiated single bacterial colony and inoculate in 1 mL of LB broth. 3. Subculture by inoculating 1 mL of fully grown culture in 100 mL of LB broth containing 1 mM MgCl2 and allow the culture to grow at 37  C, 225 rpm. 4. Meanwhile, prepare TFB-I (KCl 100 mM; MnCl2 50 mM; CH3COOK 30 mM; CaCl2 10 mM; glycerol 10%; pH 5.8) and TFB-II (KCl 10 mM; CaCl2 75 mM; MOPS 10 mM; glycerol 10%; pH 7) and filter-sterilize the buffers (see Note 4). 5. At OD600 ¼ 0.5, take out the culture and pellet at 2057  g for 10 min at 4  C. 6. Resuspend the culture in 15 mL of ice-cold TFB-I and incubate for 1.5 h with light stirring every 10 min. 7. Pellet the cell suspension again at 2057  g for 10 min at 4  C. 8. Resuspend the cells in 3.5 mL of ice-cold TFB-II. Make 80 μL aliquots and immediately transfer them into the deep freezer (70  C) (see Note 5).

3.1.4 Clone Development

1. PCR amplifies the gene of interest (GOI) by primers carrying the restriction enzyme cut-sites at both ends by a proof-reading DNA polymerase (see Note 6). 2. Purify the PCR product by PCR purification kit. 3. Digest the PCR product and plasmid (1.5 μg) with the same set of restriction enzymes chosen for cloning, as per manufacturer’s instructions (see Note 7).

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4. Run digested products on 0.8% agarose gel and gel-purify. 5. Set up a ligation reaction with T4 DNA ligase using different molar ratios, viz. 1:3, 1:6, 1:12 vector to insert, as per manufacturer’s instructions (see Note 8). 6. Incubate at 16  C overnight (12–16 h) and stop the reaction at 65  C for 10 min. 7. Transform the reaction mixtures in E. coli DH5α competent cells by heat shock method. 8. Pick distinct bacterial colonies after overnight incubation and inoculate in 5 mL of LB broth. 9. When the culture reached confluency, pellet cells and isolate plasmid. 10. Set up a digestion reaction and run the reaction mixture on 1% agarose gel to identify the band corresponding to insert (GOI). 11. After confirmation of successful insertion, confirm the sequence integrity of insert by single-pass DNA sequencing. 3.1.5 Ni-NTA Affinity Chromatography for Protein Purification (See Note 9)

1. Transform pet28a + (or any suitable prokaryotic expression vector) clone in E. coli BL21 (DE3/Rosetta-gami) strain by heat shock method. 2. Pick a distinct bacterial colony after overnight incubation and inoculate in 2 mL of LB broth. 3. After growing overnight at 225 rpm, 37  C with appropriate antibiotics, inoculate 200 μL from well-grown primary culture into fresh flasks (5–8) containing 20 mL LB broth. 4. Grow culture at 225 rpm, 37  C till OD600 ¼ 0.5. Meanwhile, prepare a sterile-filtered IPTG stock solution. 5. Identify the best condition for protein induction by growing culture induced by varying IPTG concentrations (0.1 mM, 0.25 mM, 0.5 mM) at different temperatures (18, 25, and 37  C). 6. Analyze the degree of protein induction by running bacterial lysates on SDS-PAGE. 7. Select the best condition for protein induction and work out the best condition for a good yield of protein by tweaking the components in lysis buffer (see Note 10). A generalized procedure for protein purification (500 mL bacterial culture) is as follows: (a) Resuspend bacterial pellet in 25 mL of lysis buffer containing required components as discussed in Note 10. (b) Add 1 mg/mL lysozyme, 250–300 U of nuclease, and incubate 30 min on ice. Afterward, sonicate the pellet at 85% amplitude, 4 s ON, and 6 s OFF cycle for 20 min.

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(c) Centrifuge the lysate at 18,514  g for 30 min at 4  C. (d) Collect a small fraction of supernatant and cell lysate to determine the presence of expressed protein by SDS-PAGE. (e) Take bead suspension volume equivalent to 350–400 μL of Ni-NTA beads. Wash the beads successively with 20 times the volume of Milli-Q water and lysis buffer (see Note 11). (f) Incubate the beads with supernatant obtained from step 7 (c) at 5 rpm for a defined period (see Note 12). (g) Spin the suspension at 804  g for 5 min and remove the supernatant (unbound fraction). (h) Wash beads with lysis buffer containing gradually increasing concentration of imidazole in successive washes (3 different wash solutions in general). (i) Finally, transfer the beads in a vertical column and allow them to settle down by gravity. (j) Add elution solution (lysis buffer containing a higher imidazole concentration than wash solutions) and collect 1 mL fractions. (k) Run all the collected fractions at different purification steps on an SDS-PAGE and analyze the efficiency of purification and purity of the recombinant protein. 3.2 Characterization of Whole-Parasite Based Vaccine 3.2.1 In Vitro Growth Kinetics of Avirulent Strain

1. Set up Leishmania cultures in vitro as described in Subheading 3.1.1 from a low-passage (LP; preferably less than 10) and a high-passage strain (HP; passage >500). 2. Grow culture at 25  C and take 1 mL inoculum each day for 11 successive days. 3. Quantify the number of promastigotes by counting under a phase-contrast microscope by Neubauer’s chamber.

3.2.2 In Vivo Infectivity of Avirulent Strain

1. Pellet a stationary phase Leishmania culture and wash 2 with 1 mL sterile HBSS. 2. Resuspend pellet in HBSS and count the number of promastigotes. 3. Take a volume equivalent to 2  106 promastigotes/mice (L. major), or, 2  107 promastigotes/mice (L. donovani) in a volume of 50 μL (see Note 13). 4. Inject 2 units (50 μL) of parasite suspension by insulin syringe (31G) subcutaneously in left hind footpad (L. major) or intracardially (L. donovani). 5. Measure footpad thickness successively for 5 weeks by screw gauge and represent the difference in thickness between the left

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and right (non-injected) progression.

footpad,

indicating

disease

6. After 5 weeks of infection, collect dLN of mice and crush it under the frosted end of slides in 3 mL supplemented RPMI1640 (Subheading 3.1.1). 7. Pass it through a cell strainer and make single-cell suspension. 8. Pellet cells and add supernatant to fresh 25 cm2 flask containing 7 mL RPMI-1640. 9. Incubate culture at 25  C and count the number of transformed promastigotes after 5 days of culture. This indicates the parasitic burden in mice. 3.2.3 Isolation of Peritoneum-Derived Primary Macrophages

1. Inject 2 mL of 3% thioglycolate medium (sterile) i.p. 2. On fifth day post-injection, collect peritoneal macrophages by flushing peritoneal cavity with 10 mL of PBS (137 mM NaCl, 5.4 mM Na2HPO4.7H2O, 3.22 mM KCl, 1.47 mM KH2PO4) under sterile conditions (see Note 14). 3. Pellet cells at 209  g for 8 min. 4. Resuspend cells in RPMI-1640 (containing 1 mM sodium pyruvate, 2 mM L-glutamine, 100 U Pen-strep, 6% FBS) and count the cells using Neubauer’s chamber. 5. Plate the cells (according to the experiment’s requirement) and incubate at 37  C, 5%CO2 for 6 h. 6. Wash the cells to remove any non-adhered material and replace the culture with fresh, supplemented RPMI-1640. 7. Keep the culture for 36–48 h, thereafter proceed for the downstream experiment (see Note 15).

3.2.4 In Vitro Infection of Macrophages by Leishmania sp. (See Note 16)

1. Pellet a stationary phase Leishmania culture at 581  g for 10 min at 25  C. 2. Resuspend the cells in supplemented RPMI-1640 (described in Subheading 3.2.3, step 1) and count the number of promastigotes/mL. 3. Infect macrophages with 10 parasite number (i.e., macrophage: parasite ratio—1:10) and incubate at either 33  C for 6 h for L. major strain or 37  C for 6 h for L. donovani strain. 4. After a specified time, remove the media and wash the cells vigorously with HBSS/PBS at least twice (see Note 17). 5. Add fresh supplemented RPMI-1640 and incubate infected cells at 37  C, 5% CO2.

3.2.5 In Vitro IntraMacrophage Survival of Leishmania

1. Isolate peritoneum-derived primary macrophages and infect with virulent and avirulent strain of Leishmania as described in Subheadings 3.2.3 and 3.2.4 in 8-well chamber slides at 5  104 cells/well.

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2. Culture infected macrophages for 72 h. 3. Remove the growth chamber and fix the slides by dipping in ice-cold methanol for 5 min. Let them air-dry overnight. 4. Stain the slides by dipping in a 1:10 dilution of Giemsa stain in distilled water for 1.5 h. 5. Wash the slides in distilled water extensively and air-dry the slides. 6. Enumerate the amastigotes at 100 using immersion oil by Nikon E6000 microscope. 3.2.6 Characterization of Immune Response Elicited by Avirulent Strain Cytokine/Indirect ELISA from the Culture Supernatant

1. Infect macrophages with Leishmania in a 24-well plate (in triplicates) and incubate for 48 h postinfection. 2. Meanwhile, coated ELISA strip wells with 100 μL of IL-10 (2 μg/mL), 50 μL of IL-12 (2 μg/mL), and 100 μL of TNF-α (2 μg/mL) capture antibody overnight at 4  C. 3. Wash wells with PBS, add 200 μL blocking solution (1% BSA), and incubate for 4 h at room temperature (RT). 4. Collect culture supernatant after completion of step 1 and centrifuge at 425  g for 5 min at 4  C. 5. Meanwhile, wash ELISA plate with 200 μL of wash buffer (PBS containing 0.05% Tween-20) and add 100 μL, 50 μL, and 100 μL of supernatant from step 4 for IL-10, IL-12, and TNF-α detection, respectively. Also, add the same volume of serially diluted purified, recombinant IL-10, IL-12, and TNF-α in remaining wells for standard curve plotting. 6. Incubate plates for 16 h at 4  C. 7. Wash plates 2 with wash buffer and add 100 μL (1 μg/mL), 50 μL (1 μg/mL), and 100 μL (1 μg/mL) of IL-10, IL-12, and TNF-α detection antibody (biotinylated), respectively. 8. Incubate for 2 h at RT. 9. Wash plates 3 with wash buffer and add 100 μL of streptavidin-POD dilution (1:20,000) in each well. Incubate for 30 min at RT. 10. Meanwhile, prepare a 1:1 mix of TMB substrate A and B and 1 M H2SO4. 11. Wash plates 3 with wash buffer and add 100 μL of TMB mix/well and allow the color to develop. 12. Stop reaction by adding 50 μL of 1 M H2SO4 and read absorbance at 450 nm.

Cytokine Profiling of Lymphocytes

1. Isolate draining popliteal lymph node after 5 weeks of L. major LP and HP infection. 2. Crush lymph node, pass through 60 μM cell strainer, and make a single-cell suspension.

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3. Plate 0.5  106 lymphocytes/well (in triplicates). 4. Culture the lymphocytes for 48 h, either unstimulated or in presence of 25 μg/mL leishmanial CSA (added in culture medium). 5. Collect culture supernatant and proceed for cytokine estimation by ELISA as described in Subheading 3.2.5, step 1. Macrophage T-Cell CoCulture for Assessing the Antileishmanial Function of T-Cells

1. Plate 5  104 macrophages/well in an 8-well chamber slide. 2. Infect macrophages with 5  105 Leishmania and incubate for 36 h. 3. Add lymphocytes from dLN of LP and HP L. major infected mice in a macrophage: the T-cell ratio of 1:3. 4. Culture for a period of 72 h and collect supernatant for ELISA (Subheading 3.2.7.1). 5. Fix the slide in chilled methanol, stained with Giemsa (Subheading 3.2.5, steps 3–6), and proceed for amastigote count inside macrophages.

3.2.7 Proteome Characterization

1. Harvest 2  108 promastigotes from three individual cultures of each Leishmania strain at 908  g, 10 min at 25  C. Wash 3 with PBS (pH 7.4).

SELDI Analyses of LP and HP Strain of L. major and L. donovani

2. Resuspend pellet in 300 μL of lysis buffer containing 20 mM Tris (pH 7.4), 150 mM NaCl, 0.1% Triton-X 100, 1 mM EDTA, protease inhibitor cocktail. Incubate at 4  C for 1 h. 3. Centrifuge lysed samples at 18,514  g, 1 h at 4  C. 4. Collect supernatant and estimate protein concentration using a BCA-based kit. 5. Take 10 μg of each protein sample and dilute to 50 μL by binding buffer (50 mM Tris-HCl, pH 9). 6. The sample is applied in duplicates to Q10 anion ProteinChip array and incubated for 1 h, 200 rpm. 7. Wash chip 3 with 150 μL binding buffer and 2 with deionized water for 5 min, RT at 200 rpm. 8. Air-dry the spot and 1 μL of the sinapinic acid mixture is applied (see Note 18). 9. After running the SELDI experiment, peaks are automatically detected and normalized to total ion current. Collect mass spectra over a range of 2000–20,000 Da. 10. After nullifying background, select all peaks with signal/noise >5 and valley depth >3 within a cluster mass window of 0.3% of the mass and perform the analyses.

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1. Take 25 μL protein sample and reduce with 5 mM TCEP, follow with alkylation by 50 mM iodoacetamide. 2. Digest protein sample by trypsin (trypsin/lysate ratio 1:20) by incubating at 37  C for 16 h. 3. Desalt digested sample by using C18 silica cartridge and dry the sample using speed vac. 4. Resuspend the pellet in buffer A (5% acetonitrile, 0.1% formic acid). 5. Load and resolve 1 μg peptide mixture with buffer A in a 25 cm PicoFrit column filled with 1.8 μm of C-18 resin (see Note 19). 6. Elute proteins with 0–40% gradient of buffer B (95% acetonitrile, 0.1% formic acid) flowing at 300 nL/min for 100 min. 7. Acquire data using data-dependent top 10 methods. 8. Analyze RAW files against UniProt L. major reference proteome database by SEQUEST search (see Note 20).

3.3 Immunization of Mice: (See Note 21)

Immunization protocol for various vaccination strategies is as follows: 1. DN mutant L. donovani: Intravenous (i.v.) injection of 2  107 promastigotes. 2. HP (avirulent) L. major: s.c. injection of 5  103 L. major promastigotes in the left hind footpad. 3. DNA-based vaccination: i.m. injection of 100 μg DNA vector (control or cloned) in left thigh muscle on 0, 15th, and 30th day. 4. Recombinant protein-based vaccination: i.m. injection of 15 μg of recombinant, purified protein as 1:1 (v:v) o/w-based emulsion with incomplete Freund’s adjuvant (see Note 22).

3.4 Immunogenicity Assessment PostImmunization 3.4.1 Antibody/Sandwich ELISA for Antileishmanial IgG Determination

1. Collect blood by retro-orbital vein puncture from naı¨ve, control, and vaccinated mice after 4 weeks of last vaccination dose. 2. Keep blood at RT for 30 min and centrifuge it afterward at 15,294  g for 30 min at 25  C. 3. Carefully take out the liquid supernatant, leaving behind the clotted portion. 4. On the previous day, coat ELISA plates with 30 μg/mL CSA (or 10 μg/mL purified protein, if available) in PBS overnight at 4  C. 5. Wash plates with PBS and add 200 μL of blocking solution (1% BSA in PBS)/well and incubate at RT for 4 h. 6. Make serial dilutions of each blood serum sample obtained in step 3, viz. 1:100, 1:300, 1:900, 1:2700, and 1:8100 in blocking buffer.

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7. Wash plates from step 5 with 200 μL of wash buffer and add 100 μL of each sample (in duplicate)/well. Incubate the plates for 16 h at 4  C. 8. Wash 3 with 200 μL wash buffer and add 100 μL of antiIgG1, IgG2a (0.25 μg/mL), and anti-IgM (1 μg/mL) in each well. Incubate for 2 h at RT. 9. Wash 3 with wash buffer and add 100 μL of streptavidinPOD conjugate solution (diluted 1:20,000 in wash buffer) in each well. Incubate for 30 min at RT. 10. Wash 3 with wash buffer and add 100 μL of TMB substrate mix (A:B—1:1) in each well. 11. After color develops, stop the reaction by adding 50 μL of 1 M H2SO4/well and read absorbance at 450 nm. 3.4.2 Probing Native Protein with Mouse Sera

1. Lyse 108 stationary phase promastigotes in 500 μL of lysis buffer 20 mM Tris, 0.15 M NaCl, 1 mM EDTA, 1 mM EGTA, 10% glycerol, protease inhibitor added). 2. Centrifuge cell lysate at 19,000  g for 30 min at 4  C. 3. Collect the supernatant and estimate protein concentration by BCA assay kit. 4. Take 20 μg of protein and add a 4 Laemmli sample buffer. Heat at 95  C for 5 min. 5. Run protein samples on an SDS-PAGE at 70 V and transfer the resolved protein bands on to 0.22 μM PVDF membrane by western transfer (see Note 23). 6. Incubate the PVDF membrane with blood sera dilution (1:10,000) prepared in TBS (1.37 M NaCl, 0.2 M Tris) containing 1% BSA overnight at 4  C. 7. Wash blot 3 with wash buffer (TBS containing 0.05% tween20) and incubate with horseradish peroxidase-conjugated antimouse IgG (1:5000) for 1.5 h at RT. 8. Wash blot 3 with wash buffer and develop the blot by the chemiluminescent reagent.

3.5 Protection Studies

1. Harvest stationary phase promastigotes from a LP Leishmania culture at 581  g, 10 min, 25  C.

3.5.1 In Vivo Challenge Infection

2. Wash the pellet two times in 1 mL of sterile HBSS. 3. Resuspend the pellet in HBSS and count the number of promastigotes. 4. Adjust final suspension to 4  107 promastigotes/mL for L. major, and 4  108 promastigotes/mL for L. donovani challenge, respectively.

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5. After 4 weeks of last vaccination dose or 60 days after HP parasite priming, inject 50 μL of L. major suspension s.c. in left hind footpad for cutaneous disease and 50 μL of L. donovani suspension i.c. for the visceral disease. 3.5.2 Footpad Thickness Measurement for Assessing CL

1. Tare the perfectly closed screw gauge to 0.00 mm. 2. Unscrew the gauge and take measurement of an injected hind footpad (left) and the contralateral footpad (see Note 24). 3. Starting from first week, measure the thickness successively for 5 weeks post-challenge infection for mice in control and vaccinated groups.

3.5.3 Ldu in Affected Organs for Assessing VL

1. Isolate spleen and liver after euthanizing the mice. 2. Cut transversely through the center of the organ by a clean scalpel blade. 3. On a clear microscopic slide, make multiple impression smears from each organ collected from the mice. 4. Air-dry the slides overnight. 5. Fix the slides by dipping in ice-cold methanol for 5 min. Air-dry them completely. 6. Giemsa stain the slides as described in Subheading 3.2.5, steps 3–6) and count the number of amastigotes (see Note 25).

3.5.4 In Vivo Parasite Load in Draining Lymph Node

1. After 5 weeks of L. major challenge infection, dLN was isolated and processed for parasite burden as discussed in Subheading 3.2.2, steps 6–9).

3.5.5 In Vitro Processing of Splenic Tissue

1. Isolate spleen and crush the organ with a pair of frosted-end slides in 3 mL of supplemented RPMI-1640. 2. Pass the suspension through a 70 μM cell strainer and centrifuge at 581  g, 8 min and 4  C. 3. Discard the supernatant and add 1 mL of PBS. 4. Add 5 mL lysis buffer (15 mM NH4Cl, 10 mM NaHCO3, 1 mM disodium EDTA), invert the tube a few times, and incubate. 5. When the pellet changes the color from red to white, neutralize the solution by adding 4 mL of PBS (see Note 26). 6. Resuspend the pellet and spin at 581  g for 8 min at 4  C. 7. Wash the pellet with 2  5 mL of PBS.

3.5.6 Antigen-Specific Immune Response Profiling by Indirect ELISA

1. Isolate dLN from the control and vaccinated groups after 5 weeks of challenge infection. 2. Proceed for quantitative estimation and antigen-specific cytokine secretion profiling as discussed in Subheadings 3.2.5, steps 1 and 2 (see Note 27).

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3.5.7 Cytokine Profiling by qPCR

1. Take 5  106 lymphocytes/group from a pooled single-cell suspension of dLN. 2. Wash cells in 2 1 mL of HBSS and pellet the cell suspension at 425  g for 5 min. 3. Discard the supernatant and tap the bottom of the tube containing cells vigorously to dislodge and loosen the pellet. 4. Add 1 mL of TRI®-reagent to lyse the cells followed by vigorous mixing and vortexing of the pellet, till no cellular lumps are visible. 5. Extract total RNA from cell lysate according to the manufacturer’s instructions. 6. Estimate RNA concentration on the Nano-drop spectrophotometer (see Note 28). 7. Take 2 μg RNA, add 6 μg (in 2 μL) random primer at heat at 65  C for 5 min. 8. In another tube, add first-strand synthesis buffer, 2 mM dNTP, 4 μM DTT, 100 units of MuMLV reverse transcriptase. 9. Mix solutions from steps 7 and 8 and incubate at 37  C for 1 h. 10. Inactivate the reaction mixture at 65  C for 15 min. 11. Set up a qPCR reaction containing 15 ng of cDNA, 0.2 μM each of forward and reverse primer, and 5 μL of SYBR® Premix Ex Taq ™ (Tli RNase H Plus). 12. Run the reaction at the following reaction conditions: 95  C for 30 s, 40–45 cycles of 95  C for 5 s, 60  C for 34 s. A systemspecific melt curve cycle can be run afterward (see Note 29). 13. Calculate the relative fold change in the Ct value of test samples concerning housekeeping genes by the ΔΔCt method.

3.5.8 Antibody ELISA for Post-Challenge Antileishmanial IgG Determination

1. Collect blood sera from various study groups and proceed with determining antileishmanial IgG titer as described in Subheading 3.4.1.

3.5.9 The Leishmanicidal Activity of Ag-Primed TCells by the Co-Culture Assay System

1. After 5 weeks of L. major/L. donovani challenge infection, isolate dLN from control and test groups and make a singlecell suspension.

3.5.10 The Subset Analysis by MultiChromatic Flow Cytometry

1. Isolate draining lymph node (CL study)/spleen (VL study) from control and test groups after 5 weeks of challenge infection.

2. Set up the co-culture system as described in Subheading 3.2.6.3.

2. Process the lymph node and spleen as described in section.

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3. Take 5  106 lymphocytes/group and pellet the cells. 4. Resuspend cells in 400 μL RPMI-1640 and incubate with either 20 ng/mL PMA and 1 μg/mL ionomycin for 6 h, or 30 μg/mL leishmanial CSA overnight at 37  C, 5% CO2. 5. Add brefeldin as per supplier’s instructions for the last 3 h before the incubation period ends. 6. Meanwhile, prepare FACS buffer (1 mM HEPES, 3% FBS dissolved in PBS). 7. Pellet cells and incubate with 50% FBS solution for 20 min at 4  C. 8. Wash cells with 1 mL of FACS buffer. 9. Resuspend in FACS buffer and distribute cells in different tubes such that each tube contains 1.25  106 lymphocytes in 40 μL volume (see Note 30). 10. Make fluorophore-tagged antibody cocktail for surface staining of different Th memory subsets. Dilute anti-CD4, antiCD44, and anti-CD62L for Th1, Th2, Th17 memory cells and anti-CD4, anti-CD25, anti-CD44, and anti-CD62L, for Treg memory cells such that each sample should receive 0.3 μL of each antibody (see Note 31). 11. Incubate for 1 h in dark at 4  C. 12. Wash cells 2 2 mL with FACS buffer at 209  g, 8 min, and 4  C. 13. Resuspend cells in 180 μL of permeabilization solution and incubate for 20 min at 4  C (see Note 32). 14. Wash cells 11 mL of Perm™/wash buffer (diluted, filtered). 15. Resuspend cells in 40 μL of Perm™/wash buffer and add fluorophore-tagged antibody cocktail for intracellular staining of various Th subsets, such that each tube receives 0.5 μL of each antibody (see Note 33). 16. Incubate for 75 min in dark at 4  C. 17. Wash with 21 mL of Perm™/wash buffer at 209  g, 8 min, 4  C. 18. Discard the supernatant and fix cells in 200 μL of 1% PFA/tube and store at 4  C till acquisition.

4

Notes 1. Alternatively, parasite culture flask can be kept in a rocker set at low rpm for a rapid growth rate. 2. Fast-growing culture can be obtained by increasing the seeding inoculum (1:10 ratio decreases the confluency time by approx.

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2 days). Avirulent parasite strain is obtained by the continuous passage of Leishmania for at least 500 generations. 3. It elaborates on the procedure for generating a double negative mutant of GCN-2 like kinase of L. donovani (LdeK1). Overexpression of the N-terminal fragment of LdeK1 keeps itself in an inactivated state by competing with its coactivator, just like GCN-2. Hence, this strategy was utilized here to generate attenuated parasites. Knock-out based strategies for attenuation can also be utilized for immunization and can be found elsewhere in the literature. 4. Transformation buffers (TFB-I and TFB-II) should be freshly prepared on the same day. Use 0.2 M CH3COOH and 0.1 N NaOH for adjusting the pH of TFB-I and TFB-II, respectively. 5. Transformation efficiency can be calculated by counting the colony-forming units obtained by the transformation of 1 μg plasmid. High-efficiency competent cells are generally preferred for cloning purposes. 6. Amplification cycles should be kept at a minimum to lower down the chances of base pair mismatch occurring due to PCR. Generally, 25–30 cycles yield favorable results. 7. GOI should not possess recognition sites for the same restriction enzymes. Generally, enzymes compatible with one another in a single reaction are selected; otherwise, sequential digestion is performed. Incubation with 10 units of each restriction enzyme at 37  C for 4 h is generally sufficient for >90% digestion; however, enzyme concentration and time may differ based on enzyme efficiency and pairs. 8. Amount (in μg) corresponding to a different molar ratio of vector and insert is calculated by the following formula: 9. The methodology outlined here describes the general workflow for recombinant protein induction and purification. A significant amount of troubleshooting may be needed with different proteins as their properties may significantly affect the induction and purification conditions. Instructions given with affinity beads manufacturer can be taken as a starting reference. 10. Generally, tris- and phosphate-based buffers are used as lysis buffers, with a working pH within 2 of the pI value of protein. Moreover, additional components such as β-ME for denaturation, Triton-X for a reduction in nonspecific protein interaction, NaCl and glycerol for protein stability are added to lysis buffer in varying concentrations based on specific proteins. The compatibility of beads with different chemicals and salt concentrations should be taken note from the manufacturer’s booklet.

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11. The amount of Ni-NTA beads for binding generally depends upon the degree of induction achieved by IPTG. A high yield of protein would require a low bead volume to achieve similar levels of affinity-based binding as compared to a low yield of recombinant protein. 12. Time and temperature of incubation of Ni-NTA beads depend upon the properties and amount of induced protein present in the solution. It can vary from 2 h to overnight, at a temperature ranging between 4 and 25  C. 13. If the parasite suspension is relatively more concentrated, then dilute with HBSS so that it corresponds to 2  106 parasites/ 50 μL. Alternatively, if the suspension is dilute than the intended concentration, pellet it once again and resuspend in the desired volume. 14. The peritoneal cavity contains the highest proportion of thioglycolate-elicited macrophages on fifth-day post-injection, as assessed by flow cytometric identification. Care should be observed while flushing the cavity so as not to puncture internal organs which may result in cross-contamination with RBCs, among others. 15. The incubation period for primary macrophages before the experiment is generally meant for switching back the activated-state like macrophages to a resting state. 16. A generalized procedure for macrophage infection is elaborated here. The incubation period postinfection varies depending upon the nature of the experiment being performed. 17. Vigorous washing should be avoided when macrophages are infected in an 8-well chamber slide, as it may result in detaching of adherent cells. A good practice is to observe the extracellular parasite removal after each wash to troubleshoot the washing intensity and time required. 18. The sinapinic acid mixture contains 100 μL of acetonitrile and 100 μL of 1% trifluoroacetic acid added to 5 mg of sinapinic acid. It is used as an energy-absorbing molecule. 19. 25-cm PicoFrit column has an outer diameter of 360 μM, inner diameter 75 μM, and 10 μM tips. 20. Set parameters during SEQUEST-based spectra analysis are as follows: (1) Precursor and fragment mass tolerance at 10 ppm and 0.5 Da. (2) Enzyme specificity set for trypsin/P, along with maximum missed cleavages value of 2. (3) 0.01 false discovery rate for peptide spectrum match and protein false discovery rate. 21. This section defines a generalized immunization protocol. However, the number of booster doses, amount of antigen/ vaccine, adjuvant, inter-dose interval, and route of

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administration may vary in different studies found elsewhere in the literature. Vaccination strategy such as heterologous primeboost (HPB) incorporates two different vaccination modalities as a part of the priming and boosting module. 22. Recombinant protein for immunization against L. donovani (VL) infection can be given via tail vein injection (i.v.) of purified protein solution. This would activate a better repertoire of antigen-specific lymphocytes in spleen. 23. The western blot transfer is performed in a buffer containing 24 mM Tris, 0.2 M glycine, 20% methanol at 4  C. It is advisable to soak the membrane in absolute methanol before setting up the transfer. 24. Thickness read-out should be taken when the footpad fits perfectly in between the spindle and anvil. Thickness is calculated as ¼ thickness (in mm) of left hind footpad—the thickness (in mm) of the contralateral footpad. 25. Leishman-Donovan Units (LDU) is calculated per organ, according to the following formula: LDU ¼ No: of amastigotes per 1000 nucleated cells  organ weight ðin mgÞ: 26. Color transition of pellet from red to white is an indication of RBC lysis. 27. If recombinant protein is available, it is best to analyze antigen-specific cytokine secretion alongside CSA. Set up separate triplicate wells containing 0.5  106 lymphocytes in media containing 10 μg/mL of purified, recombinant antigen. Working concentration of capture and detection antibodies for different cytokines can be taken as IL-4 (1 and 0.5 μg/mL), IFNγ (2 and 1 μg/mL), IL-10, and IL-12 (described in Subheading 3.2.7.1). 28. A260/280 ratio for RNA solution should be in the range of 1.8–2.0. 29. The melt curve is important as it gives us information about nonspecific amplification readouts (false-positives) in our sample, which can interfere with the correct interpretation of our results. 30. Make separate tubes for positive controls for each dye being used in the assay. Also, make a tube containing all isotypes for each fluorophore-tagged antibody being used. These are required for compensation during sample acquisition and post-acquiring analysis. 31. Antibodies diluted in FACS buffer, termed as antibody cocktail, are added to each tube to minimize the error associated

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with dispensing micro volumes. Effector memory T-cells (TEM) are gated as CD44hi CD62Llow and central memory T-cells (TCM) are gated as CD44hi CD62Lhi. However, some studies also utilize CCR7 as well where CCR7low CD44hi CD62Llow and CCR7hi CD44hi CD62Lhi are categorized as TEM and TCM, respectively. Moreover, CD25 is taken as an additional marker for gating Treg cells. Sometimes, if analysis permits, GITR is additionally taken as a surface marker for Treg gating. 32. If one does not have the commercial permeabilization kit, then make the permeabilization solution and wash buffer manually. A 0.1% saponin solution in PBS works well, but it can vary with cell type and experimental conditions. 33. Make separate antibody cocktail solution for each Th subset. Th1, Th2, Th17, and Treg are identified as t-bet+ IFNγ+, GATA-3+ IL-4+, RORγt+ IL-17+, and FOXP3+ IL-10+, respectively. References 1. Killick-Kendrick R (1990) The life-cycle of Leishmania in the sandfly with special reference to the form infective to the vertebrate host. Ann Parasitol Hum Comp 65:37–42 2. Guevara P, Pinto-Santı´ni D, Rojas A, ˜ ez N, Ramirez JL (2001) Crisante G, An Green fluorescent protein-tagged Leishmania in phlebotomine sand flies. J Med Entomol 38:39–43 3. World Health Organization (2020) Leishmaniasis. https://www.who.int/news-room/factsheets/detail/leishmaniasis. Accessed 10 Jun 2020 4. Khamesipour A, Dowlati Y, Asilian A, Hashemi-Fesharki R, Javadi A, Noazin S et al (2005) Leishmanization: use of an old method for evaluation of candidate vaccines against leishmaniasis. Vaccine 23:3642–3648 5. Handman E (2001) Leishmaniasis: current status of vaccine development. Clin Microbiol Rev 14:229–243 6. Noazin S, Modabber F, Khamesipour A, Smith PG, Moulton LH, Nasseri K et al (2008) First generation leishmaniasis vaccines: a review of field efficacy trials. Vaccine 26:6759–6767 7. De Luca PM, Mayrink W, Alves CR, Coutinho SG, Oliveira MP, Bertho AL et al (1999) Evaluation of the stability and immunogenicity of autoclaved and non autoclaved preparations of a vaccine against American tegumentary leishmaniasis. Vaccine 17:1179–1185

8. Amaral VF, Teva A, Oliveira-Neto MP, Silva AJ, Pereira MS, Cupolillo E et al (2002) Study of the safety, immunogenicity and efficacy of attenuated and killed Leishmania (Leishmania) major vaccines in a rhesus monkey (Macaca mulatta) model of the human disease. Mem Inst Oswaldo Cruz 97:1041–1048 9. Fiuza JA, Gannavaram S, Santiago Hda C, Selvapandiyan A, Souza DM, Passos LS et al (2015) Vaccination using live attenuated Leishmania donovani centrin deleted parasites induces protection in dogs against Leishmania infantum. Vaccine 33:280–288 10. Saravia NG, Escorcia B, Osorio Y, Valderrama L, Brooks D, Arteaga L et al (2006) Pathogenicity and protective immunogenicity of cysteine proteinase-deficient mutants of Leishmania mexicana in non-murine models. Vaccine 24:4247–4259 11. Dey R, Dagur PK, Selvapandiyan A, McCoy JP, Salotra P, Duncan R et al (2013) Live attenuated Leishmania donovani p27 gene knockout parasites are nonpathogenic and elicit longterm protective immunity in BALB/c mice. J Immunol 190:2138–2149 12. Palatnik de Sousa CB, Gomes EM, de Souza EP, dos Santos WR, de Macedo SR, de Medeiros LV et al (1996) The FML (Fucose mannose ligand) of Leishmania donovani. A new tool in diagnosis, prognosis, transfusional

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control and vaccination against human kalaazar. Rev Soc Bras Med Trop 29:153–163 13. Santos WR, Paraguai de Souza E, Palatnik M, Palatnik de Sousa CB (1999) Vaccination of Swiss albino mice against experimental visceral leishmaniasis with the FML antigen of Leishmania donovani. Vaccine 17:2554–2561 14. da Silva VO, Borja-Cabrera GP, Correia Pontes NN, de Souza EP, Luz KG, Palatnik M et al (2000) A phase III trial of efficacy of the FML-vaccine against canine kala-azar in an endemic area of Brazil (Sa˜o Gonc¸alo do Amaranto, RN). Vaccine 19:1082–1092 15. Abdelhak S, Louzir H, Timm J, Blel L, Benlasfar Z, Lagranderie M et al (1995) Recombinant BCG expressing the leishmania surface antigen Gp63 induces protective immunity against Leishmania major infection in BALB/c mice. Microbiology 141:1585–1592 16. Bhowmick S, Ravindran R, Ali N (2008) gp63 in stable cationic liposomes confers sustained vaccine immunity to susceptible BALB/c mice infected with Leishmania Donovani. Infect Immun 76:1003–1015 17. Jaffe CL, Rachamim N, Sarfstein R (1990) Characterization of two proteins from Leishmania Donovani and their use for vaccination against visceral Leishmaniasis. J Immunol 144:699–706 18. Dole VS, Raj VS, Ghosh A, Madhubala R, Myler PJ, Stuart KD (2000) Immunization with recombinant LD1 antigens protects against experimental Leishmaniasis. Vaccine 19:423–430 19. Ghosh A, Zhang WW, Matlashewski G (2001) Immunization with A2 protein results in a mixed Th1/Th2 and a humoral response which protects mice against Leishmania Donovani infections. Vaccine 20:59–66 20. Skeiky YA, Coler RN, Brannon M, Stromberg E, Greeson K, Crane RT et al (2002) Protective efficacy of a tandemly linked, multi-subunit recombinant leishmanial vaccine (Leish-111f) formulated in MPL adjuvant. Vaccine 20:3292–3303 21. Coler RN, Goto Y, Bogatzki L, Raman V, Reed SG (2007) Leish-111f, a recombinant polyprotein vaccine that protects against visceral Leishmaniasis by elicitation of CD4+ T cells. Infect Immun 75:4648–4654 22. Morris RV, Shoemaker CB, David JR, Lanzaro GC, Titus RG (2001) Sandfly Maxadilan exacerbates infection with Leishmania major and vaccinating against it protects against L. Major infection. J Immunol 167:5226–5230

23. Gomes R, Oliveira F, Teixeira C, Meneses C, Gilmore DC, Elnaiem DE (2012) Immunity to sand fly salivary protein LJM11 modulates host response to vector-transmitted leishmania conferring ulcer-free protection. J Invest Dermatol 132:2735–2743 24. Cunha JM, Abbehusen M, Suarez M, Valenzuela J, Teixeira CR, Brodskyn CI (2018) Immunization with LJM11 salivary protein protects against infection with Leishmania Braziliensis in the presence of Lutzomyia Longipalpis saliva. Acta Trop 177:164–170 25. Hobernik D, Bros M (2018) DNA vaccines— how far from clinical use? Int J Mol Sci 19:3605 26. St€ager S, Rafati S (2012) CD8(+) T cells in Leishmania infections: friends or foes? Front Immunol 3:5 27. Gurunathan S, Sacks DL, Brown DR, Reiner SL, Charest H, Glaichenhaus N et al (1997) Vaccination with DNA encoding the immunodominant LACK parasite antigen confers protective immunity to mice infected with Leishmania major. J Exp Med 186:1137–1147 28. Melby PC, Yang J, Zhao W, Perez LE, Cheng J (2001) Leishmania Donovani p36(LACK) DNA vaccine is highly immunogenic but not protective against experimental visceral Leishmaniasis. Infect Immun 69:4719–4725 29. Sukumaran B, Tewary P, Saxena S, Madhubala R (2003) Vaccination with DNA encoding ORFF antigen confers protective immunity in mice infected with Leishmania donovani. Vaccine 21:1292–1299 30. Tewary P, Saxena S, Madhubala R (2006) Co-administration of IL-12 DNA with rORFF antigen confers long-term protective immunity against experimental visceral Leishmaniasis. Vaccine 24:2409–2416 31. Basu R, Bhaumik S, Basu JM, Naskar K, De T, Roy S (2005) Kinetoplastid membrane protein-11 DNA vaccination induces complete protection against both pentavalent antimonial-sensitive and -resistant strains of Leishmania Donovani that correlates with inducible nitric oxide synthase activity and IL-4 generation: evidence for mixed Th1- and Th2-like responses in visceral Leishmaniasis. J Immunol 174:7160–7171 32. Bhaumik S, Basu R, Sen S, Naskar K, Roy S (2009) KMP-11 DNA immunization significantly protects against L. Donovani infection but requires exogenous IL-12 as an adjuvant for comparable protection against L. Major. Vaccine 27:1306–1316 33. Gamboa-Leo´n R, Paraguai de Souza E, BorjaCabrera GP, Santos FN, Myashiro LM,

Development of the Antileishmanial Vaccine Pinheiro RO et al (2006) Immunotherapy against visceral Leishmaniasis with the nucleoside hydrolase-DNA vaccine of Leishmania Donovani. Vaccine 24:4863–4873 34. Das S, Freier A, Boussoffara T, Das S, Oswald D, Losch FO et al (2014) Modular multiantigen T cell epitope-enriched DNA vaccine against human leishmaniasis. Sci Transl Med 6:234ra56 35. Dondji B, Pe´rez-Jimenez E, GoldsmithPestana K, Esteban M, McMahon-Pratt D (2005) Heterologous prime-boost vaccination with the LACK antigen protects against murine visceral leishmaniasis. Infect Immun 73:5286–5289 36. Ramos I, Alonso A, Peris A, Marcen JM, Abengozar MA, Alcolea PJ et al (2009) Antibiotic resistance free plasmid DNA expressing LACK protein leads towards a protective Th1 response against Leishmania infantum infection. Vaccine 27:6695–6703 37. Rafati S, Zahedifard F, Nazgouee F (2006) Prime-boost vaccination using cysteine proteinases type I and II of Leishmania infantum confers protective immunity in murine visceral leishmaniasis. Vaccine 24:2169–2175 38. Zutshi S, Kumar S, Chauhan P, Bansode Y, Nair A, Roy S et al (2019) Anti-leishmanial vaccines: assumptions, approaches, and annulments. Vaccines (Basel) 7:156 39. Dayakar A, Chandrasekaran S, Kuchipudi SV, Kalangi SK (2019) Cytokines: key determinants of resistance or disease progression in visceral Leishmaniasis: opportunities for novel diagnostics and immunotherapy. Front Immunol 10:670 40. Scott P, Novais FO (2016) Cutaneous leishmaniasis: immune responses in protection and pathogenesis. Nat Rev Immunol 16:581–592 41. Rodrigues V, Cordeiro-da-Silva A, Laforge M, Silvestre R, Estaquier J (2016) Regulation of immunity during visceral Leishmania infection. Parasit Vectors 9:118 42. Potter H (2003) Transfection by electroporation. Curr Protoc Mol Biol. Chapter 9:Unit 9.3 43. Rao SJ, Meleppattu S, Pal JK (2016) A GCN2like eIF2α kinase (LdeK1) of Leishmania donovani and its possible role in stress response. PLoS One 11:e0156032 44. Zutshi S, Kumar S, Sarode A, Roy S, Sarkar A, Saha B (2020) Leishmania major adenylate kinase immunization offers partial protection

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Chapter 24 In Silico Design of Recombinant Chimera T Cell Peptide Epitope Vaccines for Visceral Leishmaniasis Amanda Sanchez Machado, Vivian Tamietti Martins, Maria Victoria Humbert, Myron Christodoulides, and Eduardo Antonio Ferraz Coelho Abstract Visceral leishmaniasis (VL) is a neglected tropical disease caused by protozoan parasites of the genus Leishmania. Systemic VL is fatal if untreated and there are no prophylactic human vaccines available. Several studies suggest that Th1 cell-mediated immunity plays a major role in protecting against VL. In this chapter we describe a method for designing recombinant chimera vaccines in silico based on the prediction of T cell epitopes within protein antigens identified as potential protective immunogens. Development of a recombinant chimera protein (RCP) vaccine using T cell epitope peptides identified from four Leishmania proteins is used as an exemplar of this method. Key words Visceral leishmaniasis, Recombinant protein, Chimera, T cell epitopes, Peptide

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Introduction Leishmaniases are a group of tropical neglected diseases caused by protozoan parasites of the genus Leishmania. The diseases are present in 98 countries in the world and ~380 million people are exposed to the risk from infection by Leishmania parasites (https:// www.who.int/leishmaniasis/en/). Of the distinct clinical manifestations of disease, visceral leishmaniasis (VL) is systemic and fatal if left untreated [1]. There are ~700,000 to 1.0 million new human cases of VL reported annually with 50,000 deaths [2]. Leishmania can be transmitted to humans through the bite of infected female phlebotomine sandflies [3] and among the different species known, Leishmania infantum causes most of the cases of visceral leishmaniasis (VL) in the Americas, with ~90% of disease registered in Brazil. Treatment with antileishmanial drugs significantly reduces mortality, but treatments are expensive, and accompanied by toxicity,

Sunil Thomas (ed.), Vaccine Design: Methods and Protocols, Volume 1: Vaccines for Human Diseases, Methods in Molecular Biology, vol. 2410, https://doi.org/10.1007/978-1-0716-1884-4_24, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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adverse side effects, and the emergence of resistant strains. To obviate the use of such drugs, there is an urgent need to develop prophylactic vaccines for human VL. Many studies have examined the potential of different Leishmania candidate vaccines in murine and/or canine models, but none has satisfactorily progressed to human trials. These vaccines include dead parasites, genetically modified parasites, or viruses expressing Leishmania genes encoding for recombinant proteins, plasmid DNA encoding genes in eukaryotic expression vectors, and single or multiple recombinant proteins. Much effort has gone into developing single or multiple recombinant Leishmania proteins as candidate vaccines, but issues over protein instability and solubility as well as high manufacturing costs may limit their potential. This is compounded further by the fact that the repertoire of Leishmania proteins is high and variable, thus potentially increasing the antigen load within experimental vaccines. One method to obviate the production of multiple recombinant proteins and to improve solubility and stability is to design a chimera vaccine based on defined protective epitopes drawn from immunogenic Leishmania proteins. The production of experimental synthetic T cell-epitope-based vaccines is a promising approach [4]. Evidence that patients cured from leishmaniasis develop a Th1 response that protects against new infections suggests that targeting Th1 molecules, such as MHC class I (CD8+ cells) and MHC class II (CD4+ cells) could be useful tools to identify new targets to protect against the disease. Vaccine design is being refined with increased knowledge of the nature of the antigens targeted by the mammalian immune response and by exploiting distinctive algorithms to predict and select molecules with the highest probability of inducing beneficial immune responses. Selecting the most antigenic CD4+ and CD8+ T cell epitopes from immunogenic Leishmania proteins avoids other epitopes with low immunogenicity or undesirable immune dominance. Studies have demonstrated the improved protective efficacy of Leishmania chimera vaccines using T cell epitopes combined into a single molecule [5]. Multiple epitope-based vaccines offer the advantages of improved safety, reduced cost of manufacture, facile quality control and delivery, and the opportunity to rationally engineer the epitopes to further increase their immunological potency, when compared with vaccines using single or combinations of recombinant proteins [6, 7]. In this chapter, we describe a method for designing recombinant chimera vaccines in silico based on the prediction of T cell epitopes within protein antigens, which were identified as immunogenic from immuno-proteomics analysis of human serum responses to Leishmania infection. The development of a recombinant chimera protein (RCP) vaccine [8] using these selected protein antigens is used as an exemplar of the method.

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Materials

2.1 Construction of the Gene Encoding the Chimera Protein

1. UniProt program (www.uniprot.org).

2.1.1 FASTA Sequence of Target Proteins 2.1.2 Prediction of T Cell Epitopes

1. Immune Epitope Database (IEDB) and analysis resource program (www.iedb.org).

2.1.3 Prediction of B Cell Epitopes

1. Immune Epitope Database (IEDB) and analysis resource program (www.iedb.org).

2.2 Analysis of Peptide Identity

1. Basic Local Alignment Search Tool (BLAST) program (https://blast.ncbi.nlm.nih.gov/Blast.cgi).

2.3 Peptide Characterization

1. ExPASy program—ProtParam Tool (https://web.expasy.org/ protparam/).

2.4 Chimera Assembly

1. ExPASy program—ProtParam Tool (https://web.expasy.org/ protparam/). 2. Bioinformatics.org website (https://www.bioinformatics.org). 3. ExPASy program—Translate Tool (https://web.expasy.org/ translate/).

2.5 Chimera Gene Synthesis

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1. Integrated DNA Technologies website (https://www.idtdna. com/CodonOpt).

Methods

3.1 Construction of the Gene Encoding the Chimera Protein 3.1.1 FASTA Sequence of Target Proteins

In our example, the proteins used to develop the RCP vaccine are the LiHyp1 (XP_001468941.1) hypothetical amastigote-specific protein, found conserved between different Leishmania species; the LiHyp6 (XP_001568689.1) amastigote-specific protein; the IgE-dependent histamine-releasing factor (HRF) (CAJ05086.1) promastigote-specific protein; the LiHyV (XP_888524.1) hypothetical protein, also found conserved in Leishmania. All four proteins were individually protective against L. infantum, inducing a Th1 immune response in vaccinated hosts that was primed by high levels of IFN-γ release and antileishmanial nitrite production [8].

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Fig. 1 UniProt website, showing the output results

1. Visit the UniProt website at www.uniprot.org. 2. Fill the search unit with the access number of the protein and click search (see Note 1). 3. The results will appear showing the entry, name of the protein, gene name, to which organism it belongs and its length (Fig. 1). 4. Click on the link under “Entry Identification.” 5. All the protein information will appear on the screen, at “Display,” on the left-hand side, show the topics. Click in “Sequence” and open or download the FASTA information (Fig. 2). 3.1.2 Prediction of T Cell Epitopes

The major histocompatibility complex (MHC) are genes present in every mammalian species and are more associated with infectious and autoimmune diseases than any other region of the genome. The MHC is referred to as the human leukocyte antigen (HLA) in humans and as the histocompatibility system (H-2) in mice. MHC class I peptides are present on nucleated cells and are recognized by cytotoxic CD8+ T cells, while MHC class II peptides are present in antigen-presenting cells (APC), such as dendritic cells and macrophages, and activate CD4+ T cells, leading to the coordination and regulation of effector cells. In all cases, MHC are clonotypic T cell receptors that interact with a given MHC peptide complex, potentially leading to sustained cell–cell contact formation and T cell activation [9]. A cell-mediated immune response is believed to protect against infection and is dependent on the generation of Th1 cells and the production of pro-inflammatory cytokines, for example, interferon-γ (IFN-γ) and interleukin-12 (IL-12), which activate macrophages to kill internalized parasite [10]. In addition,

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Fig. 2 UniProt website, showing output results and FASTA sequence of protein

the Th1 profile is associated with the production of IgG2a isotype antibodies [11]. 1. Visit the Immune Epitope Database (IEDB) and analysis resource program at www.iedb.org. 2. On the right-hand side, click on “T cell Epitope Prediction”; first of all, select “MHC I Binding,” perform all the searches, and then look for “MHC II Binding,” as it can only search one MHC class type at a time (Fig. 3). 3. Fill in the “Specify Sequence” window with the FASTA sequence of the protein (see Subheading 3.1.1) (entry >Protein name, Enter, FASTA sequence), or download the file, clicking on “Choose file” (Fig. 4). 4. Prediction method: IEDB recommended (see Note 2). 5. MHC source species: select which species to look for (see Note 3). 6. Select the Allele and Length (see Note 4). 7. Specify output: sort peptides by “Percentile rank”; show “All predictions” and press “Submit” (Fig. 4). 8. As output data, the IEDB program will show which parameter was used for each peptide found. Search in the “Percentile Rank” for the numbers 1.0 (see Note 7). 7. Compare the results from the chosen predicted T cell epitopes (see Subheading 3.1.2) and predicted B cell epitopes and select the ones that are unique for T cell epitopes. 3.2 Analysis of Peptide Identity

1. Visit the BLAST website at https://blast.ncbi.nlm.nih.gov/ Blast.cgi, select Protein BLAST (BLAST-p) and insert the peptide sequence (one at a time) (Fig. 8). 2. Select “Protein Data Bank proteins (pdb)” in the section “Choose Search Set.”

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Fig. 5 The IEDB website showing results from MHC I epitope search

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Fig. 6 The IEDB Program website showing input information for B cell epitope prediction

3. Type in “Homo sapiens” in the “Organism” section. 4. Select “blastp” (protein–protein BLAST) in the “Program selection” section and press “BLAST” (Fig. 8). 5. Select the peptides that share >85% identity with the human genome, since this will ensure cell–cell recognition and that there is a response to the vaccine. 3.3 Peptide Characterization

1. Visit the ExPASy program at https://web.expasy.org/pro tparam/ and select the ProtParam Tool. 2. Fill in the box with your peptide’s sequence and press “Compute parameters.” 3. The information referring to the number of amino acids, molecular weight, theoretical isoelectric point (pI), and amino acid composition will appear at the beginning of the page (see Note 8) (Fig. 9). 4. Scroll down to the end of the page to find the Instability Index, Aliphatic Index, and Grand Average of Hydropathicity (GRAVY) values (see Notes 9, 10, and 11).

3.4 Chimera Assembly

1. Allocate the peptides in the final sequence mimicking their original arrangement in the source protein. For example, if

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Fig. 7 The IEDB Program website showing results from B cell epitope prediction as the Kolaskar and Tongaonkar antigenicity results

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Fig. 8 BLAST-P program website showing input information

one peptide was found at the beginning of a protein sequence, it should be the first one in your chimera (see Note 12) (Fig. 10a). 2. Insert two glycine residues (GG) between the peptides, which will favor protein stability (see Note 13) (Fig. 10b). 3. Once the chimera is assembled, analyze the chimeric protein characteristics using the ExPASy ProtParam Tool, as previously described for the individual peptides (see Subheading 3.3, step 1). 4. If the chimera displays high GRAVY values, insert a Lysine residue (K) between the peptide sequences, until it is predicted to be soluble (see Note 14). 5. Visit the Bioinformatics website at https://www.bioinformat ics.org to reverse translate the amino acid sequence of the chimera to its nucleotide sequence (see Note 15). 6. On the left-hand side, scroll down to “Online Analysis Tool,” and select “SMS 2—Sequence manipulation” (Fig. 11). 7. The menu will appear on the left-hand side. Select “Reverse Translate” within the “Sequence analysis” section (see Note 16) (Fig. 11). 8. Insert the FASTA sequence of the chimera and press “Submit.” 9. Finally, add a Start Codon (ATG) at the beginning of the chimera nucleotide sequence and a Stop Codon at the end (TGA, TAA, or TAG) (see Note 17). 3.5 Chimera Gene Synthesis

After design in silico, the chimera can be synthesized in-house by polymerase chain reaction (PCR) [12] in the form of a DNA string,

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Fig. 9 Expasy Program website showing ProtParam output information

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Fig. 10 (a) Amino acid sequence of the four chosen protein for the exemplar RCP chimera. The selected epitopes to assemble the chimeric protein are highlighted by the different colors. (b) Assembly of the exemplar RCP chimera protein. Each peptide sequence is separated by a GG amino acid spacer

which will need to be cloned into an appropriate vector, for example, pET28(+)TEV for protein expression. Alternatively, the chimera can be synthesized and cloned commercially (Fig. 12). Codon optimization for protein expression in Escherichia coli can be done commercially in silico with the Codon Optimization Tool at https://www.idtdna.com/CodonOpt (see Note 18).

4

Notes 1. The Access Number of the protein is provided by the National Center for Biotechnology Information (NCBI), and, for example, it can be reported as XP, SUZ, or AYU. When proteins are

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Fig. 11 The bioinformatics program showing pathways to reverse translate

Bpu1102 I(80) Dra III(5127)

Pvu I(4426) Sgf I(4426) Sma I(4300)

Xba l(335) Bgl II(401) SgrA I(442)

Mlu I(1123)

pET-28a(+) (5369bp)

Iacl (773-1852)

Cla I(4117) Nru I(4083)

Kan (3 995 -48 07 )

-5358) (4903 gin i r o f1

Xho l(158) Not l(166) Eag l(166) Hind lll(173) Sal l(179) Sac l(190) EcoR l(192) BamH l(198) Nhe l(231) Nde l(238) Nco l(296)

Bcl I(1137) BstE II(1304) Apa I(1334) BssH II(1534) EcoR V(1573)

Eco57 I(3772)

Hpa I(1629) AlwN I(3640)

or

i

(3

BssS I(3397)

28

6)

BsPLU1 I(3224) Sap I(3108) Bst1107 I(2995) Tth111 I(2969)

PshA I(1968) Bgl I(2187) Fsp I(2205) Psp5 II(2230)

Fig. 12 Expression clone vector pET-28a(+)TEV from the Novagen website. Multiple cloning sites are detailed within the black arrow

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identified with an immuno-proteomics approach, they are listed according to NCBI information, so they can be easily found [13]. 2. The IEDB uses the best possible prediction method for a given MHC molecule, among all the available ones in this website, and is recommended. 3. MHC source species: it is preferable to look first in mouse and for Leishmania peptide searches, the most commonly selected MHC class I molecules are H-2Db, H-2Dd, H-2Kb, H-2Kd, H-2Kk, and H-2Ld, and the most commonly selected MHC class II molecules are H-2IAb, H-2IAd, H-2Ias, H-2IEd. Second, look for MHC I and MHC II molecules in humans. The most popular searches for Leishmania peptide sequences are HLA*A02, HLA*A03:01, and HLA*B27 for MHC I, and HLA DRB*0101 and HLA DRB1*07 for MHC II. These MHC molecules are predicted to be present in 90% of the human population of any ethnic group. The peptides that are found both in the mouse and human are preferred. 4. The allele length depends on the characteristics of a protein. When searching for Leishmania peptides, the length usually varies between 8 and 10 amino acids for MHC I and between 12 and 15 amino acids for MHC II (predicted to cover 82% and 89% of epitope frequency, respectively). 5. The Kolaskar and Tongaonkar Antigenicity Scale method predicts antigenic determinants with ~75% accuracy and is one of the more generally preferred methods [14]. 6. When searching for B cell epitopes, the number of peptides may vary according to characteristics of the protein; choose from 10 to 15 amino acids for a better window size. 7. Threshold values >1.0 show that predicted epitopes are more accessible and, therefore, probably more antigenic. 8. The isoelectric point (pI) is the pH of the protein at which its net charge becomes zero. 9. The Instability Index is used to determine whether the protein will be stable in vitro. Values 40 predict instability. Preference should be given to stable peptides. 10. The Aliphatic Index refers to the thermostability factor of proteins. The index is defined as the relative volume occupied by aliphatic side chains (alanine, valine, isoleucine, and leucine). A globular protein is more thermostable with higher index numbers.

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11. The GRAVY Index is used to evaluate protein hydrophobicity. This index score is defined by the sum of hydropathy values of all amino acids divided by the protein length. Smaller GRAVY values predict more soluble proteins. However, at this point, no peptide should be excluded based on the GRAVY parameter, since the GRAVY value will probably change for the final chimera protein. 12. If two peptides are at the beginning of the sequence of their corresponding source protein, try different chimera designs and evaluate which one gives you better results when analyzing the parameters of the chimera in its final form (by comparing the results with the ProtParam Tool of the ExPASy program). 13. For the RCP chimera exemplar, since the chosen epitopes were all in sequence according to their position within their respective source protein, the GG spacer was not inserted to separate adjacent epitopes belonging to the same protein, but to separate epitopes coming from different proteins. 14. Adding lysine (K) residues to the protein will not change the protein’s characteristics; this amino acid is inert, and it enhances protein solubility. 15. Reverse translation of the protein sequence can also be done manually according to the Standard Genetic Code Map. 16. To confirm that the reverse translated sequence is error-free, translate it back to the amino acid sequence using the Translate Tool at ExPASy website (https://web.expasy.org/translate/) and compare it with your original chimera protein sequence. An easy way to compare sequences is by alignment, which can be done with the BLAST Tool at the NCBI website (https:// blast.ncbi.nlm.nih.gov/Blast.cgi), or with any other online program for sequence alignment, such as Clustal Omega (https://www.ebi.ac.uk/Tools/msa/clustalo/), among others. Most of these websites are access-free. 17. It is important to check that there are no Stop Codons inside the chimera nucleotide sequence (TGA; TAA; TAG), as internal Stop Codons will stop translation and will therefore result in a truncated protein. If they are present, change one nucleotide of the codon so that it encodes an amino acid residue. 18. Protocols for transformation of E. coli with chimera gene vectors, expression of the RCP and subsequent purification are described comprehensively elsewhere [8, 15].

Acknowledgements VL vaccine research is supported by grant MR/R005850/1 from the Medical Research Council (VAccine deveLopment for complex

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Intracellular neglecteD pAThogEns—VALIDATE), UK, and grant APQ-408675/2018-7 from the Conselho Nacional de Desenvolvimento Cientı´fico e Tecnolo´gico (CNPq), Brazil. References 1. Alvar J, Velez ID, Bern C, Herrero M, Desjeux P, Cano J, Jannin J, den Boer M, Team WHOLC (2012) Leishmaniasis worldwide and global estimates of its incidence. PLoS One 7(5):e35671. https://doi.org/10. 1371/journal.pone.0035671 2. Burza S, Croft SL, Boelaert M (2018) Leishmaniasis. Lancet 392(10151):951–970. https://doi.org/10.1016/S0140-6736(18) 31204-2 3. Grimaldi G Jr, Tesh RB (1993) Leishmaniases of the New World: current concepts and implications for future research. Clin Microbiol Rev 6(3):230–250. https://doi.org/10.1128/ cmr.6.3.230 4. Dikhit MR, Kumar A, Das S, Dehury B, Rout AK, Jamal F, Sahoo GC, Topno RK, Pandey K, Das VNR, Bimal S, Das P (2017) Identification of potential MHC class-II-restricted epitopes derived from Leishmania donovani antigens by reverse vaccinology and evaluation of their CD4+ T-cell responsiveness against visceral Leishmaniasis. Front Immunol 8:1763. https://doi.org/10.3389/fimmu.2017. 01763 5. Martins VT, Lage DP, Duarte MC, Carvalho AM, Costa LE, Mendes TA, Vale DL, Menezes-Souza D, Roatt BM, Tavares CA, Soto M, Coelho EA (2017) A recombinant fusion protein displaying murine and human MHC class I- and II-specific epitopes protects against Leishmania amazonensis infection. Cell Immunol 313:32–42. https://doi.org/10. 1016/j.cellimm.2016.12.008 6. Joshi S, Yadav NK, Rawat K, Kumar V, Ali R, Sahasrabuddhe AA, Siddiqi MI, Haq W, Sundar S, Dube A (2019) Immunogenicity and protective efficacy of T-cell epitopes derived from potential Th1 stimulatory proteins of Leishmania (Leishmania) donovani. Front Immunol 10:288. https://doi.org/10. 3389/fimmu.2019.00288 7. Agallou M, Athanasiou E, Koutsoni O, Dotsika E, Karagouni E (2014) Experimental validation of multi-epitope peptides including promising MHC class I- and II-restricted epitopes of four known Leishmania infantum proteins. Front Immunol 5:268. https://doi. org/10.3389/fimmu.2014.00268

8. Martins VT, Duarte MC, Lage DP, Costa LE, Carvalho AM, Mendes TA, Roatt BM, Menezes-Souza D, Soto M, Coelho EA (2017) A recombinant chimeric protein composed of human and mice-specific CD4(+) and CD8(+) T-cell epitopes protects against visceral leishmaniasis. Parasite Immunol 39(1). https://doi.org/10.1111/pim.12359 9. Wieczorek M, Abualrous ET, Sticht J, AlvaroBenito M, Stolzenberg S, Noe F, Freund C (2017) Major histocompatibility complex (MHC) class I and MHC class II proteins: conformational plasticity in antigen presentation. Front Immunol 8:292. https://doi.org/ 10.3389/fimmu.2017.00292 10. Rodrigues V, Cordeiro-da-Silva A, Laforge M, Silvestre R, Estaquier J (2016) Regulation of immunity during visceral Leishmania infection. Parasit Vectors 9:118. https://doi.org/10. 1186/s13071-016-1412-x 11. Koarada S, Wu Y, Olshansky G, Ridgway WM (2002) Increased nonobese diabetic Th1:Th2 (IFN-gamma:IL-4) ratio is CD4+ T cell intrinsic and independent of APC genetic background. J Immunol 169(11):6580–6587. https://doi.org/10.4049/jimmunol.169.11. 6580 12. Machado AS, Ramos FF, Oliveira-da-Silva JA, Santos TTO, Ludolf F, Tavares GSV, Costa LE, Lage DP, Steiner BT, Chaves AT, ChavezFumagalli MA, de Magalhaes-Soares DF, Silveira JAG, Napoles KMN, Tupinambas U, Duarte MC, Machado-de-Avila RA, Bueno LL, Fujiwara RT, Moreira RLF, Rocha MOC, Caligiorne RB, Coelho EAF (2020) A Leishmania infantum hypothetical protein evaluated as a recombinant protein and specific B-cell epitope for the serodiagnosis and prognosis of visceral leishmaniasis. Acta Trop 203:105318. https://doi.org/10.1016/j. actatropica.2019.105318 13. Machado AS, Ramos FF, Oliveira-da-Silva JA, Santos TTO, Tavares GSV, Costa LE, Lage DP, Teixeira-Ferreira A, Perales J, Fernandes AP, Moreira RLF, Duarte MC, Tupinambas U, Caligiorne RB, Cota GF, Coelho EAF, Ludolf F (2020) An immuno-proteomics approach to identify Leishmania infantum proteins to be applied for the diagnosis of visceral leishmaniasis and human immunodeficiency virus co-infection. Parasitology 147:932–939.

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https://doi.org/10.1017/ S0031182020000578 14. Kolaskar AS, Tongaonkar PC (1990) A semiempirical method for prediction of antigenic determinants on protein antigens. FEBS Lett 276(1–2):172–174. https://doi.org/10. 1016/0014-5793(90)80535-q

15. Humbert MV (2019) Cloning, expression, and purification of recombinant Neisseria gonorrhoeae proteins. Methods Mol Biol 1997:233–266. https://doi.org/10.1007/ 978-1-4939-9496-0_15

Chapter 25 Preclinical Assessment of the Immunogenicity of Experimental Leishmania Vaccines Vivian Tamietti Martins, Amanda Sanchez Machado, Maria Victoria Humbert, Myron Christodoulides, and Eduardo Antonio Ferraz Coelho Abstract Leishmaniases are neglected diseases caused by Leishmania parasites and affect millions of people worldwide. The induction of protective immunity against infection by some species of Leishmania has stimulated the development of vaccine candidates against the disease. In this chapter we describe protocols for immunizing mice with a recombinant chimera vaccine containing selected epitopes that specifically stimulate a Th1-type immune response. We describe protocols for challenging mice with live Leishmania parasite and for measuring parameters of the immune response to vaccination and parasite infection, including the production of cytokines, nitric oxide, and IgG antibodies, and the contribution of CD4+ and CD8+ T cells. We also provide protocols for isolating mouse organs for cell culture and for quantifying parasite loads in unvaccinated control animals and in vaccine-protected animals. These protocols can form the basis of immunological studies of candidate Leishmania vaccines in the mouse, as a step toward further vaccine development for human use. Key words Leishmania, Vaccines, Th1 type immunity, Cytokines, Nitric oxide, Mouse model

1

Introduction Leishmaniases are globally important diseases caused by protozoa of the genus Leishmania [1]. The disease complex is endemic in 98 countries and 3 territories ranging across the Mediterranean Basin, the Middle East, the Indian subcontinent, and the tropical regions from America and Africa. Of these 98 countries, 72 are classified as developing nations. Cutaneous leishmaniasis (CL) presents as dermatological and/or mucosal disorders that can lead to severe deformities, morbidity, and consequent social exclusion. Visceral leishmaniasis (VL) is an infection of internal organs with typical parasite tropism for the spleen, lymph nodes,

Sunil Thomas (ed.), Vaccine Design: Methods and Protocols, Volume 1: Vaccines for Human Diseases, Methods in Molecular Biology, vol. 2410, https://doi.org/10.1007/978-1-0716-1884-4_25, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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liver, and bone marrow. If VL is untreated, disease progression is fatal for most patients [2]. Disease severity is determined by the interaction of multiple factors encompassing virulence of the infecting parasite, the biology of the host and insect vector and their immune responses. This interplay could result in a wide range of clinical outcomes, varying from resistance to development of acute disease [3]. Mammalian host responses to Leishmania infection are innate—mediated by monocytes, macrophages, dendritic cells, and neutrophils—and adaptive—mediated by T and B cells. Cytokine signaling molecules allow these complex networks of interactions to take place, since they can induce or regulate an immune response profile [4]. Susceptibility and clinical progression of the disease are typically characterized by a classical Th2 response profile, defined by high expression of anti-inflammatory cytokines such as IL-4, IL-13, IL-10, and TGF-β, immunomodulation of Leishmania-specific T cells, and/or by the presence of high titers of specific IgG1 antibodies to the parasite. In contrast, induced proliferation of peripheral blood mononuclear cells (PBMCs) after stimulation with Leishmania antigens, expression of high levels of pro-inflammatory cytokines such as IFN-ɣ, IL-12, and TNF-α, proliferation of CD4+ and CD8+ T cells, and increased levels of antileishmanial IgG2a isotype antibody production are associated with a Th1 response profile and resistance to infection [4, 5]. Resistance to infection also involves activation of the inducible Nitric Oxide Synthase (iNOS) enzyme through the action of IFN-ɣ, which induces the production of nitric oxide (NO) by circulating macrophages that can control intracellular parasite multiplication [6, 7]. Understanding the basis of protective immune responses to Leishmania infection guides the research and development of candidate vaccines. Several vaccine candidates against VL have been evaluated mainly in murine and/or canine models [8–13]. They include dead parasites administered with or without adjuvants, genetically modified parasites, or viruses expressing Leishmania genes encoding for recombinant proteins, plasmid DNA-based vaccines that encode genes in eukaryotic expression [14, 15], and protein-based vaccines using whole recombinant proteins and chimera proteins containing T cell epitopes [16–19]. However, none of these experimental vaccine candidates has progressed satisfactorily in human trials. Preclinical evaluation of these vaccines involves animal vaccination studies and examination of parameters of the immune response including generation of a Th1-type response, production of cytokines and NO, and a reduction in parasite load within internal organs of vaccinated animals that are subsequently infected with live parasites. In this chapter, we describe the protocols for assessing the immunogenicity of candidate Leishmania vaccines in the BALB/c mouse. These protocols are appropriate

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for testing not only the recombinant chimera protein vaccines described in the accompanying chapter from Machado et al., but also other experimental vaccines. Fulfillment of these immune response criteria would endorse vaccines for future human trials.

2

Materials

2.1 Leishmania spp. Culture

1. Schneider’s Complete Medium: Schneider’s insect medium supplemented with 20% (v/v) of heat-inactivated fetal bovine serum (FBS) (see Note 1), 20 mM of L-glutamine, 100 U/mL of penicillin, and 50 μg/mL of streptomycin, pH 7.4. Filtersterilize using a 0.2 μm filter and a sterile syringe and store at 4  C until needed. 2. Leishmania spp. strains stored in sterile Schneider’s Complete Medium plus 15% (v/v) glycerol in liquid nitrogen cryotubes. 3. Phosphate-Buffered Saline (PBS) pH 7.4 (see Note 2). Filtersterilize through a 0.2 μm filter with a sterile syringe and store at 4  C until needed. 4. Equipment: Incubator set to 24  C, biological class II safety cabinet, water bath set to 37  C, polystyrene sterile flasks with growth areas of 25 cm2, high-speed centrifuge, Neubauer chamber, optical microscope.

2.2 Soluble Leishmania Antigen (SLA)

1. Sterile cold PBS, pH 7.4. 2. Isoton solution: 0.05 M of citric acid, 0.12 M of NaCl, 0.5% (v/v) of formaldehyde, pH 7.2. 3. Liquid nitrogen. 4. Equipment: 37  C water bath, ice bucket, high-speed refrigerated centrifuge, 15 mL conical polypropylene centrifuge tubes, probe sonicator, Neubauer chamber.

2.3 Evaluation of Antigen Concentration

1. BCA protein assay, for example, Pierce™ Kit (see Note 3).

2.4 Preparation of Injections

1. Recombinant chimera protein (RCP): 25 μg of RCP per dose in PBS, pH 7.4.

2. PBS, pH 7.4. 3. Equipment: 96-well flat-bottom microtiter plates, microplate spectrophotometer.

2. Sterile PBS, pH 7.4. 3. 5 mg/mL of a saponin stock solution in sterile PBS: 25 μg of saponin per dose in PBS, pH 7.4 (see Note 4). 4. 25 μg of RCP plus 25 μg of saponin in PBS, pH 7.4 (see Note 5).

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5. Equipment: Sterile polypropylene microtubes, ice bucket, biological class II safety cabinet. 2.5

Immunization

1. Animal model: BALB/c mice (16 per group), female, 6 to 8 weeks of age (approximate weight 20 g). 2. Sterile PBS, pH 7.4. 3. 5 mg/mL of saponin in sterile PBS, pH 7.4. 4. RCP in sterile PBS. 5. Equipment: 1 mL syringe with 30 g  ½ in. needle, acrylic mouse restrainer.

2.6 Mouse Challenge with Leishmania spp.

1. Injected and immunized BALB/c mice (8 per group). 2. Leishmania spp. culture in stationary promastigotes phase (see Note 6). 3. Sterile PBS, pH 7.4. 4. Isoton solution: 0.05 M of citric acid, 0.12 of M NaCl, 0.5% (v/v) of formaldehyde, pH 7.2. 5. Equipment: 1 mL syringe with 30 g  ½ in. needle, acrylic mouse restrainer, Neubauer chamber, inverse microscope.

2.7 Preparation of Mouse Serum After Immunization or After Immunization and Challenge

1. Mouse anesthetic cocktail: 87.5 mg/kg of ketamine and 12.5 mg/kg of xylazine in sterile PBS, pH 7.4 [20].

2.8 Isolation of Mouse Splenocytes for Cell Culture

1. RMPI 1640 medium.

2. Equipment: Surgical materials (forceps, scalpel, and scissors), biological class II safety cabinet, polyethylene Pasteur pipettes, polypropylene centrifuge microtubes, high-speed centrifuge.

2. RPMI 1640 Complete Medium: RPMI 1640 medium supplemented with 10% (v/v) of heat-inactivated FBS, 20 mM of Lglutamine, 200 U/mL of penicillin, and 100 μg/mL of streptomycin, pH 7.4. Filter-sterilize using a 0.2 μm filter and a sterile syringe and store at 4  C until needed. 3. Erythrocyte Lysis Solution: 17 mM of Tris-HCl, 144 mM of NH4Cl, pH 7.4. Filter-sterilize using a 0.2 μm filter and a sterile syringe and store at 4  C until needed. 4. 70% (v/v) ethanol. 5. Equipment: Surgical materials (forceps, scalpel, and scissors), biological class II safety cabinet, 1.5 mL and 50 mL sterile polypropylene tubes, 70 μm cell strainer, ice bucket, highspeed refrigerated centrifuge, Neubauer chamber and optical microscope, 24-well flat-bottom culture plates. 1. RPMI 1640 Complete Medium: RPMI 1640 medium supplemented with 10% (v/v) of heat-inactivated FBS, 20 mM of L-

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glutamine, 200 U/mL of penicillin, and 100 μg/mL of streptomycin, pH 7.4. Filter-sterilize using a 0.2 μm filter and a sterile syringe and store at 4  C until needed. 2. SLA [(25 μg per well) (see Note 7)]. 3. RCP [(20 μg per well) (see Note 7)]. 4. Individual recombinant proteins [source of the peptide epitopes used to produce the RCP (20 μg per well) (see Note 7)]. 5. 1 mg/mL of concanavalin A in sterile RPMI 1640 medium (see Note 7). 6. Monoclonal anti-IL-12, anti-CD4, and anti-CD8 antibodies (see Note 7).

2.10 Isolation of Spleen, Liver, and Lymph Nodes for Parasite Culture

1. Complete Schneider’s Medium: Schneider’s insect medium supplemented with 20% (v/v) of heat-inactivated fetal bovine serum (FBS), 20 mM of L-glutamine, 100 U/mL of penicillin, and 50 μg/mL of streptomycin, pH 7.4. Filter-sterilize using a 0.2 μm filter and a sterile syringe and store at 4  C until needed. 2. 70% (v/v) ethanol. 3. Equipment: Surgical materials (forceps, scalpel, and scissors), biological class II safety cabinet, glass tissue grinder, 1.5 mL, 15 mL, and 50 mL sterile polypropylene tubes, analytical balance, ice bucket.

2.11 Isolation of Bone Marrow for Parasite Culture

1. Sterile PBS, pH 7.4.

2.12 Evaluation of the Parasite Load in Mouse Organs

1. Complete Schneider’s Medium: Schneider’s insect medium supplemented with 20% (v/v) of heat-inactivated fetal bovine serum (FBS), 20 mM of L-glutamine, 100 U/mL of penicillin, and 50 μg/mL of streptomycin, pH 7.4. Filter-sterilize using a 0.2 μm filter and a sterile syringe and store at 4  C until needed.

2. Equipment: Surgical materials (forceps, scalpel, and scissors), 18 g needle, biological class II safety cabinet, 0.5 mL and 1.5 mL sterile microcentrifuge polypropylene tubes, highspeed refrigerated centrifuge.

2. Macerated organs samples (spleen, liver, lymph nodes, and bone marrow). 3. Equipment: Biological class II safety cabinet, 96-well flat-bottom polystyrene cell culture plates, parafilm, incubator set to 24  C. 2.13 Flow Cytometry for Analysis of Cytokines

1. SLA [(25 μg per well) (see Note 7)]. 2. RCP [(20 μg per well) (see Note 7)]. 3. Individual recombinant proteins [source of the peptide epitopes used to produce the RCP (20 μg per well) (see Note 7)].

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4. 10 mg/mL of Phorbol-12-myristate-13-acetate (PMA) in sterile dimethyl sulfoxide (DMSO) (1000 stock solution). Store the sterile solution protected from light at 20  C. 5. 10 μg/mL of PMA in sterile RPMI 1640 medium. Prepare by diluting the PMA stock solution 1/1000 in medium (see Note 7). 6. 200 μg/mL of ionomycin in sterile DMSO. Store protected from light at 20  C (see Note 7). 7. CM-BLAST medium: RPMI 1640 medium supplemented with 20% (v/v) of heat-inactivated FBS, 1% (w/v) of gentamicin, 1% (w/v) of L-glutamine, 0.1% (v/v) of β-mercaptoethanol, 200 U/mL of penicillin, and 100 μg/mL of streptomycin, pH 7.4. Filter-sterilize using a 0.2 μm filter and a sterile syringe and store at 4  C until needed. 8. 1 mg/mL of Brefeldin A in RPMI 1640 medium stock solution. 9. 200 μg/mL of Brefeldin A in RPMI 1640 medium. Add 200 μL of 1 mg/mL Brefeldin A to 800 μL of RPMI 1640 medium. 10. PBS, pH 7.4. 11. 20 mM of ethylenediaminetetraacetic acid (EDTA) in RPMI 1640 medium. Filter-sterilize using a 0.2 μm filter and a sterile syringe and store at 4  C until needed. 12. Fixable Viability Stain 450 [(FVS450—BD Horizon™) (see Note 8)]. 13. PBS-Wash (PBS-W): PBS supplemented with 0.5% (w/v) of bovine serum albumin (BSA), pH 7.4. Filter-sterilize using a 0.2 μm filter and a sterile syringe. Store at 4  C until needed. 14. Anti-CD3, anti-CD4, and anti-CD8 monoclonal antibodies diluted in PBS-W plus 5% (v/v) of heat-inactivated FBS (see Note 8). 15. 10 Cytometry Lysis Solution: 5.7 g of sodium citrate dehydrate, 800 μL of 5000 UI/mL heparin, 108 mL of formaldehyde, 60 mL of diethylene glycol in 200 mL of distilled H2O, pH 7.85 (see Note 9). Filter-sterilize using a 0.2 μm filter and a sterile syringe and store in a polypropylene bottle. 16. 1 Cytometry Lysis Solution: Dilute 10 Cytometry Lysis Solution in distilled H2O to use. 17. PBS-Permeabilization (PBS-P): Add 0.5% (w/v) of saponin to PBS-W, pH 7.4. Filter-sterilize using a 0.2 μm filter and a sterile syringe and store at 4  C until needed.

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18. Anti-IFN-ɣ, anti-TNF-α and anti-IL-10 monoclonal antibodies diluted in PBS-P plus 5% (v/v) of heat-inactivated FBS (see Note 8). 19. Compensation beads. 20. Equipment: High-speed refrigerated centrifuge, 96-well round-bottom polystyrene cell culture plates, 5 mL polystyrene tubes, BD LSR Fortessa Cell Analyzer™ cytometer (or equivalent), BD FACSDiva™ software, and FlowJo™ software package (or equivalent) for analyzing flow cytometry data (Becton Dickinson—BD, USA). 2.14 Enzyme-Linked Immunosorbent Assay for Cytokines

1. BD OptEIA™ Set Mouse Kits for IFN-ɣ, IL-12, GM-CSF, IL-4, and IL-10 (BD-Biosciences Pharmingen™), or equivalent. 2. Coating Buffer for IFN-ɣ, IL-4 and GM-CSF: Add 7.13 g of NaHCO3, 1.59 g of Na2CO3 in 1 L of distilled H2O, pH 9.5 (see Note 10). 3. Coating Buffer for IL-10 and IL-12: Add 12.49 g of Na2HPO4, 15.47 g of NaH2PO4 in 1 L of distilled H2O, pH 6.5 (see Note 10). 4. Assay Diluent: PBS supplemented with 10% (v/v) of heatinactivated FBS, pH 7.0 (see Note 10). 5. Wash Buffer: 0.05% (v/v) of Tween-20 in PBS, pH 7.4 (see Note 10). 6. Spleen cell culture supernatant. 7. Citrate Buffer: 24 mM of Na2HPO4 and 17.5 mM of citric acid in 1 L of distilled H2O, pH 5.0. 8. Substrate Solution: 3 mg of O-phenylenediamine (OPD), 10 mL of citrate buffer, and 3 μL of H2O2. Prepare before use (see Note 11). 9. Stop Solution: 2 N H2SO4 in distilled H2O. 10. Equipment: 96-well flat-bottom polystyrene microtiter plate, microplate spectrophotometer, and absorbent paper.

2.15 Enzyme-Linked Immunosorbent Assay for Antibodies

1. Coating Buffer: 0.1 M of Na2CO3, 0.05 M of NaHCO3 prepared in distilled H2O, pH 9.6 (see Note 10). 2. Blocking Solution: 5% (w/v) of nonfat dry milk and 0.05% (v/v) of Tween-20 in PBS, pH 7.4. 3. Wash Buffer: 0.05% (v/v) of Tween-20 in PBS, pH 7.4. 4. Incubation Buffer: 0.05% (w/v) of nonfat dry milk and 0.05% (v/v) of Tween-20 in PBS, pH 7.4. 5. Serum samples.

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6. Anti-mouse IgG, IgG1, and IgG2a horseradish-peroxidase conjugated antibodies. 7. Citrate Buffer: 24 mM of Na2HPO4 and 17.5 mM of citric acid in 1 L of distilled H2O, pH 5.0. 8. Substrate Solution: 2 mg of O-phenylenediamine (OPD), 10 mL of citrate buffer, and 2 μL of H2O2. Prepare before use. 9. Stop Solution: 2 N H2SO4 in distilled H2O. 10. Equipment: 96-well flat-bottom polystyrene microtiter plate, microplate spectrophotometer, Parafilm, and absorbent paper. 2.16 Measurement of Nitrite Using Griess Reaction 2.16.1 Preparation of Nitrite Standard Reference Curve 2.16.2 Nitrite Measurement (Griess Reaction)

1. 0.1 M Nitrite Standard stock solution. 2. RPMI 1640 Medium. 3. 100 μM of sodium nitrite in Milli-Q water. Prepare 1 mL of a 100 μM nitrite solution by diluting the 0.1 M Nitrite Standard stock solution 1/1000 in complete RPMI 1640.

1. 1% (w/v) of sulfanilamide in 5% (v/v) of phosphoric acid, in water (Sulfanilamide Solution). 2. 0.1% (w/v) of N-1-napthylethylenediamine dihydrochloride in water (NED Solution). 3. Spleen cell culture supernatant. 4. Equipment: 96-well flat-bottom polystyrene microtiter plate and microplate spectrophotometer.

2.16.3 Determination of Nitrite Concentrations in Experimental Samples

3

1. Nitrite Standard reference curve

Methods

3.1 Leishmania spp. Culture

1. On day 1, thaw the parasites stored in liquid nitrogen by gently moving the cryotube back and forth in a 37  C water bath. 2. When only a small piece of ice remains, suspend the entire cryotube content of parasites in 10 mL of sterile PBS in a 15 mL sterile polypropylene tube. 3. Centrifuge at 2000  g for 15 min at room temperature. 4. Aspirate the supernatant and suspend the parasites with 3 mL of Complete Schneider’s Medium. 5. Transfer the parasites to a polystyrene sterile flask with a growth area of 25 cm2 and incubate at 24  C. 6. On day 2, if the parasites are starting to move, add 7 mL of Complete Schneider’s Medium to give a final volume of

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Table 1 Morphological criteria used to discriminate between promastigote stages of Leishmania parasite Procyclic

Body length: 6.5–11.5 μm Body width: >1.5 μm Flagellum 1.5 μm Flagellum  body length is variable

Leptomonad

Body length: 6.5–11.5 μm Body width: >1.5 μm Flagellum > body length

Metacyclic

Body length: 6.5–11.5 μm Body width: 1.5 μm Flagellum > body length

10 mL. Evaluate the growth of the parasites daily (Table 1) in order to choose the desired moment to prepare the soluble Leishmania antigen or to prepare suspensions to challenge the immunized mice (see Note 6). 3.2 Soluble Leishmania Antigen (SLA)

1. Initially, dilute the promastigote forms in Isoton solution according to the culture concentration. Count the parasites in the stationary phase of growth using a Neubauer chamber and adjust the concentration to 2  108 promastigotes/mL (see Note 12). 2. Transfer the parasites to a 15 mL sterile polypropylene tube, centrifuge them at 2000  g for 15 min at 4  C, and suspend the pellet in 10 mL of cold sterile PBS. 3. Wash the parasite culture 3 times with PBS with centrifugation (2000  g for 15 min at 4  C between wash steps) to remove traces of glycerol. After the third centrifugation, discard the supernatant and suspend the pellet containing the parasites in 10 mL of sterile PBS. 4. Immerse the tube containing the parasites in liquid nitrogen and wait until the sample freezes. Then place the tube at 37  C in a water bath to thaw the sample. Repeat this freeze/thaw cycle for 6 times. 5. Use a probe sonicator to improve parasite lysis. Do 6 cycles of ultrasonication for 30 s each, at 38 MHz, with intervals of 30 s. Always keep the sample on ice to avoid heat denaturation. 6. Finally, centrifuge the lysate at 8000  g for 20 min, collect the supernatant containing the soluble antigens, and analyze the protein concentration by BCA protein assay. Prepare aliquots containing 500 μL of SLA per 1.5 mL sterile polypropylene microtube and store at 80  C until needed (see Note 13).

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3.3 Evaluation of Antigen Concentration

1. Measure the concentration of recombinant proteins or SLA preparation using the “Pierce™ BCA Protein Assay” Kit (or equivalent), according to the manufacturer’s instructions.

3.4 Preparation of Injections

1. Prepare the injections as shown in Table 2 (see Note 5).

3.5

1. Hold gently the left hind foot of the mouse to be injected and pull it through the acrylic mouse restrainer. On day 0, inject the footpad subcutaneously with the needle bevel up (50 μL/ footpad).

2. Keep all the samples on ice until needed.

Immunization

2. Repeat the injections on day 14 and on day 28. 3. Evaluate the immune response induced by the injections at 30 days after the third injection (Fig. 1).

Table 2 Preparation of materials for animal injections Preparation

Antigen

Adjuvant

Buffer

Final volume for injection per mouse

Vaccine

RCP (25 μg) Saponin (25 μg) PBS, pH 7.4 50 μL

Vaccine

RCP (25 μg) –

PBS, pH 7.4 50 μL

Control adjuvant –

Saponin (25 μg) PBS, pH 7.4 50 μL

Control

–

–

PBS, pH 7.4 50 μL

Fig. 1 Experimental scheme for mouse immunization with vaccine and subsequent challenge with Leishmania parasites and the sampling time points for serum antibody, spleen cell culture, and other organ removal for determining parasite load

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1. Grow the Leishmania parasites as described in Subheading 3.1. 2. Analyze the Leishmania culture using an inverse microscope to identify promastigotes (see Note 6). 3. Transfer the parasites to a 15 mL sterile polypropylene tube, centrifuge them at 2000  g for 15 min at 4  C, and suspend the pellet in sterile PBS (see Note 14). 4. According to the culture concentration, dilute the promastigote forms in Isoton solution. Count the parasites in the stationary phase of growth using a Neubauer chamber (see Note 12). 5. Adjust the concentration of the inoculum with PBS according to the Leishmania species used (see Note 15). 6. Hold gently the right hind foot of the mouse to be injected and pull it through the acrylic mouse restrainer. 7. At 30 days after the last injection/immunization, challenge 8 animals subcutaneously in the footpad, with a volume of 50 μL/footpad. Evaluate vaccine protection 45 days later (Fig. 1).

3.7 Preparation of Mouse Serum After Immunization or After Immunization and Challenge

1. Restrain the mouse by the scruff method. Expose the ventral side of the animal, tilting the head down at a slight angle. Place the sterile needle, bevel up and at 30 angle, in the lower right or left quadrant of the animals’ abdomen. Aspirate to ensure proper placement and inject 100 μL per 20 g of mouse weight of anesthetic cocktail intraperitoneally. 2. Lay the anesthetized mouse on its back. Stretch out a forelimb, pin the front foot, and swab the site with 70% (v/v) ethanol. Make a deep incision in the armpit at the side of the thorax using a scalpel blade and scissors. Hold the skin at the posterior part of the incision using forceps to create a cupped area. 3. Incise the blood vessels in the area with a scalpel blade and collect blood as it pools using a polyethylene Pasteur pipette (Fig. 2). 4. Place the blood in a sterile polypropylene microtube and incubate for 30 min at room temperature. 5. Centrifuge the blood at 1500  g for 10 min at 4  C. 6. Transfer the supernatant (serum) to a new sterile polypropylene microtube and store it at 20  C until needed.

3.8 Isolation of Mouse Splenocytes for Cell Culture

1. Euthanize the mice by decapitation using a heavy pair of scissors at 30 days after the last injection or at 45 days after parasite challenge. 2. Make a 2 cm incision at the peritoneal wall with surgical scissors and identify each organ (Fig. 2). Grasp as much of the spleen as

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Fig. 2 Necropsy of the mouse showing main internal organs sampled for parasite loads. (Adapted from https://veteriankey.com/necropsy-of-the-mouse/. X marks the position where pooled blood is collected)

possible with dressing forceps. Gently pull the spleen free of the peritoneum, tearing the connective tissue behind the spleen. Store the spleen in a 50 mL conical sterile polypropylene tube containing 1 mL of RPMI 1640 medium. Keep the organ on ice until needed. 3. Transfer the spleen into a 70 μm cell strainer fitted in a 50 mL sterile polypropylene tube. Gently cut the spleen into small pieces using a pair of sterile scissors. 4. Use a 1 mL syringe plunger to gently press the spleen tissue pieces through the strainer, while continuously adding RPMI 1640 medium. After complete maceration, adjust the final volume to 10 mL with RPMI 1640 medium (see Note 16). 5. Centrifuge the cells at 400  g for 10 min and discard the supernatants. Disaggregate the cell pellet by flicking the tube gently with fingers, suspend the cells in 3 mL of Erythrocyte Lysis Solution per tube, and incubate at room temperature for

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4 min. Stop the lysis by adding 7 mL of RPMI 1640 medium (see Note 17). 6. Centrifuge the cells at 400  g for 10 min and suspend them in 1 mL of RPMI 1640 Complete Medium. 7. Dilute the cells 100 in RPMI 1640 medium using 990 μL of medium to 10 μL of cells. Mix the diluted cells using a pipette and add 10 μL of this dilution into a Neubauer chamber to count them using a microscope. 8. Adjust the concentration of cells to 1  107 cells/mL using complete RPMI 1640 medium. 9. The splenocytes are now ready for in vitro analyses. 3.9 Stimuli for Splenocytes Culture

Spleen cell cultures are used for two experimental evaluations: (1) of cytokine production in the supernatants, for example, IFN-γ, IL-4, IL-10, IL-12, GM-CSF and (2) of IFN-γ production and the cellular source of this cytokine from CD4+ and/or CD8+ T cells. These evaluations are done with flow cytometry and ELISA protocols as described in Subheadings 3.13 and 3.14, respectively. 1. Add 100 μL/well of spleen cell suspension (1  106 cells/well) to 24-well flat-bottom culture plates and 900 μL of RPMI 1640 Complete Medium as the negative controls. To stimulate the cells, make a suspension containing 100 μL of the cell suspension plus 25 μg/mL of SLA or 20 μg/mL of each recombinant protein (RCP or the individual proteins that provided the peptide epitopes used to produce the RCP) and complete the final volume to 1 mL with RPMI 1640 Complete Medium. For the positive control, add 100 μL of the cell suspension, 5 μg/mL of ConA, and complete the final volume to 1 mL with complete RPMI 1640 medium. 2. As IFN-ɣ cytokine is very important to describe the profile of the immune response, it is necessary to analyze the cellular source of this cytokine in vaccinated mice. To determine the main source of IFN-ɣ, add separately 1  106 spleen cells/well from immunized mice or from immunized and infected animals. Add 25 μg/mL of SLA and 5 μg/mL of anti-CD4+, antiCD8+, or anti-IL12 monoclonal antibodies per well. Complete the final volume to 1 mL with RPMI 1640 Complete Medium. 3. Incubate the plates at 37  C with 5% (v/v) CO2 for 48 h. 4. Remove the supernatant to analyze general cytokine production and the involvement of CD4+, CD8+ T cells, and IL-12 dependence on IFN-ɣ production, using flow cytometry (Subheading 3.13) and cytokine ELISA (Subheading 3.14).

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3.10 Isolation of Spleen, Liver, and Lymph Nodes for Parasite Culture

These organs are removed at 45 days after parasite challenge. 1. Decapitate the mouse and make a 2 cm incision at the peritoneal wall with surgical scissors as described in Steps 1 and 2 of Subheading 3.8. Grasp as much of the liver as possible with dressing forceps (Fig. 2). Gently pull the liver free of the peritoneum, tearing the connective tissue behind. Place the liver in a 15 mL conical sterile polypropylene tube and weigh the organ using an analytical balance. Add 10 mL of Complete Schneider’s Medium for each gram of organ. Macerate the organ completely with a tissue grinder and return the sample to the 15 mL tube. Keep on ice until needed. 2. To isolate the lymph node (Fig. 2), extend the incisions down the infected leg (right side) and past the ankle joint to expose the popliteal lymph node behind the knee (see Note 18). Grasp and gently pull the lymph node. Place the popliteal lymph node in a 1.5 mL sterile polypropylene tube and weigh the organ using an analytical balance. Add 10 mL of Complete Schneider’s Medium for each gram of organ. Macerate the organ completely with a tissue grinder and return the sample to the 1.5 mL tube. Keep on ice until needed. 3. For the infected spleen, remove one-quarter of each previously isolated organ, place it in a 1.5 mL sterile polypropylene tube and weigh the organ using an analytical balance. Add 10 mL of Complete Schneider’s Medium for each gram of organ. Macerate the organ completely with a tissue grinder and return the sample to the 1.5 mL tube. Keep on ice until needed.

3.11 Isolation of Bone Marrow for Parasite Culture

Bone marrows are isolated at 45 days after parasite challenge from the same decapitated mouse carcasses (Fig. 2). 1. Cut the quadriceps muscle anchored to proximal end of the femur to expose the anterior side of the femur and pin out from the leg, placing the pin at a 45-degree angle from the board. 2. With the blade of the scissors against the posterior side of the femur, cut the hamstrings away from the knee joint. 3. Pull back the skin and the hamstring muscles anchored to proximal end of the femur to expose the posterior side of the femur and pin out from the leg, placing the pin at a 45-degree angle from the board. 4. With the forceps, hold the distal end of the femur, just above the knee joint. Guide the blades of the scissors on either side of the femoral shaft toward the hip joint. Be careful not to cut into the femur itself. 5. After reaching the femoral head, indicated by the scissors opening slightly, twist the scissors with the top blade of the

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scissors moving directly over the femoral head to dislocate the femur. Be careful not to snap the bone below the femoral head. 6. Grasp the top of the femoral shaft with the forceps and use scissors to cut the soft tissue away from the femoral head to release it from the acetabulum. 7. Pull the entire leg bone, including femur, knee, and tibia, up and away from the body, carefully cutting away, with scissors, the connective tissue and muscle connecting the leg to the skin. 8. Overextend the ankle joint and use the scissors in a twisting motion to dislocate the tibia. 9. Grasp the distal end of the tibia with forceps, taking care not to sever the tendons, pull the tibia up and away from the body and the pinboard. 10. Cut, with scissors, any remaining connective tissue attaching the long bone to the mouse at the knee. 11. Remove any additional muscle or connective tissue attached to the femur and the tibia, using forceps and scissors. 12. Using the forceps, grasp the femur with the anterior side facing away and the proximal end (femoral head end) down. Guide the scissors up the femoral shaft to the condyles. Gently rotate the scissors back and forth to remove the condyles, the patella, and the epiphysis to expose the metaphysis. 13. Using the forceps, grasp the tibia with the anterior side facing away and the distal end (ankle end) down. If the tibial epiphysis is intact, guide the scissors up the tibia shaft to the condyles. Gently rotate the scissors back and forth to remove the condyles and epiphysis to expose the metaphysis. 14. Push an 18 g needle through the bottom of a 0.5 mL sterile polypropylene microcentrifuge tube and place the long bones into the tube, knee-end down and close the lid. 15. Nest the 0.5 mL microcentrifuge tube in a 1.5 mL sterile polypropylene microcentrifuge tube and centrifuge the nested tubes at 10,000  g for 15 s. Visually inspect to confirm if the bone marrow has been centrifuged out of the bones (see Note 19). 16. Discard the 0.5 mL microcentrifuge tube with the bones and suspend the bone marrow in 1 mL of sterile cold PBS [21]. 3.12 Evaluation of the Parasite Load in Mouse Organs

1. Place 20 μL of each macerated organ sample (spleen, liver, lymph node, and bone marrow) into wells of the first column of the 96-well flat-bottom polystyrene cell culture plate. 2. Add 180 μL of Complete Schneider’s Medium into all wells, including in the macerated samples. Homogenize the samples with medium using a pipette and serially dilute tenfold (i.e.,

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20 μL into 180 μL across the rows) from 10 Discard the last 20 μL remaining in the pipette.

1

to 10

12

.

3. Incubate the plates at 24  C and quantify parasite by visual counting by microscopy after 5 to 7 days (see Note 20). 4. Express the data as the negative log of the titer (i.e., the dilution corresponding to the last positive well) adjusted per milligram of organ. 3.13 Flow Cytometry for Analysis of Cytokines

1. Add 50 μL/well of spleen cells suspension (5  105 cells/well) to 96-well round-bottom culture plates. Add CM-BLAST medium to make the final volume of each well to 200 μL. Add 50 μL of the cell suspension and 150 μL of CM-BLAST medium as the negative control. To stimulate the cells, make a suspension containing 50 μL of the cell suspension plus 25 μg/ mL of SLA or 20 μg/mL of each recombinant protein and complete the final volume to 200 μL with CM-BLAST medium. For the positive control, initially, during 44 h of culture, add just 50 μL of the cell suspension and 150 μL of CM-BLAST medium. 2. Incubate the plates at 37  C with 5% (v/v) CO2. 3. At the 44-h time point, add 25 ng/mL of PMA and 1 μg/mL of ionomycin into the positive control wells. Also, add 10 μL of 200 μg/mL of Brefeldin A into all wells, including the controls and experimental groups, and homogenize using a pipette to obtain a final concentration of 2 μg of Brefeldin A per well. Return the plates to incubation at 37 with 5% (v/v) CO2. 4. Approximately 15 min before the end of the 48 h incubation, add 20 μL of 20 mM EDTA solution into all wells to give a final concentration of 2 mM/well and return the plates to incubate at 37  C with 5% (v/v) CO2. 5. Remove the plates from the incubator at the 48-h timepoint and centrifuge at 400  g for 5 min at 4  C (acceleration 7; deceleration 5). Discard the supernatant and suspend the cells carefully by vortex-mixing. Add 200 μL of cold PBS and centrifuge the plate at 400  g for 5 min at 4  C (acceleration 7; deceleration 5) (see Note 21). Discard the supernatant by pouring out of the plate. Use a pipette to carefully suspend and homogenize the cells in 50 μL/well of FVS450 diluted 1/500 in cold PBS. Incubate for 15 min in the dark at room temperature. 6. Add 100 μL/well of PBS-W buffer and centrifuge at 400  g for 5 min at 4  C (acceleration 7; deceleration 5). 7. Discard the supernatant by pouring out of the plate and suspend the cells carefully by gentle vortex-mixing. Add 30 μL/

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well of the surface antibody mix and incubate for 30 min in the dark at room temperature (see Note 22). 8. Add 150 μL/well of cold Cytometry Lysis Solution (1) to lyse any remaining erythrocytes in the sample, mix thoroughly using a pipette, and incubate for 5 min at room temperature in the dark. Homogenize carefully by pipette and incubate for another 5 min at room temperature in the dark. 9. Centrifuge at 400  g for 5 min at 4  C (acceleration 7; deceleration 5). Discard the supernatant by pouring from the plate and suspend the cells carefully by gentle vortex-mixing. Add 200 μL/well of cold PBS-P, homogenize 3 times by pipette, and incubate the plates for 10 min in the dark at room temperature. 10. Centrifuge at 400  g for 5 min at 4  C (acceleration 7; deceleration 5). Discard the supernatant by pouring from the plate and suspend the cells carefully by gentle vortex-mixing. Add 30 μL/well of intracellular anti-cytokine antibody mix and vortex-mix with care (see Note 22). Incubate for 30 min in the dark at room temperature. 11. Add 200 μL/well of cold PBS-W buffer and centrifuge the cells at 450  g for 5 min at 4  C (acceleration 7; deceleration 5). Discard the supernatant by pouring the plate and suspend the cells by gentle vortex-mixing. Add 200 μL of cold PBS and transfer to flow cytometry tubes. 12. Use compensation beads to set voltages and gating parameters to obtain accurate fluorescence signals. Read 30,000 events per sample using a BD LSR Fortessa Cell Analyzer™ cytometer (or equivalent) and BD FACSDiva™ software package (or equivalent). Analyze the cytometry data by FlowJo™ software package (or equivalent). 3.14 Enzyme-Linked Immunosorbent Assay for Cytokines

1. Remove 900 μL of supernatant from each spleen cell culture wells incubated for 48-h described in Subheading 3.9. 2. Quantify the cytokines using the “BD OptEIA TM Set Mouse” Kits (or equivalents) following the manufacturer’s instructions (see Note 23). 3. Analyze the data using GraphPad Prism™.

3.15 Enzyme-Linked Immunosorbent Assay for Antibodies

1. Determine through titration curves what are the ideal antigen concentrations required to coat each well of a 96-well ELISA plate(s) (see Note 24). 2. Dilute the specific antigens (recombinant proteins and SLA) in Coating Buffer (according to the concentrations determined in step 1) and add 100 μL per well. Seal the plate with parafilm and store overnight at 4  C.

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3. Aspirate the wells and wash 3 times with a minimum of 300 μL/well of Wash Buffer. After the last wash, invert the plate and blot on absorbent paper to remove any residual buffer. 4. Add 200 μL/well of Blocking Solution to block the plastic spaces in the wells. Incubate at 37  C for 1 h. 5. Repeat step 3. 6. Dilute the mouse sera 1/50 in Incubation Buffer and add 100 μL/well. Incubate the plates at 37  C for 1 h. 7. Repeat step 3. 8. Dilute the anti-mouse IgG, IgG1, and IgG2a horseradishperoxidase conjugated antibodies in Incubation Buffer using the manufacturer’s recommended dilutions. Add 100 μL/well and incubate for 1 h at 37  C. 9. Repeat step 3, but wash 5 times in total. 10. Add 100 μL of Substrate Solution to each well. Incubate the plate for 30 min at room temperature in the dark. 11. Add 25 μL of Stop Solution to each well. 12. Read absorbance at λ 450 nm within 30 min of stopping the reaction. If wavelength correction is available, subtract the absorbance at λ 570 nm from the absorbance at λ 450 nm. 3.16 Measurement of Nitrite Using Griess Reaction 3.16.1 Preparation of a Nitrite Standard Reference Curve

3.16.2 Nitrite Measurement (Griess Reaction)

1. Designate 2 columns (16 wells) in the 96-flat-bottom well plate for the Nitrite Standard reference curve. Dispense 50 μL of complete RPMI 1640 medium into the wells in rows B–H. 2. Add 100 μL of the 100 μM nitrite solution to the remaining 2 wells in row A. 3. Immediately prepare 6 serial twofold dilutions (50 μL/well) in duplicate down the plate to generate the Nitrite Standard reference curve (100, 50, 25, 12.5, 6.25, 3.13, and 1.56 μM), discarding 50 μL from the final 1.56 μM set of wells. Do not add any nitrite solution to the last set of wells (0 μM). 1. Equilibrate the Sulfanilamide Solution and NED Solution to room temperature (15–30 min). 2. Add 50 μL of each experimental sample to wells in duplicate or triplicate. 3. Dispense 50 μL of the Sulfanilamide Solution with a pipette to all experimental samples and wells containing the dilution series for the Nitrite Standard reference curve. 4. Incubate for 5–10 min at room temperature, protected from light. 5. Dispense 50 μL of the NED Solution with a pipette to all wells.

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6. Incubate at room temperature for 5–10 min, protected from light. 7. Measure absorbance within 30 min in a plate reader with a filter between λ 520 nm and λ 550 nm. 3.16.3 Determination of Nitrite Concentrations in Experimental Samples

1. Generate a Nitrite Standard reference curve by plotting the average absorbance value of each concentration of the Nitrite Standard as a function of “Y” with nitrite concentration as a function of “X.” 2. Determine the average absorbance value of each experimental sample. Determine its concentration by comparison to the Nitrite Standard reference curve.

4

Notes 1. The fetal bovine serum must be heated for 30 min at 56  C in a water bath with mixing to inactivate complement proteins. 2. To prepare 1 L of Phosphate-Buffered Saline, dissolve 8.0 g of NaCl, 1.16 g of Na2HPO4, 0.2 g of KH2PO4, 0.2 g of KCL in water. pH falls within the range 7.0–7.4. 3. The “Pierce™ BCA Protein Assay” Kit is a popular method for colorimetric detection and quantitation of total protein. The kit provides greater protein-to-protein uniformity than manual quantitation methods and it is therefore recommended. However, if the kit is not available, alternative protein quantification methods can be used. 4. Different Th1 response adjuvants can be used in vaccine formulations. The choice of adjuvant will depend on the application in different animal models or in humans. Examples of Th1 adjuvants are MonoPhosphoryl Lipid A (MPLA), Glucopyranosyl Lipid A (GLA), small OligoDeoxyNucleotides (ODN) with unmethylated CpG (CpG-ODN) or Complete Freund’s Adjuvant. 5. The murine immunization protocol described in this chapter resembles those in the literature, since the concentrations and doses of the immunogen are well described for mice. However, there may be variations in the concentrations of the immunogenic proteins plus adjuvants, as well as differences in the intervals between each dose. 6. Leishmania parasites can change their morphology during the incubation period (Table 1) [22]. Metacyclic forms are the most infectious phase of the parasite and are best for carrying out experimental infections. They are frequently observed after 4 to 5 days of cultivation. Therefore, evaluation of the parasite culture must be done, aiming to find a high concentration of

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these forms to prepare the inoculum. Also, it is important to use parasites from the initial culture passages to infect mice. Do not use parasites that have gone through more than 5 passages. 7. All cell stimuli (SLA, recombinant proteins, concanavalin A, PMA, ionomycin) must be kept sterile until needed. 8. Flow cytometry is widely used and several antibodies against a wide variety of other cellular markers are available for use. 9. To prepare the 10 Cytometry Lysis Solution, add the heparin while the citrate is dissolving in water and then add the formaldehyde slowly. To finish the solution, add the diethylene glycol by dripping. 10. Coating buffer, assay diluent, and wash buffer, used in the ELISA test, must be freshly prepared or used within 3 days of preparation, if stored at 2–8  C. 11. Substrate solution using O-phenylenediamine, hydrogen peroxide, and Citrate Buffer is an alternative to TetraMethylBenzidine (TMB) and hydrogen peroxide, indicated by the manufacturer. 12. The method for using a Neubauer chamber (hemocytometer) can be found at https://laboratoryinfo.com/manual-cellcounting-neubauer-chamber. 13. If the SLA is to be stored for more than 15 days, add a protease inhibitor cocktail according to the manufacturer’s recommendations. 14. The PBS volume to suspend the parasites for challenge will depend on the culture concentration. To 10 mL of culture with high number of parasites, suspend the pellet in 100 μL of PBS. 15. The parasite inoculum to challenge mice will depend on the Leishmania species to be used. The usual inoculum for infection is between 1  106 to 1  107 parasites/mouse. 16. This is a very critical step: if the operator is not gentle, there is a risk of damaging the cells and releasing DNA, which makes the cells stick to each other and reduce their viability. 17. The incubation time is very critical. If the cells are incubated for longer, the white blood cells will lyse. Thus, it must be exactly 4 min. 18. Lymph nodes can be difficult to find. They are generally “pearly white” and/or translucent in color and can be blended into the surrounding fat. The popliteal lymph node is generally small in healthy mice, but it is enlarged in immunized or infected animals. 19. After centrifugation, the bones without the bone marrow should appear white and the bone marrow cells should be a visual pellet in the largest tube.

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20. The incubation period will depend on the Leishmania species used in the infection, but the majority can be evaluated within 5 to 7 days. 21. Washing cells with PBS is very important to remove the CM-BLAST, as it reacts with the FVS450 dye and changes the reliability of the results. 22. The dilution of each antibody is done according to the manufacturer’s instructions in cold PBS-W plus 5% (v/v) FBS. 23. We do ELISA to evaluate the production of cytokines, for example, IFN-γ, IL-4, IL-10, IL-12, and GM-CSF, in general samples and to analyze the nature of the cells mainly responsible for IFN-γ secretion. To do this, we block the spleen cells with anti-CD4+ or anti-CD8+ antibodies and measure the IFN-γ secreted into the supernatant after 48 h of culture. Addition of anti-IL-12 antibody is used as a positive control for IFN-γ production. For recombinant proteins, normally we have CD4+ as the main source of IFN-γ. When we block CD4+, the levels of IFN-γ decrease, which we measure using the specific ELISA. 24. To generate the titration curve, place different concentrations of antigens (SLA or recombinant proteins) in a 96-well flatbottom microtiter plate. A range of 0.5 to 2 μg/well is recommended.

Acknowledgements VL vaccine research is supported by grant MR/R005850/1 from the Medical Research Council (VAccine deveLopment for complex Intracellular neglecteD pAThogEns—VALIDATE), UK, and grant APQ-408675/2018-7 from the Conselho Nacional de Desenvolvimento Cientı´fico e Tecnolo´gico (CNPq), Brazil. References 1. World Health Organisation (2018). http:// www.who.int/topics/leishmaniasis/en/. Accessed 30 Mar 2020 2. Desjeux P (2004) Leishmaniasis. Nat Rev Microbiol 2(9):692. https://doi.org/10. 1038/nrmicro981 3. Kane MM et al (2000) Leishmania parasites and their ploys to disrupt macrophage activation. Curr Opin Hematol 7(1):26–31. https:// doi.org/10.1097/00062752200001000-00006 4. Reis AB et al (2010) Immunity to Leishmania and the rational search for vaccines against canine leishmaniasis. Trends Parasitol 26

(7):341–349. https://doi.org/10.1016/j.pt. 2010.04.005 5. Martins VT et al (2017) A recombinant chimeric protein composed of human and micespecific CD4(+) and CD8(+) T-cell epitopes protects against visceral leishmaniasis. Parasite Immunol 39(1). https://doi.org/10.1111/ pim.12359 6. Green SJ et al (1990) Leishmania major amastigotes initiate the L-arginine-dependent killing mechanism in IFN-gamma-stimulated macrophages by induction of tumor necrosis factor-alpha. J Immunol 145(12):4290–4297

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7. Blackwell JM (1996) Genetic susceptibility to leishmanial infections: studies in mice and man. Parasitology 112(Suppl):S67–S74 8. Pirdel L et al (2017) A non-pathogenic recombinant Leishmania expressing Lipophosphoglycan 3 against experimental infection with Leishmania infantum. Scand J Immunol 86 (1):15–22. https://doi.org/10.1111/sji. 12557 9. Banerjee A et al (2018) Live attenuated Leishmania donovani Centrin gene-deleted parasites induce IL-23-dependent IL-17protective immune response against visceral Leishmaniasis in a murine model. J Immunol 200(1):163–176. https://doi.org/10.4049/ jimmunol.1700674 10. Fiuza JA et al (2016) Intradermal immunization of Leishmania donovani Centrin Knockout parasites in combination with salivary protein LJM19 from sand Fly vector induces a durable protective immune response in hamsters. PLoS Negl Trop Dis 10(1):e0004322. https://doi.org/10.1371/journal.pntd. 0004322 11. Keerti et al (2018) Immunotherapeutic potential of Leishmania (Leishmania) donovani Th1 stimulatory proteins against experimental visceral leishmaniasis. Vaccine 36 (17):2293–2299. https://doi.org/10.1016/j. vaccine.2018.03.027 12. Lage DP et al (2015) Prophylactic properties of a Leishmania-specific hypothetical protein in a murine model of visceral leishmaniasis. Parasite Immunol 37(12):646–656. https://doi. org/10.1111/pim.12287 13. Ribeiro PAF et al (2019) Immunogenicity and protective efficacy of a new Leishmania hypothetical protein applied as a DNA vaccine or in a recombinant form against Leishmania infantum infection. Mol Immunol 106:108–118. https://doi.org/10.1016/j.molimm.2018. 12.025

14. Moafi M et al (2019) Leishmania vaccines entered in clinical trials: a review of literature. Int J Prev Med 10:95. https://doi.org/10. 4103/ijpvm.IJPVM_116_18 15. Ratnapriya S et al (2019) Visceral leishmaniasis: an overview of vaccine adjuvants and their applications. Vaccine 37(27):3505–3519. https://doi.org/10.1016/j.vaccine.2019.04. 092 16. Jain K et al (2015) Vaccines for visceral leishmaniasis: a review. J Immunol Methods 422:1–12. https://doi.org/10.1016/j.jim. 2015.03.017 17. Duarte MC et al (2016) A vaccine combining two Leishmania braziliensis proteins offers heterologous protection against Leishmania infantum infection. Mol Immunol 76:70–79. https://doi.org/10.1016/j.molimm.2016. 06.014 18. Garde E et al (2018) Analysis of the antigenic and prophylactic properties of the Leishmania translation initiation factors eIF2 and eIF2B in natural and experimental Leishmaniasis. Front Cell Infect Microbiol 8:112. https://doi.org/ 10.3389/fcimb.2018.00112 19. Lage DP et al (2020) Liposomal formulation of ChimeraT, a multiple T-cell epitope-containing recombinant protein, is a candidate vaccine for human visceral Leishmaniasis. Vaccines (Basel) 8(2):289. https://doi.org/10.3390/ vaccines8020289 20. https://animal.research.uiowa.edu/iacucguidelines-anesthesia. AGAo 21. Amend SR et al (2016) Murine hind limb long bone dissection and bone marrow isolation. J Vis Exp 110:e53936. https://doi.org/10. 3791/53936 22. Lei SM et al (2010) Population changes in Leishmania chagasi promastigote developmental stages due to serial passage. J Parasitol 96 (6):1134–1138. https://doi.org/10.1645/ ge-2566.1

Chapter 26 Production of Oral Vaccines Based on Virus-Like Particles Pseudotyped with Protozoan-Surface Proteins Lucı´a Lara Rupil, Marianela del Carmen Serradell, and Hugo Daniel Luja´n Abstract Giardia lamblia is the only known parasite that can inhabit the harsh upper gastrointestinal tract, where most of the digestive proteases are secreted. Intestinal and free-living protozoa express surface proteins containing an extraordinarily high percentage of cysteine. These cysteine-rich variant-specific surface proteins (VSPs) form a dense coat on the entire surface of Giardia trophozoites, that coat protects the parasite inside the host intestine. VSPs not only are resistant to proteolytic digestion, extreme pH and temperatures, but also stimulate host immune responses. These properties can be used to protect as well as to increase the immunogenicity of vaccine antigens for oral administration. The incorporation of VSPs onto virus-like particles bearing viral antigens allows oral administration of these vaccines, protecting the antigens from degradation and generating robust and protective immune responses. In this chapter we describe the development of this versatile vaccine platform for the generation of safe, stable, and efficient oral vaccines, including their production and validation, as well as the characterization of immune response to oral immunization. Key words Oral vaccines, Virus-like particles, Protozoan-surface proteins, Giardia lamblia, Variantspecific surface proteins (VSPs)

1

Introduction Successful vaccination against infectious diseases is considered one of the major accomplishments of medical sciences in history. Communicable diseases remain the second cause of death worldwide, despite the efficacy of vaccines in their control [1]. Development of vaccines that can be orally administered is preferable to traditional injection-based formulations for numerous reasons, including painless and safe administration. Additionally, vaccination via mucosal surfaces enables stimulation of humoral and cellular immune responses at both systemic and mucosal sites to establish broader

Lucı´a Lara Rupil and Marianela del Carmen Serradell contributed equally to this work. Sunil Thomas (ed.), Vaccine Design: Methods and Protocols, Volume 1: Vaccines for Human Diseases, Methods in Molecular Biology, vol. 2410, https://doi.org/10.1007/978-1-0716-1884-4_26, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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protection. Although the gastrointestinal tract (GIT) is readily accessible via oral administration and is the gold standard for drug delivery, only a few oral vaccines have been successfully developed [2]. One main reason for this failure is the nature of the digestive system; there the ingested antigen is degraded due to the low gastric pH and digestive enzymes of the small intestine, losing immunogenicity [1]. Consequently, for oral immunization to be effective, protein antigens must be protected from the harsh environment of the upper GIT. The early branching eukaryote Giardia lamblia, hereafter Giardia, is perhaps the only protozoan capable of colonizing the lumen of the upper small intestine of humans and many other vertebrates. In fact, this parasite is covered with a protein coat that prevents its degradation by intestinal enzymes or extreme pHs. Furthermore, to evade the host immune response, Giardia undergoes antigenic variation by continuously switching its main surface molecules [3]. These antigens belong to a family of cysteine-rich proteins called variable-specific surface proteins (VSPs), which cover the entire surface of trophozoites and constitute the interface between the parasite and the host. VSPs are integral membrane proteins consisting of an extracellular variable region rich in cysteine (mainly as CXXC motifs), a single hydrophobic transmembrane domain, and a highly conserved cytoplasmic tail. Surface proteins with the VSP signature (protein family database PF03302) are also present in other parasitic protozoa such as Entamoeba histolytica and Entamoeba dispar, which colonize the large intestine and in the free-living ciliates Paramecium tetraurelia and Tetrahymena thermophila [4]. Importantly, the adjuvant-free oral administration of VSPs, either recombinantly produced or purified from trophozoites, affords efficient vaccination against Giardia without causing any signs of giardiasis [5, 6]. Thus, VSPs remain stable and immunogenic after passage through the GIT environment and are not toxic to cells or animals. Subunit vaccines based on specific components of pathogens have the advantage of safety, allowing a more universal administration than inactivated or attenuated vaccines. Nevertheless, they tend to be less immunogenic than vaccines based on whole microorganisms. This obstacle can be overcome by using virus-like particles (VLPs), which are highly organized nanostructures that resemble a virus and self-assemble from virus-derived structural antigens, involving matrix and envelope proteins, but lacking genomic material [7]. It is well known that viruses can incorporate foreign glycoproteins to form infectious particles through a process known as pseudotyping. Pseudotyped particles or pseudovirions are thus chimeric virions that consist of a surrogate viral core with a heterologous viral envelope protein at their surface. Pseudotyped particles are usually derived from parental model viruses, such as retroviruses (murine leukemia virus, MLV, and the lentivirus

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human immunodeficiency virus, HIV) and rhabdoviruses (vesicular stomatitis virus, VSV) [8]. The retroviral Gag precursor protein drives the assembly of the retrovirus and has been used to give rise to a vast variety of chimeric particles, showing its versatility and opening the possibility of using VLPs as a delivery system. In our study, antigens of the influenza virus, which enter the body through mucosal surfaces of the respiratory tract, were used as model antigens. Influenza are enveloped viruses in which hemagglutinin (HA) is responsible for virion attachment to the target cells through recognition and binding to terminal sialic acid groups on membrane-bound proteins of the host cell, being the main target of neutralizing antibodies (NAbs). The neuraminidase (NA) catalyzes the removal of terminal sialic acid residues from viral and cellular glycoconjugates and thus facilitates virus release and efficient spread of the progeny [9]. These glycoproteins can be efficiently pseudotyped onto retrovirus-derived VLPs. It has been shown that many heterologous antigens can be addressed at the surface of retrovirus VLPs by fusing their extracellular region with the transmembrane domain and the cytoplasmic tail of the G protein of the vesicular stomatitis virus (VSV-G) [10]. To confer protease and pH-protection from digestive tract conditions, VSPs were targeted to the VLPs by fusing them to the VSV-G [4]. This allows oral administration of these vaccines and achieves an efficient immune response to flu antigens, which can protect mice from live Influenza virus challenges. We generated a versatile and efficient vaccine platform for oral administration. In this chapter we present a step-by-step development of the particles and the evaluation of the immune response to influenza antigens. It must be taken into account that by a simple cloning step, which changes HA into another antigen, a myriad of vaccines can be generated to provide protection against multiple pathogens.

2 2.1

Materials VLP Production

2.1.1 Plasmids

A four-plasmid system was designed for the construction of the VLPs: pGagYFP: The Gag protein precursor is a polyprotein consisting of matrix protein, protein p12, capsid protein, and nucleocapsid protein and has a fundamental role in retrovirus assembly at the plasma membrane [11]. Successful generation of several types of Gag-based VLPs was reported. Gag VLPs can be generated by sole expression of Gag polyprotein, Gag-Pol fusion protein, Gag fused to specific viral enzymes or accessory proteins, or by co-expression of Gag along with an envelope protein. The retrovirus VLPs used in our study for effective oral vaccination consist of pseudotyped particles whose core is composed

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of MLV-Gag fused to a fluorescent protein. The plasmid pGagYFP was constructed by cloning the cDNA sequence encoding the Gag capsid protein of the MLV-Gag without its C-terminal Pol (UniProt: P03355, amino acids 1–538), followed by a SGGGGS linker, fused to the enhanced yellow fluorescent protein (UniProt P42212, T203Y). The vector used was phCMV.pHA and pNA: The other types of viral structural determinants that compose the VLPs are the envelope proteins. For the construction of the plasmids encoding influenza glycoproteins, the full length Influenza A virus (A/Puerto Rico/8-SV14/ 1934(H1N1)) HA gene (UniProt: S5FRG0) was cloned in the phCMV vector. This is the pHA plasmid. In addition, the full length Influenza A virus (A/Puerto Rico/8-SV14/1934(H1N1)) NA gene (UniProt: H2KJ06) was cloned in the phCMV vector, resulting in the pNA plasmid. pVSP-G: Finally, we generated a plasmid containing the G. lamblia VSP. In this case, given that VSPs are type I integral membrane proteins, to locate them onto the particles, the ectodomain of a VSP protein from G. lamblia was fused to the transmembrane and cytoplasmic tail of VSV glycoprotein (VSV-G) [4]. The latter glycoprotein has been extensively used to generate chimeric viruses, pseudoviruses, and VLPs due to its broad tissue tropism, which has been exploited in many studies [10]. For the construction of the fusion protein, the ectodomain of the variant-specific surface protein 1267 (GenBank: AAA29159.1, UniProt: Q07317, amino acids 1–566) was cloned in an expression plasmid fused to a c-Myc tag (EQKLISEEDL), a six amino acids linker (SGGGGS) and the transmembrane and cytosolic domains of the glycoprotein of Vesicular stomatitis Indiana virus (strain San Juan) (UniProt: P03522, amino acids: 462–511). This fusion protein will be named VSP-G. For the generation of the stable cell line expressing VSP1267, we used the Flp-In™ system (described in Subheading 2.1.2). For this, the VSP-G construct was cloned into the expression vector pcDNA™ 5/FRT (plasmid pcDNA™ 5/FRT-1267, Subheading 2.1.2). Otherwise, if a transient transfection system is selected, the sequence can be cloned in the phCMV plasmid. When generating the vaccine vectors, the following considerations must be taken into account: – The plasmids used must be expression vectors for mammalian cells, such that the proteins are under the control of a strong promoter like cytomegalovirus (CMV). We recommend the phCMV plasmid expression vector, which is engineered to produce significantly higher expression levels than traditional human CMV promoter based constitutive expression vectors such as pcDNA3.1 (Invitrogen). The phCMV vectors contain

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a specially modified promoter/enhancer sequence that has been systematically engineered for high-level expression. In addition, the plasmid backbones of the phCMV vectors have also been optimized to allow higher plasmid yield and smaller vector size. – It should be noted that to obtain optimal results, all the sequences were synthesized with a codon optimization corresponding to Homo sapiens codon usage, matching the cell line source organism. – Bear in mind that the envelope antigens must contain a signal peptide that directs them to the plasma membrane. 2.1.2 Generation of a Cell Line That Stably Expresses VSP-G (HEK293–1267)

1. Flp-In™ 293 cell line (Invitrogen). 2. Dulbecco’s modified Eagle’s medium (DMEM) (Gibco-BRL, Bethesda, MD). 3. Complete DMEM, DMEM supplemented with 10% fetal calf serum (FCS, Gibco-BRL, Bethesda, MD), heat inactivated at 56
C for 30 min, 2 mM L-glutamine, 100 units/mL penicillin G, 100 μg/mL streptomycin. 4. 100 mg/mL hygromycin B (see Note 1). 5. pcDNA™5/FRT-1267 Invitrogen).

expression

6. pOG44 Flp-Recombinase Invitrogen).

vector

expression

vector

(V601020 (V600520

7. 1 mg/mL transfection grade linear polyethylenimine (PEI) hydrochloride solution (MW 40,000) (Polysciences Inc., Warrington, PA) (see Notes 2 and 3). 8. 6-well culture plate. 9. 0.25% trypsin-EDTA. 10. Tissue culture incubator, 37
C, 5% CO2. 11. Refrigerated microcentrifuge tube. 12. Vortex. 2.1.3 Immunofluorescence Microscopy to Validate the Correct Expression of the Antigens

1. 24-well culture plate. 2. HEK293–1267 cells. The procedure to generate this cell line that stably expresses VSP-G is described in Subheading 2.1.2. 3. Complete DMEM. 4. Treated glass coverslips (12 mm in diameter). 5. Serum-free DMEM supplemented with 2 mM glutamine. 6. Plasmid vectors encoding viral proteins: (1) pGagYFP; (2) pHA; (3) pNA (see Note 4). 7. 1 mg/mL PEI solution. 8. Tissue culture incubator, 37
C, 5% CO2.

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9. 1 PBS. 10. 4% paraformaldehyde solution. 11. 1% BSA in PBS. 12. 3% BSA in PBS 13. Primary antibody dilutions prepared in 1% BSA in PBS: mAb 7F5 for VSP1267 (described in Serradell et al. 2019), commercial mAb or pAb for HA-H1N1, commercial mAb or pAb for NA-H1N1 (see Note 5). 14. PE-conjugated secondary antibodies or other similar ones for immunofluorescence analysis diluted in 1% BSA in PBS. 15. 1 μg/mL 40 ,6-diamidino-2-phenylindole (DAPI) solution. 16. DABCO or mounting reagent. 17. Leica IRBE inverted fluorescence microscope (N.A.1.40) or a similar one. 2.1.4 Cell Transfection to Produce VLPs

1. HEK293–1267 cells or HEK293 cells. If you choose to produce the VLPs by transient transfection of the four plasmids encoding the antigens, HEK-293 cells should be used instead and the pVSP-G must be included in the transfection mix. 2. 175 cm2 T flasks. 3. Complete DMEM. 4. Sterile conical 50-mL tubes. 5. Serum-free DMEM supplemented with 2 mM glutamine. 6. Plasmid vectors encoding viral proteins: (1) pGagYFP; (2) pHA; (3) pNA (see Note 6). 7. 1 mg/mL PEI solution. 8. Tissue culture incubator, 37
C, 5% CO2.

2.1.5 VLP Purification

1. Sterile conical 50 mL tubes. 2. Table-top refrigerated centrifuge. 3. Sterile PES 0.45-μm pore filter. 4. Ultra centrifugal units Centricon® Plus-70-100 K (Millipore Cat. #UFC710008). 5. Refrigerated ultracentrifuge with a swinging bucket rotor. 6. TNE buffer: 50 mM Tris-HCl pH 7.4, 100 mM NaCl, 0.1 mM EDTA. 7. 20% sucrose solution in TNE. 8. Ultracentrifuge tubes. 9. Bradford reagent (Pierce Coomassie Plus (Bradford) Protein Assay/Thermo Cat # 23238). 10. Microcentrifuge tubes.

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VLP Validation

2.2.1 Western Blotting

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1. Equipment and buffers for western blot analysis. 2. 10% SDS-polyacrylamide gel electrophoresis (PAGE) gel. 3. 5 loading buffer (0.5 M Tris-HCl, pH 6.8; 30% v/v glycerol; 5% SDS; 5% 2-mercaptoethanol; 0.002% bromophenol blue). 4. PVDF membranes. 5. Tris-buffered saline (TBS) (50 mM Tris-HCl, 150 mM NaCl, pH 7.4). 6. 5% skim milk in TBS. 7. 0.05% v/v Tween-20 in TBS (TBS-T). 8. Primary antibody dilutions in 1% skim milk in TBS-T: antibodies described in Subheading 2.1.3 and commercial mAb or pAb anti-MLV-Gag. 9. Horseradish peroxidase (HRP)-conjugated secondary antibody dilutions in 1% skim milk in TBS-T. 10. Western blotting HRP chemiluminescent substrates (see Note 7). 11. Charged-coupled device (CCD) cameras or similar devices.

2.2.2 Hemagglutination Assay (HA)

1. 1 PBS. 2. V-bottomed 96-well plate. 3. VLPs samples. 4. Chicken red blood cells (RBC). 5. Alsever’s solution (see Note 8). 6. 0.5% chicken RBCs cell suspension. 7. Multichannel pipette.

2.2.3 Nanoparticle Tracking Analysis (NTA)

1. VLPs samples. 2. 1 PBS. 3. 0.45-μm filter unit. 4. NanoSight NS300 equipment or similar.

2.2.4 Immunoelectron Microscopy (Immuno-EM)

1. Purified VLPs. 2. 0.45-μm pore filter unit. 3. Primary specific mAb anti-VLP antigens diluted in 0.3% BSA in PBS: Antibodies described in Subheading 2.1.3 and commercial mAb anti-MLV-Gag. 4. Isotype control antibody. 5. TNE buffer (described in Subheading 2.1.5). 6. Ultracentrifuge tubes. 7. Refrigerated ultracentrifuge with a swinging bucket rotor.

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8. Glow-discharged formvar/carbon-coated cupron grids. 9. 1% uranyl acetate (pH 4.5). 10. Colloidal gold particles conjugated-secondary antibody (gold beads of 5–10-nm in diameter) diluted 0.3% BSA in PBS. 11. JEOL 100 CX electron microscope or another similar. 2.3 VLP Immune Response

1. 8-week-old BALB/c mice.

2.3.1 VLP Immunization

3. Stainless steel bulb tipped gavage needle or a flexible cannula.

2. 1 mL syringes and 25 G needles for animal injection. 4. VLPs suspension in PBS (see Note 9).

2.3.2 Ag-Specific Humoral Immune Response Analysis

1. 25 G needles for animal bleeding.

Fluid Collection

4. 18 G catheter for BAL.

2. 20 G needles for bronchoalveolar lavage (BAL). 3. 1 mL syringe. 5. Microcentrifuge tubes. 6. Several plastic cages for fecal pellet collection. 7. PBS containing protease inhibitor (Complete® Protease Inhibitor Cocktail; Roche) and 0.1% sodium azide.

Enzyme-Linked Immunosorbent Assay (ELISA) Tests

1. 96-well ELISA microtiter plate. 2. Multichannel pipette. 3. Paper towel. 4. Freshly prepared coating buffer (prepare a solution of 1.5 g/L Na2CO3, 2.93 g/L NaHCO3 pH 9.6 in distilled water). 5. Specific peptides or proteins solutions to coat the plate (5–10 μg/mL in coating buffer). In our study we employed recombinant H1N1 HA. 6. Wash solution (0.05% v/v Tween-20 in PBS 1). 7. Blocking buffer (10% w/v BSA in PBS). 8. HRP-conjugated antibodies specific for the different immunoglobulin subtypes (IgG, IgG1, IgG2, IgA) at appropriate dilutions in blocking buffer. 9. Substrate solution (3,30 ,5,50 -tetramethylbenzidine—TMB, BD Cat # 555214). 10. Stop solution (0.16 M sulfuric acid). 11. Microplate reader with a 450 nm filter.

Antibody Microneutralization Assays

1. Viable stock of A/Puerto Rico/8/1934 (H1N1) virus (see Notes 10 and 11). 2. MDCK cells (ATCC, catalog number: CCL-34).

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3. 1 PBS. 4. Complete DMEM. 5. 1 mg/mL tosyl phenylalanyl chloromethyl ketone (TPCK)treated trypsin (“TPCK-trypsin”) solution. Note: TPCK treated trypsin preparations impair the activity of contaminating chymotrypsin, which does not promote infectivity but can cause damage to cell monolayers. 6. Fresh virus diluents (see Note 12). 7. Multichannel pipette. 8. 96-well culture plate. 9. Tissue culture incubator, 37
C, 5% CO2. 10. Inactivated serum samples. 11. Inverted light microscope. 2.3.3 Ag-Specific Cellular Immune Response Analysis Splenocytes Isolation

1. Immunized mice. 2. Chamber for CO2 euthanasia. 3. 70% ethanol. 4. Styrofoam block. 5. Scissors and forceps. 6. Complete RPMI, RPMI supplemented with 10% heat inactivated FCS, 2 mM L-glutamine, 100 units/mL penicillin G, 100 μg/mL streptomycin. 7. 70-μm cell strainer. 8. Sterile conical 50 mL tubes. 9. 5 mL syringe plunger. 10. ACK lysis buffer (ammonium–chloride–potassium; 150 mM NH4Cl, 10 mM KHCO3, 0.1 mM Na2EDTA, pH 7.2–7.4). Store for up to 6 months at room temperature (RT). 11. Refrigerated centrifuge. 12. Trypan blue solution. 13. Hemocytometer. 14. Light microscope.

Cytokines Determination by CBA

1. Splenocytes from immunized mice. 2. 96-well U-bottom culture plate. 3. Complete RPMI. 4. 2 (10 μg/mL) concanavalin A (Con A) in complete RPMI. 5. 2 (2 μg/mL) recombinant HA (or other specific vaccinepeptide/protein) in complete RPMI. 6. Tissue culture incubator, 37
C, 5% CO2.

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7. CBA Mouse Th1/Th2/Th17 kit (BD Biosciences Cat. # 560485). 8. Flow cytometer. IFN-γ ELISPOT Assay

1. Standard IFN-γ ELISPOT assay kit (MabTech, Cat. #3321-2A or another similar). 2. 96-well plate for ELISpot (Millipore, Cat. MAIPSWU10 or another similar). 3. 35% ethanol. 4. Sterile 1 PBS. 5. Tissue culture incubator, 37
C, 5% CO2. 6. Blocking solution (10% FCS in PBS). 7. Purified splenocytes from immunized mice. 8. Complete RPMI. 9. 2 stimuli solutions: 2–20 μg/mL recombinant HA (or another specific vaccine antigen stimulus) and 10 μg/mL ConA (or other positive control stimuli) prepared in complete RPMI (see Note 13). 10. Wash buffer (0.1% Tween-20 in PBS). 11. Antibody and enzyme dilution buffer (0.5% FCS in PBS). 12. ELISPOT reader (AID® ELISPOT reader or other), or light microscope for manual count.

Flow Cytometry-Based Cytotoxic Assay

1. Murine AB1 malignant mesothelioma cells that express HA from Influenza A H1N1 (AB1-HA). 2. Murine AB1 malignant mesothelioma cells. 3. Complete RPMI. 4. 4 μM carboxyfluorescein succinimidyl ester (CFSE) in PBS. 5. Splenocytes from immunized mice. 6. U-bottom 96-well culture plates. 7. Tissue culture incubator, 37
C, 5% CO2. 8. 0.25% trypsin-EDTA. 9. 2% FCS in PBS. 10. 0.5 mg/mL propidium iodide. 11. Flow cytometer.

2.3.4 Challenge with Live Virus

1. Groups of vaccinated and control mice (described in Subheading 2.3.1). 2. A mouse-adapted A/Puerto Rico/8/1934 influenza virus. 3. Ketamine (75 mg/kg) + xylazine (50 mg/kg) anesthetic. 4. Infrared thermometer.

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5. Balance adequate to perform measurements of mouse weight. 6. Chamber for CO2 euthanasia.

3 3.1

Methods VLP Production

Given that chimeric VLPs can be engineered, vaccine-specific antigens may be incorporated by genetic fusion allowing immunization against peptides, peptide strings, or even whole proteins. Retroviruses utilize the cellular vesiculation pathway for virus budding/ assembly and the retrovirus Gag protein induces the spontaneous formation of microvesicles or virus-like particles when expressed in eukaryotic cells. The structural core protein Gag is synthesized on cytoplasmic polysomes and is posttranslationally targeted to the plasma membrane, where it oligomerizes. The structural protein core exits the cell through a budding process, during which Gag particles acquire their lipid envelope from the producer cell and the envelope proteins embedded in it [12]. In the present study, we generated VLPs using the budding properties of MLV-Gag protein and incorporating recombinant proteins into VLP: the protozoan derived-protein VSP1267 plus the influenza vaccine antigens HA and NA. The selection of the cell line for VLP production is very important, since enveloped-VLPs will contain the proteins expressed on its membrane. In particular, we chose mammalian cells as expression system.

3.1.1 Plasmids (Described in Subheading 2.2.1)

The chimeric VLPs that were used for oral vaccination are composed of the proteins: MLV-Gag, influenza HA and NA, and Giardia VSP1267. The sequences of the four antigens as described in the materials section were synthesized by a bioscience company and included a codon optimization step. Alternatively, the sequences can be cloned into expression vectors by standard molecular cloning techniques. Briefly, the RNA is reverse transcribed, amplified by PCR, and inserted in an expression vector. Alternatively, one or more of the plasmids can be inserted definitively in the cellular genome, creating a stable cell line. This step avoids the risk associated with introducing many plasmids during the transfection process (see Subheading 3.1.2).

3.1.2 Generation of a Cell Line That Stably Expresses VSP-G (HEK293–1267)

The hallmark of stably transfected cells is that the foreign gene becomes part of the genome and is therefore replicated. Descendants of these transfected cells will therefore also express the new gene, resulting in a stably transfected cell line. When a stable transfection is developed, selectable markers are used to distinguish transient from stable transfections; a gene for antibiotic resistance is generally used (such as the neomycin resistance gene, neo). Co-expressing the marker with the gene of interest enables the

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identification and selection of cells that have the new gene integrated into their genome. The Flp-In™ System (Invitrogen) allows integration of a gene of interest at a specific genomic location by introducing an Flp Recombination Target (FRT) site into the genome of the mammalian cell line of choice. Then, an expression vector containing the gene of interest is integrated into the genome via Flp recombinase-mediated DNA recombination at the FRT site [13]. In our study, we used the Flp-In™ 293 cell line (Invitrogen); this line is designed for rapid generation of isogenic stable cell lines that ensure high-level expression of a protein of interest. These cells contain a single stably integrated FRT site at a transcriptionally active genomic locus. There are several other options for engineering stable expression of exogenous genes in cultured cells. The classic methodology involves transfecting with a plasmid. When generating a stable cell line, the transfected plasmid undergoes recombination during chromosomal integration. A transfected circular plasmid will be linearized by a random cut within the cell. Since the recombination event can occur within any region of the plasmid, sensitive parts, such as the gene of interest, can be interrupted. Conversely, linearizing a plasmid before transfection with a restriction enzyme that cuts within the nonessential regions ensures integrity of all the necessary gene elements of the plasmid [14]. Additionally, the CRISPR-Cas9 system and transduction with retroviruses or lentiviruses can integrate genetic material into the host genome and are other methods used to achieve lifetime expression. Here, we detail the steps for the targeted integration of the expression vector pcDNA™5/FRT containing the extracellular portion of VSP1267 of G. lamblia fused to the transmembrane domain and the cytoplasmic tail of the envelope G protein of Vesicular Stomatitis Virus (VSP-G). 1. Prior to transfection, test the sensitivity of the Flp-In™ cell line to hygromycin B to more accurately determine the hygromycin B concentration required to kill untransfected cells. A suggested range of hygromycin B for Flp-In™-293 cells is 100–200 μg/mL. For effective selection, cells should be subconfluent, since confluent, nongrowing cells are resistant to the effects of antibiotics (see Note 14). Once the antibiotic concentration has been established, the transfection of the plasmids to generate the transgenic cells can be done. The ratio of pOG44 and pcDNA™5/FRT expression plasmid should be 9:1 (w/w); nevertheless, the ratio can be modified to optimize the recombination efficiency. A plate with no pOG44 as an Flp recombination control, a plate of untransfected cells as a negative control, and a plate with the pcDNA™5/FRT/CAT (Invitrogen) plasmid as a positive control should be included.

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2. Plate 800,000–1,000,000 cells per well in complete DMEM in two 6-well culture plates. Allow cells to adhere overnight. 3. After 1 day, place 9 μg pOG44 vector and 1 μg of pcDNA™5/ FRT-1267 vector in a total volume of 500 μL of serum-free DMEM in a microcentrifuge tube and mix by vortexing. 4. Add 30 μg of PEI reagent and vortex quickly. Incubate the mixture at RT for 15 min to allow the formation of the DNA: PEI complex. 5. Add 200 μL of the transfection mix to each well dropwise. Mix the content by swirling the plate. 6. Put the plate in the incubator and allow transfection to take place for 6 h. 7. After incubation, discard the medium and replace with complete medium. 8. 48 h after transfection, split the cells such that they are no more than 25% confluent. From this stage on, the culture medium must contain hygromycin in the concentration determined above. 9. Feed the cells with selective medium every 3–4 days until foci can be identified. 10. Because every Flp-In™ cell line contains a single integrated FRT site, all of the hygromycin-resistant foci that is obtained after co-transfection with the Flp-In™ expression vector and pOG44 should be isogenic, allowing polyclonal selection (i.e., the Flp-In™ expression vector should integrate into the same genomic locus in every clone; therefore, all clones should be identical). After hygromycin B selection, simply pool the hygromycin-resistant foci and screen the entire population of cells for the following phenotypes: hygromycin-resistant, Zeocin™-sensitive, and lacZ negative, then assay for expression of VSP-G. The latter can be done by immunofluorescence as detailed in the next item. Drug-resistant clones can appear in 2–5 weeks. Cell death should occur after 3–9 days in cultures transfected with the negative control plasmid. 3.1.3 Immunofluorescence Microscopy to Validate the Correct Expression of the Antigens

Analysis of the correct design of the plasmids, the expression of the proteins of interest, and the transfection conditions is recommended before producing the VLPs. Some techniques appropriate for this purpose are immunofluorescence, flow cytometry, western blotting, and transmission electron microscopy of transfected cells. Here, we describe the plaque immunofluorescence microscopy technique, since it allows us to determine the presence and subcellular localization of the antigens, simultaneously. 1. Day 0: Place the cleaned coverslips in each well and add enough sterile poly-L-lysine solution to submerge the coverslip.

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Incubate for 5 min and aspirate the solution. Wash 4–5 times with distilled water and allow to air dry in the safety cabinet. Coating with poly-L-lysine provides a positive charge for cell attachment and is used for a wide variety of cell types (see Note 15). 2. Day 0: For a 24-well plate, seed 2.5–5.0  104 HEK293–1267 cells in 0.5 mL of complete DMEM; make sure that the cells are distributed evenly and that no bubbles remain under the coverslips. Culture the cells at least 12 h before transfection (see Note 16). 3. Day 1: Transfect cells: Transfer 0.8 μg of total plasmid vector encoding the viral proteins to a microcentrifuge tube in a final volume of 50 μL of serum-free DMEM and mix well. Add 2.4 μg of PEI and mix immediately by vortexing or pipetting. Incubate at RT for 15 min. Add the mix to each well, swirl the plate to mix well, and incubate for 5–7 h. After that, replace the medium by complete DMEM. Culture the cells for 48 h for protein expression. This period depends on the expression of the protein and can vary between 24 and 72 h (see Note 17). 4. Day 3: Remove the supernatant and wash the monolayer gently with warmed PBS. Fix cells with 1 mL of a 4% paraformaldehyde solution, for 15 min at RT. Wash twice with PBS (repeat this step after each following incubation). 5. Permeabilize cells with 0.1% Triton, 100 mM glycine in PBS at RT for 15 min. 6. Blocking: 1 mL of 3% BSA in PBS at RT for 45 min. 7. Incubate the cells with the primary antibody at appropriate dilution prepared in 1% BSA in PBS at 37
C for 1 h. A volume of 0.3 mL is the minimal amount to cover the cells, you may use a larger volume if desired. 8. Incubate the cells with the corresponding fluorochromeconjugated secondary antibody prepared in 1% BSA in PBS at appropriate dilution at 37
C for 1 h. Perform this and the following steps in the dark. When choosing the antibody fluorochrome, bear in mind that the Gag polyprotein is fused to YFP. 9. Dye the nuclei with a DAPI solution in a final concentration of 1 μg/mL for 10 min. 10. Mount with mounting medium containing anti-fading agent and seal with nail polish. 11. Analyze subcellular localization of the target proteins with a fluorescence microscope.

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1. Split 8.0—10  106 HEK293T-1267 cells in 175 cm2 T flasks 1 day before transfection in 25 mL of complete DMEM. We recommend using 10 culture flasks in the production of each batch. The amounts detailed in this protocol correspond to one flask. 2. Prior to transfection, bring all reagents to RT. Cells should be transfected at 70% confluence. 3. Dilute total plasmid DNA (70 μg) in a sterile tube containing 2.5 mL serum-free DMEM (volume of transfection media should be 10% of the volume in culture T flask). Use 23.3 μg of pGagYFP, 23.3 μg of pHA, and 23.3 μg of pNA (see Note 18). 4. Add 210 μg of PEI (210 μL of PEI 1 μg/μL) to the diluted DNA. Mix immediately by vortexing or pipetting. The volume of PEI used is based on a 3:1 ratio of PEI (μg): total DNA (μg). Incubate at RT for 15 min. 5. Add the DNA/PEI mixture to the cells gently and swirl the plate to mix. 6. Return the T flasks to the incubator and allow the transfection to proceed for 5–7 h. 7. After incubation, discard the medium and replace with 25 mL of serum-free DMEM supplemented with glutamine. Add the medium gently to avoid cell detachment. 8. Incubate with 5% CO2 at 37
C for 72 h.

3.1.5 VLP Purification

Work with refrigerated equipment and keep the materials on ice. 1. Harvest the medium containing the VLPs in sterile conical tubes 72 h post-transfection. The cells are expected to be rounded or detached, since the medium lacks serum. 2. Centrifuge at 10,000  g for 10 min to remove any cellular debris. 3. Filter the medium using a 0.45-μm filter unit. 4. Concentrate the medium in Centricon® Plus-70 centrifugal filter device by placing the solution in the sample filter cup and centrifuge at 3500  g. A final volume of 8–10 mL should be obtained (see Note 19). 5. Layer the clarified harvest over a sterile 20% sucrose in TNE cushion and ultracentrifuge at 100,000  g in a swinging bucket rotor for 4 h. 6. Discard the supernatant and resuspend the VLP pellet with 1 mL sterile TNE buffer.

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7. Measure the proteins using the Bradford method. An expected yield varies between 0.5 and 2 mg of VLPs for 10 175-cm2 T flasks. 8. Aliquot in sterile microcentrifuge tubes and store at 80
C. VLP Validation

To ensure the correct and total expression of the different proteins on the VLPs, as well as the structural characteristics of the particles, it is necessary to perform different validation tests. These tests include simple techniques, such as western blotting and nanoparticle tracking analysis, and a bit more complex ones, such as immunoelectron microscopy.

3.2.1 Western Blotting

Western blotting is a technique by which an individual protein is visualized among thousands of other proteins in a given sample. The technique uses sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) to separate thousands of proteins present in a sample and specific antibody–antigen recognition to identify a particular protein. It is possible to determine the molecular weight of protein antigens and detect antigens in crude mixtures of proteins [15]. When performing the VLP analyses, the sample must be tested for the presence of all antigens, VSP, Gag, HA, and NA (see Note 20).

3.2

1. Mix 20 μL of the obtained VLP suspension containing 10 μg of proteins with 5 μL SDS-PAGE loading buffer 5 for every well to be seeded. 2. Load on a 10% SDS-PAGE gel and run. 3. Transfer onto PVDF membranes. 4. Block with 5 skim milk in TBS buffer at RT for 1 h. 5. Incubate with the primary antibodies overnight at 4
C. Wash three times with TBS-T for 5 min. 6. Incubate with the HRP-conjugated secondary antibodies at RT for 2 h and wash three times with TBS-T for 5 min. 7. Reveal by HRP chemiluminescent substrates according to the manufacturer’s instructions. 8. Obtain images from charged-coupled device cameras or similar devices. These imagers and their accompanying analytical software tools can adjust the background signal and perform densitometry. 3.2.2 Hemagglutination Assay

The hemagglutination assay is a tool used to screen and titer influenza viruses, among others, based on their ability to attach to molecules present on the surface of RBCs. A viral suspension may agglutinate the RBCs, thus preventing them from settling out of suspension. The amount of virus present can be estimated by serially diluting a virus in a 96-well plate and adding a consistent

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amount of RBCs. Specifically, hemagglutinin (HA), an envelope glycoprotein of some enveloped viruses, confers this property [16]. Here, we applied this method to evaluate the correct conformation of HA onto the chimeric retro VLPs, as well as to quantify this viral protein; thus, the HA allows validation of the VLP lot. 1. Place 50 μL 1 PBS per well in a V-bottomed 96-well plate. 2. In the first column, add 50 μL of VLP sample containing 10 μg of total proteins; if necessary, dilute in PBS. If possible, use a commercial flu vaccine as a positive control. Leave some wells with PBS only as negative controls. 3. Mix each well content and transfer 50 μL to the next well on its right (twofold serial dilutions). Repeat mixing and transferring 50 μL down the length of the plate. 4. Prepare 0.5% chicken RBCs cell suspension in Alsever’s solution (see Note 21). 5. Add 50 μL of 0.5% chicken RBCs cell suspension to each well. Mix gently. 6. Incubate at RT for 1–2 h. Do not move the plate. 7. Negative results will appear as dots in the center of V-bottomed plates. Positive results will form a uniform reddish color across the well. 8. Calculate the HA titer as the highest dilution of VLPs agglutinating RBCs. 3.2.3 Nanoparticle Tracking Analysis (NTA)

The NTA technology utilizes both light scattering and Brownian motion properties to obtain measurements of concentration and particle size distribution in a liquid suspension [17]. The NanoSight NS300 is able to analyze the size distribution and concentration of all types of nanoparticles, from 10 to 2000 nm in diameter. A laser beam is passed through the sample chamber, and the suspended particles in the path of this beam scatter the light in such a way that they can be easily seen under a 20 microscope on which a camera is mounted. The camera captures a video of the particles in their natural Brownian motion. The software tracks the particles individually and, using the Stokes–Einstein equation, calculates the hydrodynamic diameter of each one. 1. Dilute non-fixed samples of VLPs in 1 PBS at a final volume of 500 μL. 2. Filter through a 0.45-μm pore filter. 3. Run consecutive measurements of a single sample using the specific equipment NanoSight NS300 in a light scatter mode. Analyze each sample in triplicate, and measure twice each triplicate. 4. Analyze the hydrodynamic diameter and the distribution of the purified particles using the analysis software (NTA3.1).

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3.2.4 Immunoelectron Microscopy (Immuno-EM)

The high resolving power of electron microscopy (EM) permits studies at nanometer scale; therefore, scanning and transmission EM have been used to observe the surface detail of cells, tissues, and viruses. The incorporation of immunoconjugates or other ligands into electron microscopic studies permits the highresolution study of the antigenic composition of cell organelles and surfaces along with ultrastructural analyses. Singularly, or in combination, these two types of microscopic analysis have proved invaluable in the correlative study of morphological structures and their antigenic composition [18]. Immuno-EM staining of VLPs enables high resolution of virion structure, allowing us to identify and localize the correct expression of different proteins along the VLPs. Here we describe the steps for an indirect immuno-gold labeling. 1. Mix the purified and filtered VLPs with 0.1 mL of primary specific mAb or irrelevant monoclonal antibody of the same isotype (IgG1) (negative control). Determine the antibody dilution empirically to identify the best antibody concentration. Serial tenfold dilutions of antibody can be tested. 2. Incubate at 37
C for 1 h, then at 4
C for 16 h. 3. Dilute the protein–mAb complexes in 10 mL of TNE buffer and precipitate by centrifugation at 48,000  g for 1 h. 4. Discard the excess antibody present in the supernatant. 5. Resuspend the immune complexes as pellet in 0.1 mL of TNE buffer. 6. Adsorb the immune complexes onto glow-discharged formvar/carbon-coated cupron grids for 5 min. Alternatively, the VLPs can be loaded first onto the grids and the incubation with the primary antibody can be done afterwards. 7. Incubate the grids with a 1:50 dilution of colloidal gold particles conjugated-secondary antibody (gold beads of 5–10-nm in diameter). 8. Wash the grids five times with PBS and three times with distilled water. 9. Negatively stain with 1% uranyl acetate (pH 4.5) for 4 min. Remove excess liquid and allow the grid to fully air dry. 10. Visualize the preparations using a JEOL 100 CX electron microscope or a similar one.

3.3 VLP Immune Response

The type of immune response induced by a vaccine largely depends on the design of the vaccine and the type of adjuvant used. Factors such as vaccine type, product, adjuvant, and dose, as well as administration factors, such as schedule, site, route, and time of

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vaccination, are important to determine the type and intensity of the humoral and cellular vaccine responses [19]. T and B cells are crucial players of adaptive immune response, T-cells are involved in cell-mediated immune response, whereas B cells regulate humoral immunity [20]. The monitoring of both immune responses is key to vaccine validation. 3.3.1 VLP Immunization

1. Fast BALB/c mice for 4 h before immunization. At least five animals per group are recommended and two or more independent experiments should be performed (see Note 22). 2. Immunize by orogastric administration with 100 μg of VLPs diluted in 100 μL of PBS (see Note 23). 3. Administer vehicle alone (PBS) orally to the negative control group (naive). 4. A positive control group can be included, which should receive the VLPs by subcutaneous injection. 5. More doses may be administered at a two-week interval. The follow-up of the elicitation of the immune response can be done by ELISA to detect serum antigen-specific antibodies. We recommend giving a minimum of two doses.

3.3.2 Antigen-Specific Humoral Immune Response Analysis

Fluid Collection

The strength, type, and location of the humoral immune response following vaccination contribute to the vaccine protection efficacy. We examined the peripheral antibody response along with the mucosal humoral response. After immunization, we measured the anti-HA specific serum total IgG, IgG1, and IgG2 and secretory IgA levels at intestinal and respiratory mucosa at different times. 1. Collect blood weekly (before each immunization) by submandibular bleeding of mice. Allow blood to clot at RT for 30 min; separate serum by centrifugation at 3000  g and store at 80
C until analysis (see Note 24). 2. Before each immunization, place each mouse in a cage and collect 2–3 fecal pellets. With the help of a tip or a toothpick, resuspend the pellets in PBS containing protease inhibitor (Complete® Protease Inhibitor Cocktail; Roche) and 0.1% sodium azide, at a ratio of 0.1 g per 500 μL, and vortex. Leave the tubes on ice for 30 min, centrifuge at 10,000  g, and store the supernatant at 80
C until analysis. 3. Euthanize mice and collect the BAL through the trachea by injection-aspiration of 1 mL PBS with protease inhibitors. Centrifuge the liquid obtained 10,000  g and store the supernatant at 80
C until analysis (see Note 25).

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Enzyme-Linked Immunosorbent Assay (ELISA)

The ELISA is a highly versatile, sensitive, and quantitative technique. We describe an indirect ELISA to detect specific antibodies. In this protocol, solid-phase reactants are prepared by adsorbing the antigen onto plastic microtiter plates; then the solid-phase antigens are incubated with the test solution containing antibodies and a secondary reactant covalently coupled to an enzyme. Unbound conjugates are washed out and a chromogenic substrate is added [21]. ELISA was used to determine the levels of IgG, IgG1, IgG2a, or IgA antibodies against HA. Use a multichannel pipette. 1. Coat the microtiter plate with 100 μL of antigen (5 μg/mL recombinant HA) in freshly prepared coating buffer, seal the plate, and incubate overnight at 4
C. To calculate the volume of antigen, consider that each sample must be run in duplicate or triplicate (see Notes 26 and 27). 2. Remove the coating solution by flicking the plate over a sink. The remaining drops are removed by patting the plate on a paper towel. 3. Wash the plate three times by filling the wells with 200 μL of wash solution (0.05% Tween-20 in PBS). 4. Block the remaining protein-binding sites in the coated wells by adding 200 μL of blocking buffer (10% BSA in PBS) and incubating at RT for 1 h. 5. Wash the plate three times with 200 μL of wash solution. 6. Add 100 μL of serially diluted samples (serum, fecal extract or BAL) prepared in blocking buffer. For serum samples five- to tenfold dilutions are recommended, whereas for fecal extracts or BAL twofold dilutions should be prepared. Incubate at RT for 2 h. 7. Wash six times with 200 μL of wash solution. 8. Add 100 μL of HRP-conjugated specific antibodies diluted in blocking buffer and incubate at RT for 1 h. 9. Wash six times with 200 μL of wash solution. 10. Prepare the substrate solution according to the manufacturer’s instructions. 11. Dispense 100 μL of the substrate solution per well and incubate the plate at RT in the dark for 15–30 min. 12. After sufficient color development, add 50 μL of stop solution. 13. Read the absorbance at 450 nm with a plate reader. 14. The antibody titer is determined as the reciprocal of the dilution that resulted in an absorbance value that was twice that of the cut off.

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15. The baseline is determined as the mean plus twice the standard deviation of 20 absorbance values of a negative serum pool in a range of dilutions (1:100 to 1:100,000). Antibody Microneutralization Assays

The microneutralization assay using MDCK cells is a highly sensitive and specific assay for detecting virus-specific neutralizing antibodies to influenza viruses in human and animal sera. The neutralization of virus activity is based on directly visualizing the suppression of cytopathic effect under an inverted microscope. Neutralization tests with influenza differ from other viruses in that they require the use of serum-free medium and the addition of trypsin to allow productive replication of viruses. Here a microneutralization assay with a TCID50 of A/Puerto Rico/8/1934 (H1N1) was performed. For each virus to be tested once, 50 μL of sera are needed. Sera should be tested in quadruplicate, so 3 sera can be tested in each plate [22]. 1. Obtain a working dilution of virus containing 100 TCID50 (tissue culture infectious dose 50%) of A/Puerto Rico/8/ 1934 (H1N1) (see Note 28). 2. Seed MDCK cells in complete DMEM in a 96-well plate 24–48 h before the assay until confluent cell growth is obtained. Cells must be in log-phase growth for maximum virus sensitivity. 3. Obtain the complement-inactivated serum of the immunized animals to be tested by heating at 56
C for 30 min. 4. In a 96-well plate prepare the serial dilutions of serum: Pour 108 μL of virus diluent in row A and 60 μL of virus diluent in rows B–H. Add 12 μL of each serum to row A (1:10 dilution) and perform subsequent twofold dilutions by transferring 60 progressively from row to row. Discard the final 60 μL. To test one serum a number of four columns should be prepared. 5. Prepare four additional wells for each control: Control cells (CC) should contain only 120 μL virus diluent; trypsin control (TC) should contain 120 μL virus diluent supplemented with 1 μg/mL TPCK-trypsin; control virus (CV) should contain 60 μL virus diluent. 6. Dilute the virus suspension in virus diluent supplemented with 2 μg/mL TPCK-trypsin so that 50 μL contains 100 TCID. Approximately 6 mL/plate are needed. 7. Add 60 μL diluted virus to all wells (except to CC and TC). 8. Incubate the virus–serum mixtures at 37
C in 5% CO2 for 1 h. 9. Wash the cells with warm PBS three times.

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10. Inoculate 100 μL of samples of each mixture into each well containing the cells. Use a separate pipette tip for each transfer. Incubate plates in a CO2 incubator at 37
C for 2 h. 11. Remove inoculum using a multichannel pipette. Wash wells by adding and removing 250 μL of virus diluent. Add 200 μL of virus diluent supplemented with 1 μg/mL TPCK-trypsin to each well and incubate the plates at 37
C in 5% CO2 for 3–4 days. Virus diluent without trypsin (200 μL) should be added to the CC wells. 12. Examine the presence of cytopathic effect under inverted microscope and determine the HA titer as the reciprocal of the highest dilution of serum at which the virus infectivity was completely neutralized in at least two of the wells (50%). 3.3.3 Ag-Specific Cellular Immune Response Analysis

Most current vaccines mediate protection through highly specific serum antibodies and their functional characteristics as well as quantity are important for protection. However, vaccines may protect through multiple mechanisms and vaccine-induced cellmediated immune memory may be crucial for long-term protection. Similarly to the neutralizing antibodies production, the T-cellmediated immunity is a very important effector arm during the immunological response to virus [20]. Therefore, a key challenge in developing new vaccines that are effective involves the induction of optimal memory T-cell responses.

Isolation of Splenocytes

Use aseptic techniques and sterile media. 1. Euthanize mice by rapid cervical dislocation or in CO2 chamber. 2. Place the mouse on the back on a styrofoam block and spray the ventral abdomen with 70% ethanol. 3. Cut the outer skin of the peritoneum with scissors and forceps, and make a small incision in the inner skin of the peritoneum with a sterile scissor. 4. Isolate the spleen into sterile complete RPMI using sterile scissors. 5. Transfer the spleen into a 70-μm cell strainer fitted on a 50 mL polypropylene tube. 6. With a 5-mL syringe plunger, gently press the spleen through the strainer while continuously adding medium. When only the white color-associated splenic connective tissue and no red color-associated lymphoid tissue is left on the strainer, the preparation of cell suspension is completed. 7. Add media up to 25–30 mL and spin at 300  g at 4
C for 10 min.

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8. Discard the supernatant, disaggregate the cell pellet immediately, resuspend in 5 mL ACK lysis buffer, and incubate at RT for 5 min to lyse red blood cells (see Note 29). 9. Add cold complete RPMI up to 40 mL and spin at 300  g 4
C for 10 min. 10. Discard the supernatant, disaggregate the pellet, resuspend the pellet in 5 mL of media, and mix well by pipetting. 11. Leave on ice for 10 min and count at 1:20 dilution with trypan blue on a hemocytometer on a light microscope to determine their number and viability. You should obtain 95% viability or higher. 12. For the downstream processes, dilute cells immediately at a desired concentration (no more than 5  106 cell/mL, cells should not be kept at higher concentrations because they will lose viability). 13. Keep cells on ice until they are used. Cytokine Determination by Cytokine Bead Array (CBA)

Cytokines are critical mediators of both the innate and the adaptive immune responses. Detection of the spectrum of cytokines has therefore become increasingly important in the study of immune response to vaccination. There are several methods for measuring in vivo or in vitro cytokine production in mice, such as real-time quantitative polymerase chain reaction (qPCR), ELISA, CBA, detection of intracellular cytokines by flow cytometry, and immunohistochemistry. Each of these methods has advantages as well as limitations. We used the CBA technique. This flow cytometry application enables analysis of up to 30 proteins using only 25 to 50 μL of sample; thus, sample requirements and time to results are reduced in comparison to other methods such as ELISA, which enables only one protein to be analyzed per sample. The phenotype of a polarized T-cell that differentiates from a naı¨ve precursor is determined by the complex interaction of antigen-presenting cells with naı¨ve T-cells and involves a multitude of factors. To date, at least four distinct effector CD4 T-cell subsets have been described, Th1, Th2, Th17, and Treg cells, which play a critical role in orchestrating adaptive immune responses to various microorganisms. They can be distinguished by their cytokine production profiles and their functions. The elucidation of T-cell phenotype is useful for the characterization and development of immunization strategies [23]. CBATM kits provide preconfigured panels for ease of use and enable multiplex analysis of complex biological samples. The CBA Th1/Th2/Th17 cytokine kit can be used to measure IL-2, IL-4, IL-6, IFN-γ, TNF, IL-17A, and IL-10 protein levels using capture beads that contain unique amounts of a single red dye.

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1. Culture the purified splenocytes on 96-well U-bottom plates by plating 100 μL of cell suspension (2  106 cell/mL) per well. 2. Add 100 μL of the different stimuli at 2 study concentrations prepared in complete RPMI. The recommended stimuli are recombinant HA (1 μg/mL), PBS or vehicle as control, and ConA (5 μg/mL) as the positive control. The concentration of the specific antigen can be previously titrated 1–10 μg/mL. 3. Culture at 37
C and 5% CO2 for 72 h. 4. On the day of the harvest, centrifuge the plate at 1000  g for 10 min, collect the supernatant carefully to avoid touching the cell pellet, and keep at 80
C until analysis. 5. Measure the cytokines in supernatants using the CBA Mouse Th1/Th2/Th17 kit (BD Biosciences Cat. # 560485) according to the manufacturer’s instructions. 6. The unstimulated and the positive control supernatants can be pooled for each group for analysis. IFN-γ Enzyme-Linked Immunospot (ELISPOT) Assay

The ELISPOT assay is a widely used method to measure antigenspecific cytokine-producing or antibody-secreting immune cells. It is an extremely sensitive membrane-based assay that is useful for detecting secreted proteins in cellular immune responses in human and animal tissues, such as peripheral blood, lymph nodes, and spleen. The basic principle is the adsorption of the analyte to a membrane and its detection. In the IFN-γ ELISPOT assay, the cells are plated in a monolayer into 96-well PVDF membrane microtiter plates, previously coated with analyte-specific capture antibodies. The addition of the antigen to select assay wells results in the antigen presentation by APCs. Subsequent activation of the antigen-specific T-cells results in the secretion of lineage-specific cytokines which are captured by the membrane-bound antibody, resulting in a cytokine “spot” around the secreting T-cell. These membrane-bound spots are detected by adding cytokine-specific detection antibodies and a chromogenic substrate. Counting these spots, also called “spot forming units,” SFU, allows the count of the antigen-specific IFN-γ secreting T-cells within the cells plated in each well [24]. A strong-point of the ELISPOT assay is its sensitivity due to the direct contact between the secreting T-cell and the detection surface. This assay captures the presence of cytokines immediately after secretion before it diffuses into the supernatant or is captured by high-affinity receptors present on bystander cells or degraded by proteases. The limit of detection typically achieved can be 1 in 100,000 cells and even a single antigen-specific T-cell can be reliably detected within one million peripheral blood mononuclear cells, which is one hundred times more sensitive than flow

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cytometry with intracytoplasmic cytokine staining [24]. The high sensitivity of the assay makes it particularly useful for studies of the small population of cells found in specific immune responses. All the steps up to number 8 should be done under sterile conditions. 1. On day 1: Prepare the PVDF membranes in the 96-well plates by incubating with 25 μL 35% ethanol for 30 s. Immediately after that, remove the ethanol by washing thoroughly with sterile PBS. 2. The next steps are described according to the manufacturer’s instructions and should be modified appropriately if another commercial kit is employed. Coat the 96-well plate with 100 μL of capture antibody diluted at 15 μg/mL in PBS. Incubate overnight at 4
C. 3. On day 2: Empty the wells, tapping them dry, and wash five times with PBS (ELISPOT plates are more delicate than ELISA plates and should be handled with care. When tapping dry, do so gently). 4. Add 200 μL per well of blocking solution and incubate the plate at RT for 2 h. 5. Wash the plate three times in PBS. 6. Prepare the stimuli solutions at a 2 concentration in complete medium and add 100 μL to each well. Include negative controls, which consist of cells cultured without stimuli. Assay the conditions in duplicate for maximal reproducibility. The specific antigen concentration should be titrated for optimal results. 7. Isolate the splenocytes, prepare a suspension in complete RPMI, and add 100 μL (5  105 cells per well, viability should be over 95%) to each well. If optimizing the assay for cell number, use a 1:2 dilution series. Wrap the plate in aluminum foil to avoid excessive evaporation. Do not shake the plates. 8. Culture overnight at 37
C in CO2 incubator. Do not move the plates while the cells are culturing. This will lead to “snail trail” spots that will not be well defined. During the overnight incubation the cells will secrete cytokine, which will bind to the primary antibody. 9. On day 3: Empty plates and wash the cells and the unbound cytokine by incubating with 200 μL of wash buffer (0.1% Tween-20 in PBS) five times. 10. Dilute the conjugated detection antibody to 1 μg/mL in PBS with 0.5% FCS. Add 100 μL of the solutions to wells and incubate at RT for 2 h. Wash plate five times with wash buffer.

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11. Dilute the streptavidin-alkaline phosphatase 1:1000 in PBS with 0.5% FCS, add 100 μL of the solutions to wells, and incubate at RT for 1 h. Wash plate five times with wash buffer (see Note 30). 12. Add the enzyme substrate solution to each well and carefully monitor spot formation. Stop the reaction by gently washing the plate with wash buffer once development appears to slow down. Take the base off the plates and wash both sides of the membrane with distilled water to stop the spot formation. 13. Dry the plates and allow the membranes to dry at RT. 14. Measure the spots with an ELISPOT reader (AID® ELISPOT reader or other) (see Note 31). If you do not have a commercial ELISPOT reader, you can count the spots manually under the light microscope. Flow Cytometry-Based Cytotoxic Assay

Cell-mediated cytotoxicity is a mechanism used by immune cells for defense against intracellular pathogens, tumor cells, and allogeneic tissue grafts. Lymphocyte effector cells, including natural killer (NK) cells, NKT cells, and cytotoxic T lymphocytes (CTLs), recognize and kill targets by direct cell-to-cell interactions, cytokine production, and/or granule exocytosis [25]. NK and NKT cells are responsible for immune surveillance, contributing to regulation of tumor growth and metastasis as well as autoimmune disorders. CTLs, which require antigen priming and sensitization, mediate acquired immunity. The effector functions of CTLs involved in the elimination of target cells include the release of perforin and granzyme from their granules and Fas/FasL interactions. Virus-specific CTLs recognize viral peptides through their receptor presented by MHC class I molecules on the surface of antigen-presenting cells or virus-infected cells. For the efficient induction of virus-specific CTLs, the antigen must be present in the cytosol of antigenpresenting cells, where antigen processing takes place [26]. VLPs facilitate antigen uptake, endosomal processing, and crosspresentation and, therefore, are able to cross-prime long lasting CTL responses in addition to antibody responses. Here we evaluated the generation of HA-specific cytotoxic cells in the immunized mice by evaluating the death of cells expressing HA antigen. In this protocol, the target cell population is labeled with CFSE [27] to allow the discrimination between the splenocytes isolated from vaccinated mice (effector cells) and propidium iodide to identify dead cells. The labeling distinguishes four populations of cells: (1) living target cells in green, (2) dead target cells in green and red, (3) dead effector cells in red, and (4) live effector cells, which remain unstained. Thus, the cytotoxicity of the effector cells can be evaluated quantitatively.

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1. Prepare a single-cell suspension of murine AB1 malignant mesothelioma that expresses HA from H1N1 influenza A (AB1-HA) in complete RPMI (see Note 32). Generation and characterization of the AB1 cells transfected with the influenza HA gene have been described [28] (see Note 33). 2. Label AB1-HA cells (target cells) with 2 μM carboxyfluorescein succinimidyl ester (CFSE) as follows (see Note 34). 3. Count AB1-HA cells, transfer the amount needed to a 15-mL conical tube, and wash in 14 mL warm 1 PBS (see Note 35). 4. Prepare a 4-μM CFSE working solution in warm PBS (see Note 36). 5. Resuspend cells at 20  106/mL of PBS (if the number of cells is lower than 10  106/mL, resuspend cells in 0.5 mL of PBS) and stain them by adding one equal volume of 4 μM CFSEhigh working solution (final concentration: 2 μM). Vortex immediately to obtain a homogeneous staining. For example, for 1 mL of cells, add 1 mL of the CFSE working solution. Stain cells at 37
C for 15 min in 5% CO2 atmosphere. 6. Fill the tube with warm complete RPMI to quench the labeling reaction and pellet cells at 1500 rpm for 5 min. 7. Wash CFSE-cells again in complete RPMI. Resuspend at a concentration of 2  105 cells/mL in complete RPMI. 8. Obtain a single-cell suspension of the splenocytes from immunized mice (effector cells). Resuspend at a concentration 4  106 cells/mL in complete RPMI. 9. Add 100 μL of the CFSE-target cells to the 96-well plate. Add 100 μL of effector cells from each mouse to replicate wells for a final volume of 200 μL, at an E:T ratio of 20:1. Make sure that the effector and target cell are properly mixed for more accurate results. If necessary, other E:T ratios can be assayed, since optimal concentrations of E:T cells can vary with prior activation of cells. 10. To measure the basal apoptosis, seed three wells only with target cells. 11. An additional control is to perform the assay with AB1 cells as control target cells, which do not express the HA antigen. In general, CTL activity results from antigen-specific T-cells. Therefore, this control is performed to exclude MHC-independent cytolysis resulting from NK-like activity. 12. Incubate coculture in U-bottom 96-well culture plates at 37
C, 5% CO2 for 16–20 h. 13. Transfer supernatant to a 2-mL tube, wash with 200 μL 1 PBS, collect the PBS in the tube, and harvest the adherent cells

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with 200 μL trypsin. Add 500 μL of complete medium to stop the trypsin reaction. Centrifuge at 300  g for 10 min. 14. Resuspend the cell pellet in 2% FCS in PBS and stain it with propidium iodide (see Note 37). For this purpose, add an aliquot of 1 μL of 0.5 mg/mL propidium iodide reagent to 100 μL of cells and incubate at RT for 5–10 min. 15. Determine the viability within the CFSE-positive population by flow cytometry analysis immediately following completion of the staining. Keep the cells on ice, since they are not fixed. The mean percentage of each condition can be calculated from three replicate wells. 16. Express the CTL activity as: % PI-positive cells in the CFSEgated cells of experimental condition—% PI-positive cells in the CFSE-gated cells of medium only cells. 3.4 Challenge with Live Virus

The ultimate goal of a vaccine is to instruct the immune system to elicit an effective immune response and memory against the pathogen of interest. Laboratory animal models are widely used in the preclinical evaluation of potential vaccines, mainly for the assessment of the immune response to immunization and in protection studies against infection with the pathogen of interest. The safety, dose, formulation, and route of delivery of the vaccine and its efficacy in preventing or moderating infection, disease, or secondary transmission are usually investigated in animal models [29]. In selecting an animal model for vaccine research, a number of essential factors must be considered. Ideally, the animal must be susceptible to infection and supportive of its replication, must have a sensitive and specific “read-out” for infection, must mimic pathophysiology of human disease, and must sustain immune responses that closely resemble human immune responses. The induction of potent virus-specific immune responses at mucosal surfaces where virus transmission occurs is a major challenge for vaccination strategies. We have developed a vaccine platform that effectively induces mucosal, as well as systemic, immune responses. A lethal challenge of homologous influenza viruses (A/Puerto Rico/8/1934 (PR8) (H1N1)) can be performed to evaluate if the humoral and cellular immune responses elicited by oral immunization are able to effectively protect mice against virus infection. Most inbred laboratory mice are highly susceptible to disease and death following intranasal infection with certain influenza viruses, including PR8. The signs of disease that develop and their severity depend upon the challenge dose administered. In most cases, however, a lethal dose is used, resulting in a severe disease characterized by huddling, ruffled fur, lethargy, anorexia, which leads to weight loss and death (euthanasia at a humane endpoint). The parameters most commonly used to evaluate influenza viral pathogenicity in mice are body weight loss and mortality.

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In addition, viral titers, pathology scores, lung weights, oxygen saturation in the blood, and gross motor-activity levels may be monitored [30]. 1. Ten days after the last immunization, infect vaccinated and control anesthetized mice by the nasal route with a mouseadapted A/Puerto Rico/8/1934 influenza virus at 1  102 to 1  105 TCID50 in 50 μL of PBS per mouse (see Note 38). 2. Monitor the mice daily for disease signs (ruffled fur, dyspnea, lethargy, pyrexia) and body weight changes during 2 weeks. 3. Euthanize mice at a predetermined humane endpoint of 20% weight loss or if they showed no sign of recovery at 7 days postchallenge.

4

Notes 1. Hygromycin B is light sensitive. Store the liquid stock solution at 4
C, protected from exposure to light. 2. To prepare PEI stock solution: l

Dissolve PEI in endotoxin-free dH2O previously heated to ~80
C.

l

Let cool to room temperature.

l

Neutralize to pH 7.0, filter sterilize (0.22 μm), aliquot and store at 20
C; a working stock can be kept at 4
C.

3. PEI is a trusted, easy to use, consistent, and cost-effective transient transfection reagent. Nevertheless, other transfection reagents can be used instead, such as lipids or calcium phosphate. Check the usage recommendations for each protocol. 4. If you choose to produce the VLPs by transient transfection of the four plasmids encoding the antigens, the pVSP-G must be included in this analysis and HEK-293 cells should be used. 5. Preferably choose primary antibodies from different animal species to be able to perform colocalization assays. 6. Quality and purity of the plasmid DNA is critical for a successful transfection. The plasmid may often carry phenol, sodium chloride, and endotoxins. Phenol can kill the cells, salts interfere in the complexing with the transfection reagent, and endotoxins sharply reduce transfection efficiency in primary and other sensitive cells. In general, plasmid DNA purified using an anion-exchange resin is adequate. 7. The ECL western blotting substrate is a highly sensitive nonradioactive, enhanced luminol-based chemiluminescent substrate for the detection of HRP on immunoblots. ECL is based on the emission of light during the HRP and hydrogen peroxide-

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catalyzed oxidation of luminol. The emitted light is captured on film or by a CCD camera, for qualitative or semiquantitative analysis. It is able to detect picogram amounts of antigen, and the blots can be repeatedly exposed to obtain optimal results or stripped of the immunodetection reagents and reprobed. 8. Alsever’s solution: 8.0 g sodium citrate (C6H5Na3O7l2H2O); 0.55 g citric acid (C6H8O7lH2O); 4.2 g NaCl; 20.5 g dextrose; Q.S. to 1000 mL with distilled water. Filter sterilize with 0.22μm filter. Store at 4  2
C. 9. Volumes up to 10 mL/kg in the mouse and 20 mL/kg in the rat may be administered via oral gavage. The concentration of the VLPs must be modified accordingly, to avoid exceeding that volume. 10. Viruses in allantoic fluid must be stored at 70
C. 11. Propagation of influenza virus in hen eggs, quantitation of influenza virus titer in egg allantoic fluid as well as measurement of tissue culture influenza virus infectious dose (TCID50) can be found in a detailed description in Cottey et al. [31]. 12. Add 60 mL of bovine albumin fraction V (10%), 6 mL 100 antibiotics and 12.5 mL of 1 M HEPES to 500 mL DMEM. 13. In these positive control wells, the cells should be stimulated with an agent known to induce expression of the cytokine being detected to confirm both cell and assay functionality. Typical stimuli include: phytohemagglutinin (PHA) for IFN-γ; ConA; LPS for IL-1β and IL-6 secretion; PMA and ionomycin to stimulate IL-2 and IL-4 secretion; anti-CD3/ CD28 antibodies for IFN-γ, IL-4, IL-10, and Granzyme B. 14. Procedure for effective cell selection: l

Plate 200,000–300,000 cells per well in complete DMEM in a 6-well culture plate, prepare a set of 7 plates. Allow cells to adhere overnight at 37
C with 5% CO2. The cells should be grown and incubated under these conditions unless otherwise stated.

l

After 1 day, discard the culture medium and add medium containing varying concentrations of hygromycin B (0, 10, 50, 100, 200, 400, 600 μg/mL hygromycin B).

l

Replenish the selective media every 3–4 days, and observe the percentage of surviving cells.

l

Determine the percentage of surviving cells at regular intervals to set the appropriate concentration of hygromycin that kills the cells within 1–2 weeks after addition of hygromycin.

15. Pre-treatment of glass coverslips is recommended for HEK293 cells to avoid detachment during the procedure. Acid

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treatment is performed to neutralize silicate groups and clean the glass; for this purpose, submerge the coverslips in a beaker containing a solution of 1 M HCl at 50–60
C for 4 to 16 h, and shake frequently. Rinse thoroughly with distilled water. Place the coverslips in a glass petri dish and sterilize using the drying cycle of the autoclave. 16. Bear in mind that the cells will continue to grow after transfection and if they reach high confluence, the subcellular localization of the antigens will be more difficult to assess in the microscope. However, if you choose to seed too few cells, they may detach and not be enough for microscopy. 17. A plasmid containing a reporter gene can be used as a control of transfection. The expression of up to two antigens can be determined simultaneously, provided that each primary antibody has originated from a different animal host or that there are different fluorochrome-conjugated primary antibodies. In that case, the transfection mix can be prepared with 1.5 μg of each plasmid. 18. The ratio of each plasmid can be assayed to optimize VLP yield; to start, we recommend using an equivalent amount of each plasmid. 19. This step can be skipped if an ultracentrifuge rotor with a capacity of over 200 mL is available. The gradient can be prepared directly in the ultracentrifuge tubes with the filtered medium. 20. We used a chemiluminescent detection technique, since it allows us to make multiple exposures; thus, signal to noise is optimized and a large linear response range allows detection and quantitation of a wide range of protein concentrations. Chemiluminescence yields the greatest potential sensitivity of any detection method available for western blotting. Alternatively, colorimetric or fluorescent detection techniques can be used. 21. To prepare the erythrocyte suspension, harvest the chicken blood in a tube with Alsever’s solution. Then wash the RBC, pour 10–20 mL chicken blood in a 50-mL centrifuge tube, and fill the tube with PBS. Invert the tube slowly several times and centrifuge at 800  g for 10 min in a refrigerated centrifuge. Aspirate the PBS and the phlogistic crust from the tube. Repeat wash and spin cycle 2 more times. Finally, to prepare the 0.5% erythrocyte suspension, pour 49.75 mL of PBS and add 0.25 mL of washed RBCs, trying to wash the pipette tip in the solution. Erythrocytes can be stored at 4
C for up to 1 week. If hemolysis observed the solution should be discarded.

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22. The number of animals for each experiment and all the followed procedures must be approved by the Institutional Committee for Care and Use of Experimental Animals. 23. Liquid compounds may be administered directly into the stomach of mice and rats via oral gavage; for this purpose, a ball tip needle is used for administration to prevent damaging. For adult mice a 22 G ball tip needle is suitable [32]. Firmly restrain a conscious mouse in an upright position. The head must be immobilized for this procedure. A straight line is formed between the mouth and the cardiac sphincter through the esophageal orifice. Gently pass the needle through the mouth and pharynx into the esophagus. The substance should be administered slowly. Feeling an obstruction; or mouse coughing, choking, or beginning to struggle vigorously after the gavage begins; or fluid coming out through the nose may indicate that the needle has entered the lungs. If any of these signs is detected, immediately withdraw the needle. If there is any sign that the material has been injected into the lungs, the mouse should be euthanized. 24. For a detailed explanation of this procedure, see https://www. jove.com/science-education/10247/blood-withdrawal-ii. 25. The tracheas of animals were exposed by standard procedures [33] and cannulated with a 20 G needle attached to a 1 mL syringe. One milliliter of PBS was introduced into the lungs via the tracheal cannula and carefully extracted. This procedure was repeated once more, for a total of 2 mL of PBS for each lavage. 26. The recombinant antigen can be purchased from a bioscience company or produced in the laboratory. Bear in mind that several parameters must be taken into account in the design of the recombinant protein, such as cell expression system, glycosylation pattern, and formation of trimers or dimers. 27. The indicated reagent volumes correspond to a standard microtiter plate; if a half area plate is used, reduce the volumes by half. 28. Determine the virus working dilution before use. Never use any freeze–thawed virus other than the initial freeze–thawed aliquot required to prepare the assay. 29. Timing is very critical; if you incubate for a longer period, the white blood cells will also be lysed, so it has to be exactly 5 min. You can change it to 3 min, if your cell viability is low. 30. For enzymatic detection protocols, the base should be taken off the bottom of the plate to enable thorough washing of the membrane before adding substrate/chromogen. For example, after incubation with the streptavidin-alkaline phosphatase

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conjugate, remove the base and wash both sides of the membrane under running distilled water. This procedure helps to prevent high background, since some reagents can leak through the membrane into the bottom tray of the plate. 31. In the analysis software, set the following parameters for measurement: size/spot diameter; intensity/saturation; circularity/shape; spot development/slope. These parameters can be saved and used for subsequent experiments to obtain standardized results. We recommend reading each plate three times and averaging the results in order to minimize measurement errors. 32. Healthy cells should be used in the assay; determine the viability (which should be greater than 95%) by trypan blue exclusion before the assay. 33. Cytolytic activity can be induced by a large number of antigens, including major and minor alloantigens, haptens, and viruses. The frequency of T-cells that recognize allogeneic MHC antigens is high; therefore, for the detection of antigens not encoded by the MHC, syngeneic target and effector cells need to be employed [34]. Other cell lines that express HA can be employed as well as influenza-infected cells. 34. Staining of Target cells: CFSE passively diffuses into cells; then its acetate groups are cleaved by intracellular esterases to yield highly fluorescent carboxyfluorescein succinimidyl ester. CFSE covalently couples to intracellular molecules; thus, fluorescence can be retained within cells for long periods [35]. 35. Many cells are lost during the numerous centrifugation steps. Start with twice the number of cells needed to obtain a sufficient yield. 36. CFSE and propidium iodide are light sensitive. All the staining procedures should be done without direct exposure to intense light. Incubations should be performed in the dark. 37. Membrane integrity is the feature most often used to detect whether eukaryotic cells cultured in vitro are alive or dead. Cells that have lost membrane integrity and allow movement of otherwise non-permeable molecules are classified as nonviable or dead. A second class of molecules that serve as an indicator of dead cells is referred to as “vital dyes.” These dyes are typically not permeable to viable cells, but can enter dead cells through damaged membranes. Examples include trypan blue and propidium iodide [36]. 38. Select the viral titer based on titration that results in 100% mortality in mice by days 7–10.

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References 1. Vela Ramirez JE, Sharpe LA, Peppas NA (2017) Current state and challenges in developing oral vaccines. Adv Drug Deliv Rev 114:116–131. https://doi.org/10.1016/j. addr.2017.04.008 2. Rupil LL, del Serradell MC, Luja´n HD (2019) Using protozoan surface proteins for effective oral vaccination. Trends Parasitol 36(1):7–10. https://doi.org/10.1016/j.pt.2019.07.004 3. Prucca CG, Slavin I, Quiroga R et al (2008) Antigenic variation in Giardia lamblia is regulated by RNA interference. Nature 456:750–754. https://doi.org/10.1038/ nature07585 4. Serradell MC, Rupil LL, Martino RA et al (2019) Efficient oral vaccination by bioengineering virus-like particles with protozoan surface proteins. Nat Commun 10:361. https:// doi.org/10.1038/s41467-018-08265-9 5. Rivero FD, Saura A, Prucca CG et al (2010) Disruption of antigenic variation is crucial for effective parasite vaccine. Nat Med 16:551–557. https://doi.org/10.1038/nm. 2141 6. Serradell MC, Saura A, Rupil LL et al (2016) Vaccination of domestic animals with a novel oral vaccine prevents giardia infections, alleviates signs of giardiasis and reduces transmission to humans. NPJ Vaccines 1:16018. https:// doi.org/10.1038/npjvaccines.2016.18 7. Tan M, Jiang X (2017) Recent advancements in combination subunit vaccine development. Hum Vaccin Immunother 13:180–185. https://doi.org/10.1080/21645515.2016. 1229719 8. Millet JK, Tang T, Nathan L et al (2019) Production of Pseudotyped particles to study highly pathogenic coronaviruses in a biosafety level 2 setting. J Vis Exp:59010. https://doi. org/10.3791/59010 9. Sze´csi J, Boson B, Johnsson P et al (2006) Induction of neutralising antibodies by viruslike particles harbouring surface proteins from highly pathogenic H5N1 and H7N1 influenza viruses. Virol J 3:70. https://doi.org/10. 1186/1743-422X-3-70 10. Garrone P, Fluckiger A-C, Mangeot PE et al (2011) A prime-boost strategy using virus-like particles pseudotyped for HCV proteins triggers broadly neutralizing antibodies in macaques. Sci Transl Med 3:94ra71. https://doi. org/10.1126/scitranslmed.3002330 11. Yu Z, Beer C, Koester M, Wirth M (2006) Caveolin-1 interacts with the gag precursor of murine leukaemia virus and modulates virus

production. Virol J 3:73. https://doi.org/10. 1186/1743-422X-3-73 12. Kurg R, Reinsalu O, Jagur S et al (2016) Biochemical and proteomic characterization of retrovirus gag based microparticles carrying melanoma antigens. Sci Rep 6:29425. https://doi.org/10.1038/srep29425 13. O’Gorman S, Fox D, Wahl G (1991) Recombinase-mediated gene activation and site-specific integration in mammalian cells. Science 251:1351–1355. https://doi.org/10. 1126/science.1900642 14. Stuchbury G, Mu¨nch G (2010) Optimizing the generation of stable neuronal cell lines via pre-transfection restriction enzyme digestion of plasmid DNA. Cytotechnology 62:189–194. https://doi.org/10.1007/ s10616-010-9273-1 15. Hnasko TS, Hnasko RM (2015) The Western blot. Methods Mol Biol 1318:87–96. https:// doi.org/10.1007/978-1-4939-2742-5_9 16. Killian ML (2020) Hemagglutination assay for influenza virus. Methods Mol Biol 2123:3–10. https://doi.org/10.1007/978-1-0716-03468_1 17. Griffiths D, Carnell-Morris P, Wright M (2020) Nanoparticle tracking analysis for multiparameter characterization and counting of nanoparticle suspensions. In: Ferrari E, Soloviev M (eds) Nanoparticles in biology and medicine. Springer US, New York, NY, pp 289–303 18. Gonda MA (1998) Electron microscopy, immunological applications. In: Encyclopedia of immunology. Elsevier, Amsterdam, pp 790–795 19. Zimmermann P, Curtis N (2019) Factors that influence the immune response to vaccination. Clin Microbiol Rev 32:e00084. https://doi. org/10.1128/CMR.00084-18 20. Liu MA (1997) The immunologist’s grail: vaccines that generate cellular immunity. Proc Natl Acad Sci 94:10496–10498. https://doi.org/ 10.1073/pnas.94.20.10496 21. Sakamoto S, Putalun W, Vimolmangkang S et al (2018) Enzyme-linked immunosorbent assay for the quantitative/qualitative analysis of plant secondary metabolites. J Nat Med 72:32–42. https://doi.org/10.1007/s11418017-1144-z 22. World Health Organization (WHO) WHO manual on animal influenza diagnosis and surveillance 23. Kaiko GE, Horvat JC, Beagley KW, Hansbro PM (2008) Immunological decision-making:

Production of Oral Vaccines Based on Virus-Like Particles Pseudotyped with. . . how does the immune system decide to mount a helper T-cell response? Immunology 123:326–338. https://doi.org/10.1111/j. 1365-2567.2007.02719.x 24. Ji N, Forsthuber TG (2016) ELISPOT techniques. Methods Mol Biol 1304:63–71. https:// doi.org/10.1007/7651_2014_111 25. Kim GG, Donnenberg VS, Donnenberg AD et al (2007) A novel multiparametric flow cytometry-based cytotoxicity assay simultaneously immunophenotypes effector cells: comparisons to a 4 h 51Cr-release assay. J Immunol Methods 325:51–66. https://doi. org/10.1016/j.jim.2007.05.013 26. Hillaire MLB, Osterhaus ADME, Rimmelzwaan GF (2011) Induction of virus-specific cytotoxic T lymphocytes as a basis for the development of broadly protective influenza vaccines. J Biomed Biotechnol 2011:1–12. https://doi.org/10.1155/2011/939860 27. Noto A, Ngauv P, Trautmann L (2013) Cellbased flow cytometry assay to measure cytotoxic activity. J Vis Exp:e51105. https://doi. org/10.3791/51105 28. Marzo AL, Lake RA, Lo D et al (1999) Tumor antigens are constitutively presented in the draining lymph nodes. J Immunol 162:5838–5845 29. Gerdts V, Wilson HL, Meurens F et al (2015) Large animal models for vaccine development and testing. ILAR J 56:53–62. https://doi. org/10.1093/ilar/ilv009

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30. Bouvier NM, Lowen AC (2010) Animal models for influenza virus pathogenesis and transmission. Viruses 2:1530–1563. https://doi. org/10.3390/v20801530 31. Cottey R, Rowe CA, Bender BS (2001) Influenza Virus. Curr Protoc Immunol 42:19.11.1–19.11.32. https://doi.org/10. 1002/0471142735.im1911s42 32. Hedrich H, Gillian B (2004) The laboratory mouse. Elsevier, Amsterdam 33. Van Hoecke L, Job ER, Saelens X, Roose K (2017) Bronchoalveolar lavage of murine lungs to analyze inflammatory cell infiltration. J Vis Exp (123):55398. https://doi.org/10. 3791/55398 34. Wonderlich J, Shearer G, Livingstone A et al (2018) Induction and measurement of cytotoxic T lymphocyte activity. Curr Protoc Immunol 120:3.11.1–3.11.29. https://doi. org/10.1002/cpim.38 35. Quah BJC, Warren HS, Parish CR (2007) Monitoring lymphocyte proliferation in vitro and in vivo with the intracellular fluorescent dye carboxyfluorescein diacetate succinimidyl ester. Nat Protoc 2:2049–2056. https://doi. org/10.1038/nprot.2007.296 36. Riss T, Niles A, Moravec R et al (2004) Cytotoxicity assays: in vitro methods to measure dead cells. In: Sittampalam GS, Grossman A, Brimacombe K et al (eds) Assay guidance manual. Eli Lilly & Company and the National Center for Advancing Translational Sciences, Bethesda (MD)

Chapter 27 A Fast-Track Phenotypic Characterization of Plasmodium falciparum Vaccine Antigens through Lyse-Reseal Erythrocytes Mediated Delivery (LyRED) of RNA Interference for Targeted Translational Repression Malabika Chakrabarti, Swati Garg, Akshay Munjal, Sweta Karan, Soumya Pati, Lalit C. Garg, and Shailja Singh Abstract The minimal success of the malaria vaccine with available antigens indicates the need for intensive and accelerated research to identify and characterize new antigens that confer protection against infection, clinical manifestation, and even malaria transmission. Further, the genetic manipulation tools to characterize such antigens are very time-consuming and laborious due to the very low efficiency of transfection in the malaria parasite. Here, we report a human miRNA-mediated translational repression of antigens in Plasmodium falciparum as a fast-track method for understanding and validating their function. In this method, candidate miRNAs are designed based on favorable hybridization energy against a parasite gene, and miRNA mimics are delivered to the parasite by loading them as cargo in the erythrocytes by simple lysereseal method. Incubation of the miRNA loaded erythrocytes with purified mature trophozoites or schizonts results in the loaded erythrocytes’ infection. The miRNA mimics are translocated to parasites, and the effect of miRNA-mediated translation repression can be monitored within 48–72 h post-invasion. Unlike other transfection based methods, this method is fast, reproducible, and robust. We call this method as lyse-reseal erythrocytes for delivery (LyRED) of miRNA, which is a rapid and straight-forward method providing an efficient alternative to the existing genetic tools for P. falciparum to characterize the function of antigens or genes. The identification of crucial antigens from the different stages of the Plasmodium falciparum life cycle by the miRNA targeting approach can fuel the development of efficacious subunit vaccines against malaria. Key words Plasmodium, micro-RNA, Lyse-reseal erythrocytes, PfApicortin, Vaccine, Antigen, Translational repression

Malabika Chakrabarti and Swati Garg are joint first authors. Sunil Thomas (ed.), Vaccine Design: Methods and Protocols, Volume 1: Vaccines for Human Diseases, Methods in Molecular Biology, vol. 2410, https://doi.org/10.1007/978-1-0716-1884-4_27, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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Introduction Malaria is a life-threatening disease caused by Plasmodium spp. P. falciparum is the cause of most severe form of malaria and causes a large number of deaths [1, 2]. The emergence of resistant parasite strains against all known drugs is an alarming signal in malaria control and elimination efforts. Hence, there is an urgent need for developing a vaccine against malaria [3, 4]. In order to identify and characterize a target vaccine antigen, manipulating the desired gene using reverse genetic tools is of immense significance in understanding its role in merozoite invasion [5, 6]. However, P. falciparum’s transfection is a tricky, cumbersome, and time-consuming tool with very less success rates [7]. There are several reports regarding the modification of classic transfection techniques [8, 9]; however, the electroporation of plasmid DNA into ring stage-infected erythrocytes has continued to remain a method of choice by the parasitologists around the world [10, 11]. A recent report demonstrated that the lyse and resealed erythrocyte, loaded with plasmid DNA, could be used for parasite infection [12]. The plasmid DNA in the erythrocyte cytosol enters the parasite, and hence this technique is an alternative to classic electroporation techniques. However, with all these genetic manipulation techniques, the major disadvantage is the time (~21 days) taken for the first appearance of desired genetically modified parasites, due to extremely low efficiency of transfection in malaria parasites [9]. Then more time is exhausted in cloning and characterizing the desired clone. Moreover, if the desired genetic manipulation is deleterious to the parasite, one will never be successful in generating a genetically modified parasite clone. In this chapter, we present a novel method of studying the role of a candidate vaccine antigen in P. falciparum parasites [13] (Fig. 1). Herein, we use the micro-RNA mediated translational repression approach to reduce the target protein expression and understand its role in merozoite invasion in a reduced time frame. The candidate miRNA was designed against PfAARP based on favorable hybridization energy (Table 1). In this method, erythrocytes are lysed in a hypotonic buffer, mixed with desired micro-RNA, and resealed in a hypertonic buffer to encapsulate the micro-RNAs within themselves. The loaded erythrocytes were then incubated with purified P. falciparum trophozoites or schizonts, resulting in an infection of the cargo enriched erythrocytes and subsequent transfer of the cargo into the parasite, also called as LyRED. The application of human micro-RNA in parasite gene downregulation is a novel approach for the functional characterization of a gene showing effective downregulation [14–16]. Further, the effect can be monitored in the first and second cycles of infection (within 48–72 h post-invasion), which is not possible with

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Fig. 1 Schematic diagram showing the steps of lyse-reseal erythrocytes mediated delivery (LyRED) of RNA interference method for translational repression

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Table 1 Hybridization energy of miR150 with PfApicortin Target gene

Candidate miRNAs

Minimum free energy of hybridization

Pf_Apicortin

miR-150-3p

26.8 kcal

conventional genetic manipulation tools. Thus, this method can decode the role of the target antigen much faster and can work as an alternative to conventional genetic manipulation tools. Since this method does not need any high-end instrument, this can be used in limited-resource settings also.

2

Materials

2.1 Production and Validation of Loaded Erythrocytes

2.2 Assessment of miRNA Enriched Erythrocytes for Parasite Invasion

l

Packed erythrocytes (O+ preferably).

l

Filter sterilized PBS (137 mM NaCl, 2.7 mM KCL, 10 mM Na2HPO4, and 1.8 mM K2HPO4), 0.2% glucose solution.

l

Incomplete RPMI medium (iRPMI): RPMI 1640 supplemented with 25 mM HEPES.

l

500 mM K2HPO4.

l

100 mM ATP.

l

100 μM miRNA mimic.

l

10 kDa Dextran FITC.

l

Temperature controlled microcentrifuge for 1.5 ml tubes.

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Water bath or incubator (37
C).

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Fluorescence microscope.

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Fluorescence assisted cell sorter.

l

Complete RPMI medium (cRPMI): RPMI 1640 supplemented with 25 mM HEPES, 0.1 mM hypoxanthine, 2 g/L sodium bicarbonate, 25 mg/ml gentamicin, and 0.5% AlbuMax.

l

Incomplete RPMI medium (iRPMI): RPMI 1640 supplemented with 25 mM HEPES.

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Giemsa stain.

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Hemocytometer.

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37
C Incubator.

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Sorvall RT7 centrifuge.

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Eppendorf 5810R centrifuge.

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Inverted microscope.

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l

Complimentary primers.

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5% Saponin in PBS (137 mM NaCl, 2.7 mM KCL, 10 mM Na2HPO4, and 1.8 mM K2HPO4, pH 7.4).

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TRI reagent (Life Technologies, USA) for RNA isolation.

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RNAse inhibitor.

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E. coli Poly (A) polymerase.

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ATP.

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cDNA synthesis kit.

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SYBR green master mix for real-time PCR.

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2% Agarose gel.

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12% SDS-PAGE gel.

Methods

3.1 Production and Validation of Loaded Erythrocytes 3.1.1 Preparation of Erythrocyte Ghosts and Loading of Candidate miRNA Mimics Using LyRED Method

Lyse-resealed erythrocytes are extensively studied for their drugcarrying properties since they can carry the desired cargo. Moreover, lyse and resealed erythrocytes behave similarly as untreated erythrocytes. Hence, they can be successfully infected with malaria parasites, we combined these properties of lyse-resealed erythrocytes to deliver micro-RNA cargos to the parasites residing within them. 1. Packed erythrocytes were acquired from blood bank. 2. Take 50 μl packed erythrocytes for the ghost preparation and loading of miRNA mimic. Wash packed erythrocytes (50 μl) with 1 ml PBS-glucose solution three times (see Note 1). 3. Add an equal volume (50 μl) of lysis buffer (5 mM K2HPO4 containing, 1 mM ATP, 10 μM micro-RNA mimic; prechilled on ice; pH 7.5). Mix gently and incubate the suspension on ice for 1 h. 4. Add 20 μl of 5 resealing buffer (475 mM KOAc, 25 mM Na2HPO4, 25 mM MgCl2, 237.5 mM KCl, 1 mM ATP, pH 7.5). Mix gently and keep at 37
C for 1 h (see Note 2). 5. Centrifuge the cell suspension at 800  g for 5 min to pellet down the resealed erythrocytes. 6. Wash the resealed erythrocytes with iRPMI (3 times). Resuspend in iRPMI and store at 4
C (see Note 3, and Subheading 3.1.2).

3.1.2 The Measurement of Resealing Efficiency

Different methods of erythrocyte lyse and reseal have variable efficiencies in erythrocyte resealing and cargo loading. Therefore, it is critical to determine the resealing and cargo loading efficiency. For this purpose, FITC-conjugated dextran is used as a sample

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cargo and a loading control as its loading can be tracked with a fluorescence microscopy. With this approach, it is established that more than 90% of the erythrocytes are loaded with dextran FITC. 1. Wash the packed erythrocytes (50 μl) with 1 ml PBS-glucose solution three times (see Note 1). 2. Add an equal volume of prechilled lysis buffer (5 mM K2HPO4, 1 mM ATP) containing the tracker 10 kDa-FITC-dextran (30 μM). Mix gently and keep on ice for 1 h. 3. Add 1/5th volume of 5 resealing buffer, brought to 25
C. Mix gently and keep at 37
C for 1 h. 4. Wash erythrocytes 3 with iRPMI, and store it in 50 μl of iRPMI. Take 20 μl of resealed erythrocyte suspension and check FITCdextran loading of erythrocytes under a fluorescence microscope. The green fluorescence signal should be evenly distributed inside the erythrocyte cytosol. Count the number of erythrocytes positive for the green fluorescence and calculate the percentage of dextran positive cells as: %FITC  dextran positive cells ¼ ðFITC  Dextran positive cellsÞ=ðtotal number of cellsÞ  100 3.2 Assessment of miRNA Enriched Erythrocytes for Parasite Invasion 3.2.1 Parasite Culture of P. falciparum in miRNA Enriched Erythrocytes

1. Tightly synchronize P. falciparum 3D7 culture by treatment with 5% sorbitol in two subsequent cycles using the following procedures. Centrifuge the culture at 800  g for 5 min. Wash the obtained culture pellet once with 5 ml of iRPMI. Add 5 ml of 5% sorbitol solution (filter sterilized), mix gently, and incubate the suspension at 37
C for 10 min. Post-incubation, centrifuge the suspension at 800  g for 5 min. Discard the supernatant and wash the pellet twice with iRPMI (in order to remove the traces of sorbitol). Resuspend the pellet in 10 ml of complete RPMI and incubate at 37
C, under mixed gas composition (90% N2, 5% CO2, 5% O2) for normal growth. 2. When majority of the parasites are in the late trophozoite or schizont stage, centrifuge parasite culture at 800  g at room temperature (RT) for 5 min. 3. Discard the supernatant and resuspend the pellet at 10% hematocrit in iRPMI medium pre-warmed to 37
C. 4. Overlay the parasite suspension on 65% percoll (made in iRPMI) and centrifuge at 1200  g for 25 min. 5. Collect the enriched late staged parasite infected erythrocytes from the interface of percoll and medium. 6. Wash the infected erythrocytes 3 with iRPMI.

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7. Check the purity of culture by Giemsa staining and score the parasitemia. 8. Add micro-RNA enriched erythrocytes to make the final parasitemia at 1% and 2% hematocrit. Incubate at 37
C under mixed gas conditions. 9. Prepare smears at different time points post-infection, that is, 20, 40, 60 h, and score the parasitemia (Fig. 2a) (see Note 4).

Fig. 2 (a) Western blot showing PfApicortin and GAPDH expression in parasites infecting miRNA mimic loaded erythrocytes and that of the control and scrambled mimic loaded erythrocytes, (b) Level of change in PfApicortin protein expression due to loading of miRNAs (**p < 0.01)

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3.2.2 Monitoring Percent Merozoite Invasion

1. Perform infection of erythrocytes enriched with specific microRNA or scrambled micro-RNA (as a control) with mature parasites as described above. 2. Prepare smears at different time points, 24, 40, 54, and 72 h post-invasion (hpi) (see Note 4). Calculate invasion of parasites by calculating the number of rings formed at 72 h postinvasion. Invasion rate can be calculated as the number of rings formed per schizont as given below: Invasion rate ðnumber of rings per schizontÞ ¼ ðnumber of infected erythrocytes at 72 hpiÞ= ðnumber of infected erythrocytes at 54 hpiÞ  100 Compare the invasion rate of control versus micro-RNA enriched erythrocytes (Fig. 2b, c).

3.3 Monitoring Translocation of micro-RNA to Parasites and Repression of the Target Protein 3.3.1 Monitoring Translocation of miR-150-3p to the Parasite

P. falciparum Apicortin (PfApicortin) plays an important role in parasite invasion by interacting with parasite tubulin and stabilizing it. Host miR-150-3p targets the mRNA of PfApicortin and hence we have loaded miR-150-3p mimic in the erythrocytes to see its overall effect on PfApicortin expression and parasite growth.

1. Purify schizont stage parasites using 65% percoll enrichment method. 2. Incubate miRNA loaded erythrocytes with percoll purified schizonts at 2% parasitemia and 4% hematocrit for 48 h. 3. Harvest the infected erythrocytes after 48 h of infection. 4. Harvest and isolate parasites by saponin treatment according to the following procedure: Centrifuge the harvested culture at 800  g for 5 min. Resuspend the pellet in 500 μl PBS, add 5 μl of 5% saponin solution (prepared in PBS), and incubate on ice for 5 min. Centrifuge the suspension at 5000  g for 5 min to obtain the parasite pellet. Wash the pellet three times with PBS in order to remove the excess hemoglobin. Add 500 μl of TRI reagent to the pellet and mix well by vortexing. 5. Isolate total RNA from the parasites using the following method. Incubate the cell suspension in TRI reagent for 10 min at RT for the separation of nucleoproteins. Add 250 μl chloroform (molecular biology grade) to the suspension, shake well, and incubate the mixture at RT for 10 min. Postincubation, centrifuge the mixture at 12,000  g for 15 min at 4
C and collect the upper aqueous phase. Add 500 μl of chilled isopropanol (molecular biology grade) to the collected supernatant, mix well, and incubate the mixture at RT for 15 min followed by centrifugation at 15,000  g at 4
C for 10 min.

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Wash the RNA pellet once with 75% ethanol (molecular biology grade). Air-dry the pellet and resuspend it in 10 μl of nuclease free water. Add 0.5 μl of RNAse inhibitor (1 unit/μ l) to the RNA and store at 20
C (see Notes 5 and 6). 6. Perform poly (A) tailing of the isolated RNA using the following protocol: Take 2 μg of total RNA in 15 μl nuclease free water (DEPC-treated) in a microcentrifuge tube. Add 2 μl of poly A polymerase reaction buffer and 2 μl of 10 mM ATP. Add 1 μl of E. coli poly (A) polymerase and incubate the reaction mixture at 37
C for 30 min. For cleaning up the RNA, add 50 μl of 70% ethanol in the reaction mixture and incubate at RT for 10 min. Centrifuge the mixture at 7500  g for 5 min. Decant the supernatant, air-dry the RNA pellet, and dissolve it in 10 μl of nuclease free water (see Notes 5 and 6). Synthesize cDNA from the poly (A) tailed RNA by reverse transcription using cDNA synthesis kit according to the manufacture’s protocol. Add 1 μg polyadenylated RNA, 2 μl of random hexamer primer solution, 2 μl of 10 reverse transcription buffer, and 0.8 μl of 50 mM dNTP solution. Adjust the volume of the reaction mixture to 20 μl and add 1 μl of Multiscribe™ reverse transcriptase. Set up the reaction using the following conditions: 25
C for 10 min, 37
C for 120 min, 85
C for 5 min, hold at 4
C. Store the synthesized cDNA at 20
C. 7. Probe the synthesized cDNA with miR-150-3p primers (target gene) and Pf18s primers (as housekeeping control) by real time PCR analysis. Set up reactions of total volume 10 μl by adding 5 μl of Power Up SYBR green™ master mix, 0.5 μl of forward and reverse primer solution (each), 0.5 μl of template cDNA and 3.5 μl nuclease free water. Perform the reaction at the following conditions: Initial denaturation at 95
C for 5 min (1 cycle) followed by 40 cycles of denaturation at 95
C for 30 s, annealing at 60
C for 30 s and extension 72
C for 30 s. Set up the recording of Ct value at the primer annealing step and add melt curve analysis step post amplification. Calculate fold change in gene expression by the following formulae: Fold change in gene expression ¼ 2ΔΔCT, where ΔΔCT ¼ ΔCT(target gene)  ΔCT(reference gene). Also run the PCR product in 2% agarose gel and visualize under UV transilluminator. 3.3.2 Confirmation of the Translocation of miRNA Mimic in Parasites from the Cargo Loaded Erythrocytes Through Fluorescence Imaging

1. Load 20 μl of packed erythrocytes with 10 μl 3 biotinylated miR-197 (Thermo-Scientific, USA) mimic as mentioned in Subheading 3.1.1. 2. Set up infection of the loaded erythrocytes with purified schizonts at 1% parasitemia and 2% hematocrit.

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3. Prepare smears of the infected erythrocytes at 48 h post-invasion on clean glass slides. 4. Permeabilize the erythrocytes in the smears by dipping the smears in chilled methanol for 20 min and subsequently airdry the slides. 5. Prepare solution of Phycoerythrin conjugated streptavidin beads (Invitrogen, USA) in PBST with a concentration of 10 μg/ml. 6. Add 200 μl of PE conjugated bead suspension over control cargo and biotinylated mimic loaded erythrocyte smears and incubate for 2 h at RT. 7. Wash the smears 3 times with PBST (5 min for each wash). 8. Mount each smear with 10 μl of DAPI-Anti-fade reagent (Invitrogen, USA) and put clean cover slips over it. 9. Acquire images in confocal microscope (Olympus Corporation, Japan) at 100 magnification (DAPI: excitation wavelength 358 nm, emission wavelength 461 nm; PE: excitation wavelength 488 nm, emission wavelength 578 nm) (Fig. 3a, b). 3.3.3 Monitoring the Repression of the Target Protein

1. Purify schizont stage parasites using 65% percoll enrichment method. 2. Incubate miRNA loaded erythrocytes with percoll purified schizonts at 2% parasitemia and 4% hematocrit for 48 h. 3. Release the parasites from infected erythrocytes by saponin treatment (protocol explained above). Resuspend the parasite pellet in 50 μl RIPA buffer and incubate on ice for 30 min. 4. Centrifuge the lysate at 15,000  g for 15 min and collect the supernatant (approx. 40 μl). 5. Add 10 μl of 5 SDS loading buffer (0.25 M Tris–HCl pH 6.8, 10% SDS, 50% glycerol, 100 mM β-mercaptoethanol, and 0.25% bromophenol blue) to the collected supernatant, heat at 100
C for 5 min and load the samples onto a 12% SDS-polyacrylamide gel, along with prestained protein markers. Electrophorese the samples at 80 volts through the stacking gel and at 110 volts through the resolving gel (see Note 7). 6. Transfer the electrophoresed proteins from the gel onto the nitrocellulose membrane (pore size: 0.4 μm) in transfer buffer (25 mM Tris–HCl with pH 7.6, 192 mM glycine, 0.03% SDS, and 20% methanol) using semi-dry transfer method by applying a constant voltage of 17 volts for 30 min. 7. Block the membrane with 5% skimmed milk solution prepared in PBS O/N at 4
C.

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Fig. 3 (a) Images showing translocation of biotinylated miRNA mimic into the parasite (red channel) along with control erythrocytes with scale bars representing 5 μm distance. (b) graph showing increase in fluorescence intensity in parasites infecting mimic loaded erythrocytes due to binding of PE labelled streptavidin beads with the biotin group present at the 3’ end of the miRNA mimic (***p12 weeks). Strain differences may also affect yield. For reference, we typically generate 1  107 to 4  107 DCs from 5  107 plated bone marrow-derived cells. 3. Take care not to dislodge adherent cells when removing adherent cells on day 3 and harvesting DCs on day 7, as these are primarily DC progenitors and macrophages. 4. For electroporations, increasing the mRNA concentration improves expression; however, in our experience, this effect plateaus at around 50 pmol mRNA/106 DCs. 5. Be sure to use non-BSA-containing formulations of recombinant CCL3 protein, or any formulations without a carrier protein additive. There can be unwanted effects of carrier protein-stimulation on the cellular incubation assay, and stability of CCL3 over a long period of time is not required within this relatively brief assay. 6. Phenotypically, DCs grown in serum-free RPMI and AIMV are very similar; however, yields from serum-free RPMI are extremely poor (