Fields Virology, 7th ed., Volume 3 - RNA Viruses [7 ed.] 9781975112608

Table of contents :
Title
Contributors
Preface
Introduction
Contents
1 - Rhinoviruses
2 - Hepatoviruses
3 - Astroviruses
4 - Rubella Virus
5 - Arteriviruses
6 - Bornaviridae
7 - Rhabdoviridae
8 - Mumps Virus
9 - Measles Virus
10 - Respiratory Syncytial Virus and Metapneumovirus
11 - Orthoreoviruses
12 - Rotaviruses
13 - Orbiviruses
14 - Hepatitis E Virus
15 - Retroviridae
16 - Human T-Cell Leukemia Virus Types 1 and 2
17 - Human Immunodeficiency Viruses: Replication
18 - HIV-1: Pathogenesis, Clinical Manifestations, and Treatment
19 - Nonhuman Lentiviruses
20 - Foamy Viruses
21 - Severe Acute Respiratory Syndrome Coronavirus 2 (SARS-CoV-2)
22 - SARS-CoV-2/COVID-19: Clinical Characteristics, Prevention, and Treatment
Index

Citation preview

SEVENTH EDITION Fields VIROLOGY VOLUME 3: RNA Viruses Editors-in-chief Peter M. Howley, MD Shattuck Professor of Pathological Anatomy Departments of Immunology and Pathology Harvard Medical School Boston, Massachusetts David M. Knipe, PhD Higgins Professor of Microbiology and Molecular Genetics Head, Harvard Program in Virology Department of Microbiology Blavatnik Institute Harvard Medical School Boston, Massachusetts Associate Volume Editors Blossom Damania, PhD Boshamer Distinguished Professor Vice Dean for Research, School of Medicine Department of Microbiology and Immunology University of North Carolina at Chapel Hill Chapel Hill, North Carolina Jeffrey I. Cohen, MD Chief, Laboratory of Infectious Diseases National Institute of Allergy and Infectious Diseases National Institutes of Health Bethesda, Maryland Sean P. J. Whelan, PhD Marvin A. Brennecke Distinguished Professor Chair, Molecular Microbiology School of Medicine Washington University in St. Louis St. Louis, Missouri Eric O. Freed, PhD HIV Dynamics and Replication Program Center for Cancer Research National Cancer Institute

Frederick, Maryland Associate Editor Lynn Enquist, PhD Henry L. Hillman Professor of Molecular Biology Department of Molecular Biology Princeton University Princeton, New Jersey

Acquisitions Editor: Nicole Dernoski Development Editor: Ariel S. Winter Editorial Coordinator: Oliver Raj Editorial Assistant: Kristen Kardoley Marketing Manager: Kirsten Watrud Production Project Manager: Kirstin Johnson Manager, Graphic Arts & Design: Stephen Druding Manufacturing Coordinator: Beth Welsh Prepress Vendor: Straive Copyright © 2023 Wolters Kluwer All rights reserved. This book is protected by copyright. No part of this book may be reproduced or transmitted in any form or by any means, including as photocopies or scanned-in or other electronic copies, or utilized by any information storage and retrieval system without written permission from the copyright owner, except for brief quotations embodied in critical articles and reviews. Materials appearing in this book prepared by individuals as part of their official duties as U.S. government employees are not covered by the above-mentioned copyright. To request permission, please contact Wolters Kluwer at Two Commerce Square, 2001 Market Street, Philadelphia, PA 19103, via email at [email protected], or via our website at shop.lww.com (products and services). 987654321 Printed in Mexico Cataloging in Publication data available on request from publisher ISBN: 978-1-9751-1260-8 This work is provided “as is,” and the publisher disclaims any and all warranties, express or implied, including any warranties as to accuracy, comprehensiveness, or currency of the content of this work. This work is no substitute for individual patient assessment based upon healthcare professionals’ examination of each patient and consideration of, among other things, age, weight, gender, current or prior medical conditions, medication history, laboratory data and other factors unique to the patient. The publisher does not provide medical advice or guidance and this work is merely a reference tool. Healthcare professionals, and not the publisher, are solely responsible for the use of this work including all medical judgments and for any resulting diagnosis and treatments. Given continuous, rapid advances in medical science and health information, independent professional verification of medical diagnoses, indications, appropriate pharmaceutical selections and dosages, and treatment options should be made and healthcare professionals should consult a variety of sources. When prescribing medication, healthcare professionals are advised to consult the product information sheet (the manufacturer’s package insert) accompanying each drug to verify, among other things, conditions of use, warnings and side effects and identify any changes in dosage schedule or contraindications, particularly if the medication to be administered is new, infrequently used or has a narrow therapeutic range. To the maximum extent permitted under applicable law, no responsibility is assumed by the publisher for any injury and/or damage to persons or property, as a matter of products liability, negligence law or otherwise, or from any reference to or use by any person of this work. shop.lww.com

Contributors Larry J. Anderson, MD Professor of Infectious Diseases Department of Pediatrics Emory University School of Medicine and Children’s Healthcare of Atlanta Atlanta, Georgia Udeni B. R. Balasuriya, BVSc, MS, PhD, FSLCVS Professor of Virology and Director Louisiana Animal Disease Diagnostic Laboratory Department of Pathobiological Science School of Veterinary Medicine Louisiana State University Baton Rouge, Louisiana Charles R. M. Bangham, ScD, FRS, FMedSci Co-director Institute of Infection Imperial College, United Kingdom John H. Beigel, MD Associate Director for Clinical Research Division of Microbiology and Infectious Diseases National Institute of Allergy and Infectious Diseases National Institutes of Health Bethesda, Maryland Louis-Marie Bloyet, PhD Instructor Department of Molecular Microbiology School of Medicine Washington University in St. Louis St. Louis, Missouri Thomas Briese, PhD Associate Professor Department of Epidemiology Associate Director Center for Infection and Immunity Mailman School of Public Health Columbia University New York, New York Ursula J. Buchholz, PhD Chief

RNA Viruses Section Laboratory of Infectious Diseases National Institute of Allergy and Infectious Diseases National Institutes of Health Bethesda, Maryland Peter L. Collins, PhD Principal Investigator (Retired) RNA Viruses Section Laboratory of Infectious Diseases National Institute of Allergy and Infectious Diseases National Institutes of Health Bethesda, Maryland Valerie Cortez, PhD, MS Assistant Professor Department of Molecular, Cell & Developmental Biology University of California, Santa Cruz Santa Cruz, California Sue E. Crawford, PhD Assistant Professor Department of Molecular Virology and Microbiology Baylor College of Medicine Houston, Texas Terence S. Dermody, MD Vira I. Heinz Distinguished Professor and Chair of Pediatrics Professor of Microbiology and Molecular Genetics University of Pittsburgh School of Medicine Physician-in-Chief and Scientific Director UPMC Children’s Hospital of Pittsburgh Pittsburgh, Pennsylvania Ronald C. Desrosiers, PhD Professor University of Miami Miller School of Medicine Miami, Florida Siyuan Ding, PhD Assistant Professor Department of Molecular Microbiology School of Medicine Washington University in St. Louis St. Louis, Missouri W. Paul Duprex, PhD

Director, Center for Vaccine Research Professor of Microbiology and Molecular Genetics Center for Vaccine Research University of Pittsburgh Pittsburgh, Pennsylvania Mary K. Estes, PhD Distinguished Service Professor Molecular Virology and Microbiology and Medicine Baylor College of Medicine Houston, Texas David T. Evans, PhD Professor Pathology and Laboratory Medicine University of Wisconsin-Madison Madison, Wisconsin Ying Fang, PhD Professor Department of Pathobiology University of Illinois at Urbana-Champaign Urbana, Illinois Eric O. Freed, PhD HIV Dynamics and Replication Program Center for Cancer Research National Cancer Institute Frederick, Maryland James E. Gern, MD Professor of Pediatrics and Medicine University of Wisconsin School of Medicine and Public Health Madison, Wisconsin Stephen P. Goff, PhD Higgins Professor of Biochemistry Department of Biochemistry and Molecular Biophysics Department of Microbiology and Immunology Columbia University Medical Center New York, New York Stephen A. Goldstein, PhD Postdoctoral Research Associate Department of Human Genetics University of Utah School of Medicine Salt Lake City, Utah

Harry B. Greenberg, MD Professor and Associate Dean of Research Professor of Medicine and Microbiology and Immunology Stanford University Stanford, California Diane E. Griffin, MD, PhD University Distinguished Service Professor W. Harry Feinstone Department of Molecular Microbiology and Immunology Johns Hopkins Bloomberg School of Public Health Baltimore, Maryland Ottmar Herchenröder, PhD Research Scientist Institute of Experimental Gene Therapy and Cancer Research Rostock University Medical Center Rostock, Germany Christiane Herden, Prof Dr habil Full Professor Institute of Veterinary Pathology Justus-Liebig-University Giessen Giessen, Germany Tom C. Hobman, PhD, FCAHS Professor Department of Cell Biology University of Alberta Edmonton, Canada Brenda G. Hogue, MEd, PhD Professor Biodesign Institute School of Life Sciences Arizona State University Tempe, Arizona Daniel R. Kuritzkes, MD Chief, Division of Infectious Diseases; Harriet Ryan Albee Professor of Medicine Division of Infectious Diseases Brigham and Women’s Hospital and Harvard Medical School Boston, Massachusetts Robert LeDesma, PhD Department of Molecular Biology Princeton University Princeton, New Jersey

Julian L. Leibowitz, MD, PhD Professor and MD/PhD Program Director Department of Microbial Pathogenesis and Immunology Texas A&M University College of Medicine Bryan, Texas Stanley M. Lemon, MD Professor Departments of Medicine and Microbiology & Immunology UNC Lineberger Comprehensive Cancer Center The University of North Carolina at Chapel Hill Chapel Hill, North Carolina Dirk Lindemann, PhD Professor of Molecular Virology Institute of Medical Microbiology and Virology Medical Faculty Carl Gustav Carus University Hospital Carl Gustav Carus Technische Universität Dresden Dresden, Germany W. Ian Lipkin, MD John Snow Professor of Epidemiology and Director Center for Infection and Immunity Mailman School of Public Health Professor of Pathology and Neurology College of Physicians & Surgeons Columbia University New York, New York Masao Matsuoka, MD, PhD Professor Department of Hematology, Rheumatology, and Infectious Diseases Faculty of Life Sciences, Kumamoto University Kumamoto, Japan Asuncion Mejias, MD, PhD, MsCS Associate Professor of Pediatrics The Ohio State University College of Medicine Pediatrics, Division of Infectious Diseases Abigail Wexner Research Institute at Nationwide Children’s Hospital Columbus, Ohio Melanie Ott, MD, PhD Director, Senior Investigator Gladstone Institute of Virology

Senior Vice President Gladstone Institutes University of California San Francisco San Francisco, California Ann C. Palmenberg, PhD Professor Department of Biochemistry Institute for Molecular Virology University of Wisconsin-Madison Madison, Wisconsin John S. L. Parker, BVMS, PhD Associate Professor of Virology Baker Institute for Animal Health Cornell University Ithaca, New York Alexander Ploss, PhD Associate Professor of Molecular Biology Department of Molecular Biology Princeton University Princeton, New Jersey Linda J. Rennick, PhD Research Assistant Professor of Microbiology and Molecular Genetics Center for Vaccine Research University of Pittsburgh Pittsburgh, Pennsylvania Jürgen A. Richt, DVM, PhD Regents Distinguished Professor Department of Diagnostic Medicine/Pathology College of Veterinary Medicine Kansas State University Manhattan, Kansas Monica J. Roth, PhD Merck Research Laboratory Professor in Clinical Pharmacology Department of Pharmacology Rutgers-Robert Wood Johnson Medical School Piscataway, New Jersey Polly Roy, OBE, FMedSci, FRSB, PhD, MSc Professor of Virology Department of Infection Biology London School of Hygiene and Tropical Medicine

London, United Kingdom Steven A. Rubin, PhD Sr. Director, Global Regulatory Affairs GSK Rockville, Maryland Stacey Schultz-Cherry, PhD Full Member Department of Infectious Diseases St. Jude Children’s Research Hospital Senior Associate Dean St. Jude Children’s Research Hospital Graduate School of Biomedical Sciences Memphis, Tennessee Eileen P. Scully, MD, PhD Assistant Professor of Medicine Department of Medicine, Division of Infectious Diseases Johns Hopkins University Baltimore, Maryland Barbara Sherry, PhD Professor and Department Head Department of Molecular Biomedical Sciences College of Veterinary Medicine North Carolina State University Raleigh, North Carolina Eric J. Snijder, PhD Professor of Molecular Virology Department of Medical Microbiology Leiden University Medical Center Leiden, The Netherlands Timothy M. Uyeki, MD, MPH, MPP Chief Medical Officer Influenza Division, National Center for Immunization and Respiratory Diseases Centers for Disease Control and Prevention Atlanta, Georgia Susan R. Weiss, PhD Professor and Vice Chair Department of Microbiology Perelman School of Medicine University of Pennsylvania Philadelphia, Pennsylvania Sean P. J. Whelan, PhD

Marvin A. Brennecke Distinguished Professor Chair, Molecular Microbiology School of Medicine Washington University in St. Louis St. Louis, Missouri

Preface In the early 1980s, Bernie Fields originated the idea of a virology reference textbook that combined the molecular aspects of viral replication with the medical features of viral infections. This broad view of virology reflected Bernie’s own research, which applied molecular and genetic analyses to the study of viral pathogenesis, providing an important part of the foundation for the field of molecular pathogenesis. Bernie led the publication of the first three editions of Virology, but he unfortunately died soon after the third edition went into production. The third edition became Fields Virology in his memory, and it is fitting that the book continues to carry his name. A number of changes and enhancements have now been introduced with the seventh edition of Fields Virology. The publication format of Fields Virology has been changed from a twovolume book published every 5 to 6 years to an annual publication that comprises approximately one-fourth of the chapters organized by category. The annual publication provides both a physical book volume and importantly an eBook with an improved platform. Using an eBook format, our expectation is that individual chapters can be easily updated when major advances, outbreaks, etc., occur. The editorial board organized the four-volume series for the seventh edition to consist of volumes on Emerging Viruses, DNA Viruses, RNA Viruses, and Fundamental Virology, to be published on an annual basis, with the expectation that the topics will cycle approximately every 4 years creating an annualized, up-to-date publication. Each volume will contain approximately 20 chapters. The first volume of this seventh edition of Fields Virology, entitled Emerging Viruses, was published in 2020, and the second volume, DNA Viruses, was published in 2021. This third volume, RNA Viruses, has been principally edited by Eric O. Freed, Jeffrey I. Cohen, Blossom Damania, Peter M. Howley, David M. Knipe, and Sean P. J. Whelan. There have been continued rapid advances in virology since the previous edition, and all of the chapters in the RNA Viruses volume are either completely new or have been significantly updated to reflect these advances. In this seventh edition, we have chosen to highlight important references published since the last edition while maintaining older classics. The main emphasis continues to be on viruses of medical importance and interest, but other viruses are described in specific cases where more is known about their mechanisms of replication or pathogenesis. We wish to thank Patrick Waters of Harvard Medical School and all of the editorial staff members of Wolters Kluwer for all their important contributions to the preparation of this book. David M. Knipe, PhD Peter M. Howley, MD Jeffrey I. Cohen, MD Blossom Damania, PhD Lynn Enquist, PhD Eric O. Freed, PhD Sean P. J.Whelan, PhD

Introduction RNA Viruses is the third volume of the seventh edition of Fields Virology. The first two volumes, Emerging Viruses and DNA Viruses, were published in 2020 and 2021, respectively. The next volume, Fundamental Virology, will be published in 2023. There have been continued rapid advances in virology since the sixth edition that was published in 2013, and all of the chapters in the RNA Viruses volume are either completely new or have been significantly updated to reflect these advances. In this seventh edition, we have chosen to highlight important references published since the last edition while maintaining older classics. This volume covers all the RNA viruses of medical importance, including viruses that infect humans and animals. RNA viruses of plants, insects, and bacteria will be included in the fourth volume of the seventh edition, Fundamental Virology. In addition to RNA virus families, several thematic issues arise in the consideration of the RNA viruses in this volume. Similar to the DNA viruses, several of the RNA viruses are associated with persistent infections and as a consequence are associated directly or indirectly with cancer. This volume includes chapters that detail the major advances in our understanding of the molecular biology and disease pathogenesis associated with human immunodeficiency virus (HIV). Chapters on a number of RNA viruses were included in the first volume, Emerging Viruses, which with the exception of two new chapters on SARS-CoV-2, the emerging viruses in Volume 1 are not included in this third volume. We note that the Coronaviridae chapter in the Emerging Viruses volume was published just months prior to the current pandemic and therefore did not include SARS-CoV-2. The organization of the chapters in this edition is similar to that of the previous editions. For most of the viruses, there is a single chapter that combines both the basic and clinical aspects of the virus. For several of the viruses, the basic virology related to viral replication and the viral pathogenesis are split between two chapters. In the RNA volume, this includes the retroviruses, the human immunodeficiency viruses, and SARS-CoV-2. We are grateful to Eric O. Freed, Jeffrey I. Cohen, Blossom Damania, and Sean S. P. Whelan, who joined us to participate in putting this volume together. We also are thankful to the chapter authors who have updated their chapters for this volume and to the new authors who have joined us in this continued endeavor to provide a comprehensive resource in virology. Peter M. Howley David M. Knipe

Contents Contributors Preface Introduction 1 Rhinoviruses James E. Gern • Ann C. Palmenberg 2 Hepatoviruses Stanley M. Lemon 3 Astroviruses Valerie Cortez • Stacey Schultz-Cherry 4 Rubella Virus Tom C. Hobman 5 Arteriviruses Ying Fang • Eric J. Snijder • Udeni B. R. Balasuriya 6 Bornaviridae W. Ian Lipkin • Christiane Herden • Jürgen A. Richt • Thomas Briese 7 Rhabdoviridae Sean P. J. Whelan • Louis-Marie Bloyet 8 Mumps Virus Steven A. Rubin • Linda J. Rennick • W. Paul Duprex 9 Measles Virus Diane E. Griffin 10 Respiratory Syncytial Virus and Metapneumovirus Ursula J. Buchholz • Larry J. Anderson • Peter L. Collins • Asuncion Mejias 11 Orthoreoviruses Terence S. Dermody • John S. L. Parker • Barbara Sherry 12 Rotaviruses Sue E. Crawford • Siyuan Ding • Harry B. Greenberg • Mary K. Estes 13 Orbiviruses Polly Roy 14 Hepatitis E Virus Robert LeDesma • Alexander Ploss 15 Retroviridae Stephen P. Goff • Monica J. Roth 16 Human T-Cell Leukemia Virus Types 1 and 2 Charles R. M. Bangham • Masao Matsuoka 17 Human Immunodeficiency Viruses: Replication Melanie Ott • Eric O. Freed 18 HIV-1: Pathogenesis, Clinical Manifestations, and Treatment

Eileen P. Scully • Daniel R. Kuritzkes 19 Nonhuman Lentiviruses Ronald C. Desrosiers • David T. Evans 20 Foamy Viruses Dirk Lindemann • Ottmar Herchenröder 21 Severe Acute Respiratory Syndrome Coronavirus 2 (SARS-CoV-2) Stephen A. Goldstein • Brenda G. Hogue • Julian L. Leibowitz • Susan R. Weiss 22 SARS-CoV-2/COVID-19: Clinical Characteristics, Prevention, and Treatment John H. Beigel • Timothy M. Uyeki Index

CHAPTER 1 Rhinoviruses James E. Gern • Ann C. Palmenberg History Infectious Agent Classification Physical Characteristics Pathogenesis and Pathology Entry into the Host Site of Primary Replication Spread Cell and Tissue Tropism Immune Response Release from Host and Mode of Transmission Virulence Persistence Epidemiology Age Morbidity Origin and Spread of RV Infections Prevalence and Seroepidemiology Genetic Diversity of Virus Clinical Features Diagnosis Differential Laboratory Prevention and Control Perspective

HISTORY The common cold has been recognized for millennia, and both the illness and prescribed remedies have been influenced by and engendered a broad body of folklore. In fact, the name of the illness stems from the belief that being chilled caused the illness, a concept which studies using experimental inoculation techniques have been unable to verify.61 In Chinese traditional medicine, colds were considered an illness of wind and cold, and the Roman physician Galen wrote “The white-colored substance (the phlegma) collects mostly … in those who have been chilled in some way”.14 Through the ages, ideas about pathogenesis varied widely, and suggestions for common cold cures were creative, occasionally bizarre, and seldom helpful. Yet, enthusiasm for a cure for the common cold remains quite high today; a recent web-based search yielded 211,000,000 hits in response to the terms “common cold cure.”

In 1914, Kruse demonstrated that cell-free filtrates of nasal secretions from affected individuals could transmit colds.149 Dochez et al. confirmed these findings in 1930 by transmitting colds to volunteers and apes using filtered nasal secretions that were free of bacteria, indicating a viral etiology.58 Progress in finding the underlying cause for colds was accelerated immediately after World War II by the establishment of the Common Cold Unit (CCU) by the UK Medical Research Council in 1946.256 Housed in an hospital unit founded by Harvard University and the American Red Cross to support Great Britain during the war, the building was eventually donated to the British government. At that time, it was presumed the cold-causing virus was different from influenza.256 The goals of the CCU, as led by Christopher Andrewes and later David Tyrrell, were to identify the common cold virus, its means of transmission, and describe the host characteristics that sometimes promoted more severe illness. Between 1946 and 1989, when the unit closed, over 20,000 volunteers had participated in those studies. 1

The first rhinoviruses (RVs) were isolated by two independent groups,201,210 starting around 1956. It was not long before researchers realized that several different virus families actually contributed to common cold illnesses, and that among these the RV included numerous serotypes. A collaborative program in 1967 classified all then-known isolates into 55 different serotypes.218 Serotypes 56 to 89 were added in 1971,1 and the remainder of the classic 101 RV-A&B in 1987.97 More recent molecular sequencing techniques led to the description of the genetically distinct RV-C species whose isolates do not grow in standard tissue culture.155 These viruses, along with several previously unknown RV-A&B genotypes, initiated a renewed period of discovery related to RV epidemiology, their extended classification, and many additional insights into the role of specific RV not only in common colds but also otitis, sinusitis, lower respiratory infections, and acute exacerbations of chronic respiratory diseases such as asthma.

INFECTIOUS AGENT Classification The RVs (formerly Human Rhinoviruses, HRVs) comprise the RV-A, RV-B, and RV-C species of the Enterovirus genus in the Picornaviridae family. Present classification is based on overt similarities in genome organization, capsid properties, and primary sequence conservation.195 Viruses are assigned to a species if they share greater than 70% amino acid identity in the P1, 2C, and 3CD regions with other members. Within species, isolates are subdivided into numeric genotypes (Fig. 1.1). For the RV-A&B, the foundation assignments included the historic clinical panels archived by the American Type Culture Collection, which were indexed into the 101 original serotypes after assessment of antigenic cross-reactivity in rabbits or guinea pigs. RV-A87 was subsequently reassigned to the Enterovirus D (EV-D68) after reevaluation of genetic, immunogenic, and receptor use (decay-accelerating factor as a receptor) properties.225 Ultimately, the difficulty of testing clinical viruses for immunogenicity made serotyping obsolete, and such cataloging was never actively employed for any RV-C. Instead, current genotype assignments respect the historic RV-A&B index system, but now rely heavily on sequence comparisons, primarily of the VP1 region or VP4/VP2. Strains in a common genotype, including most of the historic serotypes, share a general threshold of approximately 11% nucleotide divergence in their VP1 genes.233 Full genome sequencing showed that some original types were more closely related than this and warranted reassignment (e.g., A44, A95, and A98 were abolished and the isolates combined into A29, A8, and A54, respectively), while others such as A8 and A45 defining “clade D” may be sufficiently different for eventual designation as a fourth species (Fig. 1.1). Many newly described RV-C genomes are not yet fully sequenced, so isolate genotyping is based primarily on VP1 genetic threshold data. To prevent ambiguity, RV-A (80 types), RV-B (32 types), and RV-C (57 types) nomenclature conventions require both the species letter and genotype assignment (e.g., A16, B14, C15). The RV designation (e.g., RV-A16) may be included for clarity.

FIGURE 1.1 Circle phylogram relationships for known genotypes of RV-A, RV-B, and RV-C. The tree was calculated with neighbor-joining methods from aligned, VP1 RNA sequences and rooted with data from Enterovirus (EV) A, B, and C species, similar to Ref.196 All RV-A&B are of the Major (ICAM-1) receptor group except those 10 labeled with stars (Minor receptor group, LDLR). The RV-C receptor is CDHR3. Bootstrap values (percent of 2,000 replicates) are indicated at key nodes. Genbank accession numbers for this dataset are in eTable 1.1.

Physical Characteristics As enteroviruses, the RV have genome organizations (Fig. 1.2) and capsid structures (Fig. 1.3) similar to those of polioviruses, coxsackieviruses, and ECHO viruses. But unlike other enteroviruses that remain viable at pH 3.0, RV particles are unstable below pH 5 to 6. The icosahedral capsid (~30 nm diameter) has 60 copies each of proteins VP1, VP2, VP3, and VP4, named in order of descending electrophoretic mobility. The four proteins derived from a common P1 precursor stay together during assemble as a biological subunit, or protomer. The protein shell surrounds a densely packed, single-stranded, positive-sense, RNA genome of 7,079 (C1) to 7,233 (B92) bases, a count which does not include the variable length 3′ poly(A) tail. Several RV capsids have been resolved atomic resolution, including A1 (1r1a), A2 (1fpn), A16 (1ayn), B3 (1rhi), B14 (4RV), and C15 (5k0u), with multiple other determinations showing receptor interactions (e.g., A16 with ICAM-1, 1d3e; C15 with CDHR3, 6psf)) or antiviral drug interactions (e.g., A16 with pleconaril, 1c8m). Like poliovirus, the surfaces of RV capsids are dominated by the three largest proteins (Fig. 1.3). VP4 is internal to the structure, centered near the fivefold axis. Around the 12 raised exterior fivefold VP1 2

“plateaus,” a symmetrical “canyon” provides receptor binding sites and discontinuous immunogenic surfaces230 to each RV-A&B. The C15 structure has a different surface topology, including three major VP1 deletions, which lower the fivefold plateau and remove most of the surrounding canyon features. Individual RV-C genotypes compensate for this loss of mass with characteristic unique-sequence, highly immunogenic “finger-like” surface loops contributed by a relative insertion near the C-terminus of VP1.163 Common to all RV, the VP1 cores surround a hydrophobic “pocket” or cavity. Type-1 long (e.g., B14) or type-2 short (e.g., A16) pocket shapes are defined for most RV-A&B and determine whether a particular virus is susceptible to drugs like pleconaril or WIN compounds aimed at inhibiting the uncoating process.7 The RV-C VP1 pockets have partially collapsed shapes, inaccessible to drug diffusion, rendering these isolates refractive to similar antiviral therapies.163

FIGURE 1.2 The genome of an RV encodes a single polyprotein open reading frame (A). Important RNA structural motifs include a 5′ cloverleaf (B), ORF start-site stem (C), a cis-acting replication (cre) element (D), and 3′ stem motif. All RV genomes are messenger-sense, encoding the polyprotein reading frame (ORF) and multiple important RNA structural motifs (Fig. 1.2). Adjacent to the 5′ cloverleaf, a regulatory feature for translation and replication, each RV encodes a strain-specific pyrimidine-rich tract that may be involved in suppressing innate immunity triggers.196 The type-1 IRES 3′ to this tract includes a variable-length stem structure pairing the ORF start site (AUG) with an upstream AUG. Unlike poliovirus, intervening sequences between these AUGs are probably not scanned by initiating ribosomes.121 The picornavirus VPg uridylylation reaction, required for RNA synthesis, is templated by a secondary structure called the cre (cis-acting replication element) whose location varies in every species of picornavirus. For the RV-A, the cre is in the 2A gene.243 For the RV-B, the cre is in the 2C gene.243 The RV-C cre is in the 1B gene.24 The short, 3′ untranslated sequences (UTR) are of highly variable sequence. Invariably, they configure as an inclusive stem motif displaying at least one bogus termination codon in the terminal loop. This codon may be inframe or out-of-frame with the authentic ORF stop site and has been proposed to play a role in the recruitment of translation termination factors.196

3

FIGURE 1.3 Comparative capsids A16 (1aym), B14 (4rhv), and C15 (5k0u) illustrate the icosahedral surface topography of VP1 (blue), VP2 (green), and VP3 (red) proteins. The VP4 is internal to the capsid. Antiviral capsid drugs (RVA&B) bind in a pocket, internal to VP1. All VP4 proteins have N-terminal myristate modifications, although these are rarely characterized in structure determinations. Throughout this chapter, imaged or referenced Protein Database (PDB) structure identification numbers are indicated parenthetically. Images courtesy of Dr. Jean-Yves Sgro (University of Wisconsin-Madison). In general, the RV lifecycle after infection is that of canonical enteroviruses, requiring 14 to 24 hours for cell death and lytic progeny release. The 2A and 3C proteases embedded in the polyprotein undergo monomolecular and/or bimolecular reactions to release mature enzymes or their precursors, both for further polyprotein cleavage, and for a variety of cellular proteolytic events to inactivate innate immunity functions, aid IRES-dependent translation, and rearrange cell processes advantageous to the virus. Among these, RV strain-dependent, 2A-catalyzed cleavage of multiple Phe/Gly-containing nucleoporin proteins (Nups) and eIFG translation factors may elicit different cell responses and cellular translational shutoff that directly or indirectly leads to distinct disease phenotypes.270 Also like other enteroviruses, RV infection induces significant rearrangements in cellular membranes and their lipid components, including the ER and trans-golgi network as they establish protective compartments favorable to RNA replication, genome translation, and progeny assembly.220 Proposed interference in these pathways, as essential viral replication requirements, may suggest possible novel antiviral targets.

PATHOGENESIS AND PATHOLOGY Entry Into the Host The primary portal of entry for RV infections is through inoculation of either the eyes or nose. Studies of seronegative infected volunteers have shown that very low doses of RV, substantially less than the amount needed to infect cells in vitro (1 tissue culture infectious dose50 [TCID50]), can cause infection when introduced via the conjunctiva or nasal mucosa. In contrast, approximately 10,000 times as much virus is needed to cause productive infection when the inoculation site is the tongue or external nares.50

Site of Primary Replication The primary site of infection is the airway epithelium. In studies of airway tissues from either natural or experimentally induced colds, detection of viral protein or RNA is largely confined to the epithelial layer, along with an occasional cell in the subepithelial layer.11,26,185 The RV-A and RV-C mainly replicate in ciliated epithelial cells89,125 (Fig. 1.4). RV-A&B may also replicate in specialized nonciliated epithelial cells in the adenoids that express high levels of ICAM-1.277

Spread For years it was assumed that RV infection was confined to the upper airway and did not affect the chest except under unusual circumstances. Early experiments in cell culture systems indicated that RV replicate best at 33°C to 35°C, and it was assumed that warmer temperatures in the lower airways (37°C) could limit RV infections to the cooler upper airways. Contrary to these initial assumptions, direct measurements in the lower airways have shown that large and medium size airways are at the ideal temperature for RV replication (Fig. 1.5).175 More recent replication studies demonstrate that RV-C and many RV-A types can replicate very well at 37°C.13,198 In addition, cultured lower airway epithelial cells support RV replication in vitro at least as well, and perhaps even better, than cells derived from the upper airways.167,184 Interestingly, cooler temperatures can inhibit cellular antiviral responses such as apoptosis and RNAseL, which could permit greater viral replication in the cooler temperatures of the upper airways.76

Following experimental inoculation of the upper airway, RV has been detected in the lower airways of individuals with a variety of techniques. Secretions from the lower airways sampled by bronchoscopy and bronchial lavage were analyzed by RT-PCR, and more than half of the lower airway specimens tested positive for RV.81,229 These findings were extended by subsequent studies demonstrating the presence of intracellular 4

RV RNA and protein using in situ hybridization and immunohistochemistry.185,197 Analysis of cells in sputum has been used to provide an estimate of the quantity and kinetics of viral shedding from the large lower airways. After experimental inoculation of seronegative volunteers, viral shedding from the upper airway peaks 2 to 4 days later. In contrast, peak levels of virus in the sputum occurs 3 to 7 days after inoculation.185 In addition, about half of the volunteers have viral shedding in the sputum that is equal to or exceeds that found in the nasal secretions. Notably, only small amounts of virus are detected in bronchial lavage specimens, which originate from distal airways and alveoli.185 These findings suggest that RV infections likely begin in the upper airway and can spread to the large and medium-sized airways during some illnesses.

FIGURE 1.4 RV-C15 replication within ciliated epithelial cells in tissue culture. Primary human bronchial epithelial cells were differentiated at air–liquid interface that then inoculated either RV-C15 (A) or medium alone (B). The cells were stained for C15 capsid and imaged by light microscopy. C15 capsid is represented by brown-staining cells. Scale bars indicate 20 μm. (Reprinted from Griggs TF, Bochkov YA, Basnet S, et al. Rhinovirus C targets ciliated airway epithelial cells. Respir Res 2017;18(1):84. https://creativecommons.org /licenses/by/4.0/.)

FIGURE 1.5 Temperatures in the large lower airways are ideal for RV replication. Direct measurements of temperature in lower airways have been recorded at measured distances from the nares using a bronchoscope equipped with a small thermistor.175 Even when the inspired air is at room temperature (26.7°C), airway temperatures in the medium and large airways are in the range of 33°C to 35°C. In contrast, small airway temperatures approach core temperature (37°C). Abbreviations: insp, inspiration; exp, expiration.

Natural cold studies also confirm that RV can replicate in the lower airways. Viral recovery from sputum during colds can exceed that obtained from upper airway samples.113,266 In epidemiologic studies, RV is frequently detected in children and adults with lower airway illnesses such as bronchiolitis, exacerbations of asthma, and pneumonia.122,123,179,180,204 In wheezing infants, RV has been detected in lower airway biopsies, and RV detection was associated with reduced lung function in these infants.170 In children with tracheostomies, samples of nasal mucus can be obtained directly from the lower airway without contamination from nasal secretions, and RV detection rates from upper versus lower airway specimens are similar.235

In addition to infecting the nasopharynx, conjunctiva, and lower airways, RV can be recovered in specimens obtained from the middle ear and sinuses.38,205 The respiratory epithelium in these locations is contiguous with that of the nasopharynx, and the virus presumably spreads via local extension. Rhinovirus viremia has been detected in infected children by PCR and appears to be more common during RV-C illnesses.168,281 A study of experimentally infected adults showed no evidence of circulating RV RNA (45). RVs are inactivated at pH < 6, thus preventing swallowed virus from replicating in the gastrointestinal tract.90

Cell and Tissue Tropism Biopsies of the upper airway from infected volunteers show a patchy pattern of epithelial infection with small foci of infected cells.10,206 Point cultures of the airway have demonstrated high levels of RV shedding in the nasopharynx and especially in the adenoidal region.278 Examination of biopsies obtained during experimentally induced colds suggests that a specific type of nonciliated adenoidal epithelial cell expresses high levels of ICAM-1 and supports high-level viral replication.277 It is possible that these cells play a sentinel role in the detection of viral respiratory infections.

5

FIGURE 1.6 The RV receptors. Comparative schematics of human ICAM-1, LDLR, and CDHR3 show the virus interactive domains among the various extracellular repeat units. The receptors used by RV are also expressed by airway cells other than epithelial cells. Besides epithelial cells, RV can bind to macrophages, monocytes, B cells, eosinophils, and fibroblasts.80,84,98 Macrophages and monocytes are good sources of type I and type III interferons, which may explain why there is little or no RV replication in these cells.146 Airway fibroblasts84 support RV replication in tissue culture, but it has not been established whether these cells, which are located several cell layers under the epithelial surface, are infected in vivo.

Receptors All RV-A&B use intercellular adhesion molecule–1 (ICAM-1, 101 “major” types) or alternatively, low, or very low-density lipoprotein receptor (LDLR, VLDLR, 10 “minor” types) for recognition and attachment to cells88,263 (Fig. 1.6). The RV-C instead require interactions with CDHR3, cadherin-related family member 3.27 This protein is coexpressed with FOX-J1, a transcription factor that regulates the differentiation of ciliated cells,20 but the precise native function of CDHR3 remains unknown. While the ICAM-1 gene is ubiquitous to all humans, the CDHR3 display density on the apical surfaces of ciliated pulmonary epithelial cells is dependent upon a nucleotide polymorphism (rs6967330) encoding a Tyr529 to Cys529 substitution in the protein’s extracellular repeat domain 5 (EC5), an allele change, which is also among the strongest known genetic correlates for childhood virus-induced asthma susceptibility.20,30 The asthmatic risk sequence (Tyr529), ancestral to all human and primate lineages, confers a 5- to 50-fold RV-C enhanced infection susceptibility to cultured or primary cells.20,29,271 The asthma-protective allele (Cys529), by far the dominant genotype in world-wide modern human populations, is maintained by balancing selection193 and prevents the encoded protein from properly exiting the ER after synthesis, reducing the cell surface concentration and availability to virus.

Biochemistry and structure determinations show all receptor interactions with their respective RV are through N-proximal protein domain contacts with discrete virion binding footprints (Fig. 1.7). Structure examples include B14 with soluble ICAM-1 domains 1 to 2 (1d3i), A2 with domain 3 VLDLR (3dpr), and C15 with CDHR3 EC1 (6psf). In addition, the major group virus A89 can bind to heparin sulfate in cultured cells, in the absence of ICAM-1262 as can C15 if adapted to do so through a single VP1 capsid mutation (Thr125Lys) near the fivefold axis.28 Studies utilizing atopic force spectroscopy indicate that after initial RV binding, multiple receptor proteins (i.e., ICAM-1) are rapidly recruited (within 200 milliseconds).217

Species Tropism RV primarily infect humans. Chimpanzees can be experimentally infected with RV-A&B, but while viral shedding is detected, there are few symptoms of illness.115 In contrast, a recent RV-C outbreak in a wild chimpanzee population produced severe illness in almost 100% of the individuals, several of which died.226 Given that all nonhuman primates share the susceptible Tyr529 CDHR3 receptor allele, such epizootics are presumably triggered through human contact, as these viruses are not known to circulate naturally in nonhuman populations. RV-A&B do not bind to murine ICAM-1, but the minor group viruses can bind to murine LDLR283 and replicate in murine epithelial cells.32,255 Certain RV strains have been specifically selected for this property by serial passage in murine cell lines.17,100 Epithelial cells from transgenic mice engineered to express human ICAM-1 also support replication of major group viruses.17 There are no current equivalent transgenic mice displaying human CDHR3 (Tyr529).

The demonstration that any RV-A&B can grow even marginally in murine epithelial cells has led to the development of murine models for those RV infection. The models resemble some features of human infections, including replication in the respiratory epithelium, induction of type I interferons and neutrophil chemokines, and neutrophilic airway inflammation.17,112 In addition, RV infection of mice increases inflammation in response to allergen exposure, suggesting that these models could be informative for asthma.17 Experimental limitations include the requirement for a large inoculating dose, and short duration (6 months of age) going to regions of high HAV endemicity, persons who will have close contact with children adopted from countries with high rates of hepatitis A, and those who live in group settings where a high proportion of individuals have risk factors for HAV infection.274,275 As discussed above (see “Passive Immunoprophylaxis”), immunization with hepatitis A vaccine is also recommended for postexposure prophylaxis.275 Vaccine should be administered as soon as possible after exposure and is protective if given within 2 weeks of exposure.391 Because immune response to the vaccine may be blunted in older individuals, IG may also be administered at a different anatomic site to those over the age of 40 at the discretion of the provider.275 Protection is evident within 2 weeks of primary immunization and has been demonstrated in persons immunized as late as 2 weeks after exposure to the virus.391,413 Mathematical modeling suggests that protective levels of antibody will be present in greater than 99% of persons 25 to 50 years after immunization.249 It is thus likely that vaccine immunity will persist for life without the need for booster immunization.387 Universal childhood immunization is recommended in many high-income, low-endemicity countries, and increasingly considered in middleincome countries transitioning from high-to-low endemicity with paradoxical increases in the incidence of symptomatic infection172,228,340 (see “Epidemiology”). A one-dose regimen has been used successfully for universal childhood immunization in Argentina.394 A combined hepatitis A and typhoid vaccine has been approved for use in some countries.211 Live Attenuated HAV Vaccine Cell culture passage reduces the capacity of HAV to replicate and to induce both disease and immunity to infection in nonhuman primates and humans.128,129,259,294,295 This change in the virus phenotype is associated with mutations in 2B and other nonstructural proteins.102,103,146 Poor immunogenicity of a candidate attenuated vaccine, coupled with early evidence of high immunogenicity of formaldehyde-inactivated HAV led to the abandonment of most efforts to develop attenuated vaccines.30,298 However, attenuated HAV vaccines have been successfully developed in China from cell culture–passaged virus variants31,244,245 and are used in multiple countries.73,74,228 These attenuated HAV vaccines have not been studied in the same detail as inactivated vaccines. They are somewhat less immunogenic than formalin-inactivated HAV vaccines, yet appear to have similar efficacy and to be safe.74 Reversion of these attenuated vaccines to virulence has not been noted clinically.

PERSPECTIVES Few viruses have undergone such a major change in our understanding of their biology as has HAV over the past decade. Recognition of the role played by quasi-envelopment in the pathogenesis of hepatitis A,112 the marked differences existing between hepatoviruses and enteroviruses in both capsid structure and mechanisms of assembly and cell entry,81,405 and the discovery of multiple hepatovirus species infecting a wide range of mammals96 have dramatically altered our perspectives on HAV. However, many aspects of the HAV life cycle remain to be discovered, and there are certain to be additional surprises lying ahead of us. Perhaps the biggest unknown is the potential for non-Hepatovirus A viruses to infect humans. Hepatoviruses circulating in bat populations appear to share not only antigenicity with human HAV but also similar capacities to disrupt innate immune antiviral responses in human cells.96,117 Hepatovirus phylogeny shows stark evidence of multiple trans-species jumps in the past, and this should perhaps be a warning to us for the future. Yet, the challenges are not all scientific. Recent increases in the rates of infection and numbers of HAV-associated deaths linked to homelessness and drug use in the United States (Fig. 2.14)122 highlight an urgent need to intensify efforts to immunize vulnerable populations against this vaccine-preventable disease.

ACKNOWLEDGMENTS The author gratefully acknowledges Zongdi Feng, Chris Walker, Jason Whitmire, and Kevin McKnight for critical review of the manuscript. Recent hepatovirus research in the author’s laboratory has been supported by the National Institute of Allergy and Infectious Diseases under grants R01-AI131685, R01-AI103083, and R01-AI150095.

51

References 1. Abe K, Shikata T. Fulminant type A viral hepatitis in a chimpanzee. Acta Pathol Jpn 1982;32:143–148. 2. Abernathy E, Mateo R, Majzoub K, et al. Differential and convergent utilization of autophagy components by positive-strand RNA viruses. PLoS Biol 2019;17:e2006926. 3. Advisory Committee on Immunization Practices. Prevention of hepatitis A through active or passive immunization: recommendations of the Advisory Committee on Immunization Practices (ACIP). MMWR Recomm Rep 2006;55:1–23. 4. Aggarwal R, Goel A. Hepatitis A: epidemiology in resource-poor countries. Curr Opin Infect Dis 2015;28:488–496. 5. Ali IK, McKendrick L, Morley SJ, et al. Activity of the hepatitis A virus IRES requires association between the cap-binding translation initiation factor (eIF4E) and eIF4G. J Virol 2001;75:7854–7863. 6. Allaire M, Chernala MM, Malcolm BA, et al. Picornaviral 3C cysteine proteinases have a fold similar to chymotrypsin-like serine proteinases. Nature 1994;369:72–76. 7. Amado LA, Marchevsky RS, de Paula VS, et al. Experimental hepatitis A virus (HAV) infection in cynomolgus monkeys (Macaca fascicularis): evidence of active extrahepatic site of HAV replication. Int J Exp Pathol 2010;91:87–97. 8. Anderson DA, Ross BC. Morphogenesis of hepatitis A virus: isolation and characterization of subviral particles. J Virol 1990;64:5284–5289. 9. Anderson DA, Ross BC, Locarnini SA. Restricted replication of hepatitis A virus in cell culture: encapsidation of viral RNA depletes the pool of RNA available for replication. J Virol 1988;62:4201–4206. 10. Angarano G, Trotta F, Monno L, et al. Serum IgA anti-hepatitis A virus as detected by enzyme-linked immunosorbent assay. Diagnostic significance in patients with acute and protracted hepatitis A. Diagn Microbiol Infect Dis 1985;3:521–523. 11. Anthony SJ, St Leger JA, Liang E, et al. Discovery of a novel Hepatovirus (phopivirus of seals) related to human hepatitis A virus. mBio 2015;6(4):e01180-15. 12. Aragones L, Guix S, Ribes E, et al. Fine-tuning translation kinetics selection as the driving force of codon usage bias in the hepatitis A virus capsid. PLoS Pathog 2010;6:e1000797. 13. Armstrong GL, Bell BP. Hepatitis A virus infections in the United States: model-based estimates and implications for childhood immunization. Pediatrics 2002;109:839–845. 14. Asare E, Mugavero J, Jiang P, et al. A single amino acid substitution in poliovirus nonstructural protein 2CATPase causes conditional defects in encapsidation and uncoating. J Virol 2016;90:6174–6186. 15. Asher LVS, Binn LN, Mensing TL, et al. Pathogenesis of hepatitis A in orally inoculated owl monkeys (Aotus trivirgatus). J Med Virol 1995;47:260–268. 16. Atkinson NJ, Witteveldt J, Evans DJ, et al. The influence of CpG and UpA dinucleotide frequencies on RNA virus replication and characterization of the innate cellular pathways underlying virus attenuation and enhanced replication. Nucleic Acids Res 2014;42:4527–4545. 17. Avanzino BC, Fuchs G, Fraser CS. Cellular cap-binding protein, eIF4E, promotes picornavirus genome restructuring and translation. Proc Natl Acad Sci U S A 2017;114:9611–9616. 18. Banerjee R, Dasgupta A. Interaction of picornavirus 2C polypeptide with the viral negative-strand RNA. J Gen Virol 2001;82:2621–2627. 19. Barker MH, Capps RB, Allen FW. Acute infectious hepatitis in the Mediterranean theater: including acute hepatitis without jaundice. JAMA 1945;128:997–1003. 20. Baruah V, Tiwari D, Hazam RK, et al. Prognostic, clinical, and therapeutic importance of RANTES-CCR5 axis in hepatitis A infection: a multiapproach study. J Med Virol 2021;93:3656–3665. 21. Belkaya S, Michailidis E, Korol CB, et al. Inherited IL-18BP deficiency in human fulminant viral hepatitis. J Exp Med 2019;216:1777–1790. 22. Bellou M, Kokkinos P, Vantarakis A. Shellfish-borne viral outbreaks: a systematic review. Food Environ Virol 2013;5:13–23. 23. Beneduce F, Ciervo A, Kusov Y, et al. Mapping of protein domains of hepatitis A virus 3AB essential for interaction with 3CD and viral RNA. Virology 1999;264:410–421. 24. Beneduce F, Ciervo A, Morace G. Site-directed mutagenesis of hepatitis A virus protein 3A: effects on membrane interaction. Biochim Biophys Acta 1997;1326:157–165. 25. Benenson MW, Takafuji ET, Bancroft WH, et al. A military community outbreak of hepatitis type A related to transmission in a child care facility. Am J Epidemiol 1980;112:471–481. 26. Bergmann EM, Cherney MM, McKendrick J, et al. Crystal structure of an inhibitor complex of the 3C proteinase from hepatitis A virus (HAV) and implications for the polyprotein processing in HAV. Virology 1999;265:153–163. 27. Bergmann EM, Mosimann SC, Chernaia MM, et al. The refined crystal structure of the 3C gene product from hepatitis A virus: Specific proteinase activity and RNA recognition. J Virol 1997;71:2436–2448. 28. Berthillon P, Crance JM, Leveque F, et al. Inhibition of the expression of hepatitis A and B viruses (HAV and HBV) proteins by interferon in a human hepatocarcinoma cell line (PLC/PRF/5). J Hepatol 1996;25:15–19. 29. Bienz K, Egger D, Pasamontes L. Association of polioviral proteins of the P2 genomic region with the viral replication complex and virusinduced membrane synthesis as visualized by electron microscopic immunocytochemistry and autoradiography. Virology 1987;160:220–226. 30. Binn LN, Bancroft WH, Lemon SM, et al. Preparation of a prototype inactivated hepatitis A virus vaccine from infected cell cultures. J Infect Dis 1986;153:749–756. 31. Binn LN, Lemon SM. Hepatitis A. In: Artenstein AW, ed. Vaccines, a Biography. New York: Springer-Verlag; 2010:335–346. 32. Binn LN, Lemon SM, Marchwicki RH, et al. Primary isolation and serial passage of hepatitis A virus strains in primate cell cultures. J Clin Microbiol 1984;20:28–33. 33. Binn LN, Macarthy PO, Marchwicki RH, et al. Laboratory tests and reference reagents employed in studies of inactivated hepatitis A vaccine. Vaccine 1992;10(Suppl 1):S102–S105. 34. Bishop NE, Anderson DA. RNA-dependent cleavage of VPO capsid protein in provirions of hepatitis A virus. Virology 1993;197:616–623. 35. Blank CA, Anderson DA, Beard M, et al. Infection of polarized cultures of human intestinal epithelial cells with hepatitis A virus: vectorial release of progeny virions through apical cellar membranes. J Virol 2000;74:6476–6484. 36. Boggs JD, Melnick JL, Conrad ME, et al. Viral hepatitis: clinical and tissue culture studies. JAMA 1970;214:1041–1046. 37. Borman AM, Kean KM. Intact eukaryotic initiation factor 4G is required for hepatitis A virus internal initiation of translation. Virology 1997;237:129–136. 38. Borman AM, Michel YM, Kean KM. Detailed analysis of the requirements of hepatitis A virus internal ribosome entry segment for the eukaryotic initiation factor complex eIF4F. J Virol 2001;75:7864–7871. 39. Borovec SV, Anderson DA. Synthesis and assembly of hepatitis A virus-specific proteins in BS-C-1 cells. J Virol 1993;67:3095–3102. 52

40. Boyer JL. Bile formation and secretion. Compr Physiol 2013;3:1035–1078. 41. Bozkurt H, D'Souza DH, Davidson PM. Determination of thermal inactivation kinetics of hepatitis A virus in blue mussel (Mytilus edulis) homogenate. Appl Environ Microbiol 2014;80:3191–3197. 42. Bradley DW, Schable CA, McCaustland KA, et al. Hepatitis A virus: growth characteristics of in vivo and in vitro propagated wild and attenuated virus strains. J Med Virol 1984;14:373–386. 43. Brown EA, Day SP, Jansen RW, et al. The 5′ nontranslated region of hepatitis A virus: secondary structure and elements required for translation in vitro. J Virol 1991;65:5828–5838. 44. Brown EA, Jansen RW, Lemon SM. Characterization of a simian hepatitis A virus (HAV): antigenic and genetic comparison with human HAV. J Virol 1989;63:4932–4937. 45. Brown EA, Zajac AJ, Lemon SM. In vitro characterization of an internal ribosomal entry site (IRES) present within the 5′ nontranslated region of hepatitis A virus RNA: Comparison with the IRES of encephalomyocarditis virus. J Virol 1994;68:1066–1074. 46. Cao L, Liu P, Yang P, et al. Structural basis for neutralization of hepatitis A virus informs a rational design of highly potent inhibitors. PLoS Biol 2019;17:e3000229. 47. Carette JE, Raaben M, Wong AC, et al. Ebola virus entry requires the cholesterol transporter Niemann-Pick C1. Nature 2011;477:340–343. 48. Centers for Disease Control and Prevention. Hepatitis Surveillance Statistics. 2019. https://www.cdc.gov/hepatitis/statistics /2019surveillance/index.htm. Accessed July 4, 2021. 49. Centers for Disease Control and Prevention. Hepatitis Outbreaks. https://www.cdc.gov/hepatitis/outbreaks/2017March-HepatitisA.htm. Accessed March 9. 50. Centers for Disease Control and Prevention (CDC). Positive test results for acute hepatitis A virus infection among persons with no recent history of acute hepatitis—United States, 2002–2004. MMWR Morb Mortal Wkly Rep 2005;54:453–456. 51. Chang KH, Brown EA, Lemon SM. Cell type-specific proteins which interact with the 5′ nontranslated region of hepatitis A virus RNA. J Virol 1993;67:6716–6725. 52. Chen CM, Chen SC, Yang HY, et al. Hospitalization and mortality due to hepatitis A in Taiwan: a 15-year nationwide cohort study. J Viral Hepat 2016;23:940–945. 53. Cho MW, Ehrenfeld E. Rapid completion of the replication cycle of hepatitis A virus subsequent to reversal of guanidine inhibition. Virology 1991;180:770–780. 54. Choi YS, Jung MK, Lee J, et al. Tumor necrosis factor-producing T-regulatory cells are associated with severe liver injury in patients with acute hepatitis A. Gastroenterology 2018;154:1047–1060. 55. Choi YS, Lee J, Lee HW, et al. Liver injury in acute hepatitis A is associated with decreased frequency of regulatory T cells caused by Fasmediated apoptosis. Gut 2015;64:1303–1313. 56. Choi HK, Song YG, Han SH, et al. Clinical features and outcomes of acute kidney injury among patients with acute hepatitis A. J Clin Virol 2011;52:192–197. 57. Clemens R, Safary A, Hepburn A, et al. Clinical experience with an inactivated hepatitis A vaccine. J Infect Dis 1995;171(Suppl 1):S44– S49. 58. Cockayne EA. Catarrhal jaundice, sporadic and epidemic, and its relation to acute yellow atrophy of the liver. Q J Med 1912;6:1–29. 59. Cohen L, Benichou D, Martin A. Analysis of deletion mutants indicates that the 2A polypeptide of hepatitis A virus participates in virion morphogenesis. J Virol 2002;76:7495–7505. 60. Cohen JI, Feinstone S, Purcell RH. Hepatitis A virus infection in a chimpanzee: duration of viremia and detection of virus in saliva and throat swabs. J Infect Dis 1989;160:887–890. 61. Cohen JI, Ticehurst JR, Feinstone SM, et al. Hepatitis A virus cDNA and its RNA transcripts are infectious in cell culture. J Virol 1987;61:3035–3039. 62. Cohn EJ, Oncley JL, Strong LE, et al. Chemical, clinical, and immunological studies on the products of human plasma fractionation. I. The characterization of the protein fractions of human plasma. J Clin Invest 1944;23:417–432. 63. Collier MG, Khudyakov YE, Selvage D, et al. Outbreak of hepatitis A in the USA associated with frozen pomegranate arils imported from Turkey: an epidemiological case study. Lancet Infect Dis 2014;14:976–981. 64. Corey L, Holmes KK. Sexual transmission of hepatitis A in homosexual men: incidence and mechanism. N Engl J Med 1980;302:435–438. 65. Costafreda MI, Abbasi A, Lu H, et al. Exosome mimicry by a HAVCR1-NPC1 pathway of endosomal fusion mediates hepatitis A virus infection. Nat Microbiol 2020;5:1096–1106. 66. Costafreda MI, Kaplan G. HAVCR1 (CD365) and its mouse ortholog are functional hepatitis A virus (HAV) cellular receptors that mediate HAV infection. J Virol 2018;92(9):e02065-17. 67. Costafreda MI, Pérez-Rodriguez FJ, D’Andrea L, et al. Hepatitis A virus adaptation to cellular shutoff is driven by dynamic adjustments of codon usage and results in the selection of populations with altered capsids. J Virol 2014;88:5029–5041. 68. Costafreda MI, Ribes E, Franch A, et al. A single mutation in the glycophorin A binding site of hepatitis A virus enhances virus clearance from the blood and results in a lower fitness variant. J Virol 2012;86:7887–7895. 69. Counihan NA, Anderson DA. Specific IgA enhances the transcytosis and excretion of hepatitis A virus. Sci Rep 2016;6:21855. 70. Crevat D, Crance JM, Chevrinais AM, et al. Monoclonal antibodies against an immunodominant and neutralizing epitope on hepatitis A virus antigen. Arch Virol 1990;113:95–98. 71. Cromeans T, Fields HA, Sobsey MD. Replication kinetics and cytopathic effect of hepatitis A virus. J Gen Virol 1989;70:2051–2062. 72. Cromeans T, Humphrey C, Sobsey M, et al. Use of immunogold preembedding technique to detect hepatitis A viral antigen in infected cells. Am J Anat 1989;185:314–320. 73. Cui F, Hadler SC, Zheng H, et al. Hepatitis A surveillance and vaccine use in China from 1990 through 2007. J Epidemiol 2009;19:189–195. 74. Cui F, Liang X, Wang F, et al. Development, production, and postmarketing surveillance of hepatitis A vaccines in China. J Epidemiol 2014;24:169–177. 75. Cullen JM, Lemon SM. Comparative pathology of hepatitis A virus and hepatitis E virus infection. Cold Spring Harb Perspect Med 2019;9(4):a033456. 76. Cuthbert JA. Hepatitis A: old and new. Clin Microbiol Rev 2001;14:38–58. 77. Daemer RJ, Feinstone SM, Gust ID, et al. Propagation of human hepatitis A virus in African Green Monkey kidney cell culture: primary isolation and serial passage. Infect Immun 1981;32:388–393. 78. Dagan R, Leventhal A, Anis E, et al. Incidence of hepatitis A in Israel following universal immunization of toddlers. JAMA 2005;294:202–210. 79. D’Andrea L, Pérez-Rodríguez FJ, de Castellarnau M, et al. The critical role of codon composition on the translation efficiency robustness of the hepatitis A virus capsid. Genome Biol Evol 2019;11:2439–2456. 80. Daniotti JL, Iglesias-Bartolome R. Metabolic pathways and intracellular trafficking of gangliosides. IUBMB Life 2011;63:513–520. 53

81. Das A, Barrientos RC, Shiota T, et al. Gangliosides are essential endosomal receptors for quasi-enveloped and naked hepatitis A virus. Nat Microbiol 2020;5:1069–1078. 82. Das A, Hirai-Yuki A, Gonzalez-Lopez O, et al. TIM1 (HAVCR1) is not essential for cellular entry of either quasi-enveloped or naked hepatitis A virions. mBio 2017;8(5):e00969-17. doi: 10.1128/mBio.00969-00917. 83. Das A, Maury W, Lemon SM. TIM1 (HAVCR1): an essential “receptor’’ or an “accessory attachment factor’’ for hepatitis A virus? J Virol 2019;93:e01793-01718. 84. de Jong AS, de Mattia F, Van Dommelen MM, et al. Functional analysis of picornavirus 2B proteins: effects on calcium homeostasis and intracellular protein trafficking. J Virol 2008;82:3782–3790. 85. de Oliveira Carneiro I, Sander AL, Silva N, et al. A novel marsupial hepatitis A virus corroborates complex evolutionary patterns shaping the genus hepatovirus. J Virol 2018;92(13):e00082-18. 86. Delem A, Safary A, De Namur F, et al. Characterization of the immune response of volunteers vaccinated with a killed vaccine against hepatitis A. Vaccine 1993;11:479–484. 87. D'Hondt E, Purcell RH, Emerson SU, et al. Efficacy of an inactivated hepatitis A vaccine in pre- and postexposure conditions in marmosets. J Infect Dis 1995;171(Suppl 1):S40–S43. 88. Di Cola G, Fantilli AC, Pisano MB, et al. Foodborne transmission of hepatitis A and hepatitis E viruses: A literature review. Int J Food Microbiol 2021;338:108986. 89. Dienstag JL, Feinstone SM, Purcell RH, et al. Experimental infection of chimpanzees with hepatitis A virus. J Infect Dis 1975;132:532–545. 90. Dienstag JL, Popper H, Purcell RH. The pathology of viral hepatitis types A and B in chimpanzees. Am J Pathol 1976;85:131–144. 91. Dollenmaier G, Weitz M. Interaction of glyceraldehyde-3-phosphate dehydrogenase with secondary and tertiary RNA structural elements of the hepatitis A virus 3′ translated and non-translated regions. J Gen Virol 2003;84:403–414. 92. Dotzauer A, Brenner M, Gebhardt U, et al. IgA-coated particles of hepatitis A virus are translocalized antivectorially from the apical to the basolateral site of polarized epithelial cells via the polymeric immunoglobulin receptor. J Gen Virol 2005;86:2747–2751. 93. Dotzauer A, Feinstone SM, Kaplan G. Susceptibility of nonprimate cell lines to hepatitis A virus infection. J Virol 1994;68:6064–6068. 94. Dotzauer A, Gebhardt U, Bieback K, et al. Hepatitis A virus-specific immunoglobulin A mediates infection of hepatocytes with hepatitis A virus via the asialoglycoprotein receptor. J Virol 2000;74:10950–10957. 95. Dotzauer A, Heitmann A, Laue T, et al. The role of immunoglobulin A in prolonged and relapsing hepatitis A virus infections. J Gen Virol 2012;93:754–760. 96. Drexler JF, Corman VM, Lukashev AN, et al.; Hepatovirus Ecology Consortium. Evolutionary origins of hepatitis A virus in small mammals. Proc Natl Acad Sci U S A 2015;112:15190–15195. 97. Duermeyer W, van der Veen J. Specific detection of IgM-antibodies by ELISA, applied in hepatitis-A. Lancet 1978;ii:684–685. 98. Dupzyk A, Tsai B. Bag2 is a component of a cytosolic extraction machinery that promotes membrane penetration of a nonenveloped virus. J Virol 2018;92:e00607-18. 99. Eckels KH, Summers PL, Dubois DR. Hepatitis A virus hemagglutination and a test for hemagglutination inhibition antibodies. J Clin Microbiol 1989;27:1375–1376. 100. Emerson SU, Huang YK, McRill C, et al. Mutations in both the 2B and 2C genes of hepatitis A virus are involved in adaptation to growth in cell culture. J Virol 1992;66:650–654. 101. Emerson SU, Huang YK, Nguyen H, et al. Identification of VP1/2A and 2C as virulence genes of hepatitis A virus and demonstration of genetic instability of 2C. J Virol 2002;76:8551–8559. 102. Emerson SU, Huang YK, Purcell RH. 2B and 2C mutations are essential but mutations throughout the genome of HAV contribute to adaptation to cell culture. Virology 1993;194:475–480. 103. Emerson SU, Rosenblum B, Feinstone S, et al. Identification of the hepatitis A virus genes involved in adaptation to tissue-culture growth and attenuation. In: R.A. Lerner, H. Ginsberg, R.M. Chanock, (eds). Vaccines 89: Modern Approaches to New Vaccines Including Prevention of AIDS. Cold Spring Harbor, NY: Cold Spring Harbor Laboratory Press; 1989:427–430. 104. Emerson SU, Tsarev SA, Govindarajan S, et al. A simian strain of hepatitis a virus, AGM-27, functions as an attenuated vaccine for chimpanzees. J Infect Dis 1996;173:592–597. 105. Esser-Nobis K, Harak C, Schult P, et al. Novel perspectives for hepatitis A virus therapy revealed by comparative analysis of hepatitis C virus and hepatitis A virus RNA replication. Hepatology 2015;62:397–408. 106. Feigelstock DA, Thompson P, Kaplan GG. Growth of hepatitis A virus in a mouse liver cell line. J Virol 2005;79:2950–2955. 107. Feigelstock D, Thompson P, Mattoo P, et al. Polymorphisms of the hepatitis A virus cellular receptor 1 in African green monkey kidney cells result in antigenic variants that do not react with protective monoclonal antibody 190/4. J Virol 1998;72:6218–6222. 108. Feigelstock D, Thompson P, Mattoo P, et al. The human homolog of HAVcr-1 codes for a hepatitis A virus cellular receptor. J Virol 1998;72:6621–6628. 109. Feinstone SM. History of the discovery of hepatitis A virus. Cold Spring Harb Perspect Med 2018;9(5):a031740. 110. Feinstone SM, Daemer RJ, Gust ID, et al. Live attenuated vaccine for hepatitis A. Dev Biol Stand 1983;54:429–432. 111. Feinstone SM, Kapikian AZ, Purcell RH. Hepatitis A: detection by immune electron microscopy of a viruslike antigen associated with acute illness. Science 1973;182:1026–1028. 112. Feng Z, Hensley L, McKnight KL, et al. A pathogenic picornavirus acquires an envelope by hijacking cellular membranes. Nature 2013;496:367–371. 113. Feng Z, Hirai-Yuki A, McKnight KL, et al. Naked viruses that aren’t always naked: quasi-enveloped agents of acute hepatitis. Annu Rev Virol 2014;1:539–560. 114. Feng Z, Lemon SM. Hepatitis A virus. In: Ehrenfeld E, Domingo E, Roos RP, eds. The Picornaviruses. Washington, DC: ASM Press; 2010:383–396. 115. Feng H, Lenarcic EM, Yamane D, et al. NLRX1 promotes immediate IRF1-directed antiviral responses by limiting dsRNA-activated translational inhibition mediated by PKR. Nat Immunol 2017;18:1299–1309. 116. Feng Z, Li Y, McKnight KL, et al. Human pDCs preferentially sense enveloped hepatitis A virions. J Clin Invest 2015;125:169–176. 117. Feng H, Sander AL, Moreira-Soto A, et al. Hepatovirus 3ABC proteases and evolution of mitochondrial antiviral signaling protein (MAVS). J Hepatol 2019;71:25–34. 118. Feng H, Zhang Y-B, Gui J-F, et al. Interferon regulatory factor 1 (IRF1) and anti-pathogen innate immune responses. PLoS Pathog. 2021:17(1):e1009220. 119. Fensterl V, Grotheer D, Berk I, et al. Hepatitis A virus suppresses RIG-I-mediated IRF-3 activation to block induction of beta interferon. J Virol 2005;79:10968–10977. 120. Flehmig B, Ranke M, Berthold H, et al. A solid-phase radioimmunoassay for detection of IgM antibodies to hepatitis A virus. J Infect Dis 1979;140:169–175. 121. Fleischer B, Fleischer S, Maier K, et al. Clonal analysis of infiltrating T lymphocytes in liver tissue in viral hepatitis A. Immunology 1990;69:14–19. 54

122. Foster MA, Hofmeister MG, Kupronis BA, et al. Increase in hepatitis A virus infections—United States, 2013–2018. MMWR Morb Mortal Wkly Rep 2019;68:413–415. 123. Foster M, Ramachandran S, Myatt K, et al. Hepatitis A virus outbreaks associated with drug use and homelessness—California, Kentucky, Michigan, and Utah, 2017. MMWR Morb Mortal Wkly Rep 2018;67:1208–1210. 124. Freed EO. Viral late domains. J Virol 2002;76:4679–4687. 125. Frosner GG, Deinhardt F, Scheid R, et al. Propagation of human hepatitis A virus in a hepatoma cell line. Infection 1979;7:303–305. 126. Fuchs R, Blaas D. Uncoating of human rhinoviruses. Rev Med Virol 2010;20:281–297. 127. Fujiwara K, Kojima H, Yasui S, et al. Hepatitis A viral load in relation to severity of the infection. J Med Virol 2011;83:201–207. 128. Funkhouser AW, Purcell RH, D’Hondt E, et al. Attenuated hepatitis A virus: genetic determinants of adaptation to growth in MRC-5 cells. J Virol 1994;68:148–157. 129. Funkhouser AW, Raychaudhuri G, Purcell RH, et al. Progress toward the development of a genetically engineered attenuated hepatitis a virus vaccine. J Virol 1996;70:7948–7957. 130. Funkhouser AW, Schultz DE, Lemon SM, et al. Hepatitis A virus translation is rate-limiting for virus replication in MRC-5 cells. Virology 1999;254:268–278. 131. Garriga D, Vives-Adrián L, Buxaderas M, et al. Cloning, purification and preliminary crystallographic studies of the 2AB protein from hepatitis A virus. Acta Crystallogr Sect F Struct Biol Cryst Commun 2011;67:1224–1227. 132. Gauss-Muller V, Deinhardt F. Effect of hepatitis A virus infection on cell metabolism in vitro. Proc Soc Exp Biol Med 1984;175:10–15. 133. Gauss-Muller V, Kusov YY. Replication of a hepatitis A virus replicon detected by genetic recombination in vivo. J Gen Virol 2002;83:2183–2192. 134. Gellis SS, Stokes J Jr, Brother GM, et al. The use of human immune serum globulin (gamma globulin) in infectious (epidemic) hepatitis in the Mediterranean theater of operations. I. Studies on prophylaxis in two epidemics of infectious hepatitis. JAMA 1945;128:1062–1063. 135. Glass MJ, Jia XY, Summers DF. Identification of the hepatitis A virus internal ribosome entry site: in vivo and in vitro analysis of bicistronic RNAs containing the HAV 5′ noncoding region. Virology 1993;193:842–852. 136. Glikson M, Galun E, Oren R, et al. Relapsing hepatitis A. Review of 14 cases and literature survey. Medicine (Baltimore) 1992;71:14–23. 137. Godet AC, David F, Hantelys F, et al. IRES trans-acting factors, key actors of the stress response. Int J Mol Sci 2019;20(4):924. 138. Gonzalez-Lopez O, Rivera-Serrano EE, Hu F, et al. Redundant late domain functions of tandem VP2 YPX3L motifs in nonlytic cellular egress of quasi-enveloped hepatitis A virus. J Virol 2018;92:1308–1318. 139. Goodfellow I, Chaudhry Y, Richardson A, et al. Identification of a cis-acting replication element within the poliovirus coding region. J Virol 2000;74:4590–4600. 140. Gorbalenya AE, Donchenko AP, Blinov VM, et al. Cysteine proteases of positive strand RNA viruses and chymotrypsin-like serine proteases. A distinct protein superfamily with a common structural fold. FEBS Lett 1989;243:103–114. 141. Gorbalenya AE, Koonin EV. Helicases: amino acid sequence comparisons and structure-function relationships. Curr Opin Struct Biol 1993;3:419–429. 142. Gosert R, Cassinotti P, Siegl G, et al. Identification of hepatitis A virus non-structural protein 2B and its release by the major virus protease 3C. J Gen Virol 1996;77:247–255. 143. Gosert R, Chang KH, Rijnbrand R, et al. Transient expression of cellular polypyrimidine-tract binding protein stimulates capindependent translation directed by both picornaviral and flaviviral internal ribosome entry sites in vivo. Mol Cell Biol 2000;20:1583–1595. 144. Gosert R, Egger D, Bienz K. A cytopathic and a cell culture adapted hepatitis A virus strain differ in cell killing but not in intracellular membrane rearrangements. Virology 2000;266:157–169. 145. Graff J, Cha J, Blyn LB, et al. Interaction of poly(rC) binding protein 2 with the 5′ noncoding region of hepatitis A virus RNA and its effects on translation. J Virol 1998;72:9668–9675. 146. Graff J, Emerson SU. Importance of amino acid 216 in nonstructural protein 2B for replication of hepatitis A virus in cell culture and in vivo. J Med Virol 2003;71:7–17. 147. Graff J, Richards OC, Swiderek KM, et al. Hepatitis A virus capsid protein VP1 has a heterogeneous C terminus. J Virol 1999;73:6015–6023. 148. Greninger AL, Knudsen GM, Betegon M, et al. ACBD3 interaction with TBC1 domain 22 protein is differentially affected by enteroviral and kobuviral 3A protein binding. mBio 2013;4:e00098-00013. 149. Hadler SC, Webster HM, Erben JJ, et al. Hepatitis A in day-care centers: a community-wide assessment. N Engl J Med 1980;302:1222–1227. 150. Han X, Zhou C, Jiang M, et al. Discovery of RG7834: the first-in-class selective and orally available small molecule hepatitis B virus expression inhibitor with novel mechanism of action. J Med Chem 2018;61:10619–10634. 151. Hardin CC, Sneeden JL, Lemon SM, et al. Folding of pyrimidine-enriched RNA fragments from the vicinity of the internal ribosomal entry site of hepatitis A virus. Nucleic Acids Res 1999;27:665–673. 152. Harmon SA, Emerson SU, Huang YK, et al. Hepatitis A viruses with deletions in the 2A gene are infectious in cultured cells and marmosets. J Virol 1995;69:5576–5581. 153. Harmon SA, Updike W, Jia XY, et al. Polyprotein processing in cis and in trans by hepatitis A virus 3C protease cloned and expressed in Escherichia coli. J Virol 1992;66:5242–5247. 154. Harrison SC. Viral membrane fusion. Nat Struct Mol Biol 2008;15:690–698. 155. Havens WP Jr. Period of infectivity of patients with experimentally induced infectious hepatitis. J Exp Med 1946;83:251–258. 156. Havens WP Jr, Ward R, Drill VA, et al. Experimental production of hepatitis by feeding icterogenic materials. Proc Soc Exp Biol Med 1944;57:206–208. 157. Herold J, Andino R. Poliovirus RNA replication requires genome circularization through a protein-protein bridge. Mol Cell 2001;7:581–591. 158. Hessvik NP, Llorente A. Current knowledge on exosome biogenesis and release. Cell Mol Life Sci 2018;75:193–208. 159. Hiltunen JK, Kastaniotis AJ, Autio KJ, et al. 17B-hydroxysteroid dehydrogenases as acyl thioester metabolizing enzymes. Mol Cell Endocrinol 2019;489:107–118. 160. Hindiyeh M, Li QH, Basavappa R, et al. Poliovirus mutants at histidine 195 of VP2 do not cleave VP0 into VP2 and VP4. J Virol 1999;73:9072–9079. 161. Hirai-Yuki A, Hensley L, McGivern DR, et al. MAVS-dependent host species range and pathogenicity of human hepatitis A virus. Science 2016;353:1541–1545. 162. Hirai-Yuki A, Hensley L, Whitmire JK, et al. Biliary secretion of quasi-enveloped human hepatitis A virus. mBio 2016;7:e01998-01916. 163. Hirai-Yuki A, Whitmire JK, Joyce M, et al. Murine models of hepatitis A virus infection. Cold Spring Harb Perspect Med 2019;9(1):a031674. 164. Hofmeister MG, Foster MA, Teshale EH. Epidemiology and transmission of hepatitis A virus and hepatitis E virus infections in the United States. Cold Spring Harb Perspect Med 2018;9(4):a033431. 55

165. Holmes AW, Wolfe L, Rosenblate H, et al. Hepatitis in marmosets: induction of disease with coded specimens from a human volunteer study. Science 1969;165:816–817. 166. Holzer BR, Hatz C, Schmidt-Sissolak D, et al. Immunogenicity and adverse effects of inactivated virosome versus alum-adsorbed hepatitis A vaccine: a randomized controlled trial. Vaccine 1996;14:982–986. 167. Hong S, Lee HW, Chang DY, et al. Antibody-secreting cells with a phenotype of Ki-67low, CD138high, CD31high, and CD38high secrete nonspecific IgM during primary hepatitis A virus infection. J Immunol 2013;191:127–134. 168. Hornei B, Kammerer R, Moubayed P, et al. Experimental hepatitis A virus infection in guinea pigs. J Med Virol 2001;64:402–409. 169. Hsu NY, Ilnytska O, Belov G, et al. Viral reorganization of the secretory pathway generates distinct organelles for RNA replication. Cell 2010;141:799–811. 170. Innis BL, Snitbhan R, Kunasol P, et al. Protection against hepatitis A by an inactivated vaccine. JAMA 1994;271:1328–1334. 171. Jackson WT, Giddings TH Jr, Taylor MP, et al. Subversion of cellular autophagosomal machinery by RNA viruses. PLoS Biol 2005;3:e156. 172. Jacobsen KH. Globalization and the changing epidemiology of hepatitis A virus. Cold Spring Harb Perspect Med 2018;8:a031716. 173. Jacobsen KH, Koopman JS. Declining hepatitis A seroprevalence: a global review and analysis. Epidemiol Infect 2004;132:1005–1022. 174. Jacobsen KH, Wiersma ST. Hepatitis A virus seroprevalence by age and world region, 1990 and 2005. Vaccine 2010;28:6653–6657. 175. Jansen RW, Newbold JE, Lemon SM. Complete nucleotide sequence of a cell culture-adapted variant of hepatitis A virus: comparison with wild-type virus with restricted capacity for in vitro replication. Virology 1988;163:299–307. 176. Jansen RW, Siegl G, Lemon SM. Molecular epidemiology of human hepatitis A virus defined by an antigen-capture polymerase chain reaction method. Proc Natl Acad Sci U S A 1990;87:2867–2871. 177. Jecht M, Probst C, Gauss-Muller V. Membrane permeability induced by hepatitis A virus proteins 2B and 2BC and proteolytic processing of HAV 2BC. Virology 1998;252:218–227. 178. Jewell DA, Swietnicki W, Dunn BM, et al. Hepatitis A virus 3C proteinase substrate specificity. Biochemistry 1992;31:7862–7869. 179. Jia XY, Summers DF, Ehrenfeld E. Host antibody response to viral structural and nonstructural proteins after hepatitis A virus infection. J Infect Dis 1992;165:273–280. 180. Jia XY, Tesar M, Summers DF, et al. Replication of hepatitis A viruses with chimeric 5′ nontranslated regions. J Virol 1996;70:2861–2868. 181. Jiang P, Liu Y, Ma HC, et al. Picornavirus morphogenesis. Microbiol Mol Biol Rev 2014;78:418–437. 182. Jiang W, Ma P, Deng L, et al. Hepatitis A virus structural protein pX interacts with ALIX and promotes the secretion of virions and foreign proteins through exosome-like vesicles. J Extracell Vesicles 2020;9:1716513. 183. Jiang W, Muhammad F, Ma P, et al. Sofosbuvir inhibits hepatitis A virus replication in vitro assessed by a cell-based fluorescent reporter system. Antiviral Res 2018;154:51–57. 184. Jouanguy E. Human genetic basis of fulminant viral hepatitis. Hum Genet 2020;139:877–884. 185. Jung YM, Park SJ, Kim JS, et al. Atypical manifestations of hepatitis A infection: a prospective, multicenter study in Korea. J Med Virol 2010;82:1318–1326. 186. Kankam M, Griffin R, Price J, et al. Polyvalent human immune globulin: a prospective, open-label study assessing anti-hepatitis A virus (HAV) antibody levels, pharmacokinetics, and safety in HAV-seronegative healthy subjects. Adv Ther 2020;37:2373–2389. 187. Kaplan G, Totsuka A, Thompson P, et al. Identification of a surface glycoprotein on African green monkey kidney cells as a receptor for hepatitis A virus. EMBO J 1996;15:4282–4296. 188. Karron RA, Daemer R, Ticehurst J, et al. Studies of prototype live hepatitis A virus vaccines in primate models. J Infect Dis 1988;157:338–345. 189. Keenan CM, Lemon SM, LeDuc JW, et al. Pathology of hepatitis A infection in the owl monkey (Aotus trivirgatus). Am J Pathol 1984;115:1–8. 190. Kihara A. Very long-chain fatty acids: elongation, physiology and related disorders. J Biochem 2012;152:387–395. 191. Kim J, Chang DY, Lee HW, et al. Innate-like cytotoxic function of bystander-activated CD8(+) T Cells is associated with liver injury in acute hepatitis A. Immunity 2018;48:161–173.e165. 192. Kim JD, Cho EJ, Ahn C, et al. A model to predict 1-month risk of transplant or death in hepatitis A-related acute liver failure. Hepatology 2019;70:621–629. 193. Kim HY, Eyheramonho MB, Pichavant M, et al. A polymorphism in TIM1 is associated with susceptibility to severe hepatitis A virus infection in humans. J Clin Invest 2011;121:1111–1118. 194. Kim HS, Jeong SH, Jang JH, et al. Coinfection of hepatitis A virus genotype IA and IIIA complicated with autoimmune hemolytic anemia, prolonged cholestasis, and false-positive immunoglobulin M anti-hepatitis E virus: a case report. Korean J Hepatol 2011;17:323–327. 195. Kim D, Lee YS, Jung SJ, et al. Viral hijacking of the TENT4-ZCCHC14 complex protects viral RNAs via mixed tailing. Nat Struct Mol Biol 2020;27:581–588. 196. Kim MJ, Shin JY, Oh JA, et al. Identification of transfusion-transmitted hepatitis A through postdonation information in Korea: results of an HAV lookback (2007–2012). Vox Sang 2018. https://doi.org/10.1111/vox.12672 197. Klevens RM, Denniston MM, Jiles-Chapman RB, et al. Decreasing immunity to hepatitis A virus infection among US adults: Findings from the National Health and Nutrition Examination Survey (NHANES), 1999–2012. Vaccine 2015;33:6192–6198. 198. Kolter T, Sandhoff K. Lysosomal degradation of membrane lipids. FEBS Lett 2010;584:1700–1712. 199. Konduru K, Kaplan GG. Determinants in 3Dpol modulate the rate of growth of hepatitis A virus. J Virol 2010;84:8342–8347. 200. Krawczynski KK, Bradley DW, Murphy BL, et al. Pathogenetic aspects of hepatitis A virus infection in enterally inoculated marmosets. Am J Clin Pathol 1981;76:698–706. 201. Krugman S, Giles JP, Hammond J. Infectious hepatitis: evidence for two distinctive clinical, epidemiological, and immunological types of infection. JAMA 1967;200:365–373. 202. Krugman S, Ward R, Giles JP, et al. Infectious hepatitis: detection of virus during the incubation period and in clinically inapparent infection. N Engl J Med 1959;261:729–734. 203. Kulkarni MA, Walimbe AM, Cherian S, et al. Full length genomes of genotype IIIA Hepatitis A Virus strains (1995–2008) from India and estimates of the evolutionary rates and ages. Infect Genet Evol 2009;9:1287–1294. 204. Kulsuptrakul J, Wang R, Meyers NL, et al. A genome-wide CRISPR screen identifies UFMylation and TRAMP-like complexes as host factors required for hepatitis A virus infection. Cell Rep 2021;34:108859. 205. Kusov YY, Gauss-Muller V. In vitro RNA binding of the hepatitis A virus proteinase 3C (HAV 3Cpro) to secondary structure elements within the 5′ terminus of the HAV genome. RNA 1997;3:291–302. 206. Kusov Y, Gauss-Muller V. Improving proteolytic cleavage at the 3A/3B site of the hepatitis A virus polyprotein impairs processing and particle formation, and the impairment can be complemented in trans by 3AB and 3ABC. J Virol 1999;73:9867–9878. 207. Kusov YY, Gosert R, Gauss-Muller V. Replication and in vivo repair of the hepatitis A virus genome lacking the poly(A) tail. J Gen Virol 2005;86:1363–1368. 208. Kusov YY, Morace G, Probst C, et al. Interaction of hepatitis A virus (HAV) precursor proteins 3AB and 3ABC with the 5′ and 3′ termini of the HAV RNA. Virus Res 1997;51:151–157. 56

209. Kusov Y, Weitz M, Dollenmeier G, et al. RNA-protein interactions at the 3′ end of the hepatitis A virus RNA. J Virol 1996;70:1890–1897. 210. Lanford RE, Feng Z, Chavez D, et al. Acute hepatitis A virus infection is associated with a limited type I interferon response and persistence of intrahepatic viral RNA. Proc Natl Acad Sci U S A 2011;108:11223–11228. 211. Lau CL, Streeton CL, David MC, et al. The tolerability of a combined hepatitis A and typhoid vaccine in children aged 2–16 years: an observational study. J Travel Med 2016;23:tav023. 212. Le SY, Chen JH, Sonenberg N, et al Conserved tertiary structural elements in the 5′ nontranslated region of cardiovirus, aphthovirus and hepatitis A virus RNAs. Nucleic Acids Res 1993;21:2445–2451. 213. Lednar WM, Lemon SM, Kirkpatrick JW, et al. Frequency of illness associated with epidemic hepatitis A virus infections in adults. Am J Epidemiol 1985;122:226–233. 214. LeDuc JW, Escajadillo A, Lemon SM. Transmission of hepatitis A virus among captive Panamanian owl monkeys. Lancet 1981;2(8260-61):1427–1428. 215. Lee S, Joshi A, Nagashima K, et al. Structural basis for viral late-domain binding to Alix. Nat Struct Mol Biol 2007;14:194–199. 216. Lemon SM. IgM neutralizing antibody to hepatitis A virus. J Infect Dis 1985;152:1353–1354. 217. Lemon SM. Type A viral hepatitis: new developments in an old disease. N Engl J Med 1985;313:1059–1067. 218. Lemon SM, Binn LN. Antigenic relatedness of two strains of hepatitis A virus determined by cross-neutralization. Infect Immun 1983;42:418–420. 219. Lemon SM, Binn LN. Serum neutralizing antibody response to hepatitis A virus. J Infect Dis 1983;148:1033–1039. 220. Lemon SM, Binn LN. Incomplete neutralization of hepatitis A virus in vitro due to lipid-associated virions. J Gen Virol 1985;66:2501–2505. 221. Lemon SM, Binn LN, Marchwicki R, et al. In vivo replication and reversion to wild-type of a neutralization-resistant variant of hepatitis A virus. J Infect Dis 1990;161:7–13. 222. Lemon SM, Brown CD, Brooks DS, et al. Specific immunoglobulin M response to hepatitis A virus determined by solid-phase radioimmunoassay. Infect Immun 1980;28:927–936. 223. Lemon SM, Jansen RW. A simple method for clonal selection of hepatitis A virus based on recovery of virus from radioimmunofocus overlays. J Virol Methods 1985;11:171–176. 224. Lemon SM, Jansen RW, Newbold JE. Infectious hepatitis A virus particles produced in cell culture consist of three distinct types with different buoyant densities in CsCl. J Virol 1985;54:78–85. 225. Lemon SM, Murphy PC, Provost PJ, et al. Immunoprecipitation and virus neutralization assays demonstrate qualitative differences between protective antibody responses to inactivated hepatitis A vaccine and passive immunization with immune globulin. J Infect Dis 1997;176:9–19. 226. Lemon SM, Murphy PC, Shields PA, et al. Antigenic and genetic variation in cytopathic hepatitis A virus variants arising during persistent infection: evidence for genetic recombination. J Virol 1991;65:2056–2065. 227. Lemon SM, Murphy PC, Smith A, et al. Removal/neutralization of hepatitis A virus during manufacture of high purity, solvent/detergent factor VIII concentrate. J Med Virol 1994;43:44–49. 228. Lemon SM, Ott JJ, Van Damme P, et al. Type A viral hepatitis: a summary and update on the molecular virology, epidemiology, pathogenesis and prevention. J Hepatol 2018;68:167–184. 229. Lewis JA, Armstrong ME, Larson VM, et al. Use of a live attenuated hepatitis A vaccine to prepare a highly purified, formalin-inactivated hepatitis A vaccine. In: Hollinger FB, Lemon SM, Margolis HS, (eds). Viral Hepatitis and Liver Disease. Baltimore, MD: Williams & Wilkins; 1991:94–97. 230. Li K, Foy E, Ferreon JC, et al. Immune evasion by hepatitis C virus NS3/4A protease-mediated cleavage of the Toll-like receptor 3 adaptor protein TRIF. Proc Natl Acad Sci U S A 2005;102:2992–2997. 231. Li XD, Sun L, Seth RB, et al. Hepatitis C virus protease NS3/4A cleaves mitochondrial antiviral signaling protein off the mitochondria to evade innate immunity. Proc Natl Acad Sci U S A 2005;102:17717–17722. 232. Lim J, Kim D, Lee YS, et al. Mixed tailing by TENT4A and TENT4B shields mRNA from rapid deadenylation. Science 2018;361:701–704. 233. Lin J, Lee LY, Roivainen M, et al. Structure of the Fab-labeled “breathing” state of native poliovirus. J Virol 2012;86:5959–5962. 234. Liu Y, Wang C, Mueller S, et al. Direct interaction between two viral proteins, the nonstructural protein 2C and the capsid protein VP3, is required for enterovirus morphogenesis. PLoS Pathog 2010;6:e1001066. 235. Locarnini SA, Coulepis AG, Kaldor J, et al. Coproantibodies in hepatitis A: detection of enzyme-linked immunosorbent assay and immune electron microscopy. J Clin Microbiol 1980;11:710–716. 236. Locarnini SA, Ferris AA, Stott AC, et al. Letter: pitfalls in hepatitis A? Lancet 1974;2:1007. 237. Losick VP, Schlax PE, Emmons RA, et al. Signals in hepatitis A virus P3 region proteins recognized by the ubiquitin-mediated proteolytic system. Virology 2003;309:306–319. 238. Lyoo H, van der Schaar HM, Dorobantu CM, et al. ACBD3 is an essential pan-enterovirus host factor that mediates the interaction between viral 3A protein and cellular protein PI4KB. mBio 2019;10:e02742-18. 239. MacCallum FO. Hepatitis. Br Med Bull 1953;9:221–225. 240. Maier K, Gabriel P, Koscielniak E, et al. Human gamma interferon production by cytotoxic T lymphocytes sensitized during hepatitis A virus infection. J Virol 1988;62:3756–3763. 241. Malcolm BA, Chin SM, Jewell DA, et al. Expression and characterization of recombinant hepatitis A virus 3C proteinase. Biochemistry 1992;31:3358–3363. 242. Manangeeswaran M, Jacques J, Tami C, et al. Binding of hepatitis A virus to its cellular receptor 1 inhibits T-regulatory cell functions in humans. Gastroenterology 2012;142:1516–1525.e1513. 243. Mannucci PM, Gdovin S, Gringeri A, et al. Transmission of hepatitis A to patients with hemophilia by factor VIII concentrates treated with organic solvent and detergent to inactivate viruses. Ann Intern Med 1994;120:1–7. 244. Mao JS, Dong DX, Zhang HY, et al. Primary study of attenuated live hepatitis A vaccine (H2 strain) in humans. J Infect Dis 1989;159:621–624. 245. Mao JS, Dong DX, Zhang SY, et al. Further studies of attenuated live hepatitis A vaccine (H2 strain) in humans. In: Hollinger FB, Lemon SM, Margolis HS, eds. Viral Hepatitis and Liver Disease. Baltimore, MD: Williams & Wilkins; 1991:110–111. 246. Martin A, Benichou D, Chao SF, et al. Maturation of the hepatitis A virus capsid protein VP1 is not dependent on processing by the 3Cpro proteinase. J Virol 1999;73:6220–6227. 247. Martin A, Escriou N, Chao SF, et al. Identification and site-direct mutagenesis of the primary (2A/2B) cleavage site of the hepatitis A virus polyprotein: functional impact on the infectivity of HAV transcripts. Virology 1995;213:213–222. 248. Martin A, Lemon SM. The molecular biology of hepatitis A virus. In: Ou J, ed. Hepatitis Viruses. Norwell, MA: Kluwer Academic Publishers; 2002:23–50. 249. Martin JC, Petrecz ML, Stek JE, et al. Using the power law model to predict the long-term persistence and duration of detectable hepatitis A antibody after receipt of hepatitis A Vaccine (VAQTA™). Vaccine 2021;39:2764–2771. 250. Mathiesen LR, Drucker J, Lorenz D, et al. Localization of hepatitis A antigen in marmoset organs during acute infection with hepatitis A 57

virus. J Infect Dis 1978;138:369–377. 251. Mathiesen LR, Moller AM, Purcell RH, et al. Hepatitis A virus in the liver and intestine of marmosets after oral inoculation. Infect Immun 1980;28:45–48. 252. Maynard JE, Bradley DW, Gravelle CR, et al. Preliminary studies of hepatitis A in chimpanzees. J Infect Dis 1975;131:194–197. 253. Maynard JE, Krushak DH, Bradley DW, et al. Infectivity studies of hepatitis A and B in non-human primates. Dev Biol Stand 1975;30:229–235. 254. Mbithi JN, Springthorpe VS, Boulet JR, et al. Survival of hepatitis A virus on human hands and its transfer on contact with animate and inanimate surfaces. J Clin Microbiol 1992;30:757–763. 255. Mbithi JN, Springthorpe VS, Sattar SA. Comparative in vivo efficiencies of hand-washing agents against hepatitis A virus (HM-175) and poliovirus type 1 (Sabin). Appl Environ Microbiol 1993;59:3463–3469. 256. McKnight KL, Xie L, González-López O, et al. Protein composition of the hepatitis A virus quasi-envelope. Proc Natl Acad Sci U S A 2017;114:6587–6592. 257. Melia CE, Peddie CJ, de Jong AWM, et al. Origins of enterovirus replication organelles established by whole-cell electron microscopy. mBio 2019;10(3):e00951-19. 258. Metzger WG, Ehni HJ, Kremsner PG, et al. Experimental infections in humans—historical and ethical reflections. Trop Med Int Health 2019;24:1384–1390. 259. Midthun K, Ellerbeck E, Gershman K, et al. Safety and immunogenicity of a live attenuated hepatitis A virus vaccine in seronegative volunteers. J Infect Dis 1991;163:735–739. 260. Mishra N, Fagbo SF, Alagaili AN, et al. A viral metagenomic survey identifies known and novel mammalian viruses in bats from Saudi Arabia. PLoS One 2019;14:e0214227. 261. Mishra A, Saigal S, Gupta R, et al. Acute pancreatitis associated with viral hepatitis: a report of six cases with review of literature. Am J Gastroenterol 1999;94:2292–2295. 262. Misumi I, Li Z, Sun L, et al. Iminosugar glucosidase inhibitors reduce hepatic inflammation in HAV-infected Ifnar1(−/−) mice. J Virol 2021. doi: 10.1128/JVI.00058-21. 263. Misumi I, Mitchell JE, Lund MM, et al. T cells protect against hepatitis A virus infection and limit infection-induced liver injury. J Hepatol 2021;75:1323–1334. 264. Mohamed B, Mazeaud C, Baril M, et al. Very-long-chain fatty acid metabolic capacity of 17-beta-hydroxysteroid dehydrogenase type 12 (HSD17B12) promotes replication of hepatitis C virus and related flaviviruses. Sci Rep 2020;10:4040. 265. Moller-Tank S, Maury W. Phosphatidylserine receptors: enhancers of enveloped virus entry and infection. Virology 2014;468–470:565–580. 266. Morace G, Kusov Y, Dzagurov G, et al. The unique role of domain 2A of the hepatitis A virus precursor polypeptide P1-2A in viral morphogenesis. BMB Rep 2008;41:678–683. 267. Morris TS, Frormann S, Shechosky S, et al. In vitro and ex vivo inhibition of hepatitis A virus 3C proteinase by a peptidyl monofluoromethyl ketone. Bioorg Med Chem 1997;5:797–807. 268. Murphy BL, Maynard JE, Bradley DW, et al. Immunofluorescence of hepatitis A virus antigen in chimpanzees. Infect Immun 1978;21:663–665. 269. Nainan OV, Margolis HS, Robertson BH, et al. Sequence analysis of a new hepatitis A virus naturally infecting cynomolgus macaques (Macaca fascicularis). J Gen Virol 1991;72:1685–1689. 270. Najarian R, Caput D, Gee W, et al. Primary structure and gene organization of human hepatitis A virus. Proc Natl Acad Sci U S A 1985;82:2627–2631. 271. Navarro MED, Yao CC, Whiteley A, et al. Liver transplant evaluation for fulminant liver failure due to acute hepatitis A infection: case series and literature review. Transpl Infect Dis 2021;23:e13476. 272. Ndumbi P, Freidl GS, Williams CJ, et al. Hepatitis A outbreak disproportionately affecting men who have sex with men (MSM) in the European Union and European Economic Area, June 2016 to May 2017. Euro Surveill 2018;23(33):1700641. doi: 10.2807/1560-7917. 273. Neefe JR, Gellis SS, Stokes J Jr. Homologous serum hepatitis and infectious (epidemic) hepatitis: studies in volunteers bearing on immunological and other characteristics of the etiological agents. Am J Med 1946;1:3–22. 274. Nelson NP, Link-Gelles R, Hofmeister MG, et al. Update: recommendations of the Advisory Committee on Immunization Practices for use of hepatitis A vaccine for postexposure prophylaxis and for preexposure prophylaxis for international travel. MMWR Morb Mortal Wkly Rep 2018;67:1216–1220. 275. Nelson NP, Weng MK, Hofmeister MG, et al. Prevention of hepatitis A virus infection in the United States: recommendations of the Advisory Committee on Immunization Practices, 2020. MMWR Recomm Rep 2020;69:1–38. 276. Niepmann M. Internal translation initiation of picornaviruses and hepatitis C virus. Biochim Biophys Acta 2009;1789:529–541. 277. Nikonov OS, Chernykh ES, Garber MB, et al. Enteroviruses: classification, diseases they cause, and approaches to development of antiviral drugs. Biochemistry (Mosc) 2017;82:1615–1631. 278. Ouzilou L, Caliot E, Pelletier I, et al. Poliovirus transcytosis through M-like cells. J Gen Virol 2002;83:2177–2182. 279. Parry JV, Mortimer PP. The heat sensitivity of hepatitis A virus determined by a simple tissue culture method. J Med Virol 1984;14:277–283. 280. Paul AV, Van Boom JH, Filippov D, et al. Protein-primed RNA synthesis by purified poliovirus RNA polymerase. Nature 1998;393:280–284. 281. Paul AV, Yin J, Mugavero J, et al. A “slide-back” mechanism for the initiation of protein-primed RNA synthesis by the RNA polymerase of poliovirus. J Biol Chem 2003;278:43951–43960. 282. Paulmann D, Magulski T, Schwarz R, et al. Hepatitis A virus protein 2B suppresses beta interferon (IFN) gene transcription by interfering with IFN regulatory factor 3 activation. J Gen Virol 2008;89:1593–1604. 283. Peters H, Kusov YY, Meyer S, et al. Hepatitis A virus proteinase 3C binding to viral RNA: correlation with substrate binding and enzyme dimerization. Biochem J 2005;385:363–370. 284. Peterson DA, Hurley TR, Hoff JC, et al. Effect of chlorine treatment on infectivity of hepatitis A virus. Appl Environ Microbiol 1983;45:223–227. 285. Petithory JR, Masiarz FR, Kirsch JF, et al. A rapid method for determination of endoproteinase substrate specificity: specificity of the 3C proteinase from hepatitis A virus. Proc Natl Acad Sci U S A 1991;88:11510–11514. 286. Pierce PF, Cappello M, Bernard KW. Subclinical infection with hepatitis A in Peace Corps volunteers following immune globulin prophylaxis. Am J Trop Med Hyg 1990;42:465–469. 287. Ping LH, Jansen RW, Stapleton JT, et al. Identification of an immunodominant antigenic site involving the capsid protein VP3 of hepatitis A virus. Proc Natl Acad Sci U S A 1988;85:8281–8285. 288. Ping LH, Lemon SM. Antigenic structure of human hepatitis A virus defined by analysis of escape mutants selected against murine monoclonal antibodies. J Virol 1992;66:2208–2216. 289. Pintó RM, Costafreda MI, Bosch A. Risk assessment in shellfish-borne outbreaks of hepatitis A. Appl Environ Microbiol 58

2009;75:7350–7355. 290. Pinto MA, Marchevsky RS, Baptista ML, et al. Experimental hepatitis A virus (HAV) infection in Callithrix jacchus: early detection of HAV antigen and viral fate. Exp Toxicol Pathol 2002;53:413–420. 291. Pinto RM, Perez-Rodriguez FJ, D’Andrea L, et al. Hepatitis A virus codon usage: implications for translation kinetics and capsid folding. Cold Spring Harb Perspect Med 2018;8(10):a031781. 292. Popper H, Dienstag JL, Feinstone SM, et al. The pathology of viral hepatitis in chimpanzees. Virchows Arch A Pathol Anat Histol 1980;387:91–106. 293. Probst C, Jecht M, Gauss-Muller V. Intrinsic signals for the assembly of hepatitis A virus particles. Role of structural proteins VP4 and 2A. J Biol Chem 1999;274:4527–4531. 294. Provost PJ, Banker FS, Wadsworth CW, et al. Further evaluation of a live hepatitis A vaccine in marmosets. J Med Virol 1991;34:227–231. 295. Provost PJ, Bishop RP, Gerety RJ, et al. New findings in live, attenuated hepatitis A vaccine development. J Med Virol 1986;20:165–175. 296. Provost PJ, Conti PA, Giesa PA, et al. Studies in chimpanzees of live, attenuated hepatitis A vaccine candidates. Proc Soc Exp Biol Med 1983;172:357–363. 297. Provost PJ, Giesa PA, McAleer WJ, et al. Isolation of hepatitis A virus in vitro in cell culture directly from human specimens. Proc Soc Exp Biol Med 1981;167:201–206. 298. Provost PJ, Hilleman MR. An inactivated hepatitis A virus vaccine prepared from infected marmoset liver. Proc Soc Exp Biol Med 1978;159:201–203. 299. Provost PJ, Hilleman MR. Propagation of human hepatitis A virus in cell culture in vitro. Proc Soc Exp Biol Med 1979;160:213–221. 300. Provost PJ, Villarejos VM, Hilleman MR. Tests in rufiventer and other marmosets of susceptibility to human hepatitis A virus. Primates Med 1978;10:288–294. 301. Purcell RH, D’Hondt E, Bradbury R, et al. Inactivated hepatitis A vaccine: active and passive immunoprophylaxis in chimpanzees. Vaccine 1992;10(Suppl 1):S148–S151. 302. Purcell RH, Emerson SU. Animal models of hepatitis A and E. ILAR J 2001;42:161–177. 303. Purcell RH, Feinstone SM, Ticehurst JR, et al. In: Vyas GN, Dienstag JL, Hoofnagle JH, eds. Viral Hepatitis and Liver Disease. Orlando, FL: Grune & Stratton; 1984. 304. Purcell RH, Wong DC, Shapiro M. Relative infectivity of hepatitis A virus by the oral and intravenous routes in 2 species of nonhuman primates. J Infect Dis 2002;185:1668–1671. 305. Qian M, Cai D, Verhey KJ, et al. A lipid receptor sorts polyomavirus from the endolysosome to the endoplasmic reticulum to cause infection. PLoS Pathog 2009;5:e1000465. 306. Qu L, Feng Z, Yamane D, et al. Disruption of TLR3 signaling due to cleavage of TRIF by the hepatitis A virus protease-polymerase processing intermediate, 3CD. PLoS Pathog 2011;7:e1002169. 307. Rachow A, Gauss-Muller V, Probst C. Homogeneous hepatitis A virus particles. Proteolytic release of the assembly signal 2A from procapsids by factor Xa. J Biol Chem 2003;278:29744–29751. 308. Ravindran MS, Bagchi P, Cunningham CN, et al. Opportunistic intruders: how viruses orchestrate ER functions to infect cells. Nat Rev Microbiol 2016;14:407–420. 309. Report ICTV. Picornaviridae. https://talk.ictvonline.org/ictv-reports/ictv_online_report/positive-sense-rna-viruses/w/picornaviridae. Accessed May 30, 2021. 310. Rezende G, Roque-Afonso AM, Samuel D, et al. Viral and clinical factors associated with the fulminant course of hepatitis A infection. Hepatology 2003;38:613–618. 311. Rha MS, Han JW, Kim JH, et al. Human liver CD8(+) MAIT cells exert TCR/MR1-independent innate-like cytotoxicity in response to IL-15. J Hepatol 2020;73:640–650. 312. Rivera-Serrano EE, Gonzalez-Lopez O, Das A, et al. Cellular entry and uncoating of naked and quasi-enveloped human hepatoviruses. Elife 2019;8:e43983. 313. Robertson BH, Jansen RW, Khanna B, et al. Genetic relatedness of hepatitis A virus strains recovered from different geographic regions. J Gen Virol 1992;73:1365–1377. 314. Robertson BH, Jia XY, Tian H, et al. Antibody response to nonstructural proteins of hepatitis A virus following infection. J Med Virol 1993;40:76–82. 315. Rog T, Orlowski A, Llorente A, et al. Interdigitation of long-chain sphingomyelin induces coupling of membrane leaflets in a cholesterol dependent manner. Biochim Biophys Acta 2016;1858:281–288. 316. Romero-Brey I, Bartenschlager R. Endoplasmic reticulum: the favorite intracellular niche for viral replication and assembly. Viruses 2016;8(6):160. 317. Rosenblum LS, Villarino ME, Nainan OV, et al. Hepatitis A outbreak in a neonatal intensive care unit: risk factors for transmission and evidence of prolonged viral excretion among preterm infants. J Infect Dis 1991;164:476–482. 318. Roulin PS, Lotzerich M, Torta F, et al. Rhinovirus uses a phosphatidylinositol 4-phosphate/cholesterol counter-current for the formation of replication compartments at the ER-Golgi interface. Cell Host Microbe 2014;16:677–690. 319. Rueckert RR, Wimmer E. Systematic nomenclature of picornavirus proteins. J Virol 1984;50:957–959. 320. Sachdeva H, Benusic M, Ota S, et al. Community outbreak of hepatitis A disproportionately affecting men who have sex with men in Toronto, Canada, January 2017–November 2018. Can Commun Dis Rep 2019;45:262–268. 321. Sanchez G, Aragones L, Costafreda MI, et al. Capsid region involved in hepatitis A virus binding to glycophorin A of the erythrocyte membrane. J Virol 2004;78:9807–9813. 322. Schaffner F, Dienstag JL, Purcell RH, et al. Chimpanzee livers after infection with human hepatitis viruses A and B: ultrastructural studies. Arch Pathol Lab Med 1977;101:113–117. 323. Schiff ER. Atypical clinical manifestations of hepatitis A. Vaccine 1992;10(Suppl 1):S18–S20. 324. Scholz E, Heinricy U, Flehmig B. Acid stability of hepatitis A virus. J Gen Virol 1989; 70(Pt 9):2481–2485. 325. Schulman AN, Dienstag JL, Jackson DR, et al. Hepatitis A antigen particles in liver, bile, and stool of chimpanzees. J Infect Dis 1976;134:80–84. 326. Schulte I, Hitziger T, Giugliano S, et al. Characterization of CD8+ T-cell response in acute and resolved hepatitis A virus infection. J Hepatol 2010;54:201–208. 327. Schultheiss T, Kusov YY, Gauss-Müller V. Proteinase 3C of hepatitis A virus (HAV) cleaves the HAV polyprotein P2-P3 at all sites including VP1/2A and 2A/2B. Virology 1994;198:275–281. 328. Schultheiss T, Sommergruber W, Kusov Y, Gauss-Müller V. Cleavage specificity of purified recombinant hepatitis A virus 3C proteinase on natural substrates. J Virol 1995;69:1727–1733. 329. Schultz DE, Hardin CC, Lemon SM. Specific interaction of glyceraldehyde 3-phosphate dehydrogenase with the 5′ nontranslated RNA of hepatitis A virus. J Biol Chem 1996;271:14134–14142. 330. Seganti L, Superti F, Orsi N, et al. Study of the chemical nature of Frp/3 cell recognition units for hepatitis A virus. Med Microbiol 59

Immunol 1987;176:21–26. 331. Seggewiß N, Paulmann D, Dotzauer A. Lysosomes serve as a platform for hepatitis A virus particle maturation and nonlytic release. Arch Virol 2016;161:43–52. 332. Shaffer DR, Brown EA, Lemon SM. Large deletion mutations involving the first pyrimidine-rich tract of the 5′ nontranslated RNA of human hepatitis A virus define two adjacent domains associated with distinct replication phenotypes. J Virol 1994;68:5568–5578. 333. Shaffer DR, Emerson SU, Murphy PC, et al. A hepatitis A virus deletion mutant which lacks the first pyrimidine-rich tract of the 5′ nontranslated RNA remains virulent in primates after direct intrahepatic nucleic acid transfection. J Virol 1995;69:6600–6604. 334. Shaffer DR, Lemon SM. Temperature-sensitive hepatitis A virus mutants with deletions downstream of the first pyrimidine-rich tract of the 5′ nontranslated RNA are impaired in RNA synthesis. J Virol 1995;69:6498–6506. 335. Shahbazian D, Parsyan A, Petroulakis E, et al. eIF4B controls survival and proliferation and is regulated by proto-oncogenic signaling pathways. Cell Cycle 2010;9:4106–4109. 336. Shevtsova ZV, Lapin BA, Doroshenko NV, et al. Spontaneous and experimental hepatitis A in Old World monkeys. J Med Primatol 1988;17:177–194. 337. Shimizu YK, Mathiesen LR, Lorenz D, et al. Localization of hepatitis A antigen in liver tissue by peroxidase-conjugated antibody method: light and electron microscopic studies. J Immunol 1978;121:1671–1679. 338. Shimizu YK, Shikata T, Beninger PR, et al. Detection of hepatitis A antigen in human liver. Infect Immun 1982;36:320–324. 339. Shin EC, Jeong SH. Natural history, clinical manifestations, and pathogenesis of hepatitis A. Cold Spring Harb Perspect Med 2018;8:a031708. 340. Shouval D. Immunization against Hepatitis A. Cold Spring Harb Perspect Med 2019;9(2):a031682. 341. Shubin AV, Demidyuk IV, Lunina NA, et al. Protease 3C of hepatitis A virus induces vacuolization of lysosomal/endosomal organelles and caspase-independent cell death. BMC Cell Biol 2015;16:4. 342. Shukla A, Dey D, Banerjee K, et al. The C-terminal region of the non-structural protein 2B from Hepatitis A Virus demonstrates lipidspecific viroporin-like activity. Sci Rep 2015;5:15884. 343. Shukla A, Padhi AK, Gomes J, et al. The VP4 peptide of hepatitis A virus ruptures membranes through formation of discrete pores. J Virol 2014;88:12409–12421. 344. Siciński P, Rowiński J, Warchoł JB, et al. Poliovirus type 1 enters the human host through intestinal M cells. Gastroenterology 1990;98:56–58. 345. Sickbert-Bennett EE, Weber DJ, Gergen-Teague MF, et al. Comparative efficacy of hand hygiene agents in the reduction of bacteria and viruses. Am J Infect Control 2005;33:67–77. 346. Siegl G, Frosner GG. Characterization and classification of virus particles associated with hepatitis A. I. Size, density and sedimentation. J Virol 1978;26:40–47. 347. Siegl G, Frosner GG, Gauss-Muller V, et al. The physicochemical properties of infectious hepatitis A virions. J Gen Virol 1981;57:331–341. 348. Siegl G, Weitz M, Kronauer G. Stability of hepatitis A virus. Intervirology 1984;22:218–226. 349. Sikuler E, Keynan A, Hanuka N, et al. Detection and persistence of specific IgA antibodies in serum of patients with hepatitis A by capture radioimmunoassay. J Med Virol 1983;11:287–294. 350. Silberstein E, Xing L, van de Beek W, et al. Alteration of hepatitis A virus (HAV) particles by a soluble form of HAV cellular receptor 1 containing the immunoglobulin- and mucin-like regions. J Virol 2003;77:8765–8774. 351. Sjogren MH, Tanno H, Fay O, et al. Hepatitis A virus in stool during clinical relapse. Ann Intern Med 1987;106:221–226. 352. Skinhoj P, Mikkelsen F, Hollinger FB. Hepatitis Ain Greenland: importance of specific antibody testing in epidemiologic surveillance. Am J Epidemiol 1977;105:140–147. 353. Smith DB, Simmonds P. Classification and genomic diversity of enterically transmitted hepatitis viruses. Cold Spring Harb Perspect Med 2018;8:a031880. 354. Snooks MJ, Bhat P, Mackenzie J, et al. Vectorial entry and release of hepatitis A virus in polarized human hepatocytes. J Virol 2008;82:8733–8742. 355. Stapleton JT, Frederick J, Meyer B. Hepatitis A virus attachment to cultured cell lines. J Infect Dis 1991;164:1098–1103. 356. Stapleton JT, Lange DK, LeDuc JW, et al. The role of secretory immunity in hepatitis A virus infection. J Infect Dis 1991;163:7–11. 357. Stapleton JT, Lemon SM. Neutralization escape mutants define a dominant immunogenic neutralization site on hepatitis A virus. J Virol 1987;61:491–498. 358. Stapleton JT, Raina V, Winokur PL, et al. Antigenic and immunogenic properties of recombinant hepatitis A virus 14S and 70S subviral particles. J Virol 1993;67:1080–1085. 359. Stewart DR, Morris TS, Purcell RH, et al. Detection of antibodies to the nonstructural 3C proteinase of hepatitis A virus. J Infect Dis 1997;176:593–601. 360. Stockley PG, Twarock R, Bakker SE, et al. Packaging signals in single-stranded RNA viruses: nature’s alternative to a purely electrostatic assembly mechanism. J Biol Phys 2013;39:277–287. 361. Stokes J, Jr., Farquhar JA, Drake ME, et al. Infectious hepatitis: length of protection by immune serum globulin (gamma globulin) during epidemics. JAMA 1951;147:714–719. 362. Stuart DI, Ren J, Wang X, et al. Hepatitis A virus capsid structure. Cold Spring Harb Perspect Med 2019;9(5):a031807. 363. Sun L, Li Y, Misumi I, et al. IRF3-mediated pathogenicity of human hepatitis A virus in mice. PLoS Pathog. 2021;17:e1009960. 364. Sun L, Zhang F, Guo F, et al. The dihydroquinolizinone compound RG7834 inhibits the polyadenylase function of PAPD5 and PAPD7 and accelerates the degradation of matured hepatitis B virus surface protein mRNA. Antimicrob Agents Chemother 2020;65:e00640-20. 365. Sung PS, Hong SH, Lee J, et al. CXCL10 is produced in hepatitis A virus-infected cells in an IRF3-dependent but IFN-independent manner. Sci Rep 2017;7:6387. 366. Takahashi Y, Misumi S, Muneoka A, et al. Nonhuman primate intestinal villous M-like cells: an effective poliovirus entry site. Biochem Biophys Res Commun 2008;368:501–507. 367. Taylor GM, Goldin RD, Karayiannis P, et al. In situ hybridization studies in hepatitis A infection. Hepatology 1992;16:642–648. 368. Taylor M, Goldin RD, Ladva S, et al. In situ hybridization studies of hepatitis A viral RNA in patients with acute hepatitis A. J Hepatol 1994;20:380–387. 369. Taylor KL, Murphy PC, Asher LVS, et al. Attenuation phenotype of a cell culture-adapted variant of hepatitis A virus (HM175/p16) in susceptible new world owl monkeys. J Infect Dis 1993;168:592–601. 370. Teixera MR Jr, Weller IVD, Murray A, et al. The pathology of hepatitis A in man. Liver 1982;2:53–60. 371. Teo CG. 19th-century and early 20th-century jaundice outbreaks, the USA. Epidemiol Infect 2018;146:138–146. 372. Tesar M, Jia XY, Summers DF, et al. Analysis of a potential myristoylation site in hepatitis A virus capsid protein VP4. Virology 1993;194:616–626. 373. Tesar M, Pak I, Jia XY, et al. Expression of hepatitis A virus precursor protein P3 in vivo and in vitro: polyprotein processing of the 3CD cleavage site. Virology 1994;198:524–533. 60

374. Teterina NL, Bienz K, Egger D, et al. Induction of intracellular membrane rearrangements by HAV proteins 2C and 2BC. Virology 1997;237:66–77. 375. Theamboonlers A, Abe K, Thongmee C, et al. Complete coding sequence and molecular analysis of hepatitis A virus from a chimpanzee with fulminant hepatitis. J Med Primatol 2012;41:11–17. 376. Thompson P, Lu J, Kaplan GG. The Cys-rich region of hepatitis A virus cellular receptor 1 is required for binding of hepatitis A virus and protective monoclonal antibody 190/4. J Virol 1998;72:3751–3761. 377. Ticehurst JR, Racaniello VR, Baroudy BM, et al. Molecular cloning and characterization of hepatitis A virus cDNA. Proc Natl Acad Sci U S A 1983;80:5885–5889. 378. Tjon GM, Coutinho RA, van den Hoek A, et al. High and persistent excretion of hepatitis A virus in immunocompetent patients. J Med Virol 2006;78:1398–1405. 379. Tong MJ, El-Farra NS, Grew MI. Clinical manifestations of hepatitis A: Recent experience in a community teaching hospital. J Infect Dis 1995;171(Suppl 1):S15–S18. 380. Tong MJ, Thursby M, Rakela J, et al. Studies on the maternal-infant transmission of the viruses which cause acute hepatitis. Gastroenterology 1981;80:999–1004. 381. Totsuka A, Moritsugu Y. Hepatitis A vaccine development in Japan. In: Nishioka K, Suzuki H, Mishiro S, et al., eds. Viral Hepatitis and Liver Disease. Tokyo, Japan: Springer-Verlag; 1994:509–513. 382. Treyer A, Musch A. Hepatocyte polarity. Compr Physiol 2013;3:243–287. 383. Tsarev SA, Emerson SU, Balayan MS, et al. Simian hepatitis A virus (HAV) strain AGM-27: comparison of genome structure and growth in cell culture with other HAV strains. J Gen Virol 1991;72:1677–1683. 384. Vallbracht A, Gabriel P, Maier K, et al. Cell-mediated cytotoxicity in hepatitis A virus infection. Hepatology 1986;6:1308–1314. 385. Vallbracht A, Hofmann L, Wurster KG, et al. Persistent infection of human fibroblasts by hepatitis A virus. J Gen Virol 1984;65:609–615. 386. Vallbracht A, Maier K, Stierhof YD, et al. Liver-derived cytotoxic T cells in hepatitis A virus infection. J Infect Dis 1989;160:209–217. 387. Van Damme P, Banatvala J, Fay O, et al. Hepatitis A booster vaccination: is there a need? Lancet 2003;362:1065–1071. 388. Van Effelterre T, Guignard A, Marano C, et al. Modeling the hepatitis A epidemiological transition in Brazil and Mexico. Hum Vaccin Immunother 2017;13:1942–1951. 389. Vento S, Garofano T, di Perri G, et al. Identification of hepatitis A virus as a trigger for autoimmune chronic hepatitis type 1 in susceptible individuals. Lancet 1991;337:1183–1187. 390. Vento S, Garofano T, Renzini C, et al. Fulminant hepatitis associated with hepatitis A virus superinfection in patients with chronic hepatitis C. N Engl J Med 1998;338:286–290. 391. Victor JC, Monto AS, Surdina TY, et al. Hepatitis A vaccine versus immune globulin for postexposure prophylaxis. N Engl J Med 2007;357:1685–1694. 392. Villarejos VM, Serra C J, Anderson-Visona K, et al. Hepatitis A virus infection in households. Am J Epidemiol 1982;115:577–586. 393. Vives-Adrian L, Garriga D, Buxaderas M, et al. Structural basis for host membrane remodeling induced by protein 2B of hepatitis A virus. J Virol 2015;89:3648–3658. 394. Vizzotti C, González J, Gentile A, et al. Impact of the single-dose immunization strategy against hepatitis A in Argentina. Pediatr Infect Dis J 2014;33:84–88. 395. Voegt H. Zur Aetiologie der Hepatitis epidemica. MMW Munch Med Wochenschr 1942;89:76–79. 396. Votteler J, Ogohara C, Yi S, et al. Designed proteins induce the formation of nanocage-containing extracellular vesicles. Nature 2016;540:292–295. 397. Votteler J, Sundquist WI. Virus budding and the ESCRT pathway. Cell Host Microbe 2013;14:232–241. 398. Walczak CP, Leto DE, Zhang L, et al. Ribosomal protein RPL26 is the principal target of UFMylation. Proc Natl Acad Sci U S A 2019;116:1299–1308. 399. Walker CM. Adaptive immune responses in hepatitis A virus and hepatitis E virus infections. Cold Spring Harb Perspect Med 2019;9(9):a033472. 400. Walker CM, Feng Z, Lemon SM. Reassessing immune control of hepatitis A virus. Curr Opin Virol 2015;11:7–13. 401. Walter TS, Ren J, Tuthill TJ, et al. A plate-based high-throughput assay for virus stability and vaccine formulation. J Virol Methods 2012;185:166–170. 402. Wang D, Fang L, Wei D, et al. Hepatitis A virus 3C protease cleaves NEMO to impair induction of beta interferon. J Virol 2014;88:10252–10258. 403. Wang M, Feng Z. Mechanisms of hepatocellular injury in hepatitis A. Viruses 2021;13(5):861. 404. Wang L, Jeng KS, Lai MM. Poly(C)-binding protein 2 interacts with sequences required for viral replication in the hepatitis C virus (HCV) 5′ untranslated region and directs HCV RNA replication through circularizing the viral genome. J Virol 2011;85:7954–7964. 405. Wang X, Ren J, Gao Q, et al. Hepatitis A virus and the origins of picornaviruses. Nature 2015;517:85–88. 406. Wang H, Shi Y, Song J, et al. Ebola viral glycoprotein bound to its endosomal receptor Niemann-Pick C1. Cell 2016;164:258–268. 407. Wang X, Zhu L, Dang M, et al. Potent neutralization of hepatitis A virus reveals a receptor mimic mechanism and the receptor recognition site. Proc Natl Acad Sci U S A 2017;114:770–775. 408. Wasley A, Miller JT, Finelli L. Surveillance for acute viral hepatitis—United States, 2005. MMWR Surveill Summ 2007;56:1–24. 409. Wasley A, Samandari T, Bell BP. Incidence of hepatitis A in the United States in the era of vaccination. JAMA 2005;294:194–201. 410. Wassenaar TM, Jun SR, Robeson M, et al. Comparative genomics of hepatitis A virus, hepatitis C virus, and hepatitis E virus provides insights into the evolutionary history of Hepatovirus species. Microbiologyopen 2020;9:e973. 411. Weitz M, Baroudy BM, Maloy WL, et al. Detection of a genome-linked protein (VPg) of hepatitis A virus and its comparison with other picornaviral VPgs. J Virol 1986;60:124–130. 412. Werzberger A, Kuter B, Nalin D. Six years’ follow-up after hepatitis A vaccination. N Engl J Med 1998;338:1160. 413. Werzberger A, Mensch B, Kuter B, et al. A controlled trial of a formalin-inactivated hepatitis A vaccine in healthy children. N Engl J Med 1992;327:453–457. 414. Wessels E, Duijsings D, Lanke KH, et al. Effects of picornavirus 3A proteins on protein transport and GBF1-dependent COP-I recruitment. J Virol 2006;80:11852–11860. 415. Whetter LE, Day SP, Elroy-Stein O, et al. Low efficiency of the 5′ nontranslated region of hepatitis A virus RNA in directing capindependent translation in permissive monkey kidney cells. J Virol 1994;68:5253–5263. 416. White MR, Khan MM, Deredge D, et al. A dimer interface mutation in glyceraldehyde-3-phosphate dehydrogenase regulates its binding to AU-rich RNA. J Biol Chem 2015;290:1770–1785. 417. Wiedmann M, Boehm S, Schumacher W, et al. Evaluation of three commercial assays for the detection of hepatitis a virus. Eur J Clin Microbiol Infect Dis 2003;22:129–130. 418. Winokur PL, McLinden JH, Stapleton JT. The hepatitis A virus polyprotein expressed by a recombinant vaccinia virus undergoes proteolytic processing and assembly into viruslike particles. J Virol 1991;65:5029–5036. 419. Winokur PL, Stapleton JT. Immunoglobulin prophylaxis for hepatitis A. Clin Infect Dis 1992;14:580–586. 61

420. Wong DC, Purcell RH, Rosen L. Prevalence of antibody to hepatitis A and hepatitis B viruses in selected populations of the South Pacific. Am J Epidemiol 1979;110:227–236. 421. Wright F. The ‘effective number of codons’ used in a gene. Gene 1990;87:23–29. 422. Yamane D, Feng H, Rivera-Serrano EE, et al. Constitutive expression of interferon regulatory factor 1 drives intrinsic hepatocyte resistance to multiple RNA viruses. Nat Microbiol 2019;4:1096–1104. 423. Yang Y, Liang Y, Qu L, et al. Disruption of innate immunity due to mitochondrial targeting of a picornaviral protease precursor. Proc Natl Acad Sci U S A 2007;104:7253–7258. 424. Yang Y, Rijnbrand R, McKnight KL, et al. Sequence requirements for viral RNA replication and VPg uridylylation directed by the internal cis-acting replication element (cre) of human rhinovirus type 14. J Virol 2002;76:7485–7494. 425. Yang Y, Yi M, Evans DJ, et al. Identification of a conserved RNA replication element (cre) within the 3Dpol-coding sequence of hepatoviruses. J Virol 2008;82:10118–10128. 426. Yao G. Clinical spectrum and natural history of viral hepatitis A in a 1988 Shanghai epidemic. In: Hollinger FB, Lemon SM, Margolis H, eds. Viral Hepatitis and Liver Diseases. New York: Williams & Wilkins; 1991:76–78. 427. Yi M, Lemon SM. Replication of subgenomic hepatitis A virus RNAs expressing firefly luciferase is enhanced by mutations associated with adaptation of virus to growth in cultured cells. J Virol 2002;76:1171–1180. 428. Yi M, Schultz DE, Lemon SM. Functional significance of the interaction of hepatitis A virus RNA with glyceraldehyde 3-phosphate dehydrogenase (GAPDH): opposing effects of GAPDH and polypyrimidine tract binding protein on internal ribosome entry site function. J Virol 2000;74:6459–6468. 429. Yoon EL, Sinn DH, Lee HW, et al. Current status and strategies for the control of viral hepatitis A in Korea. Clin Mol Hepatol 2017;23:196–204. 430. Young MK. The indications and safety of polyvalent immunoglobulin for post-exposure prophylaxis of hepatitis A, rubella and measles. Hum Vaccin Immunother 2019;15:2060–2065. 431. Yu JM, Li LL, Xie GC, et al. Experimental infection of Marmota monax with a novel hepatitis A virus. Arch Virol 2018;163(5):1187–1193. 432. Yu JM, Li LL, Zhang CY, et al. A novel hepatovirus identified in wild woodchuck Marmota himalayana. Sci Rep 2016;6:22361. 433. Zajac AJ, Amphlett EM, Rowlands DJ, et al. Parameters influencing the attachment of hepatitis A virus to a variety of continuous cell lines. J Gen Virol 1991;72:1667–1675. 434. Zell R. Picornaviridae—the ever-growing virus family. Arch Virol 2018;163:299–317. 435. Zell R, Delwart E, Gorbalenya AE, et al. ICTV Virus Taxonomy Profile: Picornaviridae. J Gen Virol 2017;98:2421–2422. 436. Zhang HC, Chao SF, Ping LH, et al. An infectious cDNA clone of a cytopathic hepatitis A virus: genomic regions associated with rapid replication and cytopathic effect. Virology 1995;212:686–697. 437. Zhang Y, Kaplan GG. Characterization of replication-competent hepatitis A virus constructs containing insertions at the N terminus of the polyprotein. J Virol 1998;72:349–357. 438. Zhang B, Morace G, Gauss-Muller V, et al. Poly(A) binding protein, C-terminally truncated by the hepatitis A virus proteinase 3C, inhibits viral translation. Nucleic Acids Res 2007;35:5975–5984. 439. Zhang B, Seitz S, Kusov Y, et al. RNA interaction and cleavage of poly(C)-binding protein 2 by hepatitis A virus protease. Biochem Biophys Res Commun 2007;364:725–730. 440. Zhou Y, Callendret B, Xu D, et al. Dominance of the CD4+ T helper cell response during acute resolving hepatitis A virus infection. J Exp Med 2012;209:1481–1492. 441. Zhou D, Zhao Y, Kotecha A, et al. Unexpected mode of engagement between enterovirus 71 and its receptor SCARB2. Nat Microbiol 2019;4:414–419. 442. Zhu L, Wang X, Ren J, et al. Structure of Ljungan virus provides insight into genome packaging of this picornavirus. Nat Commun 2015;6:8316. 443. Zuckerman AJ. The history of viral hepatitis from antiquity to the present. In: Deinhardt F, Deinhardt J, eds. Viral Hepatitis: Laboratory and Clinical Science. New York: Marcel Dekker; 1983:3–32.

62

CHAPTER 3 Astroviruses Valerie Cortez • Stacey Schultz-Cherry Introduction History Classification Virion structure and composition Genome structure and organization ORF1a and ORF1b ORF2 ORFX Stages of replication Attachment and entry Uncoating Translation Transcription/Replication Assembly and release Pathogenesis and pathology Entry into the host and primary site of replication Cell and tissue tropism Immune response Release from host and transmission Virulence and persistence Epidemiology Origin and spread of epidemics Prevalence and seroepidemiology Genetic diversity of virus Clinical features Diagnosis Differential Laboratory Prevention and control Treatment Vaccines Perspectives Acknowledgments

INTRODUCTION The family Astroviridae includes viruses with icosahedral morphology and is nonenveloped. Their genome is composed of positive-sense, single-stranded RNA (ssRNA) and is organized into three open reading frames (ORFs). 63

Astroviruses (AstVs) have been identified in many different animal species, with new strains isolated on a regular basis, highlighting their continued emergence and broad host range. Although they are classically known as enteric viruses associated with acute gastroenteritis, extragastrointestinal disease has become increasingly recognized. Human-infecting AstVs, in particular, are common causes of gastroenteritis in young children and contribute to sporadic outbreaks.

HISTORY The term astrovirus was coined by Madeley and Cosgrove in 1975 to describe small, round viruses with a distinctive five- or six-pointed, starlike appearance (astron, star in Greek) of about 28 to 30 nm in diameter.127,128 They were initially observed by negative-stain electron microscopy (EM) in the stools of infants hospitalized with diarrhea and in outbreaks of gastroenteritis in newborn nurseries (Fig. 3.1). Subsequently, viral particles of similar size and morphology were identified by EM in association with diarrhea in a wide variety of young mammals and birds.

An important milestone was achieved in 1981 when Lee and Kurtz reported the isolation and passage of a human isolate of astrovirus in primary cell cultures.118 This led to the recognition of five serotypes in 1984,114 collectively known as human astroviruses (HAstVs), development of an enzyme immunoassay to detect viral antigen in the late 1980s,82 and confirmation of their clinical importance.83 The molecular characterization of HAstV isolates subsequently permitted the recognition of eight serotypes based on their reactivity to hyperimmune sera and the design of molecular probes for use as diagnostic tools. Beginning in 2008, metagenomic and consensus sequencing approaches led to the identification of novel AstV from humans, commonly referred to as VA and MLB viruses, based on the location they were first identified (Virginia, United States, and Melbourne, Australia, respectively),55–57,100 in addition to numerous AstVs from a broad range of animal hosts.18,54,126,132,147,166,182,195,209 The efficient propagation of classical and novel human-infecting AstVs in cell lines91,93,198 and the use of turkey105,191,215 and murine isolates in animal models33,40,213 has further advanced our knowledge of the molecular and structural biology, as well as the pathogenesis of these viruses. Still, there is a relative paucity of data available from the murine and turkey models, highlighting the need for expanded studies as well as the development of in vivo models for human-infecting AstV.

FIGURE 3.1 Electron micrograph of astrovirus in a human fecal specimen. Bar = 100 nm. (Courtesy of T. W. Lee and J. B. Kurtz.)

CLASSIFICATION The genomic architecture of the AstV genome places the ORFs encoding the nonstructural proteins at the 5′ end and the ORF encoding the structural proteins at the 3′ end. Distinctive features of this family include their morphology,168 the lack of an RNA helicase domain encoded in the genome, and the usage of a ribosomal frameshifting mechanism to translate the RNA-dependent RNA polymerase (RdRp).96

AstVs were originally classified into genera and species based solely on the host of origin; however, recent characterization of novel AstVs has shown that isolates from different animal species can be genetically similar, while genetically diverse viruses can be isolated from the same animal species.100,126,218 In fact, AstVs that infect humans occupy four distinct phylogenetic clades and include viruses that are more similar to other animal isolates, such as those from mink and sheep, than the classical eight HAstV serotypes100 (Fig. 3.2). These highly divergent viruses found among humans, as well as pigs126 and bats,32,218 represent polyphyly that is indicative of cross-species transmission, which has been further supported by the identification of recombinant AstVs from different host species.44,46

Together, these recent findings challenged the long-held dogma that AstVs were species-specific, leading to a new classification system based on the amino acid sequence of ORF2 (Astroviruses Study Group, 9th Report ICTV, 2010), which encodes the capsid polyprotein and represents the most variable region of the genome. Based on this naming system, two genera are distinguished within the Astroviridae family: Mamastrovirus and Avastrovirus (Fig. 3.3). Viruses belonging to the genus Mamastrovirus include isolates from a number of mammals, including humans, pigs, cats, mink, sheep, cows, dogs, bats, mice, rats, deer, and marine mammals, such as sea lions and bottlenose dolphins, among others. This genus includes two genogroups, GI and GII, that encompass 10 and 9 genotypes, respectively. The HAstV, MLB, VA2-5, and VA1-3 viruses are classified into Mamastrovirus genotypes 1, 6, 8, and 9, respectively.

Viruses from the genus Avastrovirus include isolates from many avian species, including turkeys, ducks, chickens, guinea fowl, and wild aquatic birds. This genus includes two proposed genotypes in genogroup GI and one in GII. Similarly to the HAstV serotypes, members of some of these species can be distinguished by serology, indicating the existence of viral serotypes within avastrovirus genotypes.186,189

64

FIGURE 3.2 Human-infecting AstVs occupy four distinct clades. Phylogenetic analysis of ORF2 amino acid sequences highlight HAstV, MLB, VA1-3, and VA2-5 clades that have been identified in humans. Phylogenetic relationships inferred by maximum likelihood estimation (LG+F) and tree constructed based on 1,000 bootstrapped replicates using MEGA X 10.1.8. Because the AstV nomenclature is currently in flux and has yet to assign a classification for all strains discussed in this chapter, host species and commonly used names will be used as references.

VIRION STRUCTURE AND COMPOSITION Ultrastructural analysis of HAstVs propagated in cell culture by EM revealed icosahedral particles of 41 nm, with spikes protruding from the surface.168 However, the star-like form of the particles was observed only after alkaline treatment. More recent studies by cryo-EM and image processing of immature and mature (infectious) HAstV particles not only confirmed the spiked icosahedral structure of virions49 but also noted remarkable differences between the two types of particles, highlighting a maturation process. The immature virus, 46 nm in diameter, contains 180 copies of a single protein of 70 kd (VP70) arranged in a T = 3 icosahedral symmetry. Two kinds of spikes, localized at two- and fivefold vertices, can be observed in these particles. An internal protein layer forming the capsid core is almost identical in immature and mature particles, whereas the distal layer that forms the spikes shows dramatic changes after virion maturation via proteolysis (Fig. 3.4). In immature particles, the distal layer contains 90 globular spikes that are reduced to 30 spikes after proteolysis and is required for HAstV infectivity. To what extent these cleavage events mediate exposure of the receptor-binding site, promote internalization or uncoating remains to be explored. In addition, it is becoming clearer that structural findings for the classical HAstV serotypes may not be generalizable to all AstV strains, as discussed in the next sections.

FIGURE 3.3 The Astroviridae family includes two genera with two genogroups each. Virus genotypes are classified based on the ORF2 amino acid sequence distances. (Classification proposed by the Astroviruses Study Group, 9th Report ICTV, 2010.)

65

FIGURE 3.4 Three-dimensional reconstruction of HAstV. Immature and mature virus models made by fitting core and spike crystal structures into the cryo-electron microscopy. Models are colored according to structural domains: inner core (purple), outer core (yellow), and spike (green). (Courtesy of Dr. Rebecca DuBois, University of California, Santa Cruz.) Nascent AstV particles are formed by its 90-kd capsid protein surrounding the viral genome.141 Extracellular particles released from HAstVinfected cells are formed by protein VP70, which results from the intracellular processing of VP90. In cell culture, HAstV particles obtained after trypsin treatment consist of three proteins in the range of 32 to 34, 27 to 29, and 25 to 26 kd, with the last two proteins overlapping in sequence.16,139 More recently, it was shown that fully infectious virions that undergo complete proteolytic processing consist of just two proteins: VP34 and VP27.3 However, the composition of mature virions is highly dependent on the virus strain. For example, a porcine-infecting AstV was shown to contain five distinguishable proteins,179 whereas an ovine-infecting AstV was found to have only two 33-kd proteins.81 Moreover, the human-infecting strains VA1, MLB1, and MLB2 do not require trypsin treatment for infectivity.91,198 Thus, there is likely considerable heterogeneity in virion structure and composition within this virus family.

GENOME STRUCTURE AND ORGANIZATION AstVs have a positive-sense, polyadenylated ssRNA genome that varies in length from approximately 6 to 8 kilobases (kb), depending on the isolate (Fig. 3.5). The RNA extracted from AstV particles, as well as RNA transcribed from a full-length genomic copy of complementary DNA (cDNA) clone, are able to initiate a productive infection in cultured cells, although with varying efficiencies.66 Two positive-sense RNA species have been identified in AstV-infected cells: the full-length genomic RNA (gRNA) and a subgenomic RNA (sgRNA) of approximately 2.4 kb.151

FIGURE 3.5 Genome organization of human astrovirus. The genomic RNA, approximately 6 to 8 kb, contains three main open reading frames (ORF1a, ORF1b, and ORF2). ORFX has also been described among many AstVs. Key genomic characteristics of this virus family include the frameshift signal, the sequence upstream of ORF2 that acts as promoter for synthesis of the sgRNA, and the stem–loop at the 3′ end. The viral genome includes 5′ and 3′ untranslated regions (UTRs) and three main ORFs of variable length depending on the isolate. The two ORFs located toward the 5′ end of the genome, designated ORF1a and ORF1b, encode several nonstructural proteins that are involved in transcription and replication of the virus genome; however, many of their functions have not been fully characterized. ORF1a and ORF1b overlap in 10 to 148 nucleotides (nt) in the genome of mammalian viruses and between 10 and 45 nt in avian viruses. The overlapping region contains an essential signal for translation of the viral RNA polymerase encoded in ORF1b through a −1 nt frameshift mechanism at the conserved AAAAAAC motif.96,130 The third main ORF, found at the 3′ end of the genome and designated ORF2, encodes the capsid polyprotein and is expressed via a subgenomic promoter.150

Based on the transcription initiation site determined for the sgRNA in HAstV and other mammalian AstVs, ORF1b and ORF2 overlap by 6 to 11 nt. Conserved motifs within this region at the start of ORF2 shows partial identity with the 5′ end of the gRNA, suggesting that it has an important role for the synthesis of both gRNA and sgRNA.98 However, the existence, length, and homology of this ORF1b/ORF2 overlapping region vary in avian and mammalian viruses.60,98,126 Importantly, analysis of recombinant AstV isolated from different host species has identified this region as a hot spot for recombination.46,186,201

Of note, the terminal end of ORF2 and the adjacent 3′-UTR are highly conserved among many AstVs. The sequence of this region and predicted secondary structure fit a stem–loop II-like motif that is also found in members of other RNA virus families, including Coronaviridae and Caliciviridae,95,126,149 suggesting that it is relevant for AstV genome replication. However, exceptions exist, and the MLB clade of viruses lacks this conserved region of RNA secondary structure in the 3′-UTR.95,126 In silico analyses have identified putative host protein binding at conserved sequence motifs at both the 5′ and 3′ UTRs that appear to be involved with replication,45 including a polypyrimidine tract–binding protein, which was confirmed experimentally in vitro.53

ORF1a and ORF1b The polypeptide encoded by ORF1a (nsp1a) can range from 787 to 1,240 amino acids (aa) in length in avian and mammalian viruses. Five to six helical transmembrane motifs have been identified at the amino terminus followed by a viral serine protease (Pro)98 (Fig. 3.6), which has features consistent with trypsin-like proteases.185 Two predicted coiled-coil structures are present in nsp1a, one just upstream of the first helical transmembrane motif and the second one downstream of Pro, suggesting that some protein products of nsp1a form oligomers.98 A viral protein genome–linked (VPg) encoded downstream of Pro was first predicted based on its similarity with the VPg of caliciviruses,4 and its synthesis later confirmed to be essential for infectivity.61 Finally, a hypervariable region (HVR) located downstream of the VPg contains insertion/deletions that contribute to the variation of ORF1a length and is involved in replication efficiency of HAstV,75 including adaptation to 66

cultured cells.206

FIGURE 3.6 Nonstructural proteins predicted encoded by ORF1a and ORF1b. Polyproteins nsp1a and nsp1ab are translated from ORF1a and ORF1a/b. NBD, nucleotide-binding domain; CC, coiled-coil; Pro, viral protease; VPg, viral protein genome–linked; HVR, hypervariable region; RdRp, RNA-dependent RNA polymerase. The polypeptide nsp1a/b translated via a frameshift mechanism at the junction of ORF1a/ORF1b produces a polypeptide of 515 to 539 aa that encodes an RdRp, which is similar to the RdRp of picornaviruses, caliciviruses, and certain plant viruses.4,96

Regions that could encode an RNA helicase have not been identified in the AstV genome.96 The absence of an RNA helicase domain is unusual for a positive-strand RNA virus with a genome longer than 6 kb.96,99 While the amino terminus of nsp1a contains a predicted nucleosidebinding domain (NBD) with similar features of a nucleoside triphosphate–binding motif in some helicases,4 it lacks other motifs, such as the substrate-binding and hydrolysis domains present in these enzymes.99 Thus, it is likely that AstVs utilize a host helicase to promote its replication cycle, and in support of this idea, a recent study showed that knockdown of the host helicase DDX23 reduces replication efficiency.154

ORF2 The largest sequence variability in the AstV genome is found in ORF2, which codes for the viral capsid. This polyprotein varies from 672 to 851 aa in length depending on the isolate. The N-terminal half of the protein forms the capsid core of the particle,110 which is conserved among AstVs,97,203 since it is thought to interact with the gRNA within the virion.67,214 The C-terminal half of the protein contains the spike domain, which is predicted to participate in the early binding interactions of the virus with the host cell.110 The spike domain shows considerable genetic heterogeneity among AstVs isolated from different host species and also defines the eight HAstV serotypes. High-resolution structures obtained by x-ray crystallography have shown that the HAstV capsid core (residues 80 to 411 of HAstV serotype 1) forms two linear subdomains consisting of a jelly-roll β-barrel inner core and a squashed β-barrel outer core that contains multiple trypsin cleavage sites12,194,214 (Fig. 3.7). The HAstV spike domain (residues 430 to 648) in VP25 and VP27 forms the globular spikes that consist of dimeric structures containing a three-layered β-sandwich fold.12,48,214 The structure of an avian AstV capsid spike has also been resolved and has demonstrated significant structural divergence compared to HAstV.51

ORFX An alternative ORF (named ORFX) of 91 to 122 codons overlapping ORF2 in a +1 reading frame has been described in many mammalian AstVs.50,58,124 Its start site, located 41 to 50 nt downstream of the ORF2 AUG, is placed in a better Kozak sequence than that of ORF2 and is translated through a leaky scanning mechanism. The precise mechanism and oligomeric state of the encoded protein, XP, is currently unknown, but it has been shown to play a functional role in virion assembly and/or release.124 Comparative genomic analysis indicates that this alternative ORF is present in GI mamastroviruses (including all HAstV serotypes and MLB viruses), whereas it is typically absent in GII mamastroviruses (including VA viruses).124 Instead, GII viruses have a predicted ORFY in the −1 frame and are thought to be accessed via a ribosomal frameshift.124 Based on these differences in the phylogenetic topology of mamastroviruses, it is possible that these alternative ORFs evolved independently and therefore could perform different functions. It is notable that because ORFX overlaps ORF2 in a dispensable region of the capsid,194 it represents a region of flexibility, which could support its independent evolutionary history.

FIGURE 3.7 Crystal structures of the HAstV capsid core and spike domains. Dimeric spike domain. Models are colored according to structural domains: inner core (purple), outer core (yellow), and spike (green). (Courtesy of Dr. Rebecca DuBois, University of California, Santa Cruz.)

67

FIGURE 3.8 Replication cycle of HAstV. See details in the text.

STAGES OF REPLICATION Studies directed toward understanding the early interactions of AstVs with their host cells have been limited and have primarily focused on HAstV serotypes 1, 2, and 8; however, more recent studies have extended our understanding to novel human-infecting viruses that have shown a much more delayed replication cycle in comparison to the typical 24-hour cycle for HAstV.79,91,198 Thus, a general view of the replication cycle can be depicted (Fig. 3.8), although it is unclear whether the steps or timing outlined here can be ascribed to all AstVs or even to all humaninfecting AstVs.

Attachment and Entry Cell receptor molecules for AstVs have not been identified. Since different human-infecting strains show distinct tropism in cultured cells,24,91,93,198 it is likely that their initial binding interactions are different. For example, the commonly used human adenocarcinoma cell line, Caco-2, supports the replication of all HAstV serotypes and VA1 but is not permissive to MLB1 or MLB2.24,91,198 Structural analysis of the HAstV capsid protein has identified a putative receptor binding site on the highly variable spike domain48 and is supported by the fact that antibodies that bind region can block virus infectivity in Caco-2 cells.13,19 Motifs within the spike have also been predicted to bind to oligosaccharide moieties48,51; however, specific binding partners have not yet been identified. What is increasingly clear for HAstV serotypes is that infectivity is greatly enhanced (3 to 5 logs) and largely dependent on trypsin proteolysis.12,139,171 Because trypsin treatment of particles drastically changes its structure by pruning spike from the surface (Fig. 3.4), it is likely that this facilitates receptor interactions.

HAstV entry has been estimated to occur within a half-time of 10 minutes following inoculation,140 and early studies showed that binding triggered endocytosis based on ultrastructural analysis and the use of endocytosis-blocking agents.47 It was more recently shown that this endocytic pathway is clathrin dependent and that the virus uses early to late endosomes to begin the replication cycle.140

Within the first 15 minutes after HAstV inoculation, the ERK1/2 signaling pathway is activated and can be triggered during virus binding and/or entry into the cell.153 Although the mechanism for this activation is unknown, it is also triggered after VA1 infection but not until 30 minutes postinoculation.79 Accordingly, for both HAstV and VA1, ERK1/2 activation appears to be required to establish a productive infection, since inhibitors of this kinase significantly reduced virus yield. The early phosphoinositide 3-kinase pathway activation is also important for the HAstV replication, but the trigger and downstream signaling cascade is currently undefined.190

Uncoating The mechanism through which the viral genome is released from the infecting virus particle into the cytoplasm for translation, the cell site where it occurs, and the cellular and viral factors involved in this event are currently unknown, but the process of uncoating occurs within 130 minutes postinoculation.140 Host protein disulfide isomerase A4 was recently shown to be involved in HAstV uncoating and was identified by serotype-specific direct binding to the capsid spike domain,3 indicating that there may be additional mechanisms that facilitate uncoating.

FIGURE 3.9 Processing of the HAstV nonstructural proteins. Nsp1a and nsp1ab are processed by cellular and viral proteases, with cleavage sites noted (gray arrow heads). The N terminus is cleaved by a cellular protease, whereas the others are thought to be mediated by Pro cleavage and are supported by indirect evidence of the observed final products, some of which remain unconfirmed and/or have uncharacterized functions (dotted boxes).

68

Translation As do most cellular mRNAs, astroviral RNA contains a poly-A tail at the 3′ end but does not have a 5′ cap structure. Instead, a VPg is encoded in ORF1a,61,96 and this protein is thought to modulate the translation of viral mRNAs by interacting with translation initiation factors, as has been described for caliciviruses.30,43 At least 4 tyrosine residues, known for binding to ribonucleic acid, were shown to be important for HAstV replication, but the tyrosine at position 693 was found to be absolutely critical for replication to promote the covalent linkage between the VPg and viral RNA.61

Nonstructural Polyproteins Synthesis and Processing After uncoating, the gRNA is translated into nonstructural proteins that are produced as polyprotein precursors and then proteolytically cleaved into smaller proteins. ORF1a directs the synthesis of nsp1a (~100 kd), whereas nsp1ab (160 kd) is translated from both ORF1a and ORF1b through a frameshift mechanism (Fig. 3.9). Both polyproteins are processed cotranslationally at their amino termini, so that the expected fulllength proteins are not, or very rarely, observed in HAstV-infected cells. Information regarding processing of the nonstructural polyproteins has been obtained by in vitro translation, transient expression of cDNA clones, and analysis of HAstV-infected cells, using antibodies to different regions of nsp1a and nsp1ab.65,68,142,207 No specific processing of nsp1a and nsp1ab was observed by in vitro translation of cDNA-derived transcripts,68 indicating a requirement for cellular factors.

The processing of nsp1a occurs through iterative cleavage steps, including intermediate proteins, that ultimately generate at least five products ranging in size from 74 to 5.5 kd.65,74,102,142,207 These include the viral Pro, which contains a basic S1 pocket that cleaves acidic residues (Glu and Asp) at position P1185 and also the VPg, which aids in translation of the nonstructural and structural proteins as mentioned above. A 20-kd protein from the N terminus of nsp1a has been described as the sole product generated by cleavage from a cellular protease,65 as it is only produced by in vitro translation in the presence of microsomes.138 With exception of this cleavage event, all other processing of nsp1a and nsp1ab polyproteins are believed to depend on the Pro, and sites at residues 410 and 655 of nsp1a have been confirmed experimentally.65 The other nsp1a products, including the HVR downstream of the VPg described earlier, likely serve other functions that have yet to be defined.

Translation of the RdRp (~57 kd) occurs through a −1 frameshift mechanism in the overlapping region between ORF1a and ORF1b.142 The signal that modulates this event has two key features conserved among all astroviruses: a heptameric sequence (AAAAAAC) and the potential to form a downstream stem–loop structure.96,130

Structural Polyprotein Synthesis and Processing The structural proteins of HAstV encoded in ORF2 are synthesized from the sgRNA as a polyprotein precursor of 87 to 90 kd (VP90) (Fig. 3.10). VP90 is then intracellularly cleaved at Asp-657 to yield VP7010 through intermediates of 75 to 85 kd,141 whose biological relevance, if any, is not known. Processing of VP90 to VP70 is carried out by cellular enzymes (caspases) that are involved in apoptotic processes.102 Caspase activity is triggered during infection by an unknown mechanism, although nsp1a has been implicated.10,73 Pro may also be a likely candidate since it can cleave Asp residues185 and therefore could activate procaspases. The caspase recognition motifs present at the carboxy-terminal region of VP90 are conserved among AstV strains, but it is likely that different caspases can be involved in the cleavage of VP90 since the motifs vary among isolates. While activity of initiator (caspase-8, -9, and -4) and executioner (caspase-3 and -7) caspases is clearly detected at 12 hours postinfection (hpi),10 caspase-3 and caspase-9 seem particularly efficient at processing VP90. Of note, both VP90 and VP70 can assemble into viral particles.10,138 Extracellular particles formed by VP70 are weakly infectious, but its infectivity is strongly enhanced by treatment with trypsin,12,139 which is present in the intestinal lumen. It has even been shown that VP90-assembled particles generated in the presence of caspase inhibitors can be rendered infectious by trypsin treatment.141 VP70 contains cleavage sites that yield VP41 (the N-terminal product) and VP28 (the C-terminal product) polypeptides. VP41 is subsequently cleaved at its carboxy terminus to yield VP34, which forms the capsid core. A single-point mutation in VP34 (Thr-227 to Ala or Ser) can block the production of nascent HAstV, but the precise mechanism is not defined.135 VP28 is cleaved to the final products VP27 and VP25 that share their carboxy termini and form the spikes of the virion.139 More recently, it was shown that fully matured, infectious HAstV particles actually lack VP25.3 Thus, the final mature virion is composed of VP34 and VP27. Structural information on other human-infecting viruses from the VA and MLB clades as well as other mammalian AstVs remain to be determined, but it is notable that the maturation process via trypsin proteolysis may not play an essential role.

FIGURE 3.10 The HAstV structural protein VP90 domain structure and proteolytic processing. The primary ORF2 product, VP90, contains two domains that can be distinguished by their degree of conservation: the N-terminal domain is highly conserved and forms the core of the capsid; the hypervariable C-terminal domain forms the spikes of the virus particle. VP90 contains basic and acidic regions that are highly conserved. The primary product of ORF2 is sequentially processed at its carboxy terminus by caspases (open arrow heads) to generate VP70, the main protein present in extracellular particles. These particles are processed by trypsin (closed arrow heads) to generate protein intermediates and then the final products VP34, VP27, and VP25. Virion maturation is complete after VP25 disassociates from the particle.

69

Transcription/Replication Astrovirus RNA synthesis has not been well-studied. In general, the gRNA is first used as a template to synthesize a full-length negative-sense RNA (gRNA(−)) which in turn is used as a template to produce both the full-length gRNA and the sgRNA. These double-stranded RNA molecules can be detected by immunofluorescent staining by 6 hpi in HAstV-infected and 12 hpi in VA1-infected cells.79,80 Nascent gRNA and sgRNA can be detected by PCR at 8 hpi, indicating that at this time gRNA(−) is already synthesized. The gRNA(−) can start to be detected at 9 hpi, and it accumulates to 0.7% to 4% of gRNA.90 Similar to caliciviruses, the viral polymerase recognizes a cis-element that acts as a promoter on gRNA(−) to synthesize the sgRNA, which in the case of HAstV can reach 5- to 10-fold higher molar abundance than the full-length gRNA.151

Synthesis of gRNA(−) and accumulation of gRNA require protein synthesis but not cellular DNA transcription.90 Pro, VPg, RdRp, VP90, gRNA(−), and viral particles have all been found to be associated with internal cell membranes.62,74,138 This suggests that RNA replication and the first steps of morphogenesis are carried out associated with the observed membranous structures that probably derive from the ER62 based on their localization74 and the predicted hydrophobic transmembrane helixes in nsp1a may contribute to the targeting of replication complexes to these membranes. It is likely that rearrangement of internal membranes is induced by AstV infection, and this has been observed in the infected intestinal epithelial cells of lambs.70

Assembly and Release The expression of ORF2 in cultured cells using recombinant vaccinia virus42 or baculovirus28 as vectors leads to the assembly of virus-like particles, indicating that the encoded protein is able to self-assemble in the absence of gRNA. However, these particles were unstable and show atypical morphology when purified, indicating a defective assembly.28,152 Consistent with these data, virus assembly tolerates some deletion or changes in the 30 N-terminal basic amino acids of VP90/VP70, which are thought to interact with the gRNA in the particles.67

Assembly likely takes place within the same proximity of replication, but VP90-containing particles can also be detected in the cytosol.138 After their initial assembly, viral particles are thought to separate from the membrane-associated structures, exposing their carboxy termini, which would then undergo caspase cleavage.138 Proteolytic processing by caspases of VP90 to yield VP70 is required for cell egress of the virus, as caspase inhibitors can block this step, whereas proapoptotic factors, such as TRAIL, can promote it.141 In addition to its role in cleaving VP90 to VP70, the activity of caspase-3 is required for efficient viral egress.10 Despite caspase-3 activations and apoptotic markers detected at 12 hpi, cell death is typically not observed even at high multiplicities of infection.10 A nonlytic mechanism appears to be involved in the release of HAstV and also for VA1 and MLB strains.79,91,198 In support of this, XP, the viroporin coded by ORFX in a subset of mammalian AstVs, can mediate virion formation and/or release.124 It remains to be seen whether XP does so through ion channel activity or if it has a role in trafficking of nascent particles from the ER to the trans-Golgi network to the plasma membrane, much like the Matrix-2 protein encoded by influenza viruses or Viral Protein U encoded by human immunodeficiency virus-1.124 It was also recently described that a murine-infecting AstV may exit the cell through a nonlytic mechanism via the secretory pathway within goblet cells, the main epithelial cells that produce the intestinal mucous barrier, as perturbations in mucous production led to a decrease in virus replication.38 However, it is likely that AstVs use more than one mechanism to exit the cell, as a cytopathic effect has been observed upon infection with porcine179 and chicken-infecting AstV strains.14

PATHOGENESIS AND PATHOLOGY Entry Into the Host and Primary Site of Replication As enteric pathogens, AstVs can transmit via the fecal–oral route, which was originally demonstrated by experimental inoculation of human volunteers with HAstV.115,144 The virus can also transmit through fomites and contaminated food or water.2,164 In the case of avian viruses, transmission occurs through the fecal–oral route, but it has also been proposed to occur by vertical transmission.123,192 As discussed in greater detail in the following sections, extra-gastrointestinal disease associated with AstVs is becoming increasingly appreciated and may indicate alternate routes of infection and sites of replication.

Cell and Tissue Tropism Mammalian AstVs primarily infect epithelial cells of the intestinal tract. Much of what we understand about the tissue tropism of humaninfecting AstVs comes from a handful of case reports, in vitro models, and more recently, the use of epithelial only cultures derived from human intestinal epithelial crypt cells, known as enteroid and colonoids based on whether they are derived from the small intestine or colon, respectively. In 2004, a biopsy was obtained from an immunocompromised child with persistent diarrhea, which revealed HAstV present in the small intestine and within mature epithelial cells near the villus tips in the jejunum and duodenum but not in the stomach175 (Fig. 3.11). A recent study in enteroids and colonoids demonstrated a multicellular tropism for VA1, with the highest proportion of infected cells identified among mature enterocytes, goblet cells, and CD44+ progenitor cells.106 HAstV and MLB1 could also be propagated in enteroid and colonoid cultures, although their specific cell tropisms remain undefined in these models. The only immortalized cell line identified thus far that is capable of propagating HAstV, VA, and MLB viruses are A549 cells, which were derived from human alveolar basal epithelial adenocarcinoma cells.24,91,198 Consistent across these in vitro models is the lack of cell death and inflammation following infection with all human-infecting AstVs,77,79,91,133,198 which was also observed in the biopsy sample from the immunocompromised child with persistent diarrhea.175 The precise mechanism driving acute gastroenteritis after infection remains unclear. One potential mechanism may come from the ability of the HAstV capsid to increase the permeability of Caco-2 cells grown in a monolayer, disrupt the actin cytoskeleton, and reorganize occludin, a tight junction protein.152 However, it was recently shown that VA1 inoculation does not disrupt barrier permeability or reorganize occludin in Caco-2 cells.79 Thus, there may be distinct mechanism(s) of disease for human-infecting AstVs.

70

FIGURE 3.11 Electron micrograph of a jejunal enterocyte demonstrating cytoplasmic paracrystalline viral arrays of HAstV. Bar = 500 nm. (Courtesy of Glenn Anderson, Great Ormond Street Hospital for Children.) The spectrum of human enteric AstV disease is mirrored in other host species, ranging from nonpathogenic infections to severe gastroenteritis. Murine-infecting AstVs were discovered in research and commercial animal housing facilities across the globe54,157 and were established as a small animal model of asymptomatic infection since animals do not exhibit signs of gastroenteritis.33,40 Even in infected animals deficient in innate and adaptive immune responses, there is a lack of clinical disease despite detectable virus in multiple organs.213 Like HAstV, the virus selectively infects the mouse small intestine and has shown a propensity for goblet cells in wild-type C56BL/6 animals (Fig. 3.12A) and both goblet cells and enterocytes in immunodeficient Rag2−/−Il2rg−/− animals that lack B, T, and NK cells.38,40,86 Using single-cell transcriptional profiling, it was discovered that only a subset of goblet cells were infected in wild-type mice.38 This subset included goblet cells that were actively secreting mucus. Consistent with these findings, murine enteroids grown as two-dimensional monolayers support AstV replication (Fig. 3.12B) and differentiated enteroids with an air–liquid interface produce even higher amounts of virus,86 highlighting a propensity for actively secreting cells.205

Studies of other mamastroviruses indicate that ovine- and bovine-infecting AstV may have an expanded tropism beyond epithelial cells, including subepithelial cells184 as well as M cells of the small intestine.210 Infections of gnotobiotic lambs with ovine-infecting AstVs lead to diarrhea, transient villus atrophy, and crypt hypertrophy,70,184 but the data are more mixed for bovine-infecting AstVs, and the lack of disease observed following experimental infection of gnotobiotic calves has led a subset of these viruses to be deemed nonpathogenic,210 whereas others have been associated with enteritis.131

FIGURE 3.12 AstV infection in murine small intestinal goblet cells. A: Muc2 staining highlights goblet cells in brown (left) and AstV-infected cells in red (right) from a serial tissue sections collected from the duodenum of an infected C57BL/6 mouse. (Courtesy of Dr. Valerie Cortez, University of California, Santa Cruz.) B: AstV-infected cell (virus, green; nuclei, blue) within a two-dimensional murine enteroid culture. (Courtesy of Dr. Megan Baldridge, Washington University in St. Louis.) Avian AstVs primarily replicate in epithelial cells of the gut, but extra-gastrointestinal spread in the kidney, pancreas, lymphoid organs, and liver can lead to more severe diseases.15,60,188 Turkey-infecting AstVs isolated from birds with poult enteritis mortality syndrome, characterized by enteritis, growth depression, and high mortality rates, have been used as a model to study AstV pathogenesis using turkey poults.15,104 Like mammalian hosts, there are no drastic morphologic changes or cell death resulting from infection in poults. Instead, F-actin redistribution at the apical region of jejunum tight junctions, along with defects in Na+ absorption and changes in expression of sodium/hydrogen exchangers are thought to cause a secretory diarrhea.137,158 Similar to HAstV, the capsid protein of turkey-infecting AstV was also shown to induce diarrhea, indicating a shared mechanism of disease.137

Since 2010, mammalian AstVs have been increasingly identified in cases of central nervous system (CNS) disease, highlighting a potential new site of replication and distinct pathogenesis for certain strains. Eleven cases of AstV CNS disease have been identified thus far in humans,25,36,59,109,125,155,165,172,208,211 with the majority occurring in immunocompromised patients. While VA1, MLB1, MLB2, and HAstV serotypes 1 and 4 have been implicated as causative agents, VA1 was identified in five of these cases, indicating a potential propensity for the CNS. Postmortem analyses of brain tissue have indicated virus localized in astrocytes165 and neurons25,155,165 and are supported experimentally by the ability of VA1 to propagate in primary astrocytes and the immortalized neuronal cell line, SK-N-SH.93 CNS disease has also been identified in cattle, sheep, muskox, alpaca, swine, and mink hosts with genetically related AstVs.6,18,20–23,112,121,134,163,173,178 In particular, neuroinvasive bovine-infecting AstVs are frequently detected in cases of encephalitis by molecular screening, and ample histologic evidence has identified viral RNA and antigen in the brain176 (Fig. 3.13). Additionally, both mink-infecting and bovine-infecting AstVs can be transmitted to healthy animals via inoculation with infected brain homogenates, demonstrating a causal relationship. However, the mechanism by which AstVs either spread beyond the intestinal tract or if the virus enters through another anatomical site is currently unknown. Because viremia has been observed in many human cases,35,36,85,117,172,196,211,212,216 it is possible that this would allow for extra-gastrointestinal spread. Alternatively, AstV detection in nasal and nasopharyngeal swabs34,37,187,196,211,212 could implicate the respiratory tract as a potential portal of entry and dissemination. This is supported by the detection of a porcine-infecting AstV in the CNS, respiratory tract, and circulatory system but not feces of recently weaned pigs with neurologic disease.20 71

FIGURE 3.13 Bovine infection in the mesencephalon of a cow with neurological disease and nonsuppurative encephalitis. A: Perivascular cuffs involving mononuclear cells are a hallmark of viral brain infection (hematoxylin and eosin stain). B and C: Demonstration of viral RNA by in situ hybridization (brown) and of viral antigen by immunohistochemistry (capsid protein, red) in neuronal cell bodies and processes. Magnifications are indicated by scale bars. (Courtesy of Nicole Wildi DVM and Dr. Torsten Seuberlich, University of Bern.)

Immune Response Determinants of immunity to AstV are not well understood. Symptomatic infection is primarily found in two age groups: young children and elderly patients in long-term care facilities. Indirect evidence suggests that AstV-specific antibodies could play a role in limiting infection in the host.115 The biphasic age distribution of symptomatic infection indicates that antibody acquired early in life provides partial protection from illness through most of adult life and that immunity to AstV wanes, rendering the elderly at risk for reinfection. In wild-type mice, it was shown that animals were protected for 2 weeks following clearance of the initial infection, but reinfection could occur at 8 and 12 weeks postclearance, indicating that immunity is not long-lasting.40 In a study of children in eight low-income sites across the globe, prior HAstV infection was associated with reduced risk of a second infection within the first 2 years of life, which also provides evidence of at least short-term protection.169 Given the diversity of AstVs and serotype-specific immunity,44,114,189 it is also unlikely that exposure to one strain affords much antibody protection against another. While mucosal T-cell responses remain largely unexplored, human leukocyte antigen (HLA)-restricted HAstV-specific T cells were identified in the lamina propria of duodenal biopsies collected from healthy adults,148 but it is not known whether these cells mediate protection. In turkeys, it was also noted that total CD4+ and CD8+ T cells did not increase after infection in turkeys, but modest levels of AstV-specific antibodies were detected.103 Together, these data highlight a role for adaptive immune responses against these viruses, but it is unclear whether these responses are sustained and whether they offer cross-protection against different strains.

Multiple lines of evidence indicate that AstV elicits suboptimal innate immune responses, which likely precludes the induction of inflammation33,40,104,175 and robust adaptive immunity. Type I and type III interferon (IFN) responses are made early after HAstV, VA1, MLB1, and MLB2 infections, but unlike exogenous IFN treatment, endogenous levels are unable to control viral replication.77,106,133 Still, AstV drives sufficient levels of IFN-λ that mediate protection against murine norovirus coinfection in immunodeficient Rag2−/−Il2rg−/− animals.87 Collectively, these data suggest that IFNs are made in response to infection, but they are insufficient in controlling replication, and it is possible that AstVs have the means to subvert their induction and/or function. Minimal proinflammatory cytokine and chemokine production has been observed following HAstV and VA1 infection in vitro. However, VA1 infection induced notable exceptions, including IL-8 in Caco-2 cells and CXCL10 in primary astrocytes and neuronal cells,79,93 indicating cell signaling that could recruit innate and adaptive immune cells. Indeed, AstV infection in turkeys can increase nitric oxide production in macrophages, and this could mediate virus inhibition.103 It is also notable that TGF-β, an immunosuppressive cytokine,104 is activated in turkeys following AstV infection and could play a role in dampening immune responses. The HAstV capsid protein has also been shown to block complement activation through the classical and lectin pathways by binding C1q and mannose-binding lectin, respectively.72,78 By interfering with either the complement system and/or cytokine and chemokine induction, AstV can stymie its host from mounting a robust immune response.

Release From Host and Transmission Following replication in the intestine, AstVs are shed in the feces. Adult human volunteer studies and systemic reviews have established the incubation period for HAstV to be approximately 3 to 4 days, although a shorter incubation period of 24 to 36 hours was noted after an outbreak at a kindergarten.107,119,144 The highest viral shedding occurs by day 6, with levels proportional to symptom severity.144 Large numbers of HAstV particles, equivalent to approximately 1010 to 1011 virus genomes per gram of feces, have been observed in patients with diarrhea.75 In a recent study, HAstV, MLB, and VA virus was shed at a median range of 102 to 107 RNA copies/mL in children with and without diarrhea.200 It remains unknown what proportion of virus shed in feces represents infectious particles and whether this differs across strains.

Virulence and Persistence To date, there are not clear genetic demarcations that define virulent AstV strains. It is notable that CNS infections, in contrast to enteric infections, are associated with cell death and reactive gliosis.25,155,165 Currently, it is unclear whether specific AstV strains can induce differential disease or whether within-host viral dynamics drive the dissemination of the virus outside the gut and to the brain. While the subset of CNS-associated mammalian strains that infect humans and other animal hosts cluster within the same clade, there are other divergent AstVs associated with CNS disease, indicating that both scenarios could be supported.

AstV persistence has been studied in mice and humans, with a common theme of prolonged infections identified in immunocompromised hosts. HAstV has been associated with chronic diarrhea among immunocompromised pediatric and adult patients, including reports of patients persistently shedding virus for over a year.17,39,41,64 Similarly, it has been shown that animals deficient in adaptive and innate immune factors can become persistently infected and fail to clear the virus.133,213 Even wild-type mice can exhibit prolonged virus shedding for 3 to 10 weeks, which is likely virus and mouse strain dependent.33,40,133 Likewise, in a small study, persistent gastroenteritis was associated with HAstV serotype 3 in otherwise healthy children.27 MLB1 and MLB2 were also recently found to persist in culture even after cell passaging.198 Together, these data would suggest that there are likely viral and host factors that dictate virus persistence.

EPIDEMIOLOGY 72

Origin and Spread of Epidemics The precise origins of human-infecting AstVs are unknown. The phylogenetic relationships for AstVs suggest that some human-infecting viruses, particularly VA and MLB viruses, may have originated in animals. A few lines of evidence could support bats and pigs as potential reservoirs due to the high diversity of AstVs identified within these host species.126,218 As discussed in greater detail below, there is ample evidence of crossspecies transmission of mammalian and avian AstVs.

AstV can spread person-to-person but also through contaminated food and water. Person-to-person spread is an important cause of nosocomial and community-acquired infections, including outbreaks of diarrhea in childcare settings145,146 and sporadic outbreaks among elderly patients.94,120 Epidemiologic data indicate that contaminated food is also a main source for HAstV infection. Large, food-borne outbreaks, affecting thousands of individuals, have occurred among school-aged children as well as adults.159 HAstV has been found in bivalve mollusks and water from different origins.29 The capsid affords the virus resistance against chlorination and normal disinfectants116,174 and can therefore maintain its infectiousness for months.1 AstV prevalence peaks during the winter months,197 but most studies report year-round transmission.

Prevalence and Seroepidemiology The lack of active surveillance networks for AstV and comprehensive diagnostics capable of detecting HAstV, VA, and MLB viruses162 has led to an enormous underreporting and underdiagnosis of human infections. In addition, similar rates of infection in symptomatic and asymptomatic individuals have been observed across the world,11,143,161,197 highlighting what is likely a massive underestimation of total viral burden. Worldwide, human-infecting AstVs have primarily been identified in young children, but the elderly and persons of all ages who are immunocompromised are also at risk for disease.41,71,94,197 While the age distribution in the young can vary depending on several factors, such as underlying immune and nutritional status,160 studies from high-, middle-, and low-income countries indicate that the initial exposure to AstV occurs within the first 3 years of life,76,160 with the incidence highest in the first year of life.63,129,156 Antibodies to HAstV serotypes have been shown to be acquired in early childhood, quickly reaching 90% by age 5.108,111,113 Based on epidemiologic studies of AstV in humans that have primarily focused on HAstV, serotype 1 is the predominant strain found in children, but this can vary with age and geographic location.197 In general, the positivity rate for HAstV is approximately 5% but has been reported to be over 30% in some cohorts.197

Less data are available for MLB and VA viruses,26,84 but the positivity rate across cohorts is generally lower than HAstV at approximately 1.5%, although one study found over 10%.197 They have been commonly identified in young children, although strain-specific associations with gastroenteritis are mixed.88,101,143,181,199,217 Seroprevalence studies also indicate that exposure to MLB and VA viruses is more common than indicated by cohort studies of acute gastroenteritis, with positivity rates ranging from 65% to 100%.26,84 This could indicate a much higher frequency of asymptomatic or nondiarrheal infections compared to HAstV.

Systemic spread has been identified among all ages and has most frequently been detected in patients with a number of immunocompromised conditions, such as cancer as well as primary immunodeficiencies.39,41,64,71 Because this collective population is more at risk for severe disease, increased awareness and prioritization should be given to the surveillance of AstVs within this population, particularly in patients who exhibit neurologic symptoms.

Genetic Diversity of Virus Even among RNA viruses, the Astroviridae family is characterized by an incredible level of diversity. For example, within the eight serotypes of HAstV, there is 64% to 84% homology within the capsid protein.197 In comparison, there is only 23% to 24% homology between VA2-VA4, VA1VA3, and VA5 viruses and approximately 27.5% homology within the MLB clade.197 With new AstV strains frequently recognized, there is an ever-increasing appreciation for the diversity and broad host range of this virus family. Both the identification of recombinant strains in turkeys, guinea fowl, ducks, porcines, bovines, sea lions, and yaks as well as divergent strains observed within the same host species suggest low species barriers that would allow for interspecies transmission events.44 As evidence of this, brown rats from urban settings were found to harbor strains with shared homology with MLB viruses,31 and an MLB2-like virus was identified in a chimpanzee housed in a zoo.204 Avian AstVs have even been identified in mink,18 and there is serologic evidence of avian AstV-specific antibodies in poultry workers.136

CLINICAL FEATURES Gastroenteritis caused by AstV infection primarily affects young children throughout the world but is typically milder than rotavirus infection and does not lead to significant dehydration or hospitalization.202 Severe gastroenteritis resulting in death has been reported180 but is thought to be extremely rare. Table 3.1 summarizes the clinical features associated with this disease; however, these features can vary depending on the population studied. For example, in a recent study of children less than 2 years of age in eight low- and middle-income countries, HAstV disease severity exceeded all enteropathogens except for rotavirus.160 Complications such as dehydration can develop in patients with underlying gastrointestinal disease, poor nutritional status, as well as coinfections, which are incredibly common and can occur in over 30% of cases, depending on the study.197 In a study of adult volunteers, the development of gastroenteritis was limited and dependent on the size of the inoculum.144 Symptoms typically resolve within a few days and shedding ceases within a 2-week period, with longer durations for immunocompromised patients, as mentioned above. In other mammals, diarrhea as well as vomiting outbreaks have been observed, including in dogs,193 cats,122 and even cheetahs.7

In patients with neurologic involvement, clinical features include cognitive impairment, seizures, headache, uncontrolled dystonic movement, and limb weakness.36,59,125,165,172,211 In contrast to AstV gastroenteritis, CNS infections have primarily been fatal.25,36,125,155,165,211 On occasion, AstV has been associated with necrotizing enterocolitis in premature infants,8,9 as well as other with intestinal diseases, such as intussusception5,89 and celiac disease.183 AstV was also found in feces from five children with nonpoliovirus acute flaccid paralysis,100 although a direct causative association was not determined.

73

TABLE 3.1 Clinical symptoms of HAstV infection Diarrhea, average duration (2–3 days)

72%–100%

Abdominal pain

50%

Vomiting, average duration (1 day)

20%–70%

Fever

20%–25%

Dehydration to some degree

24%–30%

Severe dehydration

0%–5%

Hospitalization, average duration (6 days) 6% Severity scorea (1–20) (average)

5

Otitis

13%

Bronchiolitis

33%

aTwenty point scoring

system based on Ruuska T, Vesikari T. Rotavirus disease in Finnish children: use of numerical scores for clinical severity of diarrhoeal episodes. Scand J Infect Dis 1990;22(3):259–267. Ref.170

Adapted with permission from Walter JE, Mitchell DK. Astrovirus infection in children. Curr Opin Infect Dis 2003;16(3):247–253. A spectrum of disease has also been noted in avian species infected with AstVs. Infections in chickens have been associated with stunting and acute interstitial nephritis,177 whereas fatal hepatitis has been observed in ducklings.69 AstV infection in turkeys is associated with diarrhea as well as poult enteritis mortality syndrome, which causes high mortality.188 However, subclinical and mild infections have also been noted in turkeys, ducks, and chickens.14,123,192 It is likely that there are virus strain–specific differences that contribute to this spectrum of disease. In addition, mixed infections could also play a significant role in AstV pathogenesis, especially in commercial farm settings with dense animal populations and a plethora of pathogens.167

DIAGNOSIS Differential Astrovirus should be suspected in cases of mild to severe gastrointestinal illness, particularly among young, elderly, and immunocompromised individuals. AstV infection should also be included in a differential diagnosis in immunocompromised patients with neurological symptoms.

Laboratory After their discovery, AstVs were primarily detected by direct EM examination of negative-stained fecal specimens. Since then, technologies range from ELISA-based assays to identify virus from stool or to identify virus-specific antibodies.162 As more sequence data on AstV genomes became available, detection techniques have since moved to molecular-based probes and RT-PCR assays. However, the high sequence variability within this virus family largely precludes the use of a universal molecular assay for all members. Instead, genotype-specific methods or multiplexed RT-PCR assays have been developed recently for diagnosis of multiple human and animal viruses.162 These types of assays work well in the detection and screening process, but unbiased approaches are still widely used given the ever-expanding list of novel AstVs, and these technologies primarily employ endpoint PCR with degenerate primers or high-throughput metagenomic sequencing.

PREVENTION AND CONTROL Treatment Since the generally mild and self-limiting nature of gastroenteritis caused by AstVs, treatment is generally not needed unless a person becomes dehydrated and would need oral or intravenous fluid resuscitation. In rare instances, intravenous immunoglobulin has been successfully used to treat severely immunodeficient patients with either persistent gastroenteritis17 or CNS disease,59 but the data are mixed,41 and larger studies are needed to determine its efficacy. Recent studies of HAstV and VA1 infection in Caco-2 cells have implicated that the broad-spectrum antimicrobial, nitazoxanide, as well as the antivirals, ribavirin and favipiravir, are effective against viral replication.79,80,92 While the precise drug action is not yet known, the block for nitazoxanide appears to be prior to the formation of replication complexes and was also shown to 74

reduce virus levels in the turkey poult model.80 These major advancements offer potential treatment options for AstV CNS disease, which are unfortunately lacking.

Vaccines No vaccines have been developed for AstV infections in any host species; however, inoculation of the recombinant, baculovirus-expressed capsid protein into hens partially protected their offspring from gut lesions and weight depression after virus challenge.177 The mechanism of protection remains unclear, but capsid-based antigens could be a useful approach for future vaccine design.52

PERSPECTIVES The public health perception of AstVs has changed since the discovery of divergent viruses associated with severe CNS disease in humans. There is a clear need to characterize neuroinvasive AstVs in order to improve their detection and development of treatment strategies. In vivo and in vitro models will help to increase our appreciation of the broader spectrum of disease caused by AstVs, including host and viral factors that dictate disease outcomes. Expanded use of animal models for both mammalian and avian AstVs is greatly needed to study the elicitation of protective immunity and pathogenesis, including the interactions with coinfecting enteropathogens and commensal organisms. Increasing evidence indicates that interspecies transmission is common for this virus family, and additional study is needed to identify sources of emergent strains and investigate potential animal reservoirs. Aiding in this endeavor would be a refined understanding of the replication cycle, particularly the receptor and/or binding partners that initiate virus entry, as these can determine cellular and host tropisms. Additional structural and functional analyses of the viral proteins, particularly those that remain uncharacterized, will help to define this virus family. The cellular factors and mechanisms involved in viral replication are only beginning to be characterized, and future studies of host–virus interactions offer promising new directions.

ACKNOWLEDGMENTS We apologize to our colleagues whose work has not been cited due to space constraints. We thank Dr. Rebekah Honce for graphic design, Dr. Shaoyuan Tan for phylogenetic analyses, and our colleagues for sharing unpublished images. Our laboratories acknowledge the financial support from St. Jude Children’s Research Hospital Children’s Infectious Defense Center, ALSAC, the Hartwell Foundation, and the National Institute of Allergy and Infectious Diseases.

75

References 1. Abad FX, Pintó RM, Villena C, et al. Astrovirus survival in drinking water. Appl Environ Microbiol 1997;63(8):3119–3122. 2. Abad FX, Villena C, Guix S, et al. Potential role of fomites in the vehicular transmission of human astroviruses. Appl Environ Microbiol 2001;67(9):3904–3907. 3. Aguilar-Hernández N, López S, Arias CF. Minimal capsid composition of infectious human astrovirus. Virology 2018;521:58–61. 4. Al-Mutairy B, Walter JE, Pothen A, et al. Genome prediction of putative genome-linked viral protein (VPg) of astroviruses. Virus Genes 2005;31(1):21–30. 5. Aminu M, Ameh EA, Geyer A, et al. Role of astrovirus in intussusception in Nigerian infants. J Trop Pediatr 2009;55(3):192–194. 6. Arruda B, Arruda P, Hensch M, et al. Porcine Astrovirus type 3 in central nervous system of swine with polioencephalomyelitis. Emerg Infect Dis 2017;23(12):2097–2100. 7. Atkins A, Wellehan JFX, Childress AL, et al. Characterization of an outbreak of astroviral diarrhea in a group of cheetahs (Acinonyx jubatus). Vet Microbiol 2009;136(1–2):160–165. 8. Bagci S, Eis-Hübinger AM, Franz AR, et al. Detection of astrovirus in premature infants with necrotizing enterocolitis. Pediatr Infect Dis J 2008;27(4):347–350. 9. Bagci S, Eis-Hübinger AM, Yassin AF, et al. Clinical characteristics of viral intestinal infection in preterm and term neonates. Eur J Clin Microbiol Infect Dis 2010;29(9):1079–1084. 10. Banos-Lara Ma del R, Méndez E. Role of individual caspases induced by astrovirus on the processing of its structural protein and its release from the cell through a non-lytic mechanism. Virology 2010;401(2):322–332. 11. Barbosa G, Caetano A, Dábilla N, et al. Classical human astroviruses in symptomatic and asymptomatic children of Goiás, Brazil: positivity rates, viral loads, and molecular characterization. J Med Virol 2020;92(8):1053–1058. 12. Bass DM, Qiu S. Proteolytic processing of the astrovirus capsid. J Virol 2000;74(4):1810–1814. 13. Bass DM, Upadhyayula U. Characterization of human serotype 1 astrovirus-neutralizing epitopes. J Virol 1997;71(11):8666–8671. 14. Baxendale W, Mebatsion T. The isolation and characterisation of astroviruses from chickens. Avian Pathol 2004;33(3):364–370. 15. Behling-Kelly E, Schultz-Cherry S, Koci M, et al. Localization of astrovirus in experimentally infected turkeys as determined by in situ hybridization. Vet Pathol 2002;39(5):595–598. 16. Belliot G, Laveran H, Monroe SS. Capsid protein composition of reference strains and wild isolates of human astroviruses. Virus Res 1997;49(1):49–57. 17. Björkholm M, Celsing F, Runarsson G, et al. Successful intravenous immunoglobulin therapy for severe and persistent astrovirus gastroenteritis after fludarabine treatment in a patient with Waldenström’s macroglobulinemia. Int J Hematol 1995;62(2):117–120. 18. Blomström A-L, Widén F, Hammer A-S, et al. Detection of a novel astrovirus in brain tissue of mink suffering from shaking mink syndrome by use of viral metagenomics. J Clin Microbiol 2010;48(12):4392–4396. 19. Bogdanoff WA, Campos J, Perez EI, et al. Structure of a human astrovirus capsid-antibody complex and mechanistic insights into virus neutralization. J Virol 2017;91(2). 20. Boros Á, Albert M, Pankovics P, et al. Outbreaks of neuroinvasive astrovirus associated with encephalomyelitis, weakness, and paralysis among weaned pigs, Hungary. Emerg Infect Dis 2017;23(12):1982–1993. 21. Boujon CL, Koch MC, Kauer RV, et al. Novel encephalomyelitis-associated astrovirus in a muskox (Ovibos moschatus): a surprise from the archives. Acta Vet Scand 2019;61(1):31. 22. Boujon CL, Koch MC, Wüthrich D, et al. Indication of cross-species transmission of astrovirus associated with encephalitis in sheep and cattle. Emerg Infect Dis 2017;23(9):1604–1608. 23. Bouzalas IG, Wüthrich D, Walland J, et al. Neurotropic astrovirus in cattle with nonsuppurative encephalitis in Europe. J Clin Microbiol 2014;52(9):3318–3324. 24. Brinker JP, Blacklow NR, Herrmann JE. Human astrovirus isolation and propagation in multiple cell lines. Arch Virol 2000;145(9):1847–1856. 25. Brown JR, Morfopoulou S, Hubb J, et al. Astrovirus VA1/HMO-C: an increasingly recognized neurotropic pathogen in immunocompromised patients. Clin Infect Dis 2015;60(6):881–888. 26. Burbelo PD, Ching KH, Esper F, et al. Serological studies confirm the novel astrovirus HMOAstV-C as a highly prevalent human infectious agent. PLoS One 2011;6(8):e22576. 27. Caballero S, Guix S, El-Senousy WM, et al. Persistent gastroenteritis in children infected with astrovirus: association with serotype-3 strains. J Med Virol 2003;71(2):245–250. 28. Caballero S, Guix S, Ribes E, et al. Structural requirements of astrovirus virus-like particles assembled in insect cells. J Virol 2004;78(23):13285–13292. 29. Carter MJ. Enterically infecting viruses: pathogenicity, transmission and significance for food and waterborne infection. J Appl Microbiol 2005;98(6):1354–1380. 30. Chaudhry Y, Nayak A, Bordeleau M-E, et al. Caliciviruses differ in their functional requirements for eIF4F components. J Biol Chem 2006;281(35):25315–25325. 31. Chu DKW, Chin AWH, Smith GJ, et al. Detection of novel astroviruses in urban brown rats and previously known astroviruses in humans. J Gen Virol 2010;91(Pt 10):2457–2462. 32. Chu DKW, Poon LLM, Guan Y, et al. Novel astroviruses in insectivorous bats. J Virol 2008;82(18):9107–9114. 33. Compton SR, Booth CJ, Macy JD. Murine astrovirus infection and transmission in neonatal CD1 mice. J Am Assoc Lab Anim Sci 2017;56(4):402–411. 34. Cordey S, Brito F, Vu D-L, et al. Astrovirus VA1 identified by next-generation sequencing in a nasopharyngeal specimen of a febrile Tanzanian child with acute respiratory disease of unknown etiology. Emerg Microbes Infect 2016;5(9):e99. 35. Cordey S, Hartley M-A, Keitel K, et al. Detection of novel astroviruses MLB1 and MLB2 in the sera of febrile Tanzanian children. Emerg Microbes Infect 2018;7(1):27. 36. Cordey S, Vu D-L, Schibler M, et al. Astrovirus MLB2, a new gastroenteric virus associated with meningitis and disseminated infection. Emerg Infect Dis 2016;22(5):846–853. 37. Cordey S, Zanella M-C, Wagner N, et al. Novel human astroviruses in pediatric respiratory samples: a one-year survey in a Swiss tertiary care hospital. J Med Virol 2018;90(11):1775–1778. 38. Cortez V, Boyd DF, Crawford JC, et al. Astrovirus infects actively secreting goblet cells and alters the gut mucus barrier. Nat Commun 2020;11(1):2097. 39. Cortez V, Freiden P, Gu Z, et al. Persistent infections with diverse co-circulating astroviruses in pediatric oncology patients, Memphis, Tennessee, USA. Emerg Infect Dis 2017;23(2):288–290. 40. Cortez V, Sharp B, Yao J, et al. Characterizing a murine model for astrovirus using viral isolates from persistently infected immunocompromised mice. J Virol 2019;93(13). 76

41. Cubitt WD, Mitchell DK, Carter MJ, et al. Application of electronmicroscopy, enzyme immunoassay, and RT-PCR to monitor an outbreak of astrovirus type 1 in a paediatric bone marrow transplant unit. J Med Virol 1999;57(3):313–321. 42. Dalton RM, Pastrana EP, Sánchez-Fauquier A. Vaccinia virus recombinant expressing an 87-kilodalton polyprotein that is sufficient to form astrovirus-like particles. J Virol 2003;77(16):9094–9098. 43. Daughenbaugh KF, Wobus CE, Hardy ME. VPg of murine norovirus binds translation initiation factors in infected cells. Virol J 2006;3:33. 44. De Benedictis P, Schultz-Cherry S, Burnham A, et al. Astrovirus infections in humans and animals - molecular biology, genetic diversity, and interspecies transmissions. Infect Genet Evol 2011;11(7):1529–1544. 45. De Nova-Ocampo M, Soliman MC, Espinosa-Hernández W, et al. Human astroviruses: in silico analysis of the untranslated region and putative binding sites of cellular proteins. Mol Biol Rep 2019;46(1):1413–1424. 46. Donato C, Vijaykrishna D. The broad host range and genetic diversity of mammalian and avian astroviruses. Viruses 2017;9(5). 47. Donelli G, Superti F, Tinari A, et al. Mechanism of astrovirus entry into Graham 293 cells. J Med Virol 1992;38(4):271–277. 48. Dong J, Dong L, Méndez E, et al. Crystal structure of the human astrovirus capsid spike. Proc Natl Acad Sci U S A 2011;108(31):12681–12686. 49. Dryden KA, Tihova M, Nowotny N, et al. Immature and mature human astrovirus: structure, conformational changes, and similarities to hepatitis E virus. J Mol Biol 2012;422(5):650–658. 50. Du Y, Ji C, Liu T, et al. Identification of a novel protein in porcine astrovirus that is important for virus replication. Vet Microbiol 2021;255:108984. 51. DuBois RM, Freiden P, Marvin S, et al. Crystal structure of the avian astrovirus capsid spike. J Virol 2013;87(14):7853–7863. 52. Espinosa R, López T, Bogdanoff WA, et al. Isolation of neutralizing monoclonal antibodies to human astrovirus and characterization of virus variants that escape neutralization. J Virol 2019;93(2). 53. Espinosa-Hernández W, Velez-Uriza D, Valdés J, et al. PTB binds to the 3′ untranslated region of the human astrovirus type 8: a possible role in viral replication. PLoS One 2014;9(11):e113113. 54. Farkas T, Fey B, Keller G, et al. Molecular detection of novel astroviruses in wild and laboratory mice. Virus Genes 2012;45(3):518–525. 55. Finkbeiner SR, Holtz LR, Jiang Y, et al. Human stool contains a previously unrecognized diversity of novel astroviruses. Virol J 2009;6:161. 56. Finkbeiner SR, Kirkwood CD, Wang D. Complete genome sequence of a highly divergent astrovirus isolated from a child with acute diarrhea. Virol J 2008;5:117. 57. Finkbeiner SR, Li Y, Ruone S, et al. Identification of a novel astrovirus (astrovirus VA1) associated with an outbreak of acute gastroenteritis. J Virol 2009;83(20):10836–10839. 58. Firth AE, Atkins JF. Candidates in astroviruses, seadornaviruses, cytorhabdoviruses and coronaviruses for +1 frame overlapping genes accessed by leaky scanning. Virol J 2010;7:17. 59. Frémond M-L, Pérot P, Muth E, et al. Next-generation sequencing for diagnosis and tailored therapy: a case report of astrovirusassociated progressive encephalitis. J Pediatric Infect Dis Soc 2015;4(3):e53–e57. 60. Fu Y, Pan M, Wang X, et al. Complete sequence of a duck astrovirus associated with fatal hepatitis in ducklings. J Gen Virol 2009;90(Pt 5):1104–1108. 61. Fuentes C, Bosch A, Pintó RM, et al. Identification of human astrovirus genome-linked protein (VPg) essential for virus infectivity. J Virol 2012;86(18):10070–10078. 62. Fuentes C, Guix S, Bosch A, et al. The C-terminal nsP1a protein of human astrovirus is a phosphoprotein that interacts with the viral polymerase. J Virol 2011;85(9):4470–4479. 63. Gabbay YB, da Luz CRNE, Costa IV, et al. Prevalence and genetic diversity of astroviruses in children with and without diarrhea in São Luís, Maranhão, Brazil. Mem Inst Oswaldo Cruz 2005;100(7):709–714. 64. Gallimore CI, Taylor C, Gennery AR, et al. Use of a heminested reverse transcriptase PCR assay for detection of astrovirus in environmental swabs from an outbreak of gastroenteritis in a pediatric primary immunodeficiency unit. J Clin Microbiol 2005;43(8):3890–3894. 65. Geigenmüller U, Chew T, Ginzton N, et al. Processing of nonstructural protein 1a of human astrovirus. J Virol 2002;76(4):2003–2008. 66. Geigenmüller U, Ginzton NH, Matsui SM. Construction of a genome-length cDNA clone for human astrovirus serotype 1 and synthesis of infectious RNA transcripts. J Virol 1997;71(2):1713–1717. 67. Geigenmüller U, Ginzton NH, Matsui SM. Studies on intracellular processing of the capsid protein of human astrovirus serotype 1 in infected cells. J Gen Virol 2002;83(Pt 7):1691–1695. 68. Gibson CA, Chen J, Monroe SA, et al. Expression and processing of nonstructural proteins of the human astroviruses. Adv Exp Med Biol 1998;440:387–391. 69. Gough RE, Collins MS, Borland E, et al. Astrovirus-like particles associated with hepatitis in ducklings. Vet Rec 1984;114(11):279. 70. Gray EW, Angus KW, Snodgrass DR. Ultrastructure of the small intestine in astrovirus-infected lambs. J Gen Virol 1980;49(1):71–82. 71. Grohmann GS, Glass RI, Pereira HG, et al. Enteric viruses and diarrhea in HIV-infected patients. Enteric opportunistic infections working group. N Engl J Med 1993;329(1):14–20. 72. Gronemus JQ, Hair PS, Crawford KB, et al. Potent inhibition of the classical pathway of complement by a novel C1q-binding peptide derived from the human astrovirus coat protein. Mol Immunol 2010;48(1–3):305–313. 73. Guix S, Bosch A, Ribes E, et al. Apoptosis in astrovirus-infected CaCo-2 cells. Virology 2004;319(2):249–261. 74. Guix S, Caballero S, Bosch A, et al. C-terminal nsP1a protein of human astrovirus colocalizes with the endoplasmic reticulum and viral RNA. J Virol 2004;78(24):13627–13636. 75. Guix S, Caballero S, Bosch A, et al. Human astrovirus C-terminal nsP1a protein is involved in RNA replication. Virology 2005;333(1):124–131. 76. Guix S, Caballero S, Villena C, et al. Molecular epidemiology of astrovirus infection in Barcelona, Spain. J Clin Microbiol 2002;40(1):133–139. 77. Guix S, Pérez-Bosque A, Miró L, et al. Type I interferon response is delayed in human astrovirus infections. PLoS One 2015;10(4):e0123087. 78. Hair PS, Gronemus JQ, Crawford KB, et al. Human astrovirus coat protein binds C1q and MBL and inhibits the classical and lectin pathways of complement activation. Mol Immunol 2010;47(4):792–798. 79. Hargest V, Davis AE, Tan S, et al. Human astroviruses: a tale of two strains. Viruses 2021;13(3). 80. Hargest V, Sharp B, Livingston B, et al. Astrovirus replication is inhibited by nitazoxanide in vitro and in vivo. J Virol 2020;94(5). 81. Herring AJ, Gray EW, Snodgrass DR. Purification and characterization of ovine astrovirus. J Gen Virol 1981;53(Pt 1):47–55. 82. Herrmann JE, Nowak NA, Perron-Henry DM, et al. Diagnosis of astrovirus gastroenteritis by antigen detection with monoclonal antibodies. J Infect Dis 1990;161(2):226–229. 83. Herrmann JE, Taylor DN, Echeverria P, et al. Astroviruses as a cause of gastroenteritis in children. N Engl J Med 1991;324(25):1757–1760. 77

84. Holtz LR, Bauer IK, Jiang H, et al. Seroepidemiology of astrovirus MLB1. Clin Vaccine Immunol 2014;21(6):908–911. 85. Holtz LR, Wylie KM, Sodergren E, et al. Astrovirus MLB2 viremia in febrile child. Emerg Infect Dis 2011;17(11):2050–2052. 86. Ingle H, Hassan E, Gawron J, et al. Murine astrovirus tropism for goblet cells and enterocytes facilitates an IFN-λ response in vivo and in enteroid cultures. Mucosal Immunol 2021;14:751–761. 87. Ingle H, Lee S, Ai T, et al. Viral complementation of immunodeficiency confers protection against enteric pathogens via IFN-lambda. Nat Microbiol 2019;4:1120–1128. 88. Jacobsen S, Höhne M, Marques AM, et al. Co-circulation of classic and novel astrovirus strains in patients with acute gastroenteritis in Germany. J Infect 2018;76(5):457–464. 89. Jakab F, Péterfai J, Verebély T, et al. Human astrovirus infection associated with childhood intussusception. Pediatr Int 2007;49(1):103–105. 90. Jang SY, Jeong WH, Kim MS, et al. Detection of replicating negative-sense RNAs in CaCo-2 cells infected with human astrovirus. Arch Virol 2010;155(9):1383–1389. 91. Janowski AB, Bauer IK, Holtz LR, et al. Propagation of astrovirus VA1, a neurotropic human astrovirus, in cell culture. J Virol 2017;91(19). 92. Janowski AB, Dudley H, Wang D. Antiviral activity of ribavirin and favipiravir against human astroviruses. J Clin Virol 2020;123:104247. 93. Janowski AB, Klein RS, Wang D. Differential in vitro infection of neural cells by astroviruses. MBio 2019;10(4). 94. Jarchow-Macdonald AA, Halley S, Chandler D, et al. First report of an astrovirus type 5 gastroenteritis outbreak in a residential elderly care home identified by sequencing. J Clin Virol 2015;73:115–119. 95. Jiang H, Holtz LR, Bauer I, et al. Comparison of novel MLB-clade, VA-clade and classic human astroviruses highlights constrained evolution of the classic human astrovirus nonstructural genes. Virology 2013;436(1):8–14. 96. Jiang B, Monroe SS, Koonin EV, et al. RNA sequence of astrovirus: distinctive genomic organization and a putative retrovirus-like ribosomal frameshifting signal that directs the viral replicase synthesis. Proc Natl Acad Sci U S A 1993;90(22):10539–10543. 97. Jonassen CM, Jonassen TØ, Saif YM, et al. Comparison of capsid sequences from human and animal astroviruses. J Gen Virol 2001;82(Pt 5):1061–1067. 98. Jonassen CM, Jonassen TØ, Sveen TM, et al. Complete genomic sequences of astroviruses from sheep and turkey: comparison with related viruses. Virus Res 2003;91(2):195–201. 99. Kadaré G, Haenni AL. Virus-encoded RNA helicases. J Virol 1997;71(4):2583–2590. 100. Kapoor A, Li L, Victoria J, et al. Multiple novel astrovirus species in human stool. J Gen Virol 2009;90(Pt 12):2965–2972. 101. Khamrin P, Thongprachum A, Okitsu S, et al. Multiple astrovirus MLB1, MLB2, VA2 clades, and classic human astrovirus in children with acute gastroenteritis in Japan. J Med Virol 2016;88(2):356–360. 102. Kiang D, Matsui SM. Proteolytic processing of a human astrovirus nonstructural protein. J Gen Virol 2002;83(Pt 1):25–34. 103. Koci MD, Kelley LA, Larsen D, et al. Astrovirus-induced synthesis of nitric oxide contributes to virus control during infection. J Virol 2004;78(3):1564–1574. 104. Koci MD, Moser LA, Kelley LA, et al. Astrovirus induces diarrhea in the absence of inflammation and cell death. J Virol 2003;77(21):11798–11808. 105. Koci MD, Seal BS, Schultz-Cherry S. Molecular characterization of an avian astrovirus. J Virol 2000;74(13):6173–6177. 106. Kolawole AO, Mirabelli C, Hill DR, et al. Astrovirus replication in human intestinal enteroids reveals multi-cellular tropism and an intricate host innate immune landscape. PLoS Pathog 2019;15(10):e1008057. 107. Konno T, Suzuki H, Ishida N, et al. Astrovirus-associated epidemic gastroenteritis in Japan. J Med Virol 1982;9(1):11–17. 108. Koopmans MP, Bijen MH, Monroe SS, et al. Age-stratified seroprevalence of neutralizing antibodies to astrovirus types 1 to 7 in humans in The Netherlands. Clin Diagn Lab Immunol 1998;5(1):33–37. 109. Koukou G, Niendorf S, Hornei B, et al. Human astrovirus infection associated with encephalitis in an immunocompetent child: a case report. J Med Case Rep 2019;13(1):341. 110. Krishna NK. Identification of structural domains involved in astrovirus capsid biology. Viral Immunol 2005;18(1):17–26. 111. Kriston S, Willcocks MM, Carter MJ, et al. Seroprevalence of astrovirus types 1 and 6 in London, determined using recombinant virus antigen. Epidemiol Infect 1996;117(1):159–164. 112. Küchler L, Rüfli I, Koch MC, et al. Astrovirus-associated polioencephalomyelitis in an alpaca. Viruses 2020;13(1). 113. Kurtz J, Lee T. Astrovirus gastroenteritis age distribution of antibody. Med Microbiol Immunol 1978;166(1–4):227–230. 114. Kurtz JB, Lee TW. Human astrovirus serotypes. Lancet 1984;2(8416):1405. 115. Kurtz JB, Lee TW, Craig JW, et al. Astrovirus infection in volunteers. J Med Virol 1979;3(3):221–230. 116. Kurtz JB, Lee TW, Parsons AJ. The action of alcohols on rotavirus, astrovirus and enterovirus. J Hosp Infect 1980;1(4):321–325. 117. Lau P, Cordey S, Brito F, et al. Metagenomics analysis of red blood cell and fresh-frozen plasma units. Transfusion 2017;57(7):1787–1800. 118. Lee TW, Kurtz JB. Serial propagation of astrovirus in tissue culture with the aid of trypsin. J Gen Virol 1981;57(Pt 2):421–424. 119. Lee RM, Lessler J, Lee RA, et al. Incubation periods of viral gastroenteritis: a systematic review. BMC Infect Dis 2013;13:446. 120. Lewis DC, Lightfoot NF, Cubitt WD, et al. Outbreaks of astrovirus type 1 and rotavirus gastroenteritis in a geriatric in-patient population. J Hosp Infect 1989;14(1):9–14. 121. Li L, Diab S, McGraw S, et al. Divergent astrovirus associated with neurologic disease in cattle. Emerg Infect Dis 2013;19(9):1385–1392. 122. Li Y, Gordon E, Idle A, et al. Astrovirus outbreak in an animal shelter associated with feline vomiting. Front Vet Sci 2021;8:628082. 123. Liu N, Jiang M, Wang M, et al. Isolation and detection of duck astrovirus CPH: implications for epidemiology and pathogenicity. Avian Pathol 2016;45(2):221–227. 124. Lulla V, Firth AE. A hidden gene in astroviruses encodes a viroporin. Nat Commun 2020;11(1):4070. 125. Lum SH, Turner A, Guiver M, et al. An emerging opportunistic infection: fatal astrovirus (VA1/HMO-C) encephalitis in a pediatric stem cell transplant recipient. Transpl Infect Dis 2016;18(6):960–964. 126. Luo Z, Roi S, Dastor M, et al. Multiple novel and prevalent astroviruses in pigs. Vet Microbiol 2011;149(3–4):316–323. 127. Madeley CR, Cosgrove BP. Letter: 28 nm particles in faeces in infantile gastroenteritis. Lancet 1975;2(7932):451–452. 128. Madeley CR, Cosgrove BP. Letter: viruses in infantile gastroenteritis. Lancet 1975;2(7925):124. 129. Maldonado Y, Cantwell M, Old M, et al. Population-based prevalence of symptomatic and asymptomatic astrovirus infection in rural Mayan infants. J Infect Dis 1998;178(2):334–339. 130. Marczinke B, Bloys AJ, Brown TD, et al. The human astrovirus RNA-dependent RNA polymerase coding region is expressed by ribosomal frameshifting. J Virol 1994;68(9):5588–5595. 131. Martella V, Catella C, Capozza P, et al. Identification of astroviruses in bovine and buffalo calves with enteritis. Res Vet Sci 2020;131:59–68. 132. Martella V, Moschidou P, Pinto P, et al. Astroviruses in rabbits. Emerg Infect Dis 2011;17(12):2287–2293. 133. Marvin SA, Huerta CT, Sharp B, et al. Type I interferon response limits astrovirus replication and protects against increased barrier permeability in vitro and in vivo. J Virol 2016;90(4):1988–1996. 78

134. Matias Ferreyra FS, Bradner LK, Burrough ER, et al. Polioencephalomyelitis in domestic swine associated with porcine astrovirus type 3. Vet Pathol 2020;57(1):82–89. 135. Matsui SM, Kiang D, Ginzton N, et al. Molecular biology of astroviruses: selected highlights. Novartis Found Symp 2001;238:219–233; discussion 233–236. 136. Meliopoulos VA, Kayali G, Burnham A, et al. Detection of antibodies against Turkey astrovirus in humans. PLoS One 2014;9(5):e96934. 137. Meliopoulos VA, Marvin SA, Freiden P, et al. Oral administration of astrovirus capsid protein is sufficient to induce acute diarrhea in vivo. MBio 2016;7(6). 138. Méndez E, Aguirre-Crespo G, Zavala G, et al. Association of the astrovirus structural protein VP90 with membranes plays a role in virus morphogenesis. J Virol 2007;81(19):10649–10658. 139. Méndez E, Fernández-Luna T, López S, et al. Proteolytic processing of a serotype 8 human astrovirus ORF2 polyprotein. J Virol 2002;76(16):7996–8002. 140. Méndez E, Muñoz-Yañez C, Martín CS-S, et al. Characterization of human astrovirus cell entry. J Virol 2014;88(5):2452–2460. 141. Méndez E, Salas-Ocampo E, Arias CF. Caspases mediate processing of the capsid precursor and cell release of human astroviruses. J Virol 2004;78(16):8601–8608. 142. Méndez E, Salas-Ocampo MPE, Munguía ME, et al. Protein products of the open reading frames encoding nonstructural proteins of human astrovirus serotype 8. J Virol 2003;77(21):11378–11384. 143. Meyer CT, Bauer IK, Antonio M, et al. Prevalence of classic, MLB-clade and VA-clade astroviruses in Kenya and the Gambia. Virol J 2015;12:78. 144. Midthun K, Greenberg HB, Kurtz JB, et al. Characterization and seroepidemiology of a type 5 astrovirus associated with an outbreak of gastroenteritis in Marin County, California. J Clin Microbiol 1993;31(4):955–962. 145. Mitchell DK, Matson DO, Jiang X, et al. Molecular epidemiology of childhood astrovirus infection in child care centers. J Infect Dis 1999;180(2):514–517. 146. Mitchell DK, Van R, Morrow AL, et al. Outbreaks of astrovirus gastroenteritis in day care centers. J Pediatr 1993;123(5):725–732. 147. Mittelholzer C, Hedlund K-O, Englund L, et al. Molecular characterization of a novel astrovirus associated with disease in mink. J Gen Virol 2003;84(Pt 11):3087–3094. 148. Molberg O, Nilsen EM, Sollid LM, et al. CD4+ T cells with specific reactivity against astrovirus isolated from normal human small intestine. Gastroenterology 1998;114(1):115–122. 149. Monceyron C, Grinde B, Jonassen TO. Molecular characterisation of the 3′-end of the astrovirus genome. Arch Virol 1997;142(4):699–706. 150. Monroe SS, Jiang B, Stine SE, et al. Subgenomic RNA sequence of human astrovirus supports classification of Astroviridae as a new family of RNA viruses. J Virol 1993;67(6):3611–3614. 151. Monroe SS, Stine SE, Gorelkin L, et al. Temporal synthesis of proteins and RNAs during human astrovirus infection of cultured cells. J Virol 1991;65(2):641–648. 152. Moser LA, Carter M, Schultz-Cherry S. Astrovirus increases epithelial barrier permeability independently of viral replication. J Virol 2007;81(21):11937–11945. 153. Moser LA, Schultz-Cherry S. Suppression of astrovirus replication by an ERK1/2 inhibitor. J Virol 2008;82(15):7475–7482. 154. Murillo A, Vera-Estrella R, Barkla BJ, et al. Identification of host cell factors associated with astrovirus replication in caco-2 cells. J Virol 2015;89(20):10359–10370. 155. Naccache SN, Peggs KS, Mattes FM, et al. Diagnosis of neuroinvasive astrovirus infection in an immunocompromised adult with encephalitis by unbiased next-generation sequencing. Clin Infect Dis 2015;60(6):919–923. 156. Naficy AB, Rao MR, Holmes JL, et al. Astrovirus diarrhea in Egyptian children. J Infect Dis 2000;182(3):685–690. 157. Ng TFF, Kondov NO, Hayashimoto N, et al. Identification of an astrovirus commonly infecting laboratory mice in the US and Japan. PLoS One 2013;8(6):e66937. 158. Nighot PK, Moeser A, Ali RA, et al. Astrovirus infection induces sodium malabsorption and redistributes sodium hydrogen exchanger expression. Virology 2010;401(2):146–154. 159. Oishi I, Yamazaki K, Kimoto T, et al. A large outbreak of acute gastroenteritis associated with astrovirus among students and teachers in Osaka, Japan. J Infect Dis 1994;170(2):439–443. 160. Olortegui MP, Rouhani S, Yori PP, et al. Astrovirus infection and diarrhea in 8 countries. Pediatrics 2018;141(1). 161. Pennap G, Pager CT, Peenze I, et al. Epidemiology of astrovirus infection in Zaria, Nigeria. J Trop Pediatr 2002;48(2):98–101. 162. Pérot P, Lecuit M, Eloit M. Astrovirus diagnostics. Viruses 2017;9(1). 163. Pfaff F, Schlottau K, Scholes S, et al. A novel astrovirus associated with encephalitis and ganglionitis in domestic sheep. Transbound Emerg Dis 2017;64(3):677–682. 164. Pinto RM, Abad FX, Gajardo R, et al. Detection of infectious astroviruses in water. Appl Environ Microbiol 1996;62(5):1811–1813. 165. Quan PL, Wagner TA, Briese T, et al. Astrovirus encephalitis in boy with X-linked agammaglobulinemia. Emerg Infect Dis 2010;16(6):918–925. 166. Reuter G, Pankovics P, Delwart E, et al. Identification of a novel astrovirus in domestic sheep in Hungary. Arch Virol 2012;157(2):323–327. 167. Reynolds DL, Saif YM, Theil KW. A survey of enteric viruses of turkey poults. Avian Dis 1987;31(1):89–98. 168. Risco C, Carrascosa JL, Pedregosa AM, et al. Ultrastructure of human astrovirus serotype 2. J Gen Virol 1995;76(Pt 8):2075–2080. 169. Rogawski McQuade ET, Liu J, Kang G, et al. Protection from natural immunity against enteric infections and etiology-specific diarrhea in a longitudinal birth cohort. J Infect Dis 2020;222(11):1858–1868. 170. Ruuska T, Vesikari T. Rotavirus disease in Finnish children: use of numerical scores for clinical severity of diarrhoeal episodes. Scand J Infect Dis 1990;22(3):259–267. 171. Sanchez-Fauquier A, Carrascosa AL, Carrascosa JL, et al. Characterization of a human astrovirus serotype 2 structural protein (VP26) that contains an epitope involved in virus neutralization. Virology 1994;201(2):312–320. 172. Sato M, Kuroda M, Kasai M, et al. Acute encephalopathy in an immunocompromised boy with astrovirus-MLB1 infection detected by next generation sequencing. J Clin Virol 2016;78:66–70. 173. Schlottau K, Schulze C, Bilk S, et al. Detection of a novel bovine astrovirus in a cow with encephalitis. Transbound Emerg Dis 2016;63(3):253–259. 174. Schultz-Cherry S, King DJ, Koci MD. Inactivation of an astrovirus associated with poult enteritis mortality syndrome. Avian Dis 2001;45(1):76–82. 175. Sebire NJ, Malone M, Shah N, et al. Pathology of astrovirus associated diarrhoea in a paediatric bone marrow transplant recipient. J Clin Pathol 2004;57(9):1001–1003. 176. Selimovic-Hamza S, Boujon CL, Hilbe M, et al. Frequency and pathological phenotype of bovine astrovirus CH13/NeuroS1 infection in neurologically-diseased cattle: towards assessment of causality. Viruses 2017;9(1). 177. Sellers H, Linneman E, Icard AH, et al. A purified recombinant baculovirus expressed capsid protein of a new astrovirus provides partial 79

protection to runting-stunting syndrome in chickens. Vaccine 2010;28(5):1253–1263. 178. Seuberlich T, Wüthrich D, Selimovic-Hamza S, et al. Identification of a second encephalitis-associated astrovirus in cattle. Emerg Microbes Infect 2016;5:e71. 179. Shimizu M, Shirai J, Narita M, et al. Cytopathic astrovirus isolated from porcine acute gastroenteritis in an established cell line derived from porcine embryonic kidney. J Clin Microbiol 1990;28(2):201–206. 180. Singh PB, Sreenivasan MA, Pavri KM. Viruses in acute gastroenteritis in children in Pune, India. Epidemiol Infect 1989;102(2):345–353. 181. Siqueira JAM, de Souza Oliveira D, de Carvalho TCN, et al. Astrovirus infection in hospitalized children: molecular, clinical and epidemiological features. J Clin Virol 2017;94:79–85. 182. Smits SL, van Leeuwen M, Kuiken T, et al. Identification and characterization of deer astroviruses. J Gen Virol 2010;91(Pt 11):2719–2722. 183. Smits SL, van Leeuwen M, van der Eijk AA, et al. Human astrovirus infection in a patient with new-onset celiac disease. J Clin Microbiol 2010;48(9):3416–3418. 184. Snodgrass DR, Angus KW, Gray EW, et al. Pathogenesis of diarrhoea caused by astrovirus infections in lambs. Arch Virol 1979;60(3–4):217–226. 185. Speroni S, Rohayem J, Nenci S, et al. Structural and biochemical analysis of human pathogenic astrovirus serine protease at 2.0 A resolution. J Mol Biol 2009;387(5):1137–1152. 186. Strain E, Kelley LA, Schultz-Cherry S, et al. Genomic analysis of closely related astroviruses. J Virol 2008;82(10):5099–5103. 187. Taboada B, Espinoza MA, Isa P, et al. Is there still room for novel viral pathogens in pediatric respiratory tract infections? PLoS One 2014;9(11):e113570. 188. Tang Y, Murgia MV, Ward L, et al. Pathogenicity of turkey astroviruses in turkey embryos and poults. Avian Dis 2006;50(4):526–531. 189. Tang Y, Saif YM. Antigenicity of two turkey astrovirus isolates. Avian Dis 2004;48(4):896–901. 190. Tange S, Zhou Y, Nagakui-Noguchi Y, et al. Initiation of human astrovirus type 1 infection was blocked by inhibitors of phosphoinositide 3-kinase. Virol J 2013;10:153. 191. Thouvenelle ML, Haynes JS, Reynolds DL. Astrovirus infection in hatchling turkeys: histologic, morphometric, and ultrastructural findings. Avian Dis 1995;39(2):328–336. 192. Todd D, Wilkinson DS, Jewhurst HL, et al. A seroprevalence investigation of chicken astrovirus infections. Avian Pathol 2009;38(4):301–309. 193. Toffan A, Jonassen CM, De Battisti C, et al. Genetic characterization of a new astrovirus detected in dogs suffering from diarrhoea. Vet Microbiol 2009;139(1–2):147–152. 194. Toh Y, Harper J, Dryden KA, et al. Crystal Structure of the human astrovirus capsid protein. J Virol 2016;90(20):9008–9017. 195. Tse H, Chan W-M, Tsoi H-W, et al. Rediscovery and genomic characterization of bovine astroviruses. J Gen Virol 2011;92(Pt 8):1888–1898. 196. van der Doef HPJ, Bathoorn E, van der Linden MPM, et al. Astrovirus outbreak at a pediatric hematology and hematopoietic stem cell transplant unit despite strict hygiene rules. Bone Marrow Transplant 2016;51(5):747–750. 197. Vu D-L, Bosch A, Pintó RM, et al. Epidemiology of classic and novel human astrovirus: gastroenteritis and beyond. Viruses 2017;9(2):33. 198. Vu D-L, Bosch A, Pintó RM, et al. Human astrovirus MLB replication in vitro: persistence in extraintestinal cell lines. J Virol 2019;93(13). 199. Vu D-L, Sabrià A, Aregall N, et al. Novel human astroviruses: prevalence and association with common enteric viruses in undiagnosed gastroenteritis cases in Spain. Viruses 2019;11(7). 200. Vu D-L, Sabrià A, Aregall N, et al. A Spanish case–control study in 90%).

Extent of Sequence Variation There is detectable within-host sequence variation, resulting partly from immune-mediated selection.271 However, because HTLV-1 persists during chronic infection mainly through sustained replication of long-lived infected T-cell clones,200,394 RT contributes little to viral replication, and the extent of sequence variation, both within one host and at the population level, is consequently very small compared with HIV-1. In ATL, mutations that silence the Tax protein are observed in approximately 50% of cases, and type 2 proviruses are found in approximately 30%.172

Significance of Sequence Variation No single sequence is specifically associated with the manifestations of HTLV-1 infection: an identical sequence may result in either asymptomatic carriage, inflammatory disease, or ATL,55,189,285 although there is a small difference in the risk of HAM associated with two different HTLV-1 genotypes in southern Japan.80 Mutations that alter T-cell epitopes271 or that result in the loss of Tax expression or loss of the 5′ LTR may confer a selective advantage on the infected T cell clone and can contribute to oncogenesis in ATL.2,172,246,334

Sequence variation in HTLV-1 has been of great importance in understanding the epidemiology of the virus2,334; see “Epidemiology.”

HTLV-1 Provirus in Nonmalignant Cells and ATL Cells After infection, the HTLV-1 provirus exists in the cellular genome and causes clonal proliferation and replication of virus. Since the HTLV-1 provirus is the direct evidence of HTLV-1 infection in the cells, analyses of HTLV-1 proviruses provide important information on the pathogenesis of HTLV-1. HTLV-1 proviruses in ATL cases are divided into three subtypes: (a) complete, (b) type 1 defective, and (c) type 2 defective.358 Type 1 defective provirus retains the 5′ and 3′ LTRs but lacks internal sequences including gag, pol, and env. Type 2 defective provirus lacks the 5′ LTR and part of the internal region of the provirus. Importantly, approximately half of type 2 defective proviruses have a six-bp repeat at both ends. This short repeat is generated during integration of the viral DNA into the genome: its presence in such cases, therefore, indicates that this defective provirus is generated before proviral integration.246 This type of defective provirus is also present in HTLV-1 carriers.172,246 Since the 5′ LTR is the promoter/enhancer for plus-strand transcription of the provirus, this type of provirus lacks expression of viral genes encoded by the plus strand, including the tax gene.

APOBEC3G (A3G) is antiretroviral factor, which changes cytosine to uracil of single-stranded DNA by deamination during reverse transcription. A3G causes G-to-A mutations in the plus strand of the provirus, resulting in generation of nonsense mutations in the provirus. HTLV-1 proviruses in ATL cases frequently contain nonsense mutations, which correspond to target sequences of A3G.70 A hot-spot of nonsense mutation is observed in the tax gene. Approximately 10% of ATL cases have a nonsense mutation in the tax gene.79,356 Furthermore, this nonsense mutation of the tax gene is also observed in some HTLV-1–infected cells in asymptomatic carriers.70 These findings indicate that HTLV-1–infected cells containing the tax gene with nonsense mutation transform to ATL cells.

DNA methylation of the HTLV-1 provirus silences transcription of viral genes. In particular, DNA methylation is detected in the 5′ LTR, whereas the 3′ LTR is not methylated in ATL cells.188 DNA methylation first occurs in the gag, pol, and env regions and then extends in the 5′ and 3′ directions in vivo, and when the 5′ LTR becomes methylated, viral transcription is silenced.360 Heavy DNA methylation of the 5′ LTR silences transcription of plus-strand genes. In contrast, the 3′ LTR is not methylated, which is constant with continuous expression of the HBZ gene.

Tax expression is disrupted approximately in half of ATL cases by any of three mechanisms: (a) deletion of 5′ LTR, (b) nonsense mutation of the tax gene, and (c) DNA methylation of 5′ LTR.356 Importantly, about 25% of ATL cases do not express Tax at the stage of infection, due to nonsense mutation or deletion of 5′ LTR, which indicates that Tax is not essential for HTLV-1–induced leukemogenesis.274 The tax gene transcription is barely detected in the remaining half of ATL cases, although the 5′ LTR is not heavily methylated and the tax gene remains intact.221 It is speculated that Tax is intermittently expressed in these cases, which may, therefore, depend on both Tax and HBZ. In such cases, Tax may contribute to leukemogenesis by impairing the DNA damage response pathway and the repair of double-strand DNA breaks, resulting in genomic instability.224,425

Evolution of HTLV-1 Like the other human retroviruses, HTLV-1 had its origins in simian retroviruses in nonhuman primates190,387 and was probably introduced into humans by bushmeat hunting and animal bites.74,175 At least 4 interspecies transmission events are thought to have occurred,2 although the host nonhuman primate species has not always been clearly identified. HTLV-2 is similarly derived from closely related simian T-cell leukemia viruses (STLVs) in West Africa.388

HTLV-1 is thought to have been endemic in humans for tens of thousands of years.2,334,387 The most divergent sequences of HTLV-1 are found in Papua New Guinea and Melanesia91 and Australia41: since there are no native nonhuman primates in Australia, it is assumed that the virus was introduced by the humans who arrived on the continent between 10,000 and 60,000 years ago. The rate of evolution of HTLV-1 is significantly slower than that of other RNA viruses such as influenza A virus.137,347 The rate of evolution is constrained by the mode of withinhost persistence by clonal proliferation and by the presence of overlapping reading frames in the provirus; it is also likely that there is strong selection during transmission between individuals, that is, a transmission bottleneck.

603

A key target of the host immune response to HTLV-1 is HBZ protein (see “Cell-mediated immune response”). HBZ is expressed by a nonmalignant infected T cell approximately 50% of the time,243 and an effective host CTL response to HBZ is associated with a lower PVL and a lower risk of HAM. Consequently, HTLV-1 has evolved to minimize its exposure to CTL surveillance by restricting HBZ expression to a minimal level—typically 1 to 5 mRNA copies per HBZ-expressing cell,32 and by limiting the export of HBZ mRNA to the cytoplasm.299 As a result, HBZ protein is expressed at a very low level. The low affinity of HBZ peptides for binding to HLA class 1 proteins215 also limits the effectiveness of the CTL response.

Tax and Rex: Main Physiological Actions Most T-cell lines immortalized in vitro by HTLV-1 express high levels of Tax.216 However, the tax transcript is barely detected in vivo,356 indicating that T-cell lines with higher Tax expression do not reflect the physiological pattern of Tax expression. By contrast, several ATL cell lines derived from ATL patients (e.g., MT-1, KK-1), and HTLV-1–infected T-cell lines established from HTLV-1 carriers and HAM/TSP patients, express Tax sparsely.221 Furthermore, some ATL cell lines derived from primary ATL cells do not express Tax.356 Single-cell analyses showed that, at any given time, only a small number of T cells express tax and other viral genes encoded on the proviral plus strand,32,221 which likely corresponds with the in vivo expression pattern of the tax gene.

Tax Tax is a protein of 353 amino acids, localized in both the nucleus and cytoplasm.97 The main functions of Tax are (a) trans-activation of plusstrand transcription of viral genes and (b) modulation of cell signaling pathways through its interaction with host factors.

Tax activates plus-strand transcription of the HTLV-1 provirus Like the majority of mammalian genes, the HTLV-1 proviral plus strand is not continuously expressed but rather is expressed in intermittent, self-limiting bursts.32,221 The plus-strand bursts are infrequent, in contrast with the minus-strand (HBZ) bursts: the great majority of HTLV-1– infected cells freshly isolated from peripheral blood are tax-negative at a given instant, whereas approximately 50% express HBZ at a low level.243 Because of the positive feedback exerted by the Tax protein, the plus-strand burst is intense, producing several hundred transcripts within approximately 3 hours.243 The burst is regulated like a cellular immediate-early stress-response gene (such as c-fos): it is triggered by cellular stress, depends on p38-MAPK, and is not inhibited by protein synthesis inhibitors.194 The promoter/enhancer in the 5′ LTR is maintained in a poised state by ubiquitylation of histone H2A at lysine 119 by polycomb repressive complex 1 (PRC1); the onset of the plusstrand burst is accompanied by loss of this inhibitory epigenetic mark.194

The duration of the plus-strand burst in fresh PBMCs is approximately 6 to 10 hours243; it is likely to be similar in vivo, but direct evidence is lacking. A major factor responsible for terminating the burst is the action of Rex protein on mRNA splicing and transport; it is not known whether additional factors contribute to the active termination of the burst.

Molecular mechanism of Tax-mediated transcriptional activation Tax activates HTLV-1 transcription through the long terminal repeat (LTR)71,278,322,323,335 (Fig. 16.2). The HTLV-1 LTR is divided into three regions: U3, R, and U5. Three repetitive 21-bp sequences in the U3 region act as Tax-responsive enhancers (TREs).326 Each 21-bp TRE is in turn composed of three elements: the core element and two elements that lie, respectively, 5′ and 3′ to the core element, all three elements are indispensable for efficient Tax activation of HTLV-1 transcription. The core sequence of a TRE is highly similar to a cyclic AMP response element (CRE) and indeed acts as a binding site for the CRE-binding protein/activating transcription factor (ATF) family of transcription factors (CREB/ATFs).29,415,421,422 Thus, the TRE is also called a viral cAMP response element (vCRE) and is responsive to cAMP. The 5′ and 3′ elements within a TRE are GC-rich. While Tax itself does not interact directly with TRE DNA, it can do so in concert with DNA-bound CREB protein.421,422 When this happens, DNA-bound Tax interacts with the minor groove of the adjacent GC-rich sequences to stabilize the protein– DNA complex.205,206

CREB is ubiquitously expressed and regulates a number of cellular genes, especially cAMP-induced genes.396 A cAMP-initiated signal phosphorylates CREB at serine 133, recruiting two transcriptional coactivators, CREB-binding protein (CBP) and p300. An in vitro chromatinbased transcription study indicated that phosphorylation of CREB is essential for Tax activity, through stabilizing the Tax/CREB/CBP/DNA complex. Tax by itself induces the phosphorylation of CREB.180 CBP/p300 is a histone acetyltransferase that acetylates histone tails, thereby changing chromatin structures to activate transcription. In vitro and in vivo studies showed that p300 is crucial for Tax-dependent transcriptional activation, where p300, interacting with Tax and CREB, induces the acetylation of nucleosome histones over the TRE DNA.86,87,95,116,196,203,206 Interestingly, p300 recruitment by Tax to an integrated HTLV-1 promoter reduces the amount of histone H1 and H3 proteins on the promoter DNA and stimulates the recruitment of RNA polymerase II.204 These results suggest that recruitment of CBP/p300 to the integrated promoter acetylates histone tails, thereby removing histone octamers from the HTLV-1 promoter.

TORCs and P-TEFb (CDK-9/cyclin T1) are additional positive regulators of Tax-dependent HTLV-1 promoter activation.46,47,153,187,333,427 Both proteins interact with Tax, and the knockdown of either by siRNA reduces Tax-dependent activation. On the other hand, Bcl-3, by interacting with TORC3, acts as a negative transcriptional regulator of the HTLV-1 LTR.129

Tax-Mediated Cellular Signaling Pathways Tax not only activates plus-strand transcription of viral genes but also possesses pleiotropic effects on signaling pathways of host cells (Fig. 16.3). Tax is intermittently expressed in approximately half of ATL cases; it is probably also expressed—again intermittently—in most HTLV-1–infected cells in both asymptomatic carriers and patients with HAM. Sporadic Tax expression enhances transcription of NF-κB– associated genes and antiapoptotic genes221 but suppresses the cell cycle transition from S to G2/M.

Activation of NF-κB by Tax

604

Tax activates the transcription factor NF-κB, thereby inducing the expression of a number of cellular genes, and this activity is crucial for many aspects of the HTLV-1 life cycle.16,207,233,303,344 Several NF-κB inhibitors induce apoptosis in HTLV-1–infected T cells, indicating that NF-κB is crucial for survival of HTLV-1–infected T cells. NF-κB is a family of transcription factors including NF-κB1 (p50), p65, c-Rel, NF-κB2 (p52), RelB, and Bcl-3, and these factors are divided into two groups belonging to the canonical (NF-κB1/p50, p65, c-Rel) and the noncanonical (NFκB2/p52, RelB, Bcl-3) pathways. Both NF-κB pathways are activated by Tax in T cells.400

To activate the canonical pathway, Tax interacts with several NF-κB regulators. IKKγ is essential for Tax activation of NF-κB, since the mutation of IKKγ in fibroblasts or T cells totally abrogates Tax activation of NF-κB.48,85,114,406 IKKγ is a scaffold component of the IκB kinase (IKK) complex, IKKα/IKKβ/IKKγ. Through interacting with IKKγ, Tax activates IKKβ to induce phosphorylation and degradation of IκBs (IκBα, IκBβ), thereby allowing nuclear translocation of p50/p65.219,345

Tax also activates the noncanonical NF-κB pathway.400 Tax simultaneously binds to the IKK complex (IKKγ, IKKα) and NF-κB2/p100, but not IKKβ, and thus induces IKKα-mediated phosphorylation of p100, its processing into p52, and the subsequent translocation of p52/RelB into the nucleus. Knockdown of NF-κB2/p100 by short hairpin RNA reduces Tax-induced IL-2–independent growth induction in CTLL-2 cells.121 Thus, the noncanonical NF-κB pathway is also crucial for the transforming activity of Tax.

FIGURE 16.3 Functional domains of Tax. Regions of Tax required for activation of the cyclic adenosine monophosphate response element binding protein/activating transcription factor (CREB/ATF) pathway, activation of the NF-κB pathway, and transcriptional activation function of Tax measured by a Tax fusion protein with the yeast DNA-binding protein GAL4 are indicated. Positions of substitution mutations in the Tax mutants M22 and M47 are also indicated. M22 is active in the CREB pathway but inactive in the NF-κB pathway, whereas M47 is the reverse. LZ-2 and PDZ motifs distinguish Tax1 from Tax2. NLS, nuclear localization signal; NES, cryptic nuclear export signal; Zn, zinc finger motif; LZ-1 and LZ-2, leucine zipper-like structures. In half of ATL cases, tax is either silenced or only transiently transcribed, although the NF-κB pathway is activated in all ATL cases. This suggests that NF-κB is activated by other mechanisms. Overexpression of NF-κB–inducing kinase (NIK) is associated with constitutive activation of NF-κB.306 Furthermore, microRNA-31 (miR-31) negatively regulates the noncanonical NF-κB pathway by targeting NIK. Therefore, the suppression of miR-31 that is observed in ATL cells leads to activation of the NF-κB pathway.403

Apoptosis inhibition by Tax HTLV-1–infected T cells are resistant to various types of apoptosis, and this resistance is mostly mediated by Tax.173,258 Tax inhibits apoptosis of T cells induced by IL-2 withdrawal in an IL-2–dependent T-cell line (CTLL-2). Studies of Tax mutants (M47, M22) indicate that inhibition of apoptosis is mediated through the NF-κB pathway.146 Tax activates the expression of several antiapoptotic genes in T cells, including CTLL-2 cells, through the NF-κB pathway. These genes include Bcl-xL, xIAP, cIAP, and cFLIP.375

Activation of the PI3K/AKT pathway by Tax IL-2 activates PI3K and its downstream kinase, Akt, in normal T cells—events which are essential for apoptosis inhibition and cell growth. However, even in the absence of IL-2, HTLV-1–transformed T-cell lines show constitutive activation of the PI3K/Akt pathway. Moreover, several inhibitors of this pathway, such as LY294002 (an inhibitor for PI3K), induce growth arrest of HTLV-1–transformed T-cell lines and then induce apoptosis, suggesting that continuous activation of this pathway is essential for maintaining survival of HTLV-1–infected T cells.136,152 To do this, Tax directly interacts with PI3K. PI3K consists of two subunits: the catalytic p110α subunit and the inhibitory p85α subunit. Tax directly binds to the p85α inhibitory subunit, causing the release of the active p110α catalytic subunit.291 Activation of the PI3K/Akt pathway by Tax results in activation of the downstream target mammalian target of rapamycin (mTOR) in HTLV-1–infected T cells as well as Taxexpressing cells.416

Physiological Functions of Tax T-cell lines transformed in vitro by HTLV-1 express Tax at a high level, which does not reflect the physiological expression pattern of Tax. Tax expression in vivo is suppressed at a low level. It is speculated that Tax is intermittently expressed, so that at a given instant Tax is present in only a small number of infected cells and ATL cells.32,221 Intermittent Tax expression (Tax burst) promotes transcription of the plus-strand of the provirus, which leads to de novo infection. Simultaneously, the Tax burst generates T cells with higher expression of antiapoptotic genes, which is important to maintain the whole cell population.221 Since Tax is a highly immunogenic viral protein, restriction of Tax expression to an intermittent pattern is beneficial for survival of nonmalignant HTLV-1–infected cells and ATL cells in vivo.

Rex HTLV-1 has two posttranscriptional regulators, Rex and p30.169,412 The role of p30 in posttranscriptional mRNA regulation is described below.

In addition to unspliced genomic mRNA encoding gag/pro-pol, HTLV-1 produces singly spliced env mRNA and doubly spliced tax/rex and p30 605

mRNAs. RNAs with introns generally undergo splicing by the cellular RNA machinery; otherwise they are degraded. Thus, upon initial infection of host cells, HTLV-1 dominantly expresses doubly spliced tax/rex and p30 mRNA. Once the Rex protein is produced, Rex controls the ratio of spliced forms of HTLV-1 mRNAs119,139 (Fig. 16.2). Rex increases the amount of singly spliced (env) and unspliced (gag/pro-pol) mRNAs and reduces the amount of its own doubly spliced mRNA. Rex does this by inhibiting the splicing of singly spliced (env) and unspliced (gag/pro-pol) mRNAs, stabilizing them, and promoting their transport to the cytoplasm. In the absence of Rex, unspliced HTLV-1 mRNAs are retained in the nucleus due to two cis-acting repressive sequences (CRS) in the 5′ and 3′ LTRs,182,321 but Rex overcomes the inhibitory activity of the CRSs and induces the translocation of the unspliced RNAs into the cytoplasm.

These Rex functions are achieved through at least three activities. The first is sequence-specific RNA binding. Rex interacts specifically with the HTLV-1 RNA, at the Rex-responsive element (RxRE) located in the U3 and R regions of the 3′ LTR.3,38,379 RNA binding by Rex is accomplished by an arginine-rich highly basic region in Rex (aa1-19).109 This domain also contains a nuclear localization signal (NLS) and a nucleolus targeting signal, which allow Rex to enter the nucleolus where it can interact with the RNA.276,332 The RxRE RNA forms a long stem loop structure, and one stem loop (called stem loop D) is essential for Rex binding and function.103 The second Rex function (multimerization) is mediated through aa32-133.395 Rex mutants that cannot form multimers behave as dominant-negative proteins.37,395 The third Rex function is interaction with the nuclear export receptor, CRM1/exportin 1, which mediates the transport of viral mRNAs from the nucleus to the cytoplasm. Interaction with CRM1 is also required for multimerization of Rex, since a Rex mutant defective for CRM1 interaction failed to form a multimer. Rex contains a typical leucine-rich nuclear export signal (NES) (aa81-94) that mediates its interaction with CRM1. HTLV-1 production is less efficient in rat cells than human cells, because Rex is less active in rat cells than human cells. The rat CRM1 is impaired in promoting Rex to form multimers, which may explain the reduced activity of Rex in rat cells.106,420

Rex is activated by phosphorylation.177 While PMA (phorbol myristate acetate), an activator of PKC (protein kinase C), transiently enhances phosphorylation of Rex, treatment of HTLV-1–infected cell lines with a PKC inhibitor reduces the amount of unspliced gag-pol mRNA.1 Liquid chromatography tandem mass spectrometry analysis showed that Rex is phosphorylated at multiple sites, and mutation analysis indicated that phosphorylations at Ser-97 and Thr-174 are critical for its function.177 Rex also suppresses nonsense-mediated mRNA decay, which ensures the stability of viral genomic RNA.265

Tax Expression and Its Suppression by HBZ and Rex High expression of Tax induces senescence and suppresses the S phase of the cell cycle.195,221 Thus, Tax overexpression is harmful to infected cells. Therefore, HTLV-1 has redundant mechanisms to alleviate detrimental effects of Tax and reduce Tax expression. Rex inhibits splicing of the tax gene and promotes generation of unspliced RNA. Furthermore, HBZ suppresses Tax expression through inhibition of the binding of CREB-2 to TREs.83 In addition, HBZ RNA impairs the interaction of RNA polymerase II with LTR by displacing TATA-box binding protein (TBP), resulting in suppression of the basal transcription machinery.84 Since HBZ is transcribed from a TATA-less promoter, this suppressive effect of HBZ RNA does not influence HBZ transcription. Hyperactivation of NF-κB by Tax causes up-regulation of cyclin-dependent kinase inhibitors, p21 and p27, which leads to cellular senescence.426 HBZ protein hinders the onset of Tax-induced senescence through inhibition of the canonical NF-κB pathway.424 Thus, when Tax expression is predominant during a plus-strand burst, Rex progressively suppresses Tax expression via altered splicing. Then, HBZ (RNA and protein) maintains latency by suppression of sense-transcription.293

HBZ: Physiological Actions Transcription of the HBZ Gene Two major forms of the HBZ RNA have been reported: a spliced form (sHBZ) and an unspliced form (usHBZ). The first exon of the sHBZ gene transcript is located in the U3 and R regions of the 3′ LTR (Figs. 16.1 and 16.2). Transcriptional start sites for sHBZ are scattered in the U5 and R regions of the 3′ LTR, an observation which is consistent with the finding that the predicted promoter was TATA-less.413 Three Sp1 binding sites are critical for transcription of the HBZ gene. Since Sp1 is a well-known regulator of housekeeping genes, transcription of the sHBZ gene may be constitutive and relatively constant. The levels of HBZ gene transcript are better correlated with provirus load than those of the tax gene transcript,305 confirming that the HBZ gene is frequently expressed in HTLV-1–infected cells. Thus, transcription from the minus strand stands in contrasts to that from the plus strand, which is highly inducible by Tax. RNA-FISH analyses show that HBZ is expressed in most cells at a much lower level of transcription.32,243 Importantly, HBZ expression is strongly associated with the S and G2/M phases of the cell cycle, suggesting that HBZ promotes mitosis in expressing cells.

The difference between sHBZ and usHBZ is only a few amino acids, as shown in Figure 16.1. The spliced transcript of HBZ is translated into a polypeptide of 206 amino acids, while the protein product of unspliced HBZ is a polypeptide of 209 amino acids. The expression level of sHBZ RNA is much higher than that of usHBZ RNA,381 and the half-life of sHBZ protein is much longer than that of usHBZ.413 These data indicate that expression levels of sHBZ are much higher than those of usHBZ, suggesting that sHBZ is more important than usHBZ for HTLV-1–infected cells and ATL cells.

Functions of HBZ HBZ has important functions in both its protein and mRNA forms242,313 (Fig. 16.4). RNA-FISH showed that HBZ mRNA is mainly localized in the nucleus whereas tax is exported from the nucleus.32

Function of HBZ protein HBZ protein is localized in the nucleus with a speckled pattern.131 HBZ has three domains: the activation, central, and bZIP domains (Fig. 16.4). HBZ was originally reported to suppress Tax-mediated viral gene transcription from the 5′ LTR via interaction with CREB2 and c-Jun.24,83 Indeed, many actions of HBZ act in opposition to those of Tax (Fig. 16.5). Further, HBZ interacts with various host factors that contain a bZIP domain, including CREB, JunB, ATF-1, and ATF-3, and hinders their transcriptional activation.24,104,130,202 Conversely, interaction of HBZ with JunD activates transcription of target genes.365 As a mechanism, HBZ modulates translational control of JunD, 606

inducing expression of the truncated isoform, Delta JunD.363 Furthermore, HBZ enhances TGF-β/Smad pathway by interaction with Smad2/3 and p300.423

FIGURE 16.4 Functions of HBZ. HBZ has three domains: activation, central, and bZIP. The HBZ protein exerts a variety of functions by interacting with host factors. On the other hand, HBZ RNA promotes proliferation via up-regulation of the E2F1 gene.

FIGURE 16.5 Interplay between Tax and HBZ. Interplays between Tax and HBZ on various pathways are shown. In most pathways, HBZ has opposite effects to Tax. In most ATL cases, ATL cells express Foxp3,122 a master molecule of regulatory T (Treg) cells. Furthermore, in HTLV-1–infected individuals, the proportion of HTLV-1–infected cells is higher in Treg populations,312 indicating that HTLV-1 is associated with Treg cells. HBZ induces Foxp3 expression in T cells, and its induction is enhanced in the presence of TGF-β.314 Thus, activation of TGF-β/Smad by HBZ induces Foxp3 expression in infected cells. This accounts for why HTLV-1 is associated with Treg cells. Furthermore, HBZ interacts with Foxp3 protein and impairs its functions.314 Thus, HBZ increases the number of functionally impaired Treg cells and may lead to the development of malignancy derived from Treg cells. HBZ also induces expression of Treg-associated molecules including CCR4 and T-Cell Immunoglobulin and ITIM Domain (TIGIT).340,410 Thus, HBZ determines the immunophenotype of HTLV-1–infected cells and ATL cells.

HBZ also selectively inhibits the classical NF-κB pathway by inhibiting DNA binding of p65 and promoting the degradation of p65.424 Although Tax expression is reported to induce cell senescence by hyperactivation of NFκB, inhibition of classical pathway of NFκB by HBZ attenuates senescence caused by Tax.293

shRNA library screening identified IRF4 and BATF3 as master regulators of proliferation of ATL cells. HBZ binds to a superenhancer of the BATF3 locus, leading to up-regulation of BATF3 and its downstream target, MYC.263 Thus, HBZ promotes proliferation of expressing T cells.

Function of HBZ RNA Suppression of HBZ gene expression by shRNA inhibits proliferation of ATL cell lines.13,313 Expression of HBZ in transgenic mice increases the number of T cells, while suppression of HBZ expression decreases tumor formation and infiltration of ATL cells.13 Thus, HBZ expression is associated with proliferation of ATL cells in vivo and in vitro. Mutation analysis of the HBZ gene showed that HBZ RNA, rather than HBZ protein, has a growth promoting effect on T cells whereas HBZ protein rather suppresses proliferation.242,313 The 5′ region of the sHBZ transcript is critical for this activity.242,413 Kinetic analysis of viral mRNA shows strong nuclear retention of HBZ mRNA.299 Poor polyadenylation of HBZ mRNA causes this nuclear retention.213 An antisense transcript of HIV-1 is also poorly polyadenylated and localized in the nucleus, suggesting that this is a common feature of antisense transcripts of retroviruses.

607

FIGURE 16.6 Schematic structure, functional domains, and functions of p12/p8. The protease cleavage sites are indicated. The known functions of p12/p8 are summarized. p8 induces cellular conduit formation, which is involved in HTLV-1 transmission. TM, putative transmembrane domain; LZ, leucine zipper-like structure; SH3, Src-homology 3; ER retention, endoplasmic reticulum retention signal. APH-2 An antisense transcript similar to HBZ has been discovered in HTLV-2 and named antisense protein of HTLV-2 (APH-2).107 The APH-2 gene encodes a 183 amino acid polypeptide that is localized in the nucleus. Although APH-2 does not have a bZIP domain, APH-2 interacts with CREB and represses Tax2-mediated transcription from the 5′ LTR of HTLV-2. Both HBZ and APH-2 have suppressive effects on transcription of sense viral genes, although any other functions of APH-2 remain to be elucidated.

Accessory Genes p12/p8 p12 is encoded by the singly spliced mRNA of HTLV-1 ORFI.49,192 p12 is a highly hydrophobic membrane protein of 99 amino acids, and it is localized in the endoplasmic reticulum (ER) and Golgi complex.64,191 p12 has a noncanonical ER retention/retrieval motif, two putative transmembrane (TM) domains, four putative proline-rich (PXXP) Src-homology 3 (SH3)-binding domains, two putative leucine zipper (LZ) motifs, and a putative adaptin motif77,269 (Fig. 16.6). When ectopically expressed in HeLa cells, p12 forms a homodimer through its TM domains.374

A p12-deficient HTLV-1 mutant (p12 mutation of Met-1 to Leu and an HBZ mutation of His-151 to Gln) had reduced infectivity of DCs, and when rhesus macaques were infected with it, viral propagation and persistent infection were poorly established.383 On the other hand, this mutant HTLV-1 produced infectious viruses comparable to wild type HTLV-1, immortalized primary human T cells in vitro, and established persistent infection in rabbits. Thus, DC infection with HTLV-1 appears to be critical for persistent infection in macaques, and p12 may play a role in this process.

Effect of p12 on T-cell activation and proliferation p12 activates Ca2+ signaling in T cells.63,64 Ectopic overexpression of p12 in a T-cell line increases intracellular Ca2+ levels, stimulating the phosphatase calcineurin to dephosphorylate NFAT and then NFAT to activate transcription of genes such as IL-2.4,179 p12 increases intracellular Ca2+ concentration by interacting with two ER resident proteins, calnexin and calreticulin, which regulate Ca2+ release from the ER.64 In addition, p12 interacts directly with calcineurin. Since HTLV-1–transformed T-cell lines do not generally exhibit constitutive NFAT binding to the IL-2 promoter,272 NFAT appears to be only transiently activated by p12 at a particular phase of HTLV-1 infection such as an initial infection.

p12 in T cells can stimulate the IL-2 signaling pathway downstream of the IL-2R. p12 interacts with the beta and gamma chains of the IL-2R and enhances the phosphorylation of STAT5 and its DNA binding, thereby augmenting transcription.257 Expression of p12 in a human T-cell line transformed by a p12-defective HTLV-1 mutant augments colony formation in soft agar in conditions of low IL-2.

p12-induced attenuation of the host immune response p12 is implicated in reduced host immunity to HTLV-1–infected cells. HTLV-1–infected primary CD4+ T cells are resistant to autologous NK cell–mediated killing.16a Such killing is initiated by binding of NK to target cells via adhesion molecules such as the ICAMs. HTLV-1 infection reduces the expression of ICAM1 and ICAM2 in primary CD4+ T cells, thereby reducing NK cell binding. Lentivirus-mediated expression of p12 in primary CD4+ T cells downmodulates expression of ICAM1 and ICAM2. In addition, p12 can reduce CTL-mediated killing.157 p12, through interaction with the MHC class I heavy chain, inhibits its interaction with β2-microglobulin, thereby inducing the proteasome-dependent degradation of MHC class I.

Functions of p8 Proteolytic cleavage of p12 generates the C-terminal product p8, which may promote HTLV-1 transmission.385 The cleavage of p12 removes the ER retention/retrieval motifs, and thus p8 is localized to the T-cell membrane (Fig. 16.6). Upon T-cell receptor activation, p8 is recruited into the immunological synapse. Comparison of p8-positive and negative HTLV-1 indicates that p8 increases cell-to-cell contact and induces lymphocyte function–associated antigen-1 (LFA-1) mediated cell clustering, augmenting the number and length of conduits (filopodia-like membrane extensions) and thereby enhancing HTLV-1 transmission. 608

p13 p13 is encoded by the singly spliced monocistronic mRNA of the HTLV-1 x-II ORF.329 p13 is a highly basic protein of 87 amino acids and is identical to the C-terminal 87 amino acids of p30. p13 has the following domains: an amphipathic alpha helix (residues 20 to 35), a mitochondrial targeting signal (MTS) (residues 21 to 30), a transmembrane region (residues 30 to 40), and several PXXP motifs (Fig. 16.7).

A p13-defective HTLV-1 virus (with a mutation of the p13 initiation codon) showed comparable infectivity of rabbit PBMC to that of wild type virus, but the mutant virus could not establish infection in rabbits in vivo, as measured by viral loads and antibody responses, indicating that p13 plays a still-unknown role in establishing HTLV-1 infection in vivo.126

p13 is localized mostly in the inner mitochondrial membrane, and its ectopic expression in isolated mitochondria induces mitochondrial swelling, depolarization, increased respiratory chain activity, and production of ROS, through activation of the inward K+ current.30,330 p13 also induces ROS production in primary quiescent T cells.

p30 p30 is encoded by the doubly spliced mRNA of x-II ORF and is a protein of 241 amino acids.115,250 It is unusually rich in Ser (23%) and Arg (12%) and localized in the nucleus and the nucleolus.

Posttranscriptional regulation of viral RNA by p30 p30 inhibits the nuclear export of doubly spliced tax/rex mRNA through retaining the mRNA in the nucleus.268 p30 binds to the splice junction region (env exon) of tax/rex mRNA. Although both Rex and p30 interact with HTLV-1 RNA and decrease the amount of tax/rex mRNA exported from the nucleus, they generally have opposite functions in virus production. Whereas Rex stimulates virus production through augmenting the expression of structural proteins (Gag, Pol, Env), p30 inhibits virus production through reducing the expression of Tax and Rex. Interestingly, the Rex and p30 proteins interact with each other, and their interaction is augmented by the presence of the viral mRNAs.56,331 While p30 has little effect on Rex activity, Rex counteracts p30 activity and induces the expression of Tax and Rex proteins. Thus, the ratio of p30:Rex may control the transition between virus latency and virus production. Since Tax is transiently expressed in HTLV-1–infected cells and ATL cells, and transcription of genes encoded in the sense strand of the provirus depends on Tax, it follows that expression of p30 is also transient, which may contribute to the fine-tuning of viral replication.

FIGURE 16.7 Schematic structure, functional domains, and functions of p13. The functions of p13 are summarized. TM, putative transmembrane domain; MTS, mitochondrial targeting signal; SH3, Src-homology 3; NLS, nuclear localization signal; LZ, leucine zipper-like structure. An HTLV-1 infectious clone with a nonsense mutation in p30 was inoculated into four Rhesus macaques. Reversion to wild type was observed in two macaques, indicating that p30 is necessary for infectivity in vivo.383 Furthermore, viral replication of the HTLV-1 infectious clone with the p30 mutant was severely impaired in human dendritic cells.

Transcriptional and posttranscriptional regulation of cellular genes by p30 p30 also interacts with several cellular factors. p30 interacts with the Ets domain of PU.1, which leads to suppression of TLR4.57 Furthermore, p30 suppresses the expression of interferon-responsive genes.72 These functions of p30 account for impaired viral infectivity to myeloid cells in a p30-deficient infectious clone. p30 also suppresses proliferation of T cells by inhibiting their entry into the S phase of the cell cycle and causing accumulation of cells in the G2 and M phases.25,56 These effects are caused by p30 interacting with cyclin E and CDK2, resulting in inhibition of cyclin E-CDK2 complex. Similar to Tax, p30 also impairs homologous recombination repair.26 Thus, Tax and p30 possess similar effects on cell cycling and DNA repair of expressing cells. It is possible that these functions play critical roles in de novo infection when tax undergoes a transcriptional burst.

Genomic Instability in ATL Cells ATL cells have numerous genetic abnormalities such as DNA mutations, chromosomal translocations, deletions and duplications, and aneuploidy, and these genetic aberrations are linked to ATL development. For example, homozygous deletions of p16 (CDNN2A) and p15 (CDKN2B) genes, tumor suppressor genes that regulate the cell cycle, were frequently observed in aggressive acute and lymphoma-type ATL.117 Both Tax and HBZ are implicated in the genomic instability of ATL cells.

p53 The tumor suppressor p53 is the “guardian of the genome” in DNA damage responses. p53 is functionally inactivated by Tax.407 Tax interacts with the p53 coactivator CBP/p300 and blocks its access to p53, thereby reducing the transcriptional activation function of p53.346

Defective DNA Repair 609

DNA repair activity is attenuated by Tax in HTLV-1–infected T-cell lines.150,165,224,245,295 Tax represses transcription of the DNA polymerase β gene, which plays a crucial role in DNA repair.150 Ataxia telangiectasia mutated (ATM), a member of the PI3K-like kinase family, is activated by various DNA damaging agents, and its activation is essential in initiating signaling cascades that stop DNA replication and allow the repair of damaged DNA. ATM activation by ionizing radiation and signals downstream of ATM activation are diminished in HTLV-1–infected T-cell lines as well as cells expressing Tax alone.43 Upon treatment with ionizing radiation, both normal and Tax-expressing cells stop DNA replication, but the interval before replication resumes is shorter in Tax-expressing cells than in the parental control cells; simultaneously, the cells expressing Tax are defective for DNA repair activity. Although the precise mechanism by which Tax blocks DNA repair has not been fully elucidated, Tax is known to interact with several host factors involved in DNA repair, such as DNA-dependent protein kinase, Ku, Chk1, and Chk2.65,288,289

Constitutive NF-κB activation by Tax induces cell cycle arrest or senescence.212,293,408 Activation of NF-κB by Tax induces an R-loop structure—a three-stranded nucleic acid structure consisting of an RNA–DNA hybrid and a single-strand DNA—which induces senescence.118 Two transcription-coupled nucleotide excision-repair endonucleases, xeroderma pigmentosum F (XPF) and XPG, are important for excision of R-loops. In ATL cells, expression of these two endonucleases is suppressed, indicating that such ATL cells proliferate regardless of accumulated R-loops.

HBZ induces DNA-strand breaks, which are caused by activated miR17 and miR22. These microRNAs suppress expression of hSSB2, which encodes a single-stranded DNA-binding protein associated with genome stability.391 hSSB2 is implicated in reduced efficiency in HR-dependent repair of DSBs and defective ATM-dependent phosphorylation.211 Thus, HBZ is also associated with genome instability, which is linked to oncogenesis.

Chromosomal Instability The centrosome is a microtubule organizing center during cell division and plays a crucial role in the precise duplication and segregation of chromosomes. While normal T cells have one centrosome during interphase, in HTLV-1–infected cell lines, around 30% of the cell population has more than two centrosomes, indicating that a centrosomal aberration may contribute to the chromosomal abnormalities observed in HTLV-1–infected cells.45 Such abnormal centrosomes are induced by the ectopic expression of Tax in human T cells.

This activity of Tax is mediated through its interaction with several host proteins such as TAX1BP2 and RanBP1.45 Endogenous TAX1BP2 is localized in the centrosome, and its knockdown induces centrosome amplification. Furthermore, overexpression of Tax1BP2 reduces Taxinduced centrosome amplification. These results indicate that Tax induces centrosome aberrations by inactivating TAX1BP2. In contrast, the knockdown of RanBP1 abrogates centrosome amplification by Tax as well as centrosome localization of Tax, indicating that RanBP1 targets Tax to centrosomes. MAD1 is another Tax-interacting protein localized in the centrosome during metaphase. While the knockdown of MAD1 induces multinucleated cells, its overexpression inhibits induction of multinuclear cells by Tax.154 Thus, MAD1 is also involved in the genomic instability induced by Tax in HTLV-1–infected cells.

PATHOGENESIS AND PERSISTENCE IN VIVO Entry Into Cells and Host Entry Into the Host Transmission of HTLV-1 between individuals requires transfer of infected lymphocytes, because lymphocytes naturally infected with the virus produce few if any cell-free virions. There are three main routes of transmission: breast-feeding and sexual intercourse are the two natural routes, by which the virus is maintained in the population. Breast milk contains lymphocytes210; seminal fluid also contains lymphocytes and appears to enhance HTLV-1 replication.255 The risk of mother-to-child transmission of HTLV-1 is associated with a high PVL,380 with the presence of anti-Tax antibodies in the mother,127 and with prolonged breast-feeding.397 Transfusion of blood products that contain lymphocytes was an important source of HTLV-1 infections in countries with endemic HTLV-1 infection before screening of blood products was introduced,280 and transplantation of solid organs from HTLV-1–infected donors is a persistent but rare cause of infection.361

In a host infected by sexual intercourse, the portal of entry is the genital mucosa. In the breast-fed infant, infection probably enters in the oropharynx, which is surrounded by the lymphoid tissue of Waldeyer ring, including the tonsils. It is unlikely that the virus enters in the infant’s stomach. Although the pH of the stomach of neonates is not strongly acidic, because of ingestion of amniotic fluid, which has a pH of about 7, the pH drops quickly in the first few days after birth, reaching pH 3 by 1 week.12 The precise site and mechanism of the entry of infected T cells across the epithelial barrier in either portal of entry are not well understood.292 It is often assumed that dendritic cells play a part in the initial acquisition of infection at the portal of entry, but direct evidence is lacking.

Entry Into the Cell HTLV-1 is unusual among viruses in its almost exclusive reliance on cell-to-cell contact for transmission of infection between cells: naturally infected cells release few—if any—cell-free infectious virions, and HTLV-1 relies instead on the mobility of the host cell to spread both within and between hosts.19 Since the virus can infect a very broad range of human cell types, other mammalian cells, and some avian cells (see “Cell and “Tissue Tropism”), the cellular receptor for the virus was inferred to be widely distributed. Early attempts to identify the receptor exploited the ability of HTLV-1–producing T cell lines to induce multinucleate syncytia by fusion of the plasma membrane of the infected cell with a cell that is brought into contact.261 Sommerfelt et al.336 used human–mouse somatic cell hybrids to identify the chromosome responsible for conferring susceptibility to infection by VSV pseudotypes carrying the HTLV-1 envelope glycoprotein. Their results indicated a region on the long arm of chromosome 17. Tajima et al.351 similarly used HTLV-1 Env-induced fusion with somatic cell hybrid lines, with a recombinant vaccinia virus expression system, to localize the putative receptor to 17q21-23. However, subsequent studies identified the two key cellular receptor molecules GLUT-1 and Neuropilin-1 (NRP-1), encoded respectively on chromosomes 1 and 10. The identity of the putative additional factors encoded on chromosome 17 remains unknown.

610

The GLUT-1 glucose transporter was the first identified cellular receptor for HTLV-1 (Fig. 16.8A).54,158,181,229 GLUT-1 interacts with the receptor-binding domain (RBD) (the first 215 aa) of HTLV-1 Env and HTLV-2 Env. Knockdown of GLUT-1 by siRNA results in reduced binding of the HTLV-1 and HTLV-2 RBD to the cells as well as reduced infection by HTLV-2-env pseudotype virus, and the binding is rescued by overexpression of GLUT-1 but not GLUT-3, another glucose-transporter family member. In addition, polyclonal chicken antibody against the extracellular loop domain of GLUT-1 inhibits Env-mediated fusion as well as infection.156 Specific regions of GLUT-1 mediate the binding to HTLV-1 Env and virus entry into cells.227

NRP-1 also functions as an HTLV-1 receptor.93 Overexpression of NRP-1 in cells with a low endogenous expression of NRP-1 augments HTLV-1 Env-mediated infection, and NRP-1 knockdown by siRNA in 293T cells reduces HTLV-1 Env-mediated infection. NRP-1 colocalizes with the Env protein at the cell membrane and forms a complex with GLUT-1. This complex formation is augmented by the expression of Env protein, suggesting that NRP-1 and GLUT-1 make an HTLV-1 receptor complex that mediates HTLV-1 binding and entry (Fig. 16.8A), just as the CD4 molecule and chemokine receptors do for HIV infection. In addition, the relative importance of GLUT1 and NRP-1 for HTLV-1 infection may differ between cell types. For instance, knockdown of GLUT1 reduced Env-mediated fusion in HeLa cells but not in U87 glioblastoma/astroglyoma cells.155

FIGURE 16.8 The HTLV-1 receptor complex and the current model of HTLV-1 entry. A: (1) The HTLV-1 envelope protein (Env) attaches to heparan sulfate proteoglycans (HSPGs) on the target cell, which increases the local concentration of the viruses at the cell surface. (2) HTLV-1 Env then binds to neuropilin-1 (NRP-1), and this binding induces a conformational change of Env that facilitates its interaction with glucose transporter 1 (GLUT-1). (3) The formation of a ternary complex of Env, NRP-1, and GLUT-1 induces a conformational change of Env that triggers the fusion of the viral and cell membranes. B: The contact site between an HTLV-1–infected cell and a target cell forms a special structure referred to as the virological synapse (VS). VS formation involves polarization of the microtubule-organizing center near the site of cell-to-cell contact in the infected cell. Complete, enveloped infectious virions can pass from the infected cell to the target cell either in pockets between the two cells’ plasma membranes in the center of the VS or in an extracellular carbohydrate-rich assembly at the periphery of the VS. While the primary cellular receptors for HTLV-1 are GLUT-1 and NRP-1, infection is also enhanced by the surface adhesion molecules ICAM-1, ICAM-3, and VCAM-1. Heparan sulfate proteoglycans (HSPGs), cofactors in many viral infections, also promote HTLV-1 binding and infection of CD4+ T cells.158,296 HTLV-1 virions as well as purified gp46 proteins bind to HSPGs on CD4+ T cells, and the binding is abrogated by treatment of target cells with heparan sulfate lyase. Moreover, soluble heparin blocks HTLV-1 infection of susceptible cells.

HTLV-1 entry into susceptible cells begins with the binding of the HTLV-1 envelope proteins to a viral receptor on the membrane of the host cell, and it is followed by the fusion of viral and cell membranes.17,92,228 The HTLV-1 envelope protein (Env) is synthesized as a 68 kD polyprotein precursor and is cleaved to produce two subunits, the surface glycoprotein (SU) of 46 kD (gp46) and the transmembrane (TM) glycoprotein of 21 kD (gp21). Mature Env proteins on the virion are trimers, consisting of three SUs and three TMs linked by disulfide bonds. The cytoplasmic domain of gp21 controls envelope-mediated syncytium formation and cell-to-cell HTLV-1 entry.178,297 The C-terminus of gp21 contains a PDZdomain protein binding motif, and the deletion of this motif reduces the stability of Env.35 In addition, the Y-S-L-I motif in the cytoplasmic domain of gp21 is crucial for cell-to-cell HTLV-1 transmission.58

Virological Synapse Since the discovery of HTLV-1 in 1980, it was recognized that efficient transmission of HTLV-1 between cells required cell-to-cell contact, but the reasons for this requirement were unknown. An important clue came from the observation that HTLV-1 preferentially infects T cells that are themselves specific for HTLV-1 antigens—both CD8+ T cells112 and CD4+ T cells.102 This observation led to the hypothesis that HTLV-1 is transmitted between cells through the immunological synapse, the specialized area of cell-to-cell contact that is formed between a T cell and an antigen-presenting cell.66 Confocal microscopy of two-cell conjugates that are formed spontaneously between PBMCs from infected individuals135 indeed revealed a structure (Fig. 16.8B) that closely resembles the immunological synapse, with organized microdomains of proteins forming an annular adhesion domain surrounding a central domain containing the viral Gag protein instead of the signaling molecules and cytokines present in the center of the immunological synapse. However, there is a crucial difference between this HTLV-1–induced structure and the immunological synapse: the microtubule organizing center (MTOC), which comes to lie near the cell-to-cell contact in both HTLV-1 transmission and in the immunological synapse, is polarized inside the HTLV-1–infected cell toward the responding antigen-specific T cell. By contrast, in the immunological synapse, the MTOC is polarised in the responding T cell toward the antigen-presenting cell. It, therefore, became clear that the polarization of a microtubule cytoskeleton is driven by the virus itself, and not by T-cell receptor recognition of antigen as in the immunological synapse. The integrity of the microtubule cytoskeleton is required for cell-to-cell transmission of infection.135 The structure is, therefore, named a virological synapse135; subsequent work by other groups showed that HIV and MLV can also be transmitted between cells via a virological synapse. 611

The two chief molecular signals that trigger formation of the HTLV-1 virological synapse are engagement of ICAM-1 on the surface of the infected T cell by LFA-1 on the target cell (Barnard et al., 2005) and activation of the CREB pathway by HTLV-1 Tax protein present at the MTOC.267

Although HTLV-1–infected cells regularly cause syncytia in vitro by fusion with many different cell types, cell fusion is not involved in the virological synapse and does not appear to occur in vivo. HTLV-1, therefore, relies on the Env glycoprotein to fuse the virion envelope with the plasma membrane of the target cell.59,60 Electron tomography reveals the ultrastructure of the HTLV-1 virological synapse: large areas of the plasma membranes of the donor (infected) and target (recipient) cells are closely apposed with a separation of approximately 24 nM.225 But the closely apposed membranes are interrupted by several pockets of 100 to 200 nM, isolated from the extracellular fluid. Immunostaining reveals HTLV-1 Gag protein both lining the plasma membrane in the vicinity of the pocket and in particles resembling virions in the lumen of the pocket.

These observations led to the current picture of the mechanism of transmission of HTLV-1 between cells.19 When a cell expressing LFA-1 engages ICAM-1 on the surface of a cell expressing HTLV-1 Tax protein, the plasma membranes of the two cells form a large, close contact, with an annular adhesion structure containing LFA-1 and Talin. The microtubule cytoskeleton in the infected cell is then reoriented to bring the MTOC next to the virological synapse, and the microtubules are used to transport the viral material to the synapse. Enveloped virions are formed in the isolated pockets within the synapse225 and traverse the short distance, of the order of a few virion diameters, to fuse with the plasma membrane of the target cell. Cell-to-cell transmission of HTLV-1 can also occur at the periphery of the synapse, in carbohydrate-rich, extracellular matrix containing virions286 (Fig. 16.8B). In vitro, HTLV-1 transfer between cells has also been observed via cellular conduits named tunnelling nanotubes,385 whose formation is promoted by the HTLV-1 accessory protein p8, which is expressed by certain strains of HTLV-1.

Propagation of HTLV-1 in vitro, even by cell contact with a strong HTLV-1–producing T cell line, is surprisingly inefficient. It is likely that specific conditions in vivo, yet unidentified, make this process more efficient.

Cell and Tissue Tropism The cell types infected by HTLV-1 are determined in part by the expression of the three molecules identified as cellular receptors or cofactors: GLUT-1, NRP-1, and heparan sulphate proteoglycans (HSPGs). However, HTLV-1 can infect most types of nucleated human cells in vitro,193,261,336 whereas in vivo it is almost wholly confined to T lymphocytes: typically approximately 95% of the PVL is present in CD4+ T cells, and approximately 5% in CD8+ T cells.238 Most HTLV-1–infected T cells also carry the key marker of the memory phenotype, CD45RO300; the infected CD4+CD45RO+ and CD8+CD45RO+ cells proliferate in vivo significantly faster than uninfected cells.15 It is not known whether the CD45RO molecule itself enhances infection; it is more likely that the infected cells preferentially survive in vivo in the long-lived memory (CD45RO+) clones. HTLV-1 cannot be propagated efficiently in vitro in cell types other than T lymphocytes.

By contrast, HTLV-2 is present almost exclusively in CD8+ T cells in vivo.240 Both HTLV-1 and HTLV-2 use GLUT-1 and NRP-1 as their primary cellular receptors, and the two molecules can be expressed on both CD4+ and CD8+ T cells. In vitro, both HTLV-1 and HTLV-2 can infect both CD4+ and CD8+ T cells.163 However, the two viruses differ in their ability to transform T cells in vitro: whereas HTLV-2 transforms CD4+ T cells, HTLV-2 transforms CD8+ T cells. The distinct transforming abilities of the two viruses in vitro are due to the respective envelope glycoprotein,163,164,401 but the molecular details of this action of Env are not known. These observations imply that the cell-type specificity of both virus propagation and subsequent cellular transformation by HTLV-1 and HTLV-2 is determined by postentry factors, rather than by distinct mechanisms of cell entry.232 At present, these putative postentry factors are not well understood.

HTLV-3 uses GLUT-1 as a viral receptor, but neither NRP-1 nor HSPG is required for infection, and HTLV-3 binds not only activated primary T cells but also resting ones.401 These results suggest that these three HTLVs acquired distinct entry mechanisms after diversification from their common ancestor.

FIGURE 16.9 The genomic integration site of HTLV-1 is specified at different spatial scales. (a) Most HTLV-1–infected cells contain a single provirus. Initial integration of the provirus does not favor a particular chromosome, but a provirus integrated in an acrocentric chromosome has a significant survival advantage in vivo. (b) Integration in the host genome preferentially occurs in nucleosomal DNA in transcriptionally active regions, near certain transcription factors and other chromatin-associated proteins. (c) Protein phosphatase 2A (PP2A) facilitates the integration of the provirus, possibly by tethering the preintegration complex or intasome to the host chromatin. (d) There is a weak but reproducible preference, as in other exogenous retroviruses, to integrate at a specific, nonpalindromic DNA sequence motif. Initial targeting of the integration does not favor any one chromosome, but strong in vivo selection results in preferential survival of clones carrying a provirus in one of the five acrocentric chromosomes, especially chromosome 13. The reason for this preferential survival in vivo is not known. (Republished with permission of Annual Reviews, Inc. from Bangham CRM. Human T cell leukemia virus type 1: persistence and pathogenesis. Annu Rev Immunol 2018;36:43–71; permission conveyed through Copyright Clearance Center, Inc. Copyright © 2018 by Annual Reviews, https://www.annualreviews.org.) 612

Both HTLV-1 and HTLV-2 are exquisitely adapted to persist in vivo in T lymphocytes. Since they depend on the virological synapse to spread, it is possible that T-cell–specific factors involved in the virological synapse, such as the unusually high mobility of the MTOC in T cells, contribute to the selective propagation of these viruses in T lymphocytes.

Integration and Clonality Integration of HTLV-1 in the host cell genome as a double-stranded DNA provirus is an essential part of the viral life cycle, as in other retroviruses.19 The site of integration in host DNA is not random, but rather is determined by factors at a succession of different spatial scales (Fig. 16.9). Each of these factors imposes a statistically significant but small effect on the choice of integration site, and as a result the virus is widely dispersed throughout the genome (see “Clonality of HTLV-1 and HTLV-2”). Each infected T cell carries a single copy of the provirus53: the mechanisms of resistance to superinfection by HTLV-1 are not known. Unlike HIV-1 infection, there are no hotspots of HTLV-1 integration in the human genome.

At the smallest scale, there is preferential integration of HTLV-1 in a primary DNA sequence motif of about eight nucleotides.184 It was previously thought that retroviral insertion site motifs were palindromic, but it is now known that the palindromicity in the consensus sequence was due to the presence of the short (nonpalindromic) motif on either the plus-strand or the minus-strand of the host genome.184 The integration site motif that is used by both HTLV-1 and HTLV-2 closely resembles that of HIV-1.184

The mechanism of integration is shared by other retroviruses.218 The virus-encoded integrase forms a preintegration complex or “intasome” with the newly reverse-transcribed double-stranded DNA of the virus, the viral integrase makes a six base pair staggered cut in the host genome, and the free ends are ligated respectively to the 5′ and 3′ ends of the double-stranded viral DNA. The 6-nucleotide gap left at each end by the staggered cut in the genome is then filled in, generating a 6-base pair repeat flanking the integration provirus.

The abundant host phosphatase PP2A associates with the HTLV-1 integrase and participates in the process of proviral integration.217 The tetrameric structure of the integrase of the closely related simian virus STLV-1, in complex with the B56g regulatory subunit of PP2A, has been determined by cryo-EM.23 PP2A does not directly bind chromatin, and the molecular details of its cooperation with the integrase are not yet known. HTLV-1 also shows a bias toward integration into open chromatin near certain transcription factor binding sites, especially STAT-1, HDAC6, and TP53.239

Clonality of HTLV-1 and HTLV-2 A cardinal difference between HTLV-1 and HIV-1 lies in the direct impact of the virus on the infected T cell: HIV-1 expression usually kills the host cell, whereas HTLV-1 expression is nonlytic. This difference is reflected in the distinct modes of persistence of the viruses in vivo: while HIV-1 persists in the absence of antiretroviral therapy by constant de novo infection, HTLV-1 persists during the long chronic phase of infection chiefly by continued proliferation of clones of infected T cells.15,200 Each infected T-cell clone can be identified by the sequence of the host genome flanking the single-copy provirus; this sequence is shared by each cell in that clone.

FIGURE 16.10 Schematic depiction of the dynamics of proviral load and clone abundance in one individual over time. The trajectory of the proviral load is inferred from data on the early stage of infection and the long-term stability of the load in the chronic stage of infection. Many HTLV-1–infected clones persist for years; the number of cells in each high-abundance clone (i.e., a clone that contains many sister cells) tends to increase progressively over time, whereas each low-abundance clone decreases in number; these changes result in a progressive rise in the oligoclonality index. Each infected cell expresses Tax in uncommon, intermittent bursts32,221: the frequency of expression appears to be greater in the low-abundance clones than in the high-abundance clones, perhaps owing to cytotoxic T-lymphocyte–mediated selection. The contribution of infectious spread to the maintenance of an individual’s set point proviral load is not known. (Modified from Bangham CRM, Matsuoka M. Human T-cell leukaemia virus type 1: parasitism and pathogenesis. Philos Trans R Soc Lond B Biol Sci 2017;372(1732):20160272. https://creativecommons.org/licenses/by/4.0/.)

In chronic infection, an equilibrium is established in each host between the immune response to the virus (see “Immune Response”) and the number and abundance of HTLV-1–infected T cell clones (Fig. 16.10). At this equilibrium, a small number of infected clones can reach a high abundance in the peripheral blood: this phenomenon is known as oligoclonal expansion. Early, semiquantitative methods of detection (Southern blot and PCR-based protocols) could identify only the most abundant clones, and these techniques led to the misapprehension of two fundamental aspects. First, it appeared that oligoclonal expansion was associated with the inflammatory disease, and by implication might be involved in causing the inflammatory tissue damage. Second, the number of infected T-cell clones in each host was estimated at about 100.394 The introduction of a sensitive high-throughput method to map and quantify the proviral integration sites96 corrected these misunderstandings. First, the results showed that a typical host had over 104 distinct clones of HTLV-1–infected T cells. Second, the PVL, which is the strongest correlate of the risk of both the inflammatory and malignant diseases associated with HTLV-1, is proportional to the number of distinct clones present in the individual, not to the degree of oligoclonal expansion. The degree of oligoclonality, which is rigorously quantified by the oligoclonality index,96 does not differ between patients with HAM and asymptomatic carriers. The oligoclonal expansion is more easily detected in samples from patients with HAM simply because of the greater absolute abundance of the largest clones in these individuals.

These observations show that the risk of HTLV-1–associated disease is correlated with the number of HTLV-1–infected clones, not to the degree of oligoclonal expansion. In ATL, one or more clones undergo malignant transformation, and the oligoclonality index quickly exceeds the normal range.302 613

Although HTLV-1 persists in the chronic phase mainly by mitotic spread—proliferation of existing clones—there are two reasons why infectious spread of the virus must continue throughout the infection. First, the immune response is persistently activated, indicating constant synthesis of viral antigens. Second, the virus itself does not change between the primary infection and the persistent phase. Longitudinal data on clonal abundance in individuals with nonmalignant infection indicate that, in the chronic phase, mitotic spread exceeds infectious spread by at least 107 to 1.200 But since the number of CD4+ T cells in the body is very large, on average it can be estimated that about 5 × 109 new infected cells are produced per day, of which between 100 and 200 result from de novo infection, and the remainder from mitotic proliferation of existing clones.200 Thus, in a typical infected individual, an average of 6 × 105 new clones are created over 10 years. Since the infection (in nonmalignant cases) is in quasi-equilibrium, and the oligoclonality index and PVL change slowly (if at all) over time, this figure implies that an approximately equal number of clones must be destroyed over the same period, either by the immune response (see “Immune Response”) or by being outcompeted by faster-growing clones. Many newly infected cells are likely to fail to establish a long-lived clone. However, some newly created clones may become established, if they possess a strong growth or survival advantage.

Whereas mitotic spread of HTLV-1 dominates during the chronic phase of infection, infectious spread must have been abundant during the early phase, to establish the large number of infected clones observed in each host (Fig. 16.10). In the early stages of infection in recipients of solid organ transplants from HTLV-1–infected individuals, the PVL can double in 1.4 days,51 leading to a 1,000-fold increase in the PVL within 14 days, but the ratio of mitotic to infectious spread at this stage has not been measured.

By contrast with HTLV-1, HTLV-2—which infects virtually only CD8+ T cells in vivo—generates a much smaller number of clones of infected host cells in the steady-state chronic infection.240 As in HTLV-1 infection, the infected T-cell clones are long-lived and stable. Each infected clone, however, can reach a high abundance in the circulation. The resulting high value of the oligoclonality index96 in HTLV-2 does not carry the same implications for disease as in HTLV-1, because HTLV-2 does not cause malignant disease.

Site of Replication and Spread After transmission by breast milk, sexual contact, or blood transfusion, HTLV-1 infects DCs and T cells in vivo and spreads through cell-to-cell contact. HTLV-1–infected cells proliferate and spread in the body. In experimentally infected squirrel monkeys (Saimiri sciureus), the HTLV-1 provirus was detected first in peripheral blood mononuclear cells (PBMCs) and then in lymphoid organs including lymph node, spleen, and bone marrow.176 This shows that HTLV-1 spreads in lymphoid tissues. In HTLV-1–infected individuals, bone marrow has been identified as a reservoir of HTLV-1.208 The identification of the HTLV-1 provirus integrated in precisely the same genomic site in different hematopoietic cell types81 suggests that hematopoietic stem cells can be infected with HTLV-1.

HTLV-1 virions cannot be detected in HTLV-1–infected individuals. Instead, HTLV-1–infected cells proliferate in vivo: HTLV-1 copy number increases by mitosis of infected cells, while HTLV remains in the form of a provirus integrated in the host genome. Expression of adhesion molecules, including ICAM-1 and LFA-1, is up-regulated on HTLV-1–infected cells.76 In HAM/TSP patients and HTLV-1 carriers, infected cells not only circulate in the blood but also infiltrate into the skin, lung, and intestine. Thus, HTLV-1–infected cells persist and spread in vivo. If infected cells have proinflammatory properties, their proliferation and activation might induce inflammatory diseases.

Proviral Load: Correlation With Disease HTLV-1 infection is almost wholly intracellular, and most infected T cells in vivo do not express the provirus at a given instant, so the appropriate measure of the viral burden is the PVL, the average number of proviral copies per 100 PBMCs. Since the great majority of infected cells carry a single copy of the provirus,53 the PVL may be approximated by the percentage of infected PBMCs.

The PVL is the strongest correlate of the risk of both the inflammatory disease HAM260 and the malignant disease ATL145 caused by HTLV-1. The PVL reaches a steady state value or fixed point in each infected individual: this value can fluctuate by about fivefold over time within the individual host236 but typically does not progress unless ATL supervenes. But the PVL set point can differ between infected people by several orders of magnitude.260 What determines a person’s set point PVL? Since the virus varies little in sequence between individuals, and there is wide variation in the PVL between hosts infected with an identical virus, this variation is attributable to differences in the host, rather than the virus. Further, since the PVL in a given individual usually returns to the same set point after perturbation by drug treatment, the set point is likely to be determined at least in part by genetic mechanisms. The highly polymorphic gene complexes of the major histocompatibility complex (human leukocyte antigen [HLA]) and the killer immunoglobulin-like receptors (KIRs) play a significant part in determining a person’s PVL set point151,318 (see “Immune Response”). However, these genetic effects account for only a fraction of the observed variation between people in the PVL of HTLV-1318; it is likely that the remaining variation is due to the additive effects of many genetic polymorphisms in the host, each polymorphism contributing a small proportion to the total observed variation.

The reasons for the strong correlation observed between PVL and the risk of ATL are not known with certainty, but two factors are likely to contribute. First, the PVL is correlated with the number of different clones of HTLV-1–infected T cell,96 and the probability that one or more clones accumulates replicative mutations that lead to malignant transformation is likely to rise in proportion. Second, a high PVL results partly from an inefficient T-cell–mediated immune response to the virus18; this inefficient response will also reduce the rate at which HTLV-1– expressing cells are killed in vivo, so enhancing the survival of potentially malignant clones.

The reasons for the observed correlation between the PVL and the risk of HAM are discussed below (see “Pathogenesis of Inflammatory Diseases”).

Immune Response Innate Immune Response Primary infection with HTLV-1 is asymptomatic, and consequently it has not been possible to study the impact of the innate immune response in the early stages of infection. However, certain components of the innate immune response have been shown to affect the virus even during chronic infection. The cytidine deaminase APOBEC3G, which plays a significant part in HIV-1 infection, produces uncommon mutations in the

614

plus strand of the provirus of HTLV-1.70 A clone carrying such mutations may survive indefinitely, because the HBZ gene, which is necessary for clonal survival, is encoded on the minus strand of the provirus and so is not mutated by the APOBEC3G. The contribution of cytokines to both the innate and the acquired immune response is diminished by the HTLV-1 Tax protein, which induces the expression of the gene SOCS1, which encodes a suppressor of cytokine signaling.44,281 The deoxynucleotide triphosphate (dNTP) triphosphohydrolase SAMHD1 again has important activity against certain lentiviruses.98 The enzyme acts by depleting the intracellular concentration of dNTPs, thereby inhibiting viral replication. SAMHD1 limits HTLV-1 replication in macrophages,348 resulting in apoptosis induced by STING. However, since the macrophage carries a very small proportion of the HTLV-1 PVL, the importance of SAMHD1 in HTLV-1 infection is limited.

Type 1 interferons can suppress HTLV-1 in replication both in vitro and in vivo.160,183 However, the impact of type 1 interferon in natural infection is limited by two factors. First, Tax protein inhibits the kinase TBK1,418 which normally phosphorylates the interferon regulatory factor IRF3, so diminishing interferon production. Second, the induction of SOCS1 may reduce the impact of interferon-induced cytokines.

While type 1 interferon alone has little impact on HTLV-1 infection in vivo, sustained treatment with a combination of IFNα and the nucleoside analogue AZT has been shown to prevent the progression of ATL in certain cases28; see Treatment.

The three main cell types in the innate immune response to viruses are the dendritic cell (DC), the natural killer (NK) cell, and the regulatory T cell (Treg). DCs can be infected with HTLV-1,149,159,193,214 and these cells may play a significant part in the earliest stage of the acquisition of the infection. HTLV-1–infected DCs can immortalize CD4+ T cells, indicating that DCs can transmit the virus to CD4+ T cells. DCs can also infect other cells through the virological synapse.27 Cell-free HTLV-1 infection of DCs has been demonstrated in vitro: this process is enhanced by DC-specific ICAM-3–grabbing nonintegrin (DC-SIGN), which interacts with ICAM-2 and ICAM-3 but not HTLV-1 Env. However, since infectious cell-free HTLV-1 virions are rare or absent in vivo, the physiological importance of cell-free infection is doubtful. HSPG and NRP-1 are involved in the transfer of HTLV-1 from DCs to CD4+ T cells.

DCs are also the most important producer of type 1 interferons. The small numbers of DCs compared with T cells in both the circulation and the tissues may limit the importance of this cell type during chronic infection. However, DCs efficiently elicit a cytotoxic T-lymphocyte response, and DCs treated with HTLV-1 Tax peptides can elicit a CTL response that causes at least a temporary reduction in the PVL of HTLV-1 in both rats7 and humans.339

The role of the NK cell in HTLV-1 infection is also poorly understood. Both NK and NK-T cells are reduced in frequency in the blood in HTLV-1 infection,75,304,417 especially in patients with HAM, but the importance of this phenomenon is unknown.

HTLV-1 induces the expression of the transcription factor FoxP3, which is characteristic of Tregs312; in addition, infected cells frequently express the surface markers CD25, GITR, CTLA4, and CCR4 that are also found on Tregs. These molecules are also frequently expressed in ATL clones: FoxP3 is found in approximately 36% of cases.167 However, HTLV-1–infected FoxP3+ cells lack regulatory function,5,370 so the infected cell is not a true regulatory T cell, and ATL is not necessarily a tumor of Tregs.371 HTLV-1 also induces expression of the chemokine CCL22,120,372 which binds the receptor CCR4 that is found on true Tregs. By this means, HTLV-1 maintains a high frequency of CCR4+ cells in the host, which have Treg function. This may represent an adaptation by the virus to reduce the efficacy of the cell-mediated immune response, thereby favoring its persistence in the host.

Acquired Immune Response Antibody response In primary infection, antibodies to the core (Gag) protein of HTLV-1 are the first to appear, and they predominate the first 2 months.230 Antibodies to Env appear later, and finally antibodies to the Tax protein are detected in about 50% of individuals.230,256 The titre of HTLV-1– specific antibodies can reach very high levels in chronic infection, consistent with the notion that viral antigen synthesis persists indefinitely in vivo. It is likely that a high titre of HTLV-1–specific antibody simply results from a high PVL. But the role of antibody in either protection against infection or the pathogenesis of the associated diseases remains uncertain. Passively transferred antibody can inhibit milk-borne transmission of HTLV-1 in rabbits315 and in vitro infection of cord blood lymphocytes by co-culture with HTLV-1–infected cells can be prevented by HTLV-1 seropositive plasma from cord blood.352 However, there is no strong evidence that antibodies provide effective protection against HTLV-1 infection in the human.

Cell-mediated immune response HTLV-1 infection elicits a strong cell-mediated immune response, with abundant circulating CD8+ (cytotoxic) T lymphocytes (CTLs)148,290 and CD4+ (helper) T cells100 specific to the viral antigens. Activated CTLs kill virus-infected, MHC-matched target cells. By contrast, memory CTLs, which persist for months or years following an active infection, are not directly cytolytic but require reexposure to their cognate viral antigen to restimulate the cell and render it capable of cell-mediated killing. However, CTLs isolated from the peripheral blood of a person with nonmalignant HTLV-1 infection—both patients with HAM and asymptomatic carriers—can kill HLA-matched HTLV-1–infected target cells immediately ex vivo.112,290 This observation implied that HTLV-1 antigens are frequently expressed in vivo: the implications for the understanding of the regulation of HTLV-1 latency are discussed in Clonality of HTLV-1 and HTLV-2.

The HTLV-1 Tax protein is highly immunodominant in the CTL response to the virus99,148,162; activated CTLs specific to Gag and Pol are also frequently detectable in freshly isolated PBMCs.

CTLs are usually necessary to eradicate a transient virus from the body: in the absence of CTLs, a normally short-lived virus becomes persistent. CTLs also play a central part in limiting the replication of persistent viruses, such as Epstein-Barr virus or cytomegalovirus. Evidence that CTLs determine the outcome of HTLV-1 infection came from immunogenetic studies of a population with endemic HTLV-1 infection in Japan.151,392 In this population, individuals carrying either of the class 1 HLA genes HLA-A*02 or HLA-Cw*08 had a significant reduction in both the PVL and the risk of HAM151 compared with those who lacked both these alleles. Since class HLA proteins determine the specificity and activity of CTLs, these observations show that the genetically determined efficiency or “quality” of the HTLV-1–specific CTL response limits HTLV-1 615

replication in vivo.18,277 Calculation of the prevented fraction of disease shows that the HLA class 1 protective effect prevents about 50% of potential cases of HAM in this population in southern Japan.151

Further evidence of the protective role of CTLs came from two observations. First, heterozygosity in HLA class 1 genes, which is associated with a broader antigenic repertoire of CTLs, is associated with a lower PVL in the southern Japanese population.151 Second, it was discovered that inhibitory genes of the KIR complex enhance the effectiveness of the CTL response to persistent viruses. This effect was discovered in HTLV-1 infection and then generalized to hepatitis C virus and HIV-1 infections.36,318

Because Tax is the dominant target antigen recognized by the abundant, chronically activated HTLV-1–specific CTLs present in the circulation, it was natural to assume that the anti-Tax CTL response restricted by the common allele HLA-A*0201 was critical in this protective response. However, Boelen et al.36 showed that the protective effect of the A*02-restricted CTLs was effected via the HLA-A*0207 allele, not the more frequent HLA-A*0201 allele.

The most important determinant of the PVL and the risk of HAM is the CTL response not to the immunodominant Tax protein, but rather to the HBZ protein.123,215,341 This observation is counterintuitive, because HBZ is a weak CTL antigen, and is expressed at a very low level in the infected cell. However, it is now known that whereas each HTLV-1–infected cell expresses tax in uncommon, intermittent bursts,32,221 it expresses HBZ about 50% of the time.243 Further, HBZ expression is required for clonal persistence in vivo.359 It appears that HTLV-1 has evolved to limit the expression of this critical gene to the minimal level required for survival in vivo, while minimizing its exposure to immune surveillance.

In addition to the intermittent plus-strand expression and the low level and low immunogenicity of the HBZ protein, HTLV-1 uses further tactics that may contribute to immune evasion. First, the HTLV-1 accessory protein p12, which is expressed by some strains of HTLV-1, physically interacts with the free human MHC class I heavy chain and induces its degradation,157 facilitating the escape of HTLV-1–infected cells from destruction by CTLs. Second, Tax induces the production of CCL22 by infected T cells,372 which maintains a higher frequency of circulating FoxP3+ Tregs; in turn, the high FoxP3+ Treg frequency is correlated with reduced efficiency of CTL-mediated killing of HTLV-1–infected cells by CTLs.370

Study of the helper T-cell response to HTLV-1 is complicated by the fact that it is the dominant host cell for the virus, and expression of the virus alters the function of the infected cell. However, a short-term ELISpot assay made it possible to distinguish between activation of the cell by exogenous viral antigen from activation by the expression of Tax inside the cell.100,101 The results show that the frequency of HTLV-1–specific CD4+ T cells was significantly higher in patients with HAM than in asymptomatic carriers with a similar PVL.100 Two further observations corroborate the implication that CD4+ T cells play a part in the pathogenesis of HAM. First, these cells predominate in the early stage of the inflammatory lesion in the central nervous system.377 Second, the class 2 MHC allele DRB1*0101 is associated with a higher risk of HAM.151,185,273,337,382

Immunodeficiency and HTLV-1-Associated Diseases Thus, the host immune system controls the dynamics of HTLV-1 infection and the development of disease. This idea is supported by the clinical observation that ATL occurs more commonly in immunocompromised hosts. Among 24 patients with posttransplantation lymphoproliferative disorders (PT-LPDs) after renal transplantation in Japan, five ATL cases were reported.133 In these cases, HTLV-1 was probably transmitted by blood transfusion during hemodialysis. Of eight HTLV-1 carriers who received living-donor liver transplants and subsequent immunosuppressive treatment, three developed ATL.174 These findings indicate that the host immune system commonly suppresses the development of ATL, while an impaired immune system may allow HTLV-1–infected cells to transform into ATL cells.

The impairment of cell-mediated immunity is profound in ATL patients, who frequently succumb to opportunistic infections with various pathogens, including various fungi, cytomegalovirus, Pneumocystis jirovecii, and mycobacteria.355 Thus, immune impairment is both a cause and a consequence of ATL.

Individuals with nonmalignant HTLV-1 infection, both asymptomatic and those with inflammatory diseases such as HAM, also show a degree of impairment of cell-mediated immunity, as evidenced by a higher incidence of a range of opportunistic infections and malignancies.317 The associated infections include severe Strongyloides stercoralis infection390; tuberculosis389; infective dermatitis33; bronchiectasis67; and infections of the lung, kidney, and bladder.317 The associated malignant diseases include liver cancer, cervical cancer, and lymphoma (distinct from ATL).317

Within-Host Evolution: Selection In vivo In a persistent infection, the immune response exerts a strong selection pressure on the infectious agent, which in turn develops adaptations that lessen its susceptibility to immune-mediated destruction. In HTLV-1 infection, the strong CTL response (see “Immune Response”) selects immune escape variants,270,271 but the inability of HTLV-1 to tolerate major sequence change limits the outgrowth of such variant sequences. Instead of immune escape, HTLV-1 relies more for its persistence in vivo on the regulation of its transcriptional latency (see “Clonality of HTLV-1 and HTLV-2”) to minimize its exposure to the immune system.

Each clone of infected T cells possesses characteristics of HTLV-1 expression that depend in part on the genomic integration site of the provirus.32,239 Consequently, there is selection in the host for the survival of HTLV-1–infected clones with particular characteristics, which are only beginning to be understood. Initial integration of HTLV-1 occurs in each chromosome in proportion to the size of the chromosome, but survival of that clone in vivo is favored if the provirus is integrated in one of the acrocentric chromosomes, especially chromosome 13.52 It is possible that this selective survival is associated with the intranuclear location of acrocentric chromosomes, which are characteristically found during interphase at the periphery of the nucleolus, which is a transcriptionally repressive region.

Expression of the tax gene is lost in the malignant clone in approximately 60% of cases of ATL.356 Three mechanisms by which tax expression was disrupted were identified: (a) deletion of the 5′LTR,246,358 (b) DNA methylation of the 5′LTR,188,360 and (c) genetic changes in the tax gene 616

(nonsense mutations, deletion, and insertion).70,79 In all cases, the pX region and the 3′LTR remained intact, suggesting the importance of the HBZ gene in the survival of the malignant clone.

Analysis of whole HTLV-1 proviruses in ATL cells shows that all the viral genes can contain nonsense mutations except the HBZ gene.70 Most nonsense mutations were generated in a tryptophan codon (TGG) by G-to-A mutation (TGG to TGA, or TGG to TAG). The sequences of these G-to-A mutations in HTLV-1 proviruses correspond to the target sequences of APOBEC3G.70 APOBEC3G is a host factor that generates G-to-A mutations during reverse transcription. In the case of HIV-1, an accessory protein, Vif, promotes the degradation of APOBEC3G, which enables HIV-1 to escape fatal nonsense mutations. By contrast, HTLV-1 does not encode an accessory gene that counteracts APOBEC3G. The Gag protein of HTLV-1 suppresses the incorporation of APOBEC3G into the virion,61 but this inhibitory activity is not strong enough to completely suppress all G-to-A mutations.308 If nonsense mutations are generated in viral genes that are not essential for cellular proliferation, HTLV-1– infected cells can still proliferate and transform to leukemic cells. This result indicates that nonsense mutations of all viral genes except the HBZ gene can be generated before integration and ATL can still ensue, suggesting that HBZ is an essential gene for leukemogenesis.

Pathogenesis of ATL HTLV-1 infection is a necessary cause of ATL, but it is not sufficient: the lifetime risk of ATL in HTLV-1–infected people is about 5%,142 and the disease usually presents clinically after several decades of HTLV-1 infection. The median age at diagnosis is between 40 and 50 years in the Caribbean and South America.128 In Japan, the median age at presentation has risen progressively from 56.9 years in the mid-1980s to 67.5 years in 2020,144 suggesting that a cohort of the population developed a high prevalence of HTLV-1 infection in the 1920s and 1930s.

Two broad types of oncogenic mechanism must be considered in ATL: first, insertional mutagenesis, and second, transactivation by viral gene products. HTLV-1 usually infects approximately 104 clones in each individual host; in some cases, this number exceeds 106.199 Since each clone contains a provirus in a unique genomic location, the potential for insertional mutagenesis is great. The provirus may disrupt host gene expression in cis in three ways. First, if the provirus is inserted inside a host gene, the resulting disruption of gene expression may be oncogenic, if the gene acts as a tumor suppressor. However, there is no evidence of this mechanism in ATL. Second, the provirus can deregulate host transcription immediately adjacent to the integration site.241 Third, the provirus can alter the pattern of chromatin looping of the host genome. HTLV-1 binds the zinc finger protein CTCF,311 which plays a central part in forming the chromatin loops that regulate the structure and expression of the genome.294 CTCF binds to a DNA motif of approximately 20 nucleotides at approximately 50,000 sites in the human genome. By dimerizing with CTCF bound to a nearby site in the host chromatin, HTLV-1–bound CTCF brings the provirus near host genes that are distant (up to several Mb) from the integration site in the linear genome and can alter host transcription at these distant sites.241

Which of these three types of proviral mutagenesis might contribute to the oncogenesis of ATL? There are no hotspots of HTLV-1 integration in the host genome in ATL,52 so direct insertional mutagenesis of host genes is not a major cause of ATL. However, integration within 10 kb upstream of a group of 11 host genes was significantly overrepresented in a study of 197 ATL cases, accounting for 6% of the cases.52 These 11 genes are associated with three ontological categories: cell morphology, immune cell trafficking, and hematological system development and function, and they have been implicated in other malignancies. Thus, deregulation of nearby host genes by the provirus may contribute to oncogenesis in some cases. It is not yet known whether the observed deregulation of more distant host genes in cis that are brought into proximity by CTCF-mediated chromatin looping also plays a part in ATL oncogenesis.

Most attention in the study of ATL oncogenesis has been focused on the two chief regulatory genes of HTLV-1, tax and HBZ, because the products of these genes exert profound diverse effects on gene expression and function of the host cell (see “Tax and Rex: Main Physiological Actions”). Transgenic mice expressing either tax or HBZ develop tumors, and tax can immortalize T cells in vitro. However, neither tax nor HBZ is an acutely transforming oncogene such as myc, ras, or src. Further, as mentioned, tax is expressed in each infected cell in uncommon, intermittent bursts.32,221 In approximately 25% of ATL cases, Tax expression is disabled by nonsense mutations of the tax gene or loss of the 5′ LTR at the time of HTLV-1 infection.356 These ATL cells are transformed and persist without Tax; they depend on HBZ expression. In approximately 50% of ATL cases, Tax is transiently expressed: these ATL cells depend on both Tax and HBZ. Taken together, these findings indicate two different types of requirement for viral genes for leukemogenesis in ATL: one is HBZ dependent, and the other type needs both Tax and HBZ.274

Kataoka et al.170 identified the host genes that are most commonly mutated in ATL, in a study of 426 cases. The most frequent mutations were found in genes (including PLCG1, PRKCB, CARD11, etc.) that participate in pathways highly active in T cells, including T-cell receptor–NF-κB signaling and T-cell trafficking. These putative oncogenic driver mutations can be detected in the circulation in some cases several years before clinical presentation.301 The mutational burden in infected T cells 1 year before diagnosis of ATL is significantly greater than in age-matched HTLV-1 carriers who remained healthy during 10 years of follow-up.301

These observations suggest that the dominant mechanism of oncogenesis in ATL is the accumulation of replicative mutations due to the continued proliferation of HTLV-1–infected T-cell clones over many years. The risk of oncogenesis in a given cell type is proportional to the total number of cell divisions that a given cell lineage has undergone.367,368 HTLV-1–infected T-cell clones typically survive indefinitely96 and proliferate faster than uninfected T cells.15 While specific actions of tax and HBZ (see “Tax and Rex: Main Physiological Actions”) may exacerbate the risk of malignant transformation, their fundamental function is to promote the survival and transmission of the virus by maintaining the longevity of the infected T-cell clone. Thus, their chief contribution to oncogenesis is indirect, by promoting the accumulation of replicative mutations in long-lived clones.21

Pathogenesis of Inflammatory Diseases Up to 5% of HTLV-1–infected people develop a chronic inflammatory disease. The most frequently diagnosed of these conditions is HAM20 (see ‘History of HAM’ above). HTLV-1 also causes chronic inflammatory conditions in many other tissues, in particular uveitis,248 infective dermatitis,201 polymyositis,252 and bronchiectasis,67 which also has an infective component in its pathogenesis.

The risk of these inflammatory diseases is strongly correlated with the PVL of HTLV-1.260,317 Certain strains of HTLV-1 are associated with a slightly higher risk of HAM in Japan,78 but the bulk of the variation in individual risk is attributable to the host. The reason for the correlation 617

between PVL and the risk of HAM is likely to be the less efficient cytotoxic T-cell response to HTLV-1 in these individuals18 (see “Immune Response”), which is associated with both a higher PVL, a higher risk of HAM151 and a greater number of HTLV-1–expressing cells at a given time.14

Helper (CD4+) T cells predominate in early lesions of HAM.143,147,377 As the disease progresses, the proportion of CD8+ T cells in the lesion increases. It is not understood what initiates the infiltration of T cells across the blood–brain barrier or into other tissues. However, HTLV-1 Tax induces strong expression of ICAM-1, which increases the adhesion and migration of T cells across epithelial barriers. In HAM, a mechanism of self-perpetuating inflammation has been proposed.6 The secretion of IFNγ by T cells stimulates the secretion by astrocytes of the chemokine CXCL10 (IP-10), which attracts more T cells that express the cognate receptor CXCR3. It is not known whether a similar mechanism operates in the other inflammatory conditions associated with HTLV-1.

The mechanism of tissue damage is not known with certainty. There is no convincing evidence that the resident cells in the inflamed tissues are themselves infected with HTLV-1, and antibodies that cross-react between Tax protein and a self-antigen209 are not specific to HAM.419 The cell damage is thought to be caused by inflammatory substances, in particular IFNγ, secreted by the invading T cells. HTLV-1–infected CD4+ T cells express high levels of IFNγ,111 and the frequency of IFNγ+CD4+ T cells is especially high in patients with HAM.100 In mature lesions in HAM, it is likely that the invading CD8+ T cells contribute further inflammatory substances such as IFNγ and TNFα.

EPIDEMIOLOGY Age A high risk of mother-to-child transmission is associated with prolonged breast-feeding (>12 months), a high PVL (>1% PBMCs) and seropositivity in more than one child of the same mother.287 In adults, HTLV-1 seroprevalence rises with age, and the seropositivity rate in females is higher than that in males,34 because sexual transmission of this virus is more efficient from male to female than from female to male.134,338 Sexual transmission may occur even in old HTLV-1 discordant couples, which can account for the continuous increase in seropositivity rate in the older population. As mentioned, the average age of ATL patients at diagnosis was 63 years in Japan, which is 15 years older than the average age of ATL patients at presentation in the Caribbean islands,355 suggesting that genetic background or environmental factors—or both—influence the development of ATL. The mean age at onset of HAM/TSP patients in Japan was 43 years.283 In addition, HAM/TSP can develop within a few months after infection,369 whereas ATL typically develops after a long clinically latent period in nonimmunosuppressed individuals.

Morbidity/Mortality HTLV-1 establishes a persistent infection. There are geographical differences in the incidences of these HTLV-1–associated diseases.128 The cumulative lifetime risk of ATL among HTLV-1 carriers in Japan is about 6.6% for men and 2.1% for women; most HTLV-1 carriers remain asymptomatic throughout their lives.11 By contrast, the risk of ATL is four times lower in Jamaicans. The lifetime risk of HAM/TSP in HTLV-1 carriers has been estimated as 0.25% in southern Japan,166 but the risk in populations of Afro-Caribbean descent is about 10 times greater (1.9% to 3.7%).226,282 The reasons for these surprisingly large differences between populations in the risk of ATL and HAM/TSP are not well understood. In addition to ATL and HAM/TSP, HTLV-1 is associated with a significantly increased risk of a wide range of diseases,317 especially seborrheic dermatitis, Sjögren syndrome, and lung disease (bronchiectasis, bronchitis, and bronchiolitis). HTLV-1 infection also predisposes to a number of infections with other pathogens, notably TB and Strongyloides stercoralis, and to infections of the kidney and bladder. Finally, the risk of liver cancer, cervical cancer, and lymphoma (other than ATL) is greater in HTLV-1–seropositive persons, but the risk of gastric cancer is reduced. The adjusted risk of death from any cause is approximately 1.57 times higher in HTLV-1–infected individuals than in seronegative individuals.317

Origin and Spread of Epidemics HTLV and STLV are thought to originate from common ancestors and share molecular, virological, and epidemiological features. Therefore, they have been designated primate T-cell leukemia viruses (PTLVs). Phylogenetic analyses have revealed that HTLV-1c first diverged from STLV around 50,000 ± 10,000 years ago, while the spread of HTLV-1 in Africa is estimated to have occurred at least 27,300 ± 8,200 years ago. Subsequently, HTLV-1a, which is the most common subtype in Japan, diverged from the African strain 12,300 ± 4,900 years ago.384 Thus, these viruses have had a long history with humans after the initial interspecies transmission. By contrast, HIV-1 is thought to originate from simian immunodeficiency virus in chimpanzees (SIVCPZ),82 and the interspecies transmission to humans is estimated to have occurred about one hundred years ago.399 Recently, new retroviruses have been identified in bushmeat hunters in central Africa. HTLV-3 has been found in asymptomatic carriers in Cameroon.40,398 HTLV-3 is highly homologous to its simian counterpart, STLV-3, which was detected in monkeys in Africa. The genome organization of HTLV-3 is similar to HTLV-1.222 No diseases have been reported to be associated with HTLV-3.

Prevalence and Seroepidemiology It is estimated that at least 5 to 10 million people live with HTLV-1 worldwide.89 However, this number remains highly uncertain, because systematic epidemiological studies have not been carried out in some regions endemic for the virus. The chief known endemic areas are southwestern Japan, the Caribbean basins, Central and West Africa, and South America. In addition, epidemiological studies of HTLV-1 have revealed high seroprevalence rates in Melanesia, Papua New Guinea, and the Solomon islands, as well as among Australian aborigines.34,68 In Japan, approximately 1.08 million individuals were estimated to be infected with HTLV-1 in 2009, and among them, 1,000 carriers develop ATL each year. The rate of mother-to-child transmission of HTLV-1 in Japan has declined, following the introduction of a nationwide policy of discouraging breast-feeding by seropositive mothers. However, the rate of sexual transmission among adolescents and adults has maintained the annual number of new cases of infection at >4000.309

The epidemiology of HTLV-2 is quite distinct.108 The main populations where HTLV-2 is endemic are in North America, where it is prevalent in 618

certain Native American groups and intravenous drug user groups,259 and in Brazil.69 The total number of people infected with HTLV-2 in the world is estimated to be 6- to 12-fold lower than the number infected with HTLV-1,259 although as in HTLV-1, this estimate is somewhat uncertain, because many areas have not been systematically studied.

FIGURE 16.11 The natural course of HTLV-1 infection. After transmission of HTLV-1, infected cells proliferate owing to the actions of HBZ and Tax. The growth of HTLV-1–infected cells is suppressed by cytotoxic T lymphocytes in vivo. The number of HTLV-1–infected cells is thus determined by balance between viral gene expression and the host immune system. About 5% of HTLV carriers develop adult T-cell leukemia after a long latent period. A smaller fraction of carriers present with inflammatory diseases, such as HTLV-1–associated myelopathy/tropical spastic paraparesis (HAM/TSP), uveitis, or alveolitis. Most carriers remain asymptomatic.

Genetic Diversity of HTLV-1 Several subtypes of HTLV-1 have been identified: the cosmopolitan HTLV-1a subtype244; the Central African subtypes HTLV-1b,105,386 HTLV-1d,223 HTLV-1e,334 and HTLV-1f307; and the Australo-Melanesian subtype HTLV-1c.91,324 These subtypes are closely linked with interspecies transmission and human migration events. The sequence of HTLV-1 does not differ systematically between asymptomatic carriers and those with ATL or HAM/TSP,55 although small differences in the risk of HAM/TSP have been associated with certain genotypes.78

CLINICAL FEATURES During chronic infection with HTLV-1, the PVL remains relatively constant within an individual but can differ between infected individuals by over 1,000-fold. Most individuals remain asymptomatic during their entire lives, while a small fraction of carriers develop either ATL or one of the HTLV-1–associated inflammatory diseases (Fig. 16.11) (317; see “Morbidity/Mortality”).

ATL ATL develops more frequently in males (the male to female ratio is 1.5:1). The age at onset ranges from 25 to 94 years. The predominant physical findings are peripheral lymph node enlargement (72% of cases), hepatomegaly (47%), splenomegaly (25%), and skin lesions (53%).355 ATL cells infiltrate into various organs/tissues, including skin, liver, lung, the gastrointestinal tract, the central nervous system, and bone. Various skin lesions, such as papules, erythema, and nodules are frequently observed in ATL patients (Fig. 16.12). ATL cells densely infiltrate the dermis and epidermis, forming Pautrier microabscesses in the epidermis (Fig. 16.13).

FIGURE 16.12 Skin lesion of an adult T-cell leukemia (ATL) patient. ATL cells frequently involve skin and form various lesions. In this patient, ATL cells form a tumor lesion. ATL cells in the peripheral blood have characteristic indented or lobulated nuclei (Fig. 16.14): such cells are known as “flower cells.” Anemia and thrombocytopenia are rare since involvement of bone marrow is not severe. The typical surface phenotype of ATL cells is CD4+CD8-CD25+, which is similar to that of Treg cells. ATL cells frequently express FoxP3168 (Fig. 16.15) and other markers characteristic of Treg cells or activated T cells; however, the phenotype is somewhat variable, and ATL should not be regarded as a tumor of Treg cells.371 ATL cells often express CD25 (IL-2 receptor alpha chain) on the cell surface and secrete its soluble form; consequently, the concentration of soluble IL-2 619

receptor is abnormally high in ATL and is correlated with the tumor mass and clinical course.409 ATL cells elaborate various cytokines that can affect the immune response and influence the pathophysiology of ATL. For example, eosinophilia, caused by elevated IL-5 levels, is frequently observed in patients with ATL.

Hypercalcemia is a complication in about 70% of ATL patients at some point during the clinical course of the disease, particularly during the aggressive stage.186 Pathological studies of ATL patients with hypercalcemia have shown that high serum Ca2+ levels are due to an increased number of osteoclasts and accelerated bone resorption (Fig. 16.16). During differentiation of osteoclasts, precursor cells sequentially express c-Fms (the receptor for M-CSF) followed by receptor activator nuclear factor kB (RANK).9 M-CSF and RANK ligand (RANKL) have been shown to be critical factors for the differentiation of osteoclasts and are physiologically produced by stromal cells and osteoblasts. ATL cells from patients with hypercalcemia have high RANKL transcript levels and induce the differentiation of hematopoietic stem cells into osteoclasts in vitro in the presence of M-CSF.275 In addition, parathyroid hormone–related protein (PTH-rP) elaborated from ATL cells activates osteoclasts and promotes bone resorption.393

FIGURE 16.13 Skin involvement of adult T-cell leukemia (ATL). A: ATL cells infiltrate into epidermis and form Pautrier microabscesses (arrows). B: CD4+ cells are identified by the immunohistochemical staining.

HTLV-1–Associated Myelopathy/Tropical Spastic Paraparesis (HAM/TSP) HAM/TSP is characterized by a slow progressive spastic paraparesis, urinary dysfunction, and sensory disturbances.20,283 HAM/TSP patients also often have other organ disorders such as leukoencephalopathy (69%), abnormal findings on chest X-ray (50%), Sjögren syndrome (25%), and arthropathy (17%).262 In contrast to ATL, HAM/TSP develops predominantly in females (the male to female ratio is 1:2.9).

Pathological studies of HAM/TSP patients demonstrate severe involvement of the thoracic spinal cord, in which T cells cause perivascular cuffing and infiltrate into the parenchyma.283 In HAM/TSP, the PVL is typically higher than in asymptomatic carriers,260 and HTLV-1–infected CD4+ T cells infiltrate into the spinal cord.42,234,254 The number of HTLV-1–specific HLA class I restricted CD8+ T cells is also increased in HAM/TSP patients.22 Among viral antigens, Tax has a dominant epitope that is recognized by CTLs.99,162 Tax-reacting CD8+ T cells also migrate into the spinal cord along with HTLV-1–infected CD4+ T cells.378 These HTLV-1–infected cells and reacting CTLs produce proinflammatory cytokines (TNFα, IFNγ, IL-1β), leading to demyelination and axonal damage.

FIGURE 16.14 Adult T-cell leukemia (ATL) cells in the peripheral blood. A and B: ATL cells in an acute ATL case show the characteristic morphology.

HTLV-1 Uveitis HTLV-1 infection induces uveitis, an inflammatory disorder of the intraocular tissues. The most characteristic findings in HTLV-1 uveitis are vitreous opacities associated with mild iritis and mild retinal vasculitis: this type is classified as intermediate uveitis, since the intermediate part 620

of the eyeball is affected by the inflammation.249 Infiltrating HTLV-1–infected cells are implicated in the pathogenesis of this uveitis. The standard treatment of the condition is topical or systemic corticosteroids.

Infective Dermatitis Infective dermatitis (ID), a severe, chronic, relapsing dermatitis first described in Jamaican children, was found to be associated with HTLV-1 infection.197 ID is characterized by a severe exudative dermatitis with crusting of the scalp, neck, external ears, axillae, and groin. In the affected skin region, Staphylococcus aureus or beta-hemolytic Streptococcus infection is typically detected. Oral trimethoprim–sulfamethoxazole is the treatment of choice because of low cost and effectiveness.220 Patients with ID tend to have an elevated provirus load. ID has been reported prior to the development of ATL and HAM/TSP; ID is, therefore, implicated as an indicator for other HTLV-1–associated diseases.

FIGURE 16.15 Lymph node lesion of an adult T-cell leukemia (ATL) patient. A: Histological analysis (hematoxylin–eosin staining) shows monotonous proliferation of ATL cells. B: Immunohistochemical analysis shows that ATL cells are FoxP3 positive.

PREVENTION AND TREATMENT Prevention Transmission of HTLV-1 between individuals requires the transfer of living infected cells. All routes are preventable. Mother-to-infant transmission can be reduced by bottle-feeding or freeze–thaw processing of breast milk,8 which kills the infected cells in the breast milk. However, HTLV-1 transmission still occurs in about 5% of infants who do not drink breast milk, indicating the presence of a route of transmission other than breast-feeding.350 As a mechanism, HTLV-1 infection was found in placental villous tissues, suggesting that infected trophoblast cells serve as viral reservoirs.364 The risk of transmission by breast-feeding was found to increase with the degree of concordance of the HLA genotype between mother and child,31 suggesting that immunological responses influence the transmission from mother to child. Blood screening almost completely prevents viral transmission by blood transfusion.138 RTIs should be useful for preventing de novo infection by accidental exposure to contaminated blood, although no clinical evidence has yet been reported.

FIGURE 16.16 Increased number of osteoclasts in a hypercalcemic adult T-cell leukemia patient. In a hypercalcemic patient, the number of osteoclasts increases in the bone and accelerates bone resorption.

Treatment Diagnosis HTLV-1 infection can be diagnosed by the presence of antibody to HTLV-1. The presence of serum antibodies against HTLV-1 can be demonstrated by enzyme-linked immunosorbance assay (ELISA), gelatin particle hemagglutination, indirect immunofluorescence, and western blot assays. The provirus load, which represents the percentage of PBMCs infected with HTLV-1, is measured by real-time PCR or droplet digital PCR (ddPCR). In HTLV-1–infected individuals, the provirus load ranges from less than 0.01% up to more than 50% PBMCs (each cell typically carries a single copy of the HTLV-1 provirus53).

The diagnostic criteria for ATL have been defined as follows50,327: (a) histologically and/or cytologically proved lymphoid malignancy with T-cell surface antigens; (b) abnormal T cells present in the peripheral blood (except for lymphoma-type ATL). These abnormal T lymphocytes can include not only typical ATL cells, the so-called flower cells, but also the small and mature T lymphocytes with incised or lobulated nuclei that are characteristic of the chronic or smoldering type of ATL; (c) antibody to HTLV-1 present in the serum at diagnosis; and (d) demonstration of monoclonal integration of HTLV-1 provirus by the Southern blot method.

ATL is a poorly treatable neoplastic disease due to its resistance to anticancer drugs and the complication of immunodeficiency. Patients with acute or lymphoma-type ATL are usually treated with combination chemotherapy. With the most commonly used combination chemotherapy in Japan (VCAP-AMP-VECP therapy: vincristine, cyclophosphamide, doxorubicin, and prednisone (VCAP), doxorubicin, ranimustine, and prednisone [AMP], and vindesine, etoposide, carboplatin, and prednisone [VECP]),402 81% of the 93 eligible patients responded, with 33 621

patients obtaining complete response (35.5%) and 42 obtaining partial response (45.2%). The median survival time of patients was 13 months. Thus, the prognosis of ATL patients treated by chemotherapy remains poor.171 The major impediments in therapy for ATL patients are the drug resistance of ATL cells to chemotherapeutic agents and the profoundly immunodeficient state of patients.328 Cell-mediated immunity is impaired in ATL patients, whereas humoral immunity remains intact. The immunodeficiency of ATL patients may be caused by the immunosuppressive function of ATL cells, since they express immunosuppressive molecules,359 frequently including many molecules that are characteristic of Tregs. The immunodeficiency of ATL patients often leads to opportunistic fungal, viral, and bacterial infections, which worsens the prognosis.355 Successful allogeneic bone marrow transplantation was reported for an acute ATL patient in 1996.39,366 In a recent systematic review of 18 studies, the overall survival rate after transplantation was approximately 40% and the relapse rate was 36%.140 ATL patients with chronic graft versus host disease (GVHD) show a better prognosis than those without chronic GVHD, indicating that host immune responses play critical roles.161 Thus, immune responses are critical for treatment of ATL. The HTLV-1 provirus load decreased remarkably in many patients who received an allogeneic stem-cell transplant,279 suggesting that cellmediated immunity to HTLV-1 was enhanced in these patients, an observation which might account for the efficacy of this therapy. ATL cells express high levels of cell-surface Fas antigen and are susceptible to Fas-mediated signaling.357,411These findings suggest that ATL cells are vulnerable to CTLs, which accounts for the good therapeutic responses. In individuals who received alloSCT, CTLs to Tax peptides were activated in the recipients; the provirus load became profoundly suppressed, suggesting a role of anti-HTLV-1 immune responses in the efficacious outcome.113 Tax-peptide pulsed dendritic cells could suppress ATL.339 Thus, Tax is a promising target for treatment of ATL in the approximately 50% of cases in which Tax is expressed by the malignant clone. ATL cells express CCR4 on their surfaces.414 ATL cells resemble Treg cells, which is because HBZ induces Foxp3. Treg cells express CCR4. Furthermore, HBZ also induces transcription of the CCR4 gene,340 and HTLV-1–infected cells express CCR4.405 Mogamulizumab, a defucosylated humanized anti-CCR4 antibody, has shown to be effective in ATL patients.141,404 Mogamulizumab not only kills ATL cells via enhanced antibody-dependent cell-mediated cytotoxicity (ADCC) but also enhances antiviral immunity through suppression of Treg cells.342 Previous studies reported that the prognosis of patients with aggressive subtypes of ATL (acute and lymphoma types) was less than 1 year, whereas that of indolent types (chronic and smoldering types) was much longer.327 Later studies indicated that the prognosis of the indolent subtypes of ATL, chronic and smoldering ATL, is poorer than previously thought: the mean survival time of patients was only 4.1 years.353 Therapy using IFN-α combined with zidovudine has been reported to be highly effective for these indolent ATL patients.28 However, viral replication has not been demonstrated in ATL cells. Instead, many nonsense mutations, deletion, and insertions of HTLV-1 provirus were reported in ATL cells.70,356 These data suggest that IFN-α combined with zidovudine may suppress the growth of ATL cells without any effects on viral replication. A systematic review of therapies for HAM/TSP10 concluded that corticosteroids can provide significant benefit: pulsed, high-dose methylprednisolone may be effective in induction therapy, and low-dose oral prednisolone can be used in maintenance therapy. However, the benefit of other types of treatment is less clear. HAM/TSP is associated with a high PVL. Since nonmalignant HTLV-1–infected cells also include CCR4-like ATL cells,405 the anti-CCR4 monoclonal antibody mogamulizimab was used to treat HAM/TSP. The treatment decreased the number of HTLV-1–infected cells, with an associated reduction of spasticity.310 However, the clinical benefit of this treatment may be short lived. As expected, RTIs can suppress replication of HTLV-1 in vitro, although the efficacy of various RTIs against HTLV-1 differs from that against HIV-1.124 RTIs can also block HTLV-1 infection if they are administrated at the time of HTLV-1 exposure.235 However, RTIs cannot suppress HTLV-1 infection if they are administered 1 week later.247 This finding suggests that clonal proliferation of infected cells is predominant after the initial in vivo spread of HTLV-1200; it is consistent with the observation that RTIs do not change provirus load in patients with HAM.362 Thus, the clinical use of inhibitors against HTLV-1 replication is limited to preventive administration after accidental exposure to HTLV-1– positive blood.

Vaccines The passive transfer of immunoglobulin derived from HTLV-1–infected individuals could block HTLV-1 infection,253,315 suggesting that a vaccine to HTLV-1 is capable of inhibiting its transmission. Indeed, immunization with peptide derived from envelope protein264 and vaccines expressing envelope protein325 can block transmission of HTLV-1 in rabbits and monkeys. These findings show that a preventive vaccine for HTLV-1 is possible, in contrast with HIV-1 infection. Tax might be a target for a therapeutic vaccine due to its high immunogenicity. In a rat model, a Tax peptide-based vaccine inhibited the growth of a Tax expressing rat T-cell line in vivo.110 However, the fact that Tax expression is frequently lost in ATL cells limits the value of Tax for a therapeutic vaccine for ATL.

622

References 1. Adachi Y, Copeland TD, Takahashi C, et al. Phosphorylation of the Rex protein of human T-cell leukemia virus type I. J Biol Chem 1992;267:21977–21981. 2. Afonso PV, Cassar O, Gessain A. Molecular epidemiology, genetic variability and evolution of HTLV-1 with special emphasis on African genotypes. Retrovirology 2019;16:39. 3. Ahmed YF, Hanly SM, Malim MH, et al. Structure-function analyses of the HTLV-I Rex and HIV-1 Rev RNA response elements: insights into the mechanism of Rex and Rev action. Genes Dev 1990;4:1014–1022. 4. Albrecht B, DSouza CD, Ding W, et al. Activation of nuclear factor of activated T cells by human T-lymphotropic virus type 1 accessory protein p12(I). J Virol 2002;76:3493–3501. 5. Anderson MR, Enose-Akahata Y, Massoud R, et al. Epigenetic modification of the FoxP3 TSDR in HAM/TSP decreases the functional suppression of Tregs. J Neuroimmune Pharmacol 2014;9:522–532. 6. Ando H, Sato T, Tomaru U, et al. Positive feedback loop via astrocytes causes chronic inflammation in virus-associated myelopathy. Brain 2013;136:2876–2887. 7. Ando S, Hasegawa A, Murakami Y, et al. HTLV-1 tax-specific CTL epitope-pulsed dendritic cell therapy reduces proviral load in infected rats with immune tolerance against tax. J Immunol 2017;198:1210–1219. 8. Ando Y, Ekuni Y, Matsumoto Y, et al. Long-term serological outcome of infants who received frozen-thawed milk from human T-lymphotropic virus type-I positive mothers. J Obstet Gynaecol Res 2004;30:436–438. 9. Arai F, Miyamoto T, Ohneda O, et al. Commitment and differentiation of osteoclast precursor cells by the sequential expression of c-Fms and receptor activator of nuclear factor kappaB (RANK) receptors. J Exp Med 1999;190:1741–1754. 10. Araujo A, Bangham CRM, Casseb J, et al. Management of HAM/TSP: systematic review and consensus-based recommendations 2019. Neurol Clin Pract 2021;11:49–56. 11. Arisawa K, Soda M, Endo S, et al. Evaluation of adult T-cell leukemia/lymphoma incidence and its impact on non-Hodgkin lymphoma incidence in southwestern Japan. Int J Cancer 2000;85:319–324. 12. Armand M, Hamosh M, Mehta NR, et al. Effect of human milk or formula on gastric function and fat digestion in the premature infant. Pediatr Res 1996;40:429–437. 13. Arnold J, Zimmerman B, Li M, et al. Human T-cell leukemia virus type-1 antisense-encoded gene, Hbz, promotes T-lymphocyte proliferation. Blood 2008;112:3788–3797. 14. Asquith B, Bangham CR. Quantifying HTLV-I dynamics. Immunol Cell Biol 2007;85:280–286. 15. Asquith B, Zhang Y, Mosley AJ, et al. In vivo T lymphocyte dynamics in humans and the impact of human T-lymphotropic virus 1 infection. Proc Natl Acad Sci U S A 2007;104:8035–8040. 16. Ballard DW, Bohnlein E, Lowenthal JW, et al. HTLV-I tax induces cellular proteins that activate the kappa B element in the IL-2 receptor alpha gene. Science 1988;241:1652–1655. 16a.Banerjee P, Feuer G, Barker E. Human T-cell leukemia virus type 1 (HTLV-1) p12I down-modulates ICAM-1 and -2 and reduces adherence of natural killer cells, thereby protecting HTLV-1-infected primary CD4+ T cells from autologous natural killer cell-mediated cytotoxicity despite the reduction of major histocompatibility complex class I molecules on infected cells. J Virol 2007;81:9707–9717. 17. Bangham CRM. The immune control and cell-to-cell spread of human T-lymphotropic virus type 1. J Gen Virol 2003;84:3177–3189. 18. Bangham CRM. CTL quality and the control of human retroviral infections. Eur J Immunol 2009;39:1700–1712. 19. Bangham CRM. Human T cell leukemia virus type 1: persistence and pathogenesis. Annu Rev Immunol 2018;36:43–71. 20. Bangham CRM, Araujo A, Yamano Y, et al. HTLV-1-associated myelopathy/tropical spastic paraparesis. Nat Rev Dis Primers 2015;1:15012. 21. Bangham CRM, Matsuoka M. Human T-cell leukemia virus type 1: parasitism and pathogenesis. Philos Trans R Soc Lond B Biol Sci 2017;372:20160272. 22a.Barnard AL, Igakura T, Tanaka Y, et al. Engagement of specific T cell surface molecules regulates cytoskeletal polarization in HTLV-1infected lymphocytes. Blood 2005;106:988–995. 22. Bangham CRM, Osame M. Cellular immune response to HTLV-1. Oncogene 2005;24:6035–6046. 23. Barski MS, Minnell JJ, Hodakova Z, et al. Cryo-EM structure of the deltaretroviral intasome in complex with the PP2A regulatory subunit B56gamma. Nat Commun 2020;11:5043. 24. Basbous J, Arpin C, Gaudray G, et al. The HBZ factor of human T-cell leukemia virus type I dimerizes with transcription factors JunB and c-Jun and modulates their transcriptional activity. J Biol Chem 2003;278:43620–43627. 25. Baydoun HH, Pancewicz J, Bai X, et al. HTLV-I p30 inhibits multiple S phase entry checkpoints, decreases cyclin E-CDK2 interactions and delays cell cycle progression. Mol Cancer 2010;9:302. 26. Baydoun HH, Pancewicz J, Nicot C. Human T-lymphotropic type 1 virus p30 inhibits homologous recombination and favors unfaithful DNA repair. Blood 2011;117:5897–5906. 27. Bayliss RJ, Piguet V. Masters of manipulation: viral modulation of the immunological synapse. Cell Microbiol 2018;20:e12944. 28. Bazarbachi A, Plumelle Y, Carlos Ramos J, et al. Meta-analysis on the use of zidovudine and interferon-alfa in adult T-cell leukemia/lymphoma showing improved survival in the leukemic subtypes. J Clin Oncol 2010;28:4177–4183. 29. Beimling P, Moelling K. Direct interaction of CREB protein with 21 bp Tax-response elements of HTLV-ILTR. Oncogene 1992;7:257–262. 30. Biasiotto R, Aguiari P, Rizzuto R, et al. The p13 protein of human T cell leukemia virus type 1 (HTLV-1) modulates mitochondrial membrane potential and calcium uptake. Biochim Biophys Acta 2010;1797:945–951. 31. Biggar RJ, Ng J, Kim N, et al. Human leukocyte antigen concordance and the transmission risk via breast-feeding of human T cell lymphotropic virus type I. J Infect Dis 2006;193:277–282. 32. Billman MR, Rueda D, Bangham CRM. Single-cell heterogeneity and cell-cycle-related viral gene bursts in the human leukaemia virus HTLV-1. Wellcome Open Res 2017;2:87. 33. Bittencourt AL, de Oliveira M. Cutaneous manifestations associated with HTLV-1 infection. Int J Dermatol 2010;49:1099–1110. 34. Blattner WA, Gallo RC. Epidemiology of HTLV-I and HTLV-II infection. In Takahashi K, ed. Adult T-cell Leukemia. New York: Oxford University Press; 1994. 35. Blot V, Delamarre L, Perugi F, et al. Human Dlg protein binds to the envelope glycoproteins of human T-cell leukemia virus type 1 and regulates envelope mediated cell-cell fusion in T lymphocytes. J Cell Sci 2004;117:3983–3993. 36. Boelen L, Debebe B, Silveira M, et al. Inhibitory killer cell immunoglobulin-like receptors strengthen CD8(+) T cell-mediated control of HIV-1, HCV, and HTLV-1. Sci Immunol 2018;3:eaao2892. 623

37. Bogerd H, Greene WC. Dominant negative mutants of human T-cell leukemia virus type I Rex and human immunodeficiency virus type 1 Rev fail to multimerize in vivo. J Virol 1993;67:2496–2502. 38. Bogerd HP, Huckaby GL, Ahmed YF, et al. The type I human T-cell leukemia virus (HTLV-I) Rex trans-activator binds directly to the HTLV-I Rex and the type 1 human immunodeficiency virus Rev RNA response elements. Proc Natl Acad Sci U S A 1991;88:5704–5708. 39. Borg A, Yin JA, Johnson PR, et al. Successful treatment of HTLV-1-associated acute adult T-cell leukaemia lymphoma by allogeneic bone marrow transplantation. Br J Haematol 1996;94:713–715. 40. Calattini S, Chevalier SA, Duprez R, et al. Discovery of a new human T-cell lymphotropic virus (HTLV-3) in Central Africa. Retrovirology 2005;2:30. 41. Cassar O, Einsiedel L, Afonso PV, et al. Human T-cell lymphotropic virus type 1 subtype C molecular variants among indigenous australians: new insights into the molecular epidemiology of HTLV-1 in Australo-Melanesia. PLoS Negl Trop Dis 2013;7:e2418. 42. Cavrois M, Gessain A, Gout O, et al. Common human T cell leukemia virus type 1 (HTLV-1) integration sites in cerebrospinal fluid and blood lymphocytes of patients with HTLV-1-associated myelopathy/tropical spastic paraparesis indicate that HTLV-1 crosses the bloodbrain barrier via clonal HTLV-1-infected cells. J Infect Dis 2000;182:1044–1050. 43. Chandhasin C, Ducu RI, Berkovich E, et al. Human T-cell leukemia virus type 1 tax attenuates the ATM-mediated cellular DNA damage response. J Virol 2008;82:6952–6961. 44. Charoenthongtrakul S, Zhou Q, Shembade N, et al. Human T cell leukemia virus type 1 tax inhibits innate antiviral signaling via NF-{kappa}B-dependent induction of SOCS1. J Virol 2011;85:6955–6962. 45. Ching YP, Chan SF, Jeang KT, et al. The retroviral oncoprotein Tax targets the coiled-coil centrosomal protein TAX1BP2 to induce centrosome overduplication. Nat Cell Biol 2006;8:717–724. 46. Cho WK, Jang MK, Huang K, et al. Human T-lymphotropic virus type 1 Tax protein complexes with P-TEFb and competes for Brd4 and 7SK snRNP/HEXIM1 binding. J Virol 2010;84:12801–12809. 47. Cho WK, Zhou M, Jang MK, et al. Modulation of the Brd4/P-TEFb interaction by the human T-lymphotropic virus type 1 tax protein. J Virol 2007;81:11179–11186. 48. Chu ZL, Shin YA, Yang JM, et al. IKKgamma mediates the interaction of cellular IkappaB kinases with the tax transforming protein of human T cell leukemia virus type 1. J Biol Chem 1999;274:15297–15300. 49. Ciminale V, Pavlakis GN, Derse D, et al. Complex splicing in the human T-cell leukemia virus (HTLV) family of retroviruses: novel mRNAs and proteins produced by HTLV type I. J Virol 1992;66:1737–1745. 50. Cook L, Melamed A, Yaguchi H, et al. The impact of HTLV-1 on the cellular genome. Curr Opin Virol 2017;26:125–131. 51. Cook LB, Melamed A, Demontis MA, et al. Rapid dissemination of human T-lymphotropic virus type 1 during primary infection in transplant recipients. Retrovirology 2016;13:3. 52. Cook LB, Melamed A, Niederer H, et al. The role of HTLV-1 clonality, proviral structure, and genomic integration site in adult T-cell leukemia/lymphoma. Blood 2014;123:3925–3931. 53. Cook LB, Rowan AG, Melamed A, et al. HTLV-1-infected T cells contain a single integrated provirus in natural infection. Blood 2012;120:3488–3490. 54. Coskun AK, Sutton RE. Expression of glucose transporter 1 confers susceptibility to human T-cell leukemia virus envelope-mediated fusion. J Virol 2005;79:4150–4158. 55. Daenke S, Nightingale S, Cruickshank JK, et al. Sequence variants of human T-cell lymphotropic virus type I from patients with tropical spastic paraparesis and adult T-cell leukemia do not distinguish neurological from leukemic isolates. J Virol 1990;64:1278–1282. 56. Datta A, Silverman L, Phipps AJ, et al. Human T-lymphotropic virus type-1 p30 alters cell cycle G2 regulation of T lymphocytes to enhance cell survival. Retrovirology 2007;4:49. 57. Datta A, Sinha-Datta U, Dhillon NK, et al. The HTLV-I p30 interferes with TLR4 signaling and modulates the release of pro- and antiinflammatory cytokines from human macrophages. J Biol Chem 2006;281:23414–23424. 58. Delamarre L, Pique C, Rosenberg AR, et al. The Y-S-L-I tyrosine-based motif in the cytoplasmic domain of the human T-cell leukemia virus type 1 envelope is essential for cell-to-cell transmission. J Virol 1999;73:9659–9663. 59. Delamarre L, Rosenberg AR, Pique C, et al. A novel human T-leukemia virus type 1 cell-to-cell transmission assay permits definition of SU glycoprotein amino acids important for infectivity. J Virol 1997;71:259–266. 60. Derse D, Hill SA, Lloyd PA, et al. Examining human T-lymphotropic virus type 1 infection and replication by cell-free infection with recombinant virus vectors. J Virol 2001;75:8461–8468. 61. Derse D, Hill SA, Princler G, et al. Resistance of human T cell leukemia virus type 1 to APOBEC3G restriction is mediated by elements in nucleocapsid. Proc Natl Acad Sci U S A 2007;104:2915–2920. 62. Desrames A, Cassar O, Gout O, et al. Northern African strains of human T-lymphotropic virus type 1 arose from a recombination event. J Virol 2014;88:9782–9788. 63. Ding W, Albrecht B, Kelley RE, et al. Human T-cell lymphotropic virus type 1 p12(I) expression increases cytoplasmic calcium to enhance the activation of nuclear factor of activated T cells. J Virol 2002;76:10374–10382. 64. Ding W, Albrecht B, Luo R, et al. Endoplasmic reticulum and cis-Golgi localization of human T-lymphotropic virus type 1 p12(I): association with calreticulin and calnexin. J Virol 2001;75:7672–7682. 65. Durkin SS, Guo X, Fryrear KA, et al. HTLV-1 Tax oncoprotein subverts the cellular DNA damage response via binding to DNA-dependent protein kinase. J Biol Chem 2008;283:36311–36320. 66. Dustin ML, Cooper JA. The immunological synapse and the actin cytoskeleton: molecular hardware for T cell signaling. Nat Immunol 2000;1:23–29. 67. Einsiedel L, Fernandes L, Spelman T, et al. Bronchiectasis is associated with human T-lymphotropic virus 1 infection in an Indigenous Australian population. Clin Infect Dis 2012;54:43–50. 68. Einsiedel L, Pham H, Wilson K, et al. Human T-Lymphotropic Virus type 1c subtype proviral loads, chronic lung disease and survival in a prospective cohort of Indigenous Australians. PLoS Negl Trop Dis 2018;12:e0006281. 69. Eiraku N, Novoa P, da Costa Ferreira M, et al. Identification and characterization of a new and distinct molecular subtype of human T-cell lymphotropic virus type 2. J Virol 1996;70:1481–1492. 70. Fan J, Ma G, Nosaka K, et al. APOBEC3G generates nonsense mutations in human T-cell leukemia virus type 1 proviral genomes in vivo. J Virol 2010;84:7278–7287. 71. Felber BK, Paskalis H, Kleinman-Ewing C, et al. The pX protein of HTLV-I is a transcriptional activator of its long terminal repeats. Science 1985;229:675–679. 72. Fenizia C, Fiocchi M, Jones K, et al. Human T-cell leukemia/lymphoma virus type 1 p30, but not p12/p8, counteracts toll-like receptor 3 (TLR3) and TLR4 signaling in human monocytes and dendritic cells. J Virol 2014;88:393–402. 73. Feuer G, Green PL. Comparative biology of human T-cell lymphotropic virus type 1 (HTLV-1) and HTLV-2. Oncogene 2005;24:5996–6004. 74. Filippone C, Betsem E, Tortevoye P, et al. A severe bite from a nonhuman primate is a major risk factor for HTLV-1 infection in hunters from Central Africa. Clin Infect Dis 2015;60:1667–1676. 624

75. Fujihara K, Itoyama Y, Yu F, et al. Cellular immune surveillance against HTLV-I infected T lymphocytes in HTLV-I associated myelopathy/tropical spastic paraparesis (HAM/TSP). J Neurol Sci 1991;105:99–107. 76. Fukudome K, Furuse M, Fukuhara N, et al. Strong induction of ICAM-1 in human T cells transformed by human T-cell-leukemia virus type 1 and depression of ICAM-1 or LFA-1 in adult T-cell-leukemia-derived cell lines. Int J Cancer 1992;52:418–427. 77. Fukumoto R, Andresen V, Bialuk I, et al. In vivo genetic mutations define predominant functions of the human T-cell leukemia/lymphoma virus p12I protein. Blood 2009;113:3726–3734. 78. Furukawa Y, Bangham CR, Taylor GP, et al. Frequent reversible membrane damage in peripheral blood B cells in human T cell lymphotropic virus type I (HTLV-I)-associated myelopathy/tropical spastic paraparesis (HAM/TSP). Clin Exp Immunol 2000;120:307–316. 79. Furukawa Y, Kubota R, Tara M, et al. Existence of escape mutant in HTLV-I tax during the development of adult T-cell leukemia. Blood 2001;97:987–993. 80. Furukawa Y, Yamashita M, Usuku K, et al. Phylogenetic subgroups of human T cell lymphotropic virus (HTLV) type I in the tax gene and their association with different risks for HTLV-I-associated myelopathy/tropical spastic paraparesis. J Infect Dis 2000;182:1343–1349. 81. Furuta R, Yasunaga JI, Miura M, et al. Human T-cell leukemia virus type 1 infects multiple lineage hematopoietic cells in vivo. PLoS Pathog 2017;13:e1006722. 82. Gao F, Bailes E, Robertson DL, et al. Origin of HIV-1 in the chimpanzee Pan troglodytes troglodytes. Nature 1999;397:436–441. 83. Gaudray G, Gachon F, Basbous J, et al. The complementary strand of the human T-cell leukemia virus type 1 RNA genome encodes a bZIP transcription factor that down-regulates viral transcription. J Virol 2002;76:12813–12822. 84. Gazon H, Chauhan PS, Porquet F, et al. Epigenetic silencing of HTLV-1 expression by the HBZ RNA through interference with the basal transcription machinery. Blood Adv 2020;4:5574–5579. 85. Geleziunas R, Ferrell S, Lin X, et al. Human T-cell leukemia virus type 1 Tax induction of NF-kappaB involves activation of the IkappaB kinase alpha (IKKalpha) and IKKbeta cellular kinases. Mol Cell Biol 1998;18:5157–5165. 86. Georges SA, Giebler HA, Cole PA, et al. Tax recruitment of CBP/p300, via the KIX domain, reveals a potent requirement for acetyltransferase activity that is chromatin dependent and histone tail independent. Mol Cell Biol 2003;23:3392–3404. 87. Georges SA, Kraus WL, Luger K, et al. p300-mediated tax transactivation from recombinant chromatin: histone tail deletion mimics coactivator function. Mol Cell Biol 2002;22:127–137. 88. Gessain A, Barin F, Vernant JC, et al. Antibodies to human T-lymphotropic virus type-I in patients with tropical spastic paraparesis. Lancet 1985;2:407–410. 89. Gessain A, Cassar O. Epidemiological aspects and world distribution of HTLV-1 infection. Front Microbiol 2012;3:388. 90. Gessain A, Gallo RC, Franchini G. Low degree of human T-cell leukemia/lymphoma virus type I genetic drift in vivo as a means of monitoring viral transmission and movement of ancient human populations. J Virol 1992;66:2288–2295. 91. Gessain A, Yanagihara R, Franchini G, et al. Highly divergent molecular variants of human T-lymphotropic virus type I from isolated populations in Papua New Guinea and the Solomon Islands. Proc Natl Acad Sci U S A 1991;88:7694–7698. 92. Ghez D, Lepelletier Y, Jones KS, et al. Current concepts regarding the HTLV-1 receptor complex. Retrovirology 2010;7:99. 93. Ghez D, Lepelletier Y, Lambert S, et al. Neuropilin-1 is involved in human T-cell lymphotropic virus type 1 entry. J Virol 2006;80:6844–6854. 94. Ghorbel S, Sinha-Datta U, Dundr M, et al. Human T-cell leukemia virus type I p30 nuclear/nucleolar retention is mediated through interactions with RNA and a constituent of the 60 S ribosomal subunit. J Biol Chem 2006;281:37150–37158. 95. Giebler HA, Loring JE, van Orden K, et al. Anchoring of CREB binding protein to the human T-cell leukemia virus type 1 promoter: a molecular mechanism of Tax transactivation. Mol Cell Biol 1997;17:5156–5164. 96. Gillet NA, Malani N, Melamed A, et al. The host genomic environment of the provirus determines the abundance of HTLV-1-infected T-cell clones. Blood 2011;117:3113–3122. 97. Goh WC, Sodroski J, Rosen C, et al. Subcellular localization of the product of the long open reading frame of human T-cell leukemia virus type I. Science 1985;227:1227–1228. 98. Goldstone DC, Ennis-Adeniran V, Hedden JJ, et al. HIV-1 restriction factor SAMHD1 is a deoxynucleoside triphosphate triphosphohydrolase. Nature 2011;480:379–382. 99. Goon PK, Biancardi A, Fast N, et al. Human T cell lymphotropic virus (HTLV) type-1-specific CD8+ T cells: frequency and immunodominance hierarchy. J Infect Dis 2004;189:2294–2298. 100. Goon PK, Hanon E, Igakura T, et al. High frequencies of Th1-type CD4(+) T cells specific to HTLV-1 Env and Tax proteins in patients with HTLV-1-associated myelopathy/tropical spastic paraparesis. Blood 2002;99:3335–3341. 101. Goon PK, Igakura T, Hanon E, et al. High circulating frequencies of tumor necrosis factor alpha- and interleukin-2-secreting human T-lymphotropic virus type 1 (HTLV-1)-specific CD4+ T cells in patients with HTLV-1-associated neurological disease. J Virol 2003;77:9716–9722. 102. Goon PK, Igakura T, Hanon E, et al. Human T cell lymphotropic virus type I (HTLV-I)-specific CD4+ T cells: immunodominance hierarchy and preferential infection with HTLV-I. J Immunol 2004;172:1735–1743. 103. Grone M, Hoffmann E, Berchtold S, et al. A single stem-loop structure within the HTLV-1 Rex response element is sufficient to mediate Rex activity in vivo. Virology 1994;204:144–152. 104. Hagiya K, Yasunaga J, Satou Y, et al. ATF3, an HTLV-1 bZip factor binding protein, promotes proliferation of adult T-cell leukemia cells. Retrovirology 2011;8:19. 105. Hahn BH, Shaw GM, Popovic M, et al. Molecular cloning and analysis of a new variant of human T-cell leukemia virus (HTLV-ib) from an African patient with adult T-cell leukemia-lymphoma. Int J Cancer 1984;34:613–618. 106. Hakata Y, Yamada M, Shida H. Rat CRM1 is responsible for the poor activity of human T-cell leukemia virus type 1 Rex protein in rat cells. J Virol 2001;75:11515–11525. 107. Halin M, Douceron E, Clerc I, et al. Human T-cell leukemia virus type 2 produces a spliced antisense transcript encoding a protein that lacks a classic bZIP domain but still inhibits Tax2-mediated transcription. Blood 2009;114:2427–2438. 108. Hall WW, Ishak R, Zhu SW, et al. Human T lymphotropic virus type II (HTLV-II): epidemiology, molecular properties, and clinical features of infection. J Acquir Immune Defic Syndr Hum Retrovirol 1996;13(Suppl 1):S204–S214. 109. Hammes SR, Greene WC. Multiple arginine residues within the basic domain of HTLV-I Rex are required for specific RNA binding and function. Virology 1993;193:41–49. 110. Hanabuchi S, Ohashi T, Koya Y, et al. Regression of human T-cell leukemia virus type I (HTLV-I)-associated lymphomas in a rat model: peptide-induced T-cell immunity. J Natl Cancer Inst 2001;93:1775–1783. 111. Hanon E, Goon P, Taylor GP, et al. High production of interferon gamma but not interleukin-2 by human T-lymphotropic virus type I-infected peripheral blood mononuclear cells. Blood 2001;98:721–726. 112. Hanon E, Hall S, Taylor GP, et al. Abundant tax protein expression in CD4+ T cells infected with human T-cell lymphotropic virus type I (HTLV-I) is prevented by cytotoxic T lymphocytes. Blood 2000;95:1386–1392. 113. Harashima N, Kurihara K, Utsunomiya A, et al. Graft-versus-Tax response in adult T-cell leukemia patients after hematopoietic stem cell 625

transplantation. Cancer Res 2004;64:391–399. 114. Harhaj EW, Sun SC. IKKgamma serves as a docking subunit of the IkappaB kinase (IKK) and mediates interaction of IKK with the human T-cell leukemia virus Tax protein. J Biol Chem 1999;274:22911–22914. 115. Harrod R. Silencers of HTLV-1 and HTLV-2: the pX-encoded latency-maintenance factors. Retrovirology 2019;16:25. 116. Harrod R, Tang Y, Nicot C, et al. An exposed KID-like domain in human T-cell lymphotropic virus type 1 Tax is responsible for the recruitment of coactivators CBP/p300. Mol Cell Biol 1998;18:5052–5061. 117. Hatta Y, Hirama T, Miller CW, et al. Homozygous deletions of the p15 (MTS2) and p16 (CDKN2/MTS1) genes in adult T-cell leukemia. Blood 1995;85:2699–2704. 118. He Y, Pasupala N, Zhi H, et al. NF-kappaB-induced R-loop accumulation and DNA damage select for nucleotide excision repair deficiencies in adult T cell leukemia. Proc Natl Acad Sci U S A 2021;118(10):e2005568118. 119. Hidaka M, Inoue J, Yoshida M, et al. Post-transcriptional regulator (rex) of HTLV-1 initiates expression of viral structural proteins but suppresses expression of regulatory proteins. EMBO J 1988;7:519–523. 120. Hieshima K, Nagakubo D, Nakayama T, et al. Tax-inducible production of CC chemokine ligand 22 by human T cell leukemia virus type 1 (HTLV-1)-infected T cells promotes preferential transmission of HTLV-1 to CCR4-expressing CD4+ T cells. J Immunol 2008;180:931–939. 121. Higuchi M, Tsubata C, Kondo R, et al. Cooperation of NF-kappaB2/p100 activation and the PDZ domain binding motif signal in human T-cell leukemia virus type 1 (HTLV-1) Tax1 but not HTLV-2 Tax2 is crucial for interleukin-2-independent growth transformation of a T-cell line. J Virol 2007;81:11900–11907. 122. Higuchi Y, Yasunaga JI, Mitagami Y, et al. HTLV-1 induces T cell malignancy and inflammation by viral antisense factor-mediated modulation of the cytokine signaling. Proc Natl Acad Sci U S A 2020;117:13740–13749. 123. Hilburn S, Rowan A, Demontis MA, et al. In vivo expression of human T-lymphotropic virus type 1 basic leucine-zipper protein generates specific CD8+ and CD4+ T-lymphocyte responses that correlate with clinical outcome. J Infect Dis 2011;203:529–536. 124. Hill SA, Lloyd PA, McDonald S, et al. Susceptibility of human T cell leukemia virus type I to nucleoside reverse transcriptase inhibitors. J Infect Dis 2003;188:424–427. 125. Hinuma Y, Nagata K, Hanaoka M, et al. Adult T-cell leukemia: antigen in an ATL cell line and detection of antibodies to the antigen in human sera. Proc Natl Acad Sci U S A 1981;78:6476–6480. 126. Hiraragi H, Kim SJ, Phipps AJ, et al. Human T-lymphotropic virus type 1 mitochondrion-localizing protein p13(II) is required for viral infectivity in vivo. J Virol 2006;80:3469–3476. 127. Hirata M, Hayashi J, Noguchi A, et al. The effects of breastfeeding and presence of antibody to p40tax protein of human T cell lymphotropic virus type-I on mother to child transmission. Int J Epidemiol 1992;21:989–994. 128. Hisada M, Stuver SO, Okayama A, et al. Persistent paradox of natural history of human T lymphotropic virus type I: parallel analyses of Japanese and Jamaican carriers. J Infect Dis 2004;190:1605–1609. 129. Hishiki T, Ohshima T, Ego T, et al. BCL3 acts as a negative regulator of transcription from the human T-cell leukemia virus type 1 long terminal repeat through interactions with TORC3. J Biol Chem 2007;282:28335–28343. 130. Hivin P, Basbous J, Raymond F, et al. The HBZ-SP1 isoform of human T-cell leukemia virus type I represses JunB activity by sequestration into nuclear bodies. Retrovirology 2007;4:14. 131. Hivin P, Frederic M, Arpin-Andre C, et al. Nuclear localization of HTLV-I bZIP factor (HBZ) is mediated by three distinct motifs. J Cell Sci 2005;118:1355–1362. 132. Hjelle B, Appenzeller O, Mills R, et al. Chronic neurodegenerative disease associated with HTLV-II infection. Lancet 1992;339:645–646. 133. Hoshida Y, Li T, Dong Z, et al. Lymphoproliferative disorders in renal transplant patients in Japan. Int J Cancer 2001;91:869–875. 134. Iga M, Okayama A, Stuver S, et al. Genetic evidence of transmission of human T cell lymphotropic virus type 1 between spouses. J Infect Dis 2002;185:691–695. 135. Igakura T, Stinchcombe JC, Goon PK, et al. Spread of HTLV-I between lymphocytes by virus-induced polarization of the cytoskeleton. Science 2003;299:1713–1716. 136. Ikezoe T, Nishioka C, Bandobashi K, et al. Longitudinal inhibition of PI3K/Akt/mTOR signaling by LY294002 and rapamycin induces growth arrest of adult T-cell leukemia cells. Leuk Res 2007;31(5):673–682. 137. Ina Y, Gojobori T. Molecular evolution of human T-cell leukemia virus. J Mol Evol 1990;31:493–499. 138. Inaba S, Okochi K, Sato H, et al. Efficacy of donor screening for HTLV-I and the natural history of transfusion-transmitted infection. Transfusion 1999;39:1104–1110. 139. Inoue J, Seiki M, Yoshida M. The second pX product p27 chi-III of HTLV-1 is required for gag gene expression. FEBS Lett 1986;209:187–190. 140. Iqbal M, Reljic T, Klocksieben F, et al. Efficacy of allogeneic hematopoietic cell transplantation in human T cell lymphotropic virus type 1-associated adult t cell leukemia/lymphoma: results of a systematic review/meta-analysis. Biol Blood Marrow Transplant 2019;25:1695–1700. 141. Ishida T, Jo T, Takemoto S, et al. Dose-intensified chemotherapy alone or in combination with mogamulizumab in newly diagnosed aggressive adult T-cell leukaemia-lymphoma: a randomized phase II study. Br J Haematol 2015;169:672–682. 142. Ishitsuka K, Tamura K. Human T-cell leukaemia virus type I and adult T-cell leukaemia-lymphoma. Lancet Oncol 2014;15:e517–e526. 143. Iwai K, Mori N, Oie M, et al. Human T-cell leukemia virus type 1 tax protein activates transcription through AP-1 site by inducing DNA binding activity in T cells. Virology 2001;279:38–46. 144. Iwanaga M. Epidemiology of HTLV-1 infection and ATL in Japan: an update. Front Microbiol 2020;11:1124. 145. Iwanaga M, Watanabe T, Utsunomiya A, et al. Human T-cell leukemia virus type I (HTLV-1) proviral load and disease progression in asymptomatic HTLV-1 carriers: a nationwide prospective study in Japan. Blood 2010;116:1211–1219. 146. Iwanaga Y, Tsukahara T, Ohashi T, et al. Human T-cell leukemia virus type 1 tax protein abrogates interleukin-2 dependence in a mouse T-cell line. J Virol 1999;73:1271–1277. 147. Izumo S. Neuropathology of HTLV-1-associated myelopathy (HAM/TSP): the 50th Anniversary of Japanese Society of Neuropathology. Neuropathology 2010;30(5):480–485. 148. Jacobson S, Shida H, McFarlin DE, et al. Circulating CD8+ cytotoxic T lymphocytes specific for HTLV-I pX in patients with HTLV-I associated neurological disease. Nature 1990;348:245–248. 149. Jain P, Manuel SL, Khan ZK, et al. DC-SIGN mediates cell-free infection and transmission of human T-cell lymphotropic virus type 1 by dendritic cells. J Virol 2009;83:10908–10921. 150. Jeang KT, Widen SG, Semmes OJ IV, et al. HTLV-I trans-activator protein, tax, is a trans-repressor of the human beta-polymerase gene. Science 1990;247:1082–1084. 151. Jeffery KJ, Usuku K, Hall SE, et al. HLA alleles determine human T-lymphotropic virus-I (HTLV-I) proviral load and the risk of HTLVI-associated myelopathy. Proc Natl Acad Sci U S A 1999;96:3848–3853. 152. Jeong SJ, Pise-Masison CA, Radonovich MF, et al. Activated AKT regulates NF-kappaB activation, p53 inhibition and cell survival in HTLV-1-transformed cells. Oncogene 2005;24:6719–6728. 626

153. Jiang S, Inada T, Tanaka M, et al. Involvement of TORC2, a CREB co-activator, in the in vivo-specific transcriptional control of HTLV-1. Retrovirology 2009;6:73. 154. Jin DY, Spencer F, Jeang KT. Human T cell leukemia virus type 1 oncoprotein Tax targets the human mitotic checkpoint protein MAD1. Cell 1998;93:81–91. 155. Jin Q, Agrawal L, VanHorn-Ali Z, et al. Infection of CD4+ T lymphocytes by the human T cell leukemia virus type 1 is mediated by the glucose transporter GLUT-1: evidence using antibodies specific to the receptors large extracellular domain. Virology 2006;349:184–196. 156. Jin Q, Agrawal L, Vanhorn-Ali Z, et al. GLUT-1-independent infection of the glioblastoma/astroglioma U87 cells by the human T cell leukemia virus type 1. Virology 2006;353:99–110. 157. Johnson JM, Nicot C, Fullen J, et al. Free major histocompatibility complex class I heavy chain is preferentially targeted for degradation by human T-cell leukemia/lymphotropic virus type 1 p12(I) protein. J Virol 2001;75:6086–6094. 158. Jones KS, Akel S, Petrow-Sadowski C, et al. Induction of human T cell leukemia virus type I receptors on quiescent naive T lymphocytes by TGF-beta. J Immunol 2005;174:4262–4270. 159. Jones KS, Petrow-Sadowski C, Huang YK, et al. Cell-free HTLV-1 infects dendritic cells leading to transmission and transformation of CD4(+) T cells. Nat Med 2008;14:429–436. 160. Journo C, Mahieux R. HTLV-1 and innate immunity. Viruses 2011;3:1374–1394. 161. Kanda J, Hishizawa M, Utsunomiya A, et al. Impact of graft-versus-host disease on outcomes after allogeneic hematopoietic cell transplantation for adult T-cell leukemia: a retrospective cohort study. Blood 2012;119:2141–2148. 162. Kannagi M, Harada S, Maruyama I, et al. Predominant recognition of human T cell leukemia virus type I (HTLV-I) pX gene products by human CD8+ cytotoxic T cells directed against HTLV-I-infected cells. Int Immunol 1991;3:761–767. 163. Kannian P, Fernandez S, Jones KS, et al. Human T lymphotropic virus type 1 SU residue 195 plays a role in determining the preferential CD4+ T cell immortalization/transformation tropism. J Virol 2013;87:9344–9352. 164. Kannian P, Yin H, Doueiri R, et al. Distinct transformation tropism exhibited by human T lymphotropic virus type 1 (HTLV-1) and HTLV-2 is the result of postinfection T cell clonal expansion. J Virol 2012;86:3757–3766. 165. Kao SY, Marriott SJ. Disruption of nucleotide excision repair by the human T-cell leukemia virus type 1 Tax protein. J Virol 1999;73:4299–4304. 166. Kaplan JE, Osame M, Kubota H, et al. The risk of development of HTLV-I-associated myelopathy/tropical spastic paraparesis among persons infected with HTLV-I. J Acquir Immune Defic Syndr 1990;3:1096–1101. 167. Karube K, Aoki R, Sugita Y, et al. The relationship of FOXP3 expression and clinicopathological characteristics in adult T-cell leukemia/lymphoma. Mod Pathol 2008;21:617–625. 168. Karube K, Ohshima K, Tsuchiya T, et al. Expression of FoxP3, a key molecule in CD4CD25 regulatory T cells, in adult T-cell leukaemia/lymphoma cells. Br J Haematol 2004;126:81–84. 169. Kashanchi F, Brady JN. Transcriptional and post-transcriptional gene regulation of HTLV-1. Oncogene 2005;24:5938–5951. 170. Kataoka K, Nagata Y, Kitanaka A, et al. Integrated molecular analysis of adult T cell leukemia/lymphoma. Nat Genet 2015;47:1304–1315. 171. Katsuya H, Ishitsuka K, Utsunomiya A, et al. TL-Prognostic Index Project. Treatment and survival among 1594 patients with ATL. Blood 2015;126:2570–2577. 172. Katsuya H, Islam S, Tan BJY, et al. The nature of the HTLV-1 provirus in naturally infected individuals analyzed by the viral DNAcapture-Seq approach. Cell Rep 2019;29:724–735 e4. 173. Kawakami A, Nakashima T, Sakai H, et al. Inhibition of caspase cascade by HTLV-I tax through induction of NF-kappaB nuclear translocation. Blood 1999;94:3847–3854. 174. Kawano N, Shimoda K, Ishikawa F, et al. Adult T-cell leukemia development from a human T-cell leukemia virus type I carrier after a living-donor liver transplantation. Transplantation 2006;82:840–843. 175. Kazanji M, Mouinga-Ondeme A, Lekana-Douki-Etenna S, et al. Origin of HTLV-1 in hunters of nonhuman primates in Central Africa. J Infect Dis 2015;211:361–365. 176. Kazanji M, Ureta-Vidal A, Ozden S, et al. Lymphoid organs as a major reservoir for human T-cell leukemia virus type 1 in experimentally infected squirrel monkeys (Saimiri sciureus): provirus expression, persistence, and humoral and cellular immune responses. J Virol 2000;74:4860–4867. 177. Kesic M, Doueiri R, Ward M, et al. Phosphorylation regulates human T-cell leukemia virus type 1 Rex function. Retrovirology 2009;6:105. 178. Kim FJ, Manel N, Boublik Y, et al. Human T-cell leukemia virus type 1 envelope-mediated syncytium formation can be activated in resistant Mammalian cell lines by a carboxy-terminal truncation of the envelope cytoplasmic domain. J Virol 2003;77:963–969. 179. Kim SJ, Ding W, Albrecht B, et al. A conserved calcineurin-binding motif in human T lymphotropic virus type 1 p12I functions to modulate nuclear factor of activated T cell activation. J Biol Chem 2003;278:15550–15557. 180. Kim YM, Ramirez JA, Mick JE, et al. Molecular characterization of the tax-containing HTLV-1 enhancer complex reveals a prominent role for CREB phosphorylation in tax transactivation. J Biol Chem. 2007;282(26):18750–18757. 181. Kinet S, Swainson L, Lavanya M, et al. Isolated receptor binding domains of HTLV-1 and HTLV-2 envelopes bind Glut-1 on activated CD4+ and CD8+ T cells. Retrovirology 2007;4:31. 182. King JA, Bridger JM, Lochelt M, et al. Nucleocytoplasmic transport of HTLV-1 RNA is regulated by two independent LTR encoded nuclear retention elements. Oncogene 1998;16:3309–3316. 183. Kinpara S, Hasegawa A, Utsunomiya A, et al. Stromal cell-mediated suppression of human T-cell leukemia virus type 1 expression in vitro and in vivo by type I interferon. J Virol 2009;83:5101–5108. 184. Kirk PD, Huvet M, Melamed A, et al. Retroviruses integrate into a shared, non-palindromic DNA motif. Nat Microbiol 2016;2:16212. 185. Kitze B, Usuku K, Yamano Y, et al. Human CD4+ T lymphocytes recognize a highly conserved epitope of human T lymphotropic virus type 1 (HTLV-1) env gp21 restricted by HLA DRB1*0101. Clin Exp Immunol 1998;111:278–285. 186. Kiyokawa T, Yamaguchi K, Takeya M, et al. Hypercalcemia and osteoclast proliferation in adult T-cell leukemia. Cancer 1987;59:1187–1191. 187. Koga H, Ohshima T, Shimotohno K. Enhanced activation of tax-dependent transcription of human T-cell leukemia virus type I (HTLV-I) long terminal repeat by TORC3. J Biol Chem 2004;279:52978–52983. 188. Koiwa T, Hamano-Usami A, Ishida T, et al. 5-long terminal repeat-selective CpG methylation of latent human T-cell leukemia virus type 1 provirus in vitro and in vivo. J Virol 2002;76:9389–9397. 189. Komurian F, Pelloquin F, de The G. In vivo genomic variability of human T-cell leukemia virus type I depends more upon geography than upon pathologies. J Virol 1991;65:3770–3778. 190. Koralnik IJ, Boeri E, Saxinger WC, et al. Phylogenetic associations of human and simian T-cell leukemia/lymphotropic virus type I strains: evidence for interspecies transmission. J Virol 1994;68:2693–2707. 191. Koralnik IJ, Fullen J, Franchini G. The p12I, p13II, and p30II proteins encoded by human T-cell leukemia/lymphotropic virus type I open reading frames I and II are localized in three different cellular compartments. J Virol 1993;67:2360–2366. 192. Koralnik IJ, Gessain A, Klotman ME, et al. Protein isoforms encoded by the pX region of human T-cell leukemia/lymphotropic virus type 627

I. Proc Natl Acad Sci U S A 1992;89:8813–8817. 193. Koyanagi Y, Itoyama Y, Nakamura N, et al. In vivo infection of human T-cell leukemia virus type I in non-T cells. Virology 1993;196:25–33. 194. Kulkarni A, Taylor GP, Klose RJ, et al. Histone H2A monoubiquitylation and p38-MAPKs regulate immediate-early gene-like reactivation of latent retrovirus HTLV-1. JCI Insight 2018;3(20):e123196. 195. Kuo YL, Giam CZ. Activation of the anaphase promoting complex by HTLV-1 tax leads to senescence. EMBO J 2006;25:1741–1752. 196. Kwok RP, Laurance ME, Lundblad JR, et al. Control of cAMP-regulated enhancers by the viral transactivator Tax through CREB and the co-activator CBP. Nature 1996;380:642–646. 197. LaGrenade L, Hanchard B, Fletcher V, et al. Infective dermatitis of Jamaican children: a marker for HTLV-I infection. Lancet 1990;336:1345–1347. 198. Larocca D, Chao LA, Seto MH, et al. Human T-cell leukemia virus minus strand transcription in infected T-cells. Biochem Biophys Res Commun 1989;163:1006–1013. 199. Laydon DJ, Melamed A, Sim A, et al. Quantification of HTLV-1 clonality and TCR diversity. PLoS Comput Biol 2014;10:e1003646. 200. Laydon DJ, Sunkara V, Boelen L, et al. The relative contributions of infectious and mitotic spread to HTLV-1 persistence. PLoS Comput Biol 2020;16:e1007470. 201. Lee R, Schwartz RA. Human T-lymphotrophic virus type 1-associated infective dermatitis: a comprehensive review. J Am Acad Dermatol 2011;64:152–160. 202. Lemasson I, Lewis MR, Polakowski N, et al. Human T-cell leukemia virus type 1 (HTLV-1) bZIP protein interacts with the cellular transcription factor CREB to inhibit HTLV-1 transcription. J Virol 2007;81:1543–1553. 203. Lemasson I, Polakowski NJ, Laybourn PJ, et al. Transcription factor binding and histone modifications on the integrated proviral promoter in human T-cell leukemia virus-I-infected T-cells. J Biol Chem 2002;277:49459–49465. 204. Lemasson I, Polakowski NJ, Laybourn PJ, et al. Tax-dependent displacement of nucleosomes during transcriptional activation of human T-cell leukemia virus type 1. J Biol Chem 2006;281:13075–13082. 205. Lenzmeier BA, Baird EE, Dervan PB, et al. The tax protein-DNA interaction is essential for HTLV-I transactivation in vitro. J Mol Biol 1999;291:731–744. 206. Lenzmeier BA, Giebler HA, Nyborg JK. Human T-cell leukemia virus type 1 Tax requires direct access to DNA for recruitment of CREB binding protein to the viral promoter. Mol Cell Biol 1998;18:721–731. 207. Leung K, Nabel GJ. HTLV-1 transactivator induces interleukin-2 receptor expression through an NF-kappa B-like factor. Nature 1988;333:776–778. 208. Levin MC, Krichavsky M, Fox RJ, et al. Extensive latent retroviral infection in bone marrow of patients with HTLV-I-associated neurologic disease. Blood 1997;89:346–348. 209. Levin MC, Lee SM, Kalume F, et al. Autoimmunity due to molecular mimicry as a cause of neurological disease. Nat Med 2002;8:509–513. 210. Li HC, Biggar RJ, Miley WJ, et al. Provirus load in breast milk and risk of mother-to-child transmission of human T lymphotropic virus type I. J Infect Dis 2004;190:1275–1278. 211. Li Y, Bolderson E, Kumar R, et al. HSSB1 and hSSB2 form similar multiprotein complexes that participate in DNA damage response. J Biol Chem 2009;284:23525–23531. 212. Liu M, Yang L, Zhang L, et al. Human T-cell leukemia virus type 1 infection leads to arrest in the G1 phase of the cell cycle. J Virol 2008;82:8442–8455. 213. Ma G, Yasunaga JI, Shimura K, et al. Human retroviral antisense mRNAs are retained in the nuclei of infected cells for viral persistence. Proc Natl Acad Sci U S A 2021;118(17):e2014783118. 214. Macatonia SE, Cruickshank JK, Rudge P, et al. Dendritic cells from patients with tropical spastic paraparesis are infected with HTLV-1 and stimulate autologous lymphocyte proliferation. AIDS Res Hum Retroviruses 1992;8:1699–1706. 215. MacNamara A, Rowan A, Hilburn S, et al. HLA class I binding of HBZ determines outcome in HTLV-1 infection. PLoS Pathog 2010;6:e1001117. 216. Maeda M, Tanabe-Shibuya J, Miyazato P, et al. IL-2/IL-2 receptor pathway plays a crucial role in the growth and malignant transformation of HTLV-1-infected T cells to develop adult T-cell leukemia. Front Microbiol 2020;11:356. 217. Maertens GN. B-protein phosphatase 2A is a functional binding partner of delta-retroviral integrase. Nucleic Acids Res 2016;44:364–376. 218. Maertens GN, Hare S, Cherepanov P. The mechanism of retroviral integration from X-ray structures of its key intermediates. Nature 2010;468:326–329. 219. Maggirwar SB, Harhaj E, Sun SC. Activation of NF-kappa B/Rel by Tax involves degradation of I kappa B alpha and is blocked by a proteasome inhibitor. Oncogene 1995;11:993–998. 220. Mahe A, Chollet-Martin S, Gessain A. HTLV-I-associated infective dermatitis. Lancet 1999;354:1386. 221. Mahgoub M, Yasunaga JI, Iwami S, et al. Sporadic on/off switching of HTLV-1 Tax expression is crucial to maintain the whole population of virus-induced leukemic cells. Proc Natl Acad Sci U S A 2018;115:E1269–E1278. 222. Mahieux R, Gessain A. The human HTLV-3 and HTLV-4 retroviruses: new members of the HTLV family. Pathol Biol (Paris) 2009;57:161–166. 223. Mahieux R, Ibrahim F, Mauclere P, et al. Molecular epidemiology of 58 new African human T-cell leukemia virus type 1 (HTLV-1) strains: identification of a new and distinct HTLV-1 molecular subtype in Central Africa and in Pygmies. J Virol 1997;71:1317–1333. 224. Majone F, Jeang KT. Clastogenic effect of the human T-cell leukemia virus type I Tax oncoprotein correlates with unstabilized DNA breaks. J Biol Chem 2000;275:32906–32910. 225. Majorovits E, Nejmeddine M, Tanaka Y, et al. Human T-lymphotropic virus-1 visualized at the virological synapse by electron tomography. PLoS One 2008;3:e2251. 226. Maloney EM, Cleghorn FR, Morgan OS, et al. Incidence of HTLV-I-associated myelopathy/tropical spastic paraparesis (HAM/TSP) in Jamaica and Trinidad. J Acquir Immune Defic Syndr Hum Retrovirol 1998;17:167–170. 227. Manel N, Battini JL, Sitbon M. Human T cell leukemia virus envelope binding and virus entry are mediated by distinct domains of the glucose transporter GLUT1. J Biol Chem 2005;280:29025–29029. 228. Manel N, Battini JL, Taylor N, et al. HTLV-1 tropism and envelope receptor. Oncogene 2005;24:6016–6025. 229. Manel N, Kim FJ, Kinet S, et al. The ubiquitous glucose transporter GLUT-1 is a receptor for HTLV. Cell 2003;115:449–459. 230. Manns A, Murphy EL, Wilks R, et al. Detection of early human T-cell lymphotropic virus type I antibody patterns during seroconversion among transfusion recipients. Blood 1991;77:896–905. 231. Mansky LM. In vivo analysis of human T-cell leukemia virus type 1 reverse transcription accuracy. J Virol 2000;74:9525–9531. 232. Martinez MP, Al-Saleem J, Green PL. Comparative virology of HTLV-1 and HTLV-2. Retrovirology 2019;16:21. 233. Maruyama M, Shibuya H, Harada H, et al. Evidence for aberrant activation of the interleukin-2 autocrine loop by HTLV-1-encoded p40x and T3/Ti complex triggering. Cell 1987;48:343–350. 628

234. Matsuoka E, Takenouchi N, Hashimoto K, et al. Perivascular T cells are infected with HTLV-I in the spinal cord lesions with HTLVI-associated myelopathy/tropical spastic paraparesis: double staining of immunohistochemistry and polymerase chain reaction in situ hybridization. Acta Neuropathol 1998;96:340–346. 235. Matsushita S, Mitsuya H, Reitz MS, et al. Pharmacological inhibition of in vitro infectivity of human T lymphotropic virus type I. J Clin Invest 1987;80:394–400. 236. Matsuzaki T, Nakagawa M, Nagai M, et al. HTLV-I proviral load correlates with progression of motor disability in HAM/TSP: analysis of 239 HAM/TSP patients including 64 patients followed up for 10 years. J Neurovirol 2001;7:228–234. 237. Mazurov D, Ilinskaya A, Heidecker G, et al. Quantitative comparison of HTLV-1 and HIV-1 cell-to-cell infection with new replication dependent vectors. PLoS Pathog 2010;6:e1000788. 238. Melamed A, Laydon DJ, Al Khatib H, et al. HTLV-1 drives vigorous clonal expansion of infected CD8(+) T cells in natural infection. Retrovirology 2015;12:91. 239. Melamed A, Laydon DJ, Gillet NA, et al. Genome-wide determinants of proviral targeting, clonal abundance and expression in natural HTLV-1 infection. PLoS Pathog 2013;9:e1003271. 240. Melamed A, Witkover AD, Laydon DJ, et al. Clonality of HTLV-2 in natural infection. PLoS Pathog 2014;10:e1004006. 241. Melamed A, Yaguchi H, Miura M, et al. The human leukemia virus HTLV-1 alters the structure and transcription of host chromatin in cis. Elife 2018;7:e36245. 242. Mitobe Y, Yasunaga J, Furuta R, et al. HTLV-1 bZIP factor RNA and protein impart distinct functions on T-cell proliferation and survival. Cancer Res 2015;75: 4143–4152. 243. Miura M, Dey S, Ramanayake S, et al. Kinetics of HTLV-1 reactivation from latency quantified by single-molecule RNA FISH and stochastic modelling. PLoS Pathog 2019;15:e1008164. 244. Miura T, Fukunaga T, Igarashi T, et al. Phylogenetic subtypes of human T-lymphotropic virus type I and their relations to the anthropological background. Proc Natl Acad Sci U S A 1994;91:1124–1127. 245. Miyake H, Suzuki T, Hirai H, et al. Trans-activator Tax of human T-cell leukemia virus type 1 enhances mutation frequency of the cellular genome. Virology 1999;253:155–161. 246. Miyazaki M, Yasunaga J, Taniguchi Y, et al. Preferential selection of human T-cell leukemia virus type 1 provirus lacking the 5 long terminal repeat during oncogenesis. J Virol 2007;81:5714–5723. 247. Miyazato P, Yasunaga J, Taniguchi Y, et al. De novo human T-cell leukemia virus type 1 infection of human lymphocytes in NOD-SCID, common gamma-chain knockout mice. J Virol 2006;80:10683–10691. 248. Mochizuki M, Watanabe T, Yamaguchi K, et al. Uveitis associated with human T lymphotropic virus type I:seroepidemiologic, clinical, and virologic studies. J Infect Dis 1992;166:943–944. 249. Mochizuki M, Yamaguchi K, Takatsuki K, et al. HTLV-I and uveitis. Lancet 1992;339:1110. 250. Moles R, Sarkis S, Galli V, et al. p30 protein: a critical regulator of HTLV-1 viral latency and host immunity. Retrovirology 2019;16:42. 251. Montgomery RD, Cruickshank EK, Robertson WB, et al. Clinical and pathological observations on Jamaican neuropathy, a report on 206 cases. Brain 1964;87:425–462. 252. Morgan OS, Rodgers-Johnson P, Mora C, et al. HTLV-1 and polymyositis in Jamaica. Lancet 1989;2:1184–1187. 253. Morishita N, Ishii K, Tanaka Y, et al. Immunoglobulin prophylaxis against human T cell lymphotropic virus type II in rabbits. J Infect Dis 1994;169:620–623. 254. Moritoyo T, Reinhart TA, Moritoyo H, et al. Human T-lymphotropic virus type I-associated myelopathy and tax gene expression in CD4+ T lymphocytes. Ann Neurol 1996;40:84–90. 255. Moriuchi M, Moriuchi H. Seminal fluid enhances replication of human T-cell leukemia virus type 1: implications for sexual transmission. J Virol 2004;78:12709–12711. 256. Mueller NE, Blattner WA. Retroviruses: HTLV. In: Evans AS, Kaslow R, eds. Viral Infections of Humans: Epidemiology and Control. New York: Plenum Medical Press; 1997. 257. Mulloy JC, Crownley RW, Fullen J, et al. The human T-cell leukemia/lymphotropic virus type 1 p12I proteins bind the interleukin-2 receptor beta and gamma chains and affects their expression on the cell surface. J Virol 1996;70:3599–3605. 258. Mulloy JC, Kislyakova T, Cereseto A, et al. Human T-cell lymphotropic/leukemia virus type 1 Tax abrogates p53-induced cell cycle arrest and apoptosis through its CREB/ATF functional domain. J Virol 1998;72:8852–8860. 259. Murphy EL, Cassar O, Gessain A. Estimating the number of HTLV-2 infected persons in the world. Retrovirology 2015;12:O5. 260. Nagai M, Usuku K, Matsumoto W, et al. Analysis of HTLV-I proviral load in 202 HAM/TSP patients and 243 asymptomatic HTLV-I carriers: high proviral load strongly predisposes to HAM/TSP. J Neurovirol 1998;4:586–593. 261. Nagy K, Clapham P, Cheingsong-Popov R, et al. Human T-cell leukemia virus type I: induction of syncytia and inhibition by patients sera. Int J Cancer 1983;32:321–328. 262. Nakagawa M, Izumo S, Ijichi S, et al. HTLV-I-associated myelopathy: analysis of 213 patients based on clinical features and laboratory findings. J Neurovirol 1995;1:50–61. 263. Nakagawa M, Shaffer AL III, Ceribelli M, et al. Targeting the HTLV-I-regulated BATF3/IRF4 transcriptional network in adult T cell leukemia/lymphoma. Cancer Cell 2018;34:286–297 e10. 264. Nakamura H, Hayami M, Ohta Y, et al. Protection of cynomolgus monkeys against infection by human T-cell leukemia virus type-I by immunization with viral env gene products produced in Escherichia coli. Int J Cancer 1987;40:403–407. 265. Nakano K, Watanabe T. HTLV-1 Rex tunes the cellular environment favorable for viral replication. Viruses 2016;8:58. 266. Nakao K, Matsumoto M, Ohba N. Seroprevalence of antibodies to HTLV-I in patients with ocular disorders. Br J Ophthalmol 1991;75:76–78. 267. Nejmeddine M, Negi VS, Mukherjee S, et al. HTLV-1-Tax and ICAM-1 act on T-cell signal pathways to polarize the microtubuleorganizing center at the virological synapse. Blood 2009;114:1016–1025. 268. Nicot C, Dundr M, Johnson JM, et al. HTLV-1-encoded p30II is a post-transcriptional negative regulator of viral replication. Nat Med 2004;10:197–201. 269. Nicot C, Harrod RL, Ciminale V, et al. Human T-cell leukemia/lymphoma virus type 1 nonstructural genes and their functions. Oncogene 2005;24:6026–6034. 270. Niewiesk S, Daenke S, Parker CE, et al. The transactivator gene of human T-cell leukemia virus type I is more variable within and between healthy carriers than patients with tropical spastic paraparesis. J Virol 1994;68:6778–6781. 271. Niewiesk S, Daenke S, Parker CE, et al. Naturally occurring variants of human T-cell leukemia virus type I Tax protein impair its recognition by cytotoxic T lymphocytes and the transactivation function of Tax. J Virol 1995;69:2649–2653. 272. Niinuma A, Higuchi M, Takahashi M, et al. Aberrant activation of the interleukin-2 autocrine loop through the nuclear factor of activated T cells by nonleukemogenic human T-cell leukemia virus type 2 but not by leukemogenic type 1 virus. J Virol 2005;79:11925–11934. 273. Nishimura Y, Okubo R, Minato S, et al. A possible association between HLA and HTLV-I-associated myelopathy (HAM) in Japanese. Tissue Antigens 1991;37:230–231. 274. Nosaka K, Matsuoka M. Adult T-cell leukemia-lymphoma as a viral disease: subtypes based on viral aspects. Cancer Sci 629

2021;112(5):1688–1694. 275. Nosaka K, Miyamoto T, Sakai T, et al. Mechanism of hypercalcemia in adult T-cell leukemia: overexpression of receptor activator of nuclear factor kappaB ligand on adult T-cell leukemia cells. Blood 2002;99:634–640. 276. Nosaka T, Siomi H, Adachi Y, et al. Nucleolar targeting signal of human T-cell leukemia virus type I rex-encoded protein is essential for cytoplasmic accumulation of unspliced viral mRNA. Proc Natl Acad Sci U S A 1989;86:9798–9802. 277. Nowak MA, Bangham CR. Population dynamics of immune responses to persistent viruses. Science 1996;272:74–79. 278. Nyborg JK, Egan D, Sharma N. The HTLV-1 Tax protein: revealing mechanisms of transcriptional activation through histone acetylation and nucleosome disassembly. Biochim Biophys Acta 2010;1799:266–274. 279. Okamura J, Utsunomiya A, Tanosaki R, et al. Allogeneic stem-cell transplantation with reduced conditioning intensity as a novel immunotherapy and antiviral therapy for adult T-cell leukemia/lymphoma. Blood 2005;105:4143–4145. 280. Okochi K, Sato H. Transmission of ATLV (HTLV-I) through blood transfusion. Princess Takamatsu Symp 1984;15:129–135. 281. Oliere S, Hernandez E, Lezin A, et al. HTLV-1 evades type I interferon antiviral signaling by inducing the suppressor of cytokine signaling 1 (SOCS1). PLoS Pathog 2010;6:e1001177. 282. Orland JR, Engstrom J, Fridey J, et al. Prevalence and clinical features of HTLV neurologic disease in the HTLV Outcomes Study. Neurology 2003;61:1588–1594. 283. Osame M, Nakagawa M, Izumo S. HTLV-1-associated myelopathy/tropical spastic paraparesis (HAM/TSP). In: Takatsuki K, ed. Adult T-cell Leukemia. New York: Oxford University Press; 1994. 284. Osame M, Usuku K, Izumo S, et al. HTLV-I associated myelopathy, a new clinical entity. Lancet 1986;1:1031–1032. 285. Paine E, Garcia J, Philpott TC, et al. Limited sequence variation in human T-lymphotropic virus type 1 isolates from North American and African patients. Virology 1991;182:111–123. 286. Pais-Correia AM, Sachse M, Guadagnini S, et al. Biofilm-like extracellular viral assemblies mediate HTLV-1 cell-to-cell transmission at virological synapses. Nat Med 2010;16:83–89. 287. Paiva AM, Assone T, Haziot MEJ, et al. Risk factors associated with HTLV-1 vertical transmission in Brazil: longer breastfeeding, higher maternal proviral load and previous HTLV-1-infected offspring. Sci Rep 2018;8:7742. 288. Park HU, Jeong JH, Chung JH, et al. Human T-cell leukemia virus type 1 Tax interacts with Chk1 and attenuates DNA-damage induced G2 arrest mediated by Chk1. Oncogene 2004;23:4966–4974. 289. Park HU, Jeong SJ, Jeong JH, et al. Human T-cell leukemia virus type 1 Tax attenuates gamma-irradiation-induced apoptosis through physical interaction with Chk2. Oncogene 2006;25:438–447. 290. Parker CE, Daenke S, Nightingale S, et al. Activated, HTLV-1-specific cytotoxic T-lymphocytes are found in healthy seropositives as well as in patients with tropical spastic paraparesis. Virology 1992;188:628–636. 291. Peloponese JM Jr, Jeang KT. Role for Akt/protein kinase B and AP-1 in cellular proliferation induced by the human T-cell leukemia virus type 1 (HTLV-1) tax oncoprotein. J Biol Chem 2006;281:8927–8938. 292. Percher F, Jeannin P, Martin-Latil S, et al. Mother-to-child transmission of HTLV-1 epidemiological aspects, mechanisms and determinants of mother-to-child transmission. Viruses 2016;8(2):40. 293. Philip S, Zahoor MA, Zhi HJ, et al. Regulation of human T-lymphotropic virus type I latency and reactivation by HBZ and Rex. PLoS Pathog 2014;10(4):e1004040. 294. Phillips JE, Corces VG. CTCF: master weaver of the genome. Cell 2009;137:1194–1211. 295. Philpott SM, Buehring GC. Defective DNA repair in cells with human T-cell leukemia/bovine leukemia viruses: role of tax gene. J Natl Cancer Inst 1999;91:933–942. 296. Pinon JD, Klasse PJ, Jassal SR, et al. Human T-cell leukemia virus type 1 envelope glycoprotein gp46 interacts with cell surface heparan sulfate proteoglycans. J Virol 2003;77:9922–9930. 297. Pique C, Pham D, Tursz T, et al. The cytoplasmic domain of the human T-cell leukemia virus type I envelope can modulate envelope functions in a cell type-dependent manner. J Virol 1993;67:557–561. 298. Poiesz BJ, Ruscetti FW, Gazdar AF, et al. Detection and isolation of type C retrovirus particles from fresh and cultured lymphocytes of a patient with cutaneous T-cell lymphoma. Proc Natl Acad Sci U S A 1980;77:7415–7419. 299. Rende F, Cavallari I, Corradin A, et al. Kinetics and intracellular compartmentalization of HTLV-1 gene expression: nuclear retention of HBZ mRNAs. Blood 2011;117:4855–4859. 300. Richardson JH, Edwards AJ, Cruickshank JK, et al. In vivo cellular tropism of human T-cell leukemia virus type 1. J Virol 1990;64:5682–5687. 301. Rowan AG, Dillon R, Witkover A, et al. Evolution of retrovirus-infected premalignant T-cell clones prior to adult T-cell leukemia/lymphoma diagnosis. Blood 2020;135:2023–2032. 302. Rowan AG, Witkover A, Melamed A, et al. T cell receptor Vbeta staining identifies the malignant clone in adult t cell leukemia and reveals killing of leukemia cells by autologous CD8+ T cells. PLoS Pathog 2016;12:e1006030. 303. Ruben S, Poteat H, Tan TH, et al. Cellular transcription factors and regulation of IL-2 receptor gene expression by HTLV-I tax gene product. Science 1988;241:89–92. 304. Saito M, Braud VM, Goon P, et al. Low frequency of CD94/NKG2A+ T lymphocytes in patients with HTLV-1-associated myelopathy/tropical spastic paraparesis, but not in asymptomatic carriers. Blood 2003;102:577–584. 305. Saito M, Matsuzaki T, Satou Y, et al. In vivo expression of the HBZ gene of HTLV-1 correlates with proviral load, inflammatory markers and disease severity in HTLV-1 associated myelopathy/tropical spastic paraparesis (HAM/TSP). Retrovirology 2009;6:19. 306. Saitoh Y, Yamamoto N, Dewan MZ, et al. Overexpressed NF-kappaB-inducing kinase contributes to the tumorigenesis of adult T-cell leukemia and Hodgkin Reed-Sternberg cells. Blood 2008;111:5118–5129. 307. Salemi M, Desmyter J, Vandamme AM. Tempo and mode of human and simian T-lymphotropic virus (HTLV/STLV) evolution revealed by analyses of full-genome sequences. Mol Biol Evol 2000;17:374–386. 308. Sasada A, Takaori-Kondo A, Shirakawa K, et al. APOBEC3G targets human T-cell leukemia virus type 1. Retrovirology 2005;2:32. 309. Satake M, Iwanaga M, Sagara Y, et al. Incidence of human T-lymphotropic virus 1 infection in adolescent and adult blood donors in Japan: a nationwide retrospective cohort analysis. Lancet Infect Dis 2016;16:1246–1254. 310. Sato T, Coler-Reilly ALG, Yagishita N, et al. Mogamulizumab (Anti-CCR4) in HTLV-1-associated myelopathy. N Engl J Med 2018;378:529–538. 311. Satou Y, Miyazato P, Ishihara K, et al. The retrovirus HTLV-1 inserts an ectopic CTCF-binding site into the human genome. Proc Natl Acad Sci U S A 2016;113:3054–3059. 312. Satou Y, Utsunomiya A, Tanabe J, et al. HTLV-1 modulates the frequency and phenotype of FoxP3+CD4+ T cells in virus-infected individuals. Retrovirology 2012;9:46. 313. Satou Y, Yasunaga J, Yoshida M, et al. HTLV-I basic leucine zipper factor gene mRNA supports proliferation of adult T cell leukemia cells. Proc Natl Acad Sci U S A 2006;103:720–725. 314. Satou Y, Yasunaga J, Zhao T, et al. HTLV-1 bZIP factor induces T-cell lymphoma and systemic inflammation in vivo. PLoS Pathog 2011;7:e1001274. 630

315. Sawada T, Iwahara Y, Ishii K, et al. Immunoglobulin prophylaxis against milkborne transmission of human T cell leukemia virus type I in rabbits. J Infect Dis 1991;164:1193–1196. 316. Saxon A, Stevens RH, Quan SG, et al. Immunologic characterization of hairy cell leukemias in continuous culture. J Immunol 1978;120:777–782. 317. Schierhout G, McGregor S, Gessain A, et al. Association between HTLV-1 infection and adverse health outcomes: a systematic review and meta-analysis of epidemiological studies. Lancet Infect Dis 2020;20:133–143. 318. Seich al Basatena NK, MacNamara A, Vine AM, et al. KIR2DL2 enhances protective and detrimental HLA class I-mediated immunity in chronic viral infection. PLoS Pathog 2011;7:e1002270. 319. Seiki M, Hattori S, Hirayama Y, et al. Human adult T-cell leukemia virus: complete nucleotide sequence of the provirus genome integrated in leukemia cell DNA. Proc Natl Acad Sci U S A 1983;80:3618–3622. 320. Seiki M, Hikikoshi A, Taniguchi T, et al. Expression of the pX gene of HTLV-I: general splicing mechanism in the HTLV family. Science 1985;228:1532–1534. 321. Seiki M, Hikikoshi A, Yoshida M. The U5 sequence is a cis-acting repressive element for genomic RNA expression of human T cell leukemia virus type I. Virology 1990;176:81–86. 322. Seiki M, Inoue J, Takeda T, et al. The p40x of human T-cell leukemia virus type I is a trans-acting activator of viral gene transcription. Jpn J Cancer Res 1985;76:1127–1131. 323. Seiki M, Inoue J, Takeda T, et al. Direct evidence that p40x of human T-cell leukemia virus type I is a trans-acting transcriptional activator. EMBO J 1986;5:561–565. 324. Sherman MP, Saksena NK, Dube DK, et al. Evolutionary insights on the origin of human T-cell lymphoma/leukemia virus type I (HTLV-I) derived from sequence analysis of a new HTLV-I variant from Papua New Guinea. J Virol 1992;66:2556–2563. 325. Shida H, Tochikura T, Sato T, et al. Effect of the recombinant vaccinia viruses that express HTLV-I envelope gene on HTLV-I infection. EMBO J 1987;6:3379–3384. 326. Shimotohno K, Takano M, Teruuchi T, et al. Requirement of multiple copies of a 21-nucleotide sequence in the U3 regions of human T-cell leukemia virus type I and type II long terminal repeats for trans-acting activation of transcription. Proc Natl Acad Sci U S A 1986;83:8112–8116. 327. Shimoyama M. Diagnostic criteria and classification of clinical subtypes of adult T-cell leukaemia-lymphoma. A report from the Lymphoma Study Group (1984–87). Br J Haematol 1991;79:428–437. 328. Shimoyama M. Chemotherapy of ATL. In: Takatsuki K, ed. Adult T-cell leukemia. New York: Oxford University Press; 1994. 329. Silic-Benussi M, Biasiotto R, Andresen V, et al. HTLV-1 p13, a small protein with a busy agenda. Mol Aspects Med 2010;31:350–358. 330. Silic-Benussi M, Cavallari I, Vajente N, et al. Redox regulation of T-cell turnover by the p13 protein of human T-cell leukemia virus type 1: distinct effects in primary versus transformed cells. Blood 2010;116:54–62. 331. Sinha-Datta U, Datta A, Ghorbel S, et al. Human T-cell lymphotrophic virus type I rex and p30 interactions govern the switch between virus latency and replication. J Biol Chem 2007;282:14608–14615. 332. Siomi H, Shida H, Nam SH, et al. Sequence requirements for nucleolar localization of human T cell leukemia virus type I pX protein, which regulates viral RNA processing. Cell 1988;55:197–209. 333. Siu YT, Chin KT, Siu KL, et al. TORC1 and TORC2 coactivators are required for tax activation of the human T-cell leukemia virus type 1 long terminal repeats. J Virol 2006;80:7052–7059. 334. Slattery JP, Franchini G, Gessain A. Genomic evolution, patterns of global dissemination, and interspecies transmission of human and simian T-cell leukemia/lymphotropic viruses. Genome Res 1999;9:525–540. 335. Sodroski JG, Rosen CA, Haseltine WA. Trans-acting transcriptional activation of the long terminal repeat of human T lymphotropic viruses in infected cells. Science 1984;225:381–385. 336. Sommerfelt MA, Williams BP, Clapham PR, et al. Human T cell leukemia viruses use a receptor determined by human chromosome 17. Science 1988;242:1557–1559. 337. Sonoda S, Fujiyoshi T, Yashiki S. Immunogenetics of HTLV-I/II and associated diseases. J Acquir Immune Defic Syndr Hum Retrovirol 1996;13(Suppl 1):S119–S123. 338. Stuver SO, Tachibana N, Okayama A, et al. Heterosexual transmission of human T cell leukemia/lymphoma virus type I among married couples in southwestern Japan: an initial report from the Miyazaki Cohort Study. J Infect Dis 1993;167:57–65. 339. Suehiro Y, Hasegawa A, Iino T, et al. Clinical outcomes of a novel therapeutic vaccine with Tax peptide-pulsed dendritic cells for adult T cell leukaemia/lymphoma in a pilot study. Br J Haematol 2015;169:356–367. 340. Sugata K, Yasunaga J, Kinosada H, et al. HTLV-1 viral factor HBZ induces CCR4 to promote T-cell migration and proliferation. Cancer Res 2016;76:5068–5079. 341. Sugata K, Yasunaga J, Mitobe Y, et al. Protective effect of cytotoxic T lymphocytes targeting HTLV-1 bZIP factor. Blood 2015;126:1095–1105. 342. Sugata K, Yasunaga J, Miura M, et al. Enhancement of anti-STLV-1/HTLV-1 immune responses through multimodal effects of anti-CCR4 antibody. Sci Rep 2016;6:27150. 343. Sugimoto M, Nakashima H, Watanabe S, et al. T-lymphocyte alveolitis in HTLV-I-associated myelopathy. Lancet 1987;2:1220. 344. Sun SC, Yamaoka S. Activation of NF-kappaB by HTLV-I and implications for cell transformation. Oncogene 2005;24:5952–5964. 345. Suzuki T, Hirai H, Murakami T, et al. Tax protein of HTLV-1 destabilizes the complexes of NF-kappa B and I kappa B-alpha and induces nuclear translocation of NF-kappa B for transcriptional activation. Oncogene 1995;10:1199–1207. 346. Suzuki T, Uchida-Toita M, Yoshida M. Tax protein of HTLV-1 inhibits CBP/p300-mediated transcription by interfering with recruitment of CBP/p300 onto DNA element of E-box or p53 binding site. Oncogene 1999;18:4137–4143. 347. Suzuki Y, Gojobori T. The origin and evolution of human T-cell lymphotropic virus types I and II. Virus Genes 1998;16:69–84. 348. Sze A, Belgnaoui SM, Olagnier D, et al. Host restriction factor SAMHD1 limits human T cell leukemia virus type 1 infection of monocytes via STING-mediated apoptosis. Cell Host Microbe 2013;14:422–434. 349. Tagaya Y, Matsuoka M, Gallo R. 40 years of the human T-cell leukemia virus: past, present, and future. F1000Res 2019;8:F1000 Faculty Rev-228. 350. Tajima K, Inoue M, Takezaki T, et al. Ethnoepidemiology of ATL in Japan with special reference to the Mongoloid dispersal. In: Takatsuki K, ed. Adult T-cell Leukemia New York: Oxford University Press; 1994. 351. Tajima Y, Tashiro K, Camerini D. Assignment of the possible HTLV receptor gene to chromosome 17q21-q23. Somat Cell Mol Genet 1997;23:225–227. 352. Takahashi K, Takezaki T, Oki T, et al. Inhibitory effect of maternal antibody on mother-to-child transmission of human T-lymphotropic virus type I. The Mother-to-Child Transmission Study Group. Int J Cancer 1991;49:673–677. 353. Takasaki Y, Iwanaga M, Imaizumi Y, et al. Long-term study of indolent adult T-cell leukemia-lymphoma. Blood 2010;115:4337–4343. 354. Takatsuki K. Discovery of adult T-cell leukemia. Retrovirology 2005;2:16. 355. Takatsuki K, Yamaguchi K, Matsuoka M. ATL and HTLV-I-related diseases. In: Takatsuki K, ed. Adult T-cell Leukemia. New York: Oxford University Press; 1994. 631

356. Takeda S, Maeda M, Morikawa S, et al. Genetic and epigenetic inactivation of tax gene in adult T-cell leukemia cells. Int J Cancer 2004;109:559–567. 357. Tamiya S, Etoh K, Suzushima H, et al. Mutation of CD95 (Fas/Apo-1) gene in adult T-cell leukemia cells. Blood 1998;91:3935–3942. 358. Tamiya S, Matsuoka M, Etoh K, et al. Two types of defective human T-lymphotropic virus type I provirus in adult T-cell leukemia. Blood 1996;88:3065–3073. 359. Tanaka A, Matsuoka M. HTLV-1 alters T cells for viral persistence and transmission. Front Microbiol 2018;9:461. 360. Taniguchi Y, Nosaka K, Yasunaga J, et al. Silencing of human T-cell leukemia virus type I gene transcription by epigenetic mechanisms. Retrovirology 2005;2:64. 361. Taylor GP. Human T-lymphotropic virus type 1 infection and solid organ transplantation. Rev Med Virol 2018;28. 362. Taylor GP, Goon P, Furukawa Y, et al. Zidovudine plus lamivudine in Human T-Lymphotropic Virus type-I-associated myelopathy: a randomised trial. Retrovirology 2006;3:63. 363. Terol M, Gazon H, Lemasson I, et al. HBZ-mediated shift of JunD from growth suppressor to tumor promoter in leukemic cells by inhibition of ribosomal protein S25 expression. Leukemia 2017;31:2235–2243. 364. Tezuka K, Fuchi N, Okuma K, et al. HTLV-1 targets human placental trophoblasts in seropositive pregnant women. J Clin Invest 2020;130:6171–6186. 365. Thebault S, Basbous J, Hivin P, et al. HBZ interacts with JunD and stimulates its transcriptional activity. FEBS Lett 2004;562:165–170. 366. Tholouli E, Liu Yin JA. Successful treatment of HTLV-1-associated acute adult T-cell leukemia lymphoma by allogeneic bone marrow transplantation: a 12 year follow-up. Leuk Lymphoma 2006;47:1691–1692. 367. Tomasetti C, Li L, Vogelstein B. Stem cell divisions, somatic mutations, cancer etiology, and cancer prevention. Science 2017;355:1330–1334. 368. Tomasetti C, Vogelstein B. Cancer etiology. Variation in cancer risk among tissues can be explained by the number of stem cell divisions. Science 2015;347:78–81. 369. Toro C, Rodes B, Poveda E, et al. Rapid development of subacute myelopathy in three organ transplant recipients after transmission of human T-cell lymphotropic virus type I from a single donor. Transplantation 2003;75:102–104. 370. Toulza F, Heaps A, Tanaka Y, et al. High frequency of CD4+FoxP3+ cells in HTLV-1 infection: inverse correlation with HTLV-1-specific CTL response. Blood 2008;111:5047–5053. 371. Toulza F, Nosaka K, Takiguchi M, et al. FoxP3+ regulatory T cells are distinct from leukemia cells in HTLV-1-associated adult T-cell leukemia. Int J Cancer 2009;125:2375–2382. 372. Toulza F, Nosaka K, Tanaka Y, et al. Human T-lymphotropic virus type 1-induced CC chemokine ligand 22 maintains a high frequency of functional FoxP3+ regulatory T cells. J Immunol 2010;185:183–189. 373. Trevino A, Parra P, Bar-Magen T, et al. Antiviral effect of raltegravir on HTLV-1 carriers. J Antimicrob Chemother 2012;67:218–221. 374. Trovato R, Mulloy JC, Johnson JM, et al. A lysine-to-arginine change found in natural alleles of the human T-cell lymphotropic/leukemia virus type 1 p12(I) protein greatly influences its stability. J Virol 1999;73:6460–6467. 375. Tsukahara T, Kannagi M, Ohashi T, et al. Induction of Bcl-x(L) expression by human T-cell leukemia virus type 1 Tax through NF-kappaB in apoptosis-resistant T-cell transfectants with Tax. J Virol 1999;73:7981–7987. 376. Uchiyama T, Yodoi J, Sagawa K, et al. Adult T-cell leukemia: clinical and hematologic features of 16 cases. Blood 1977;50:481–492. 377. Umehara F, Izumo S, Nakagawa M, et al. Immunocytochemical analysis of the cellular infiltrate in the spinal cord lesions in HTLVI-associated myelopathy. J Neuropathol Exp Neurol 1993;52:424–430. 378. Umehara F, Nakamura A, Izumo S, et al. Apoptosis of T lymphocytes in the spinal cord lesions in HTLV-I-associated myelopathy: a possible mechanism to control viral infection in the central nervous system. J Neuropathol Exp Neurol 1994;53:617–624. 379. Unge T, Solomin L, Mellini M, et al. The Rex regulatory protein of human T-cell lymphotropic virus type I binds specifically to its target site within the viral RNA. Proc Natl Acad Sci U S A 1991;88:7145–7149. 380. Ureta-Vidal A, Angelin-Duclos C, Tortevoye P, et al. Mother-to-child transmission of human T-cell-leukemia/lymphoma virus type I: implication of high antiviral antibody titer and high proviral load in carrier mothers. Int J Cancer 1999;82:832–836. 381. Usui T, Yanagihara K, Tsukasaki K, et al. Characteristic expression of HTLV-1 basic zipper factor (HBZ) transcripts in HTLV-1 proviruspositive cells. Retrovirology 2008;5:34. 382. Usuku K, Nishizawa M, Matsuki K, et al. Association of a particular amino acid sequence of the HLA-DR beta 1 chain with HTLVI-associated myelopathy. Eur J Immunol 1990;20:1603–1606. 383. Valeri VW, Hryniewicz A, Andresen V, et al. Requirement of the human T-cell leukemia virus p12 and p30 products for infectivity of human dendritic cells and macaques but not rabbits. Blood 2010;116:3809–3817. 384. Van Dooren S, Salemi M, Vandamme AM. Dating the origin of the African human T-cell lymphotropic virus type-i (HTLV-I) subtypes. Mol Biol Evol 2001;18:661–671. 385. Van Prooyen N, Gold H, Andresen V, et al. Human T-cell leukemia virus type 1 p8 protein increases cellular conduits and virus transmission. Proc Natl Acad Sci U S A 2010;107:20738–20743. 386. Vandamme AM, Liu HF, Goubau P, et al. Primate T-lymphotropic virus type I LTR sequence variation and its phylogenetic analysis: compatibility with an African origin of PTLV-I. Virology 1994;202:212–223. 387. Vandamme AM, Salemi M, Desmyter J. The simian origins of the pathogenic human T-cell lymphotropic virus type I. Trends Microbiol 1998;6:477–483. 388. Vandamme AM, Salemi M, Van Brussel M, et al. African origin of human T-lymphotropic virus type 2 (HTLV-2) supported by a potential new HTLV-2d subtype in Congolese Bambuti Efe Pygmies. J Virol 1998;72:4327–4340. 389. Verdonck K, Gonzalez E, Schrooten W, et al. HTLV-1 infection is associated with a history of active tuberculosis among family members of HTLV-1-infected patients in Peru. Epidemiol Infect 2008;136:1076–1083. 390. Verdonck K, Gonzalez E, Van Dooren S, et al. Human T-lymphotropic virus 1: recent knowledge about an ancient infection. Lancet Infect Dis 2007;7:266–281. 391. Vernin C, Thenoz M, Pinatel C, et al. HTLV-1 bZIP factor HBZ promotes cell proliferation and genetic instability by activating OncomiRs. Cancer Res 2014;74:6082–6093. 392. Vine AM, Witkover AD, Lloyd AL, et al. Polygenic control of human T lymphotropic virus type I (HTLV-I) provirus load and the risk of HTLV-I-associated myelopathy/tropical spastic paraparesis. J Infect Dis 2002;186:932–939. 393. Watanabe T, Yamaguchi K, Takatsuki K, et al. Constitutive expression of parathyroid hormone-related protein gene in human T cell leukemia virus type 1 (HTLV-1) carriers and adult T cell leukemia patients that can be trans-activated by HTLV-1 tax gene. J Exp Med 1990;172:759–765. 394. Wattel E, Cavrois M, Gessain A, et al. Clonal expansion of infected cells: a way of life for HTLV-I. J Acquir Immune Defic Syndr Hum Retrovirol 1996;13(Suppl 1):S92–S99. 395. Weichselbraun I, Farrington GK, Rusche JR, et al. Definition of the human immunodeficiency virus type 1 Rev and human T-cell leukemia virus type I Rex protein activation domain by functional exchange. J Virol 1992;66:2583–2587. 396. Wen AY, Sakamoto KM, Miller LS. The role of the transcription factor CREB in immune function. J Immunol 2010;185:6413–6419. 632

397. Wiktor SZ, Pate EJ, Rosenberg PS, et al. Mother-to-child transmission of human T-cell lymphotropic virus type I associated with prolonged breast-feeding. J Hum Virol 1997;1:37–44. 398. Wolfe ND, Heneine W, Carr JK, et al. Emergence of unique primate T-lymphotropic viruses among central African bushmeat hunters. Proc Natl Acad Sci U S A 2005;102:7994–7999. 399. Worobey M, Gemmel M, Teuwen DE, et al. Direct evidence of extensive diversity of HIV-1 in Kinshasa by 1960. Nature 2008;455:661–664. 400. Xiao G, Cvijic ME, Fong A, et al. Retroviral oncoprotein Tax induces processing of NF-kappaB2/p100 in T cells: evidence for the involvement of IKKalpha. EMBO J 2001;20:6805–6815. 401. Xie L, Green PL. Envelope is a major viral determinant of the distinct in vitro cellular transformation tropism of human T-cell leukemia virus type 1 (HTLV-1) and HTLV-2. J Virol 2005;79:14536–14545. 402. Yamada Y, Tomonaga M, Fukuda H, et al. A new G-CSF-supported combination chemotherapy, LSG15, for adult T-cell leukaemialymphoma: Japan Clinical Oncology Group Study 9303. Br J Haematol 2001;113:375–382. 403. Yamagishi M, Nakano K, Miyake A, et al. Polycomb-mediated loss of miR-31 activates NIK-dependent NF-kappaB pathway in adult T cell leukemia and other cancers. Cancer Cell 2012;21:121–135. 404. Yamamoto K, Utsunomiya A, Tobinai K, et al. Phase I study of KW-0761, a defucosylated humanized anti-CCR4 antibody, in relapsed patients with adult T-cell leukemia-lymphoma and peripheral T-cell lymphoma. J Clin Oncol 2010;28:1591–1598. 405. Yamano Y, Araya N, Sato T, et al. Abnormally high levels of virus-infected IFN-gamma+ CCR4+ CD4+ CD25+ T cells in a retrovirusassociated neuroinflammatory disorder. PLoS One 2009;4:e6517. 406. Yamaoka S, Courtois G, Bessia C, et al. Complementation cloning of NEMO, a component of the IkappaB kinase complex essential for NFkappaB activation. Cell 1998;93:1231–1240. 407. Yamato K, Oka T, Hiroi M, et al. Aberrant expression of the p53 tumor suppressor gene in adult T-cell leukemia and HTLV-I-infected cells. Jpn J Cancer Res 1993;84:4–8. 408. Yang L, Kotomura N, Ho YK, et al. Complex cell cycle abnormalities caused by human T-lymphotropic virus type 1 Tax. J Virol 2011;85:3001–3009. 409. Yasuda N, Lai PK, Ip SH, et al. Soluble interleukin 2 receptors in sera of Japanese patients with adult T cell leukemia mark activity of disease. Blood 1988;71:1021–1026. 410. Yasuma K, Yasunaga J, Takemoto K, et al. HTLV-1 bZIP factor impairs anti-viral immunity by inducing co-inhibitory molecule, T cell immunoglobulin and ITIM domain (TIGIT). PLoS Pathog 2016;12:e1005372. 411. Yasunaga J, Taniguchi Y, Nosaka K, et al. Identification of aberrantly methylated genes in association with adult T-cell leukemia. Cancer Res 2004;64:6002–6009. 412. Yoshida M. Multiple viral strategies of HTLV-1 for dysregulation of cell growth control. Annu Rev Immunol 2001;19:475–496. 413. Yoshida M, Satou Y, Yasunaga J, et al. Transcriptional control of spliced and unspliced human T-cell leukemia virus type 1 bZIP factor (HBZ) gene. J Virol 2008;82:9359–9368. 414. Yoshie O, Fujisawa R, Nakayama T, et al. Frequent expression of CCR4 in adult T-cell leukemia and human T-cell leukemia virus type 1-transformed T cells. Blood 2002;99:1505–1511. 415. Yoshimura T, Fujisawa J, Yoshida M. Multiple cDNA clones encoding nuclear proteins that bind to the tax-dependent enhancer of HTLV-1: all contain a leucine zipper structure and basic amino acid domain. EMBO J 1990;9:2537–2542. 416. Yoshita M, Higuchi M, Takahashi M, et al. Activation of mTOR by human T-cell leukemia virus type 1 Tax is important for the transformation of mouse T cells to interleukin-2-independent growth. Cancer Sci 2012;103(2):369–374. 417. Yu F, Itoyama Y, Fujihara K, et al. Natural killer (NK) cells in HTLV-I-associated myelopathy/tropical spastic paraparesis-decrease in NK cell subset populations and activity in HTLV-I seropositive individuals. J Neuroimmunol 1991;33:121–128. 418. Yuen CK, Chan CP, Fung SY, et al. Suppression of type I interferon production by human T-cell leukemia virus type 1 oncoprotein tax through inhibition of IRF3 phosphorylation. J Virol 2016;90:3902–3912. 419. Yukitake M, Sueoka E, Sueoka-Aragane N, et al. Significantly increased antibody response to heterogeneous nuclear ribonucleoproteins in cerebrospinal fluid of multiple sclerosis patients but not in patients with human T-lymphotropic virus type I-associated myelopathy/tropical spastic paraparesis. J Neurovirol 2008;14:130–135. 420. Zhang X, Hakata Y, Tanaka Y, et al. CRM1, an RNA transporter, is a major species-specific restriction factor of human T cell leukemia virus type 1 (HTLV-1) in rat cells. Microbes Infect 2006;8:851–859. 421. Zhao LJ, Giam CZ. Interaction of the human T-cell lymphotrophic virus type I (HTLV-I) transcriptional activator Tax with cellular factors that bind specifically to the 21-base-pair repeats in the HTLV-I enhancer. Proc Natl Acad Sci U S A 1991;88:11445–11449. 422. Zhao LJ, Giam CZ. Human T-cell lymphotropic virus type I (HTLV-I) transcriptional activator, Tax, enhances CREB binding to HTLV-I 21-base-pair repeats by protein-protein interaction. Proc Natl Acad Sci U S A 1992;89:7070–7074. 423. Zhao T, Satou Y, Sugata K, et al. HTLV-1 bZIP factor enhances TGF-{beta} signaling through p300 coactivator. Blood 2011;118:1865–1876. 424. Zhao T, Yasunaga J, Satou Y, et al. Human T-cell leukemia virus type 1 bZIP factor selectively suppresses the classical pathway of NFkappaB. Blood 2009;113:2755–2764. 425. Zhi H, Guo X, Ho YK, et al. RNF8 dysregulation and down-regulation during HTLV-1 infection promote genomic instability in adult T-Cell leukemia. PLoS Pathog 2020;16:e1008618. 426. Zhi H, Yang L, Kuo YL, et al. NF-kappaB hyper-activation by HTLV-1 tax induces cellular senescence, but can be alleviated by the viral anti-sense protein HBZ. PLoS Pathog 2011;7:e1002025. 427. Zhou M, Lu H, Park H, et al. Tax interacts with P-TEFb in a novel manner to stimulate human T-lymphotropic virus type 1 transcription. J Virol 2006;80:4781–4791.

633

CHAPTER 17 Human Immunodeficiency Viruses: Replication Melanie Ott • Eric O. Freed Introduction Classification and origins of HIVs Genomic organization of HIVs Biology of HIV infections HIV replication and tropism HIV transmission HIV-1 animal models Chimpanzees SIV models in nonhuman primates Humanized mouse models Molecular biology of HIV-1 replication Overview Virus binding and entry: the Env glycoproteins Postentry trafficking of the HIV capsid Reverse transcription Nuclear import Postentry blocks to lentiviral infection Integration Viral gene expression Virus assembly and release: the Gag proteins Env glycoprotein incorporation into virions Protease (PR) and virus maturation The accessory proteins Perspectives Acknowledgments

INTRODUCTION In the late 1970s and early 1980s, previously healthy individuals in the United States and Europe presented with symptoms of immunologic dysfunction, including generalized lymphadenopathy, opportunistic infections, and a variety of unusual cancers (non-Hodgkin lymphoma and Kaposi’s sarcoma). A common accompanying laboratory finding in affected individuals was marked depletion of the CD4+ T lymphocyte subset in the peripheral blood. The disease was first brought to the attention of the general medical community in June 1981.489 Within several months, it became clear that a similar immunodeficiency syndrome, which came to be known as acquired immunodeficiency syndrome, or AIDS, was also affecting other groups, including hemophiliacs, blood transfusion recipients, recent Haitian immigrants, and, most significantly, sexual partners or children of members of the various risk groups.

The emerging epidemiological pattern suggested that the new disease was transmitted by a novel pathogen in contaminated blood or following sexual intercourse with an affected individual. In the spring of 1983, Montagnier, Barre-Sinoussi, and their colleagues at the Pasteur Institute in Paris reported the isolation of a virus from the lymph nodes of an individual who presented with generalized lymphadenopathy of unknown origin.30 During its replication in cultured cells, the virus, termed lymphadenopathy-associated virus (LAV), released high titers of progeny virions that contained magnesium-dependent reverse transcriptase (RT) activity and exhibited electron microscopic (EM) features typical of retroviruses. However, unlike the commonly studied retroviruses, LAV was highly cytopathic in human peripheral blood mononuclear cells (PBMCs), specifically killing CD4+ T lymphocytes in cell cultures.493 Gallo and colleagues at the National Institutes of Health subsequently reported the isolation of a nearly identical retrovirus from tissue culture samples obtained from the Pasteur Institute group. Gallo and colleagues 634

named this virus human T-cell leukemia virus type III (HTLV-III) to distinguish it from the noncytopathic HTLV-1 and HTLV-2 and obtained the first serological evidence linking exposure to LAV-like retroviruses and immunodeficient individuals from the various groups at risk.565,623 The new retrovirus, associated with AIDS in the United States, Europe, and central Africa, and exhibiting morphologic and genetic characteristics typical of the lentivirus genus (Fig. 17.1), was named human immunodeficiency virus, or HIV136 (and subsequently HIV-1). In 1986, a related but immunologically distinct human retrovirus (now called HIV-2) was recovered from individuals residing in several West African countries.134

CLASSIFICATION AND ORIGINS OF HIVS The observation of particle-associated RT activity placed the new agent in the Retroviridae family. EM analysis showed that the mature HIV-1 particles contained a cone-shaped, cylindrical core reminiscent of that previously described for visna virus (Fig. 17.1).259 The cloning and sequencing of proviral DNA, initially purified from productively infected cultures of PBMC/T-cell leukemia lines (T-cell lines), indicated that HIV-1 not only possessed a genomic organization related to that of other replication–competent retroviruses but placed it taxonomically in the lentivirus genus.584,724 This relationship is shown diagrammatically in Figure 2 of Chapter 15. As their name suggests, lentiviruses (lenti, slow) were known to cause slow, unremitting disease in sheep, goats, and horses, and to target various lineages of hematopoietic cells, particularly monocytes/macrophages and lymphocytes.

FIGURE 17.1 Morphology of HIV-1 particles. Electron micrograph showing an HIV-1 particle in the process of budding from infected cultured human peripheral blood mononuclear cells (PBMCs) (arrowhead) and several particles containing the conical core characteristic of mature, infectious HIV-1 virions (arrows) (100,000× magnification). (Courtesy of Dr. Jan Orenstein.). After the isolation, molecular cloning, and initial classification of HIV-1, several genetically distinct primate lentiviruses were discovered and their phylogenetic relationships to HIV-1 were determined. For example, viruses isolated from captive macaques or feral monkey species in Africa were shown to possess particle morphologies and genomic organizations similar to those of HIV-1 (described in Chapter 19). Because inoculation of Asian macaque species, such as rhesus monkeys, with some of these newly discovered viruses recovered from African monkeys induced an AIDS-like illness,157 these viruses were named simian immunodeficiency virus (SIV) to distinguish them from the human viruses, HIV-1 and HIV-2. The detailed genetic interrelationships of members of the primate lentivirus genus are presented in Figure 1 of Chapter 19. HIV-2 is more closely related to SIVsmm,299 a virus indigenous to African sooty mangabey monkeys, than to HIV-1; HIV-2 likely arose through a zoonotic transmission of SIVsmm from monkeys to humans.248,649

Based on previously studied replication–competent retroviruses, it was originally anticipated that HIV-1 would be genetically homogeneous. However, early comparisons of proviral DNAs from Europe, North America, and Africa revealed extensive genetic heterogeneity even among sequences derived from a single individual.41 Although nucleotide changes were distributed throughout the HIV-1 genome, the greatest variability occurred in the gene encoding the envelope (Env) glycoprotein, gp160. The term “quasispecies” was subsequently coined to describe the pool of diverse and changing populations of virus present in an HIV-1–infected individual.487 Several factors contribute to the extraordinary genetic heterogeneity of HIV-1: (a) error-prone viral DNA synthesis during reverse transcription, (b) high recombination frequencies accompanying reverse transcription, (c) large population size in vivo, and (d) continuous pressure from the host to select for new variants.

The earliest phylogenetic analyses of HIV-1 isolates focused on samples from Europe, North America, and Africa; discrete clusters of viruses were identified from these geographical regions. Distinct genetic subtypes or clades of HIV-1 were subsequently defined and classified into four groups: M (main); O (outlier); N (non-M, non-O); and P (Fig. 17.2A).291 The M group of HIV-1, which comprises greater than 99% of the global virus isolates, consists of nine subtypes (A, B, C, D, F, G, H, J, and K) and several dozen circulating recombinant forms (CRFs), which arose by the intermixing of viruses cocirculating in a particular geographical locale.443,480,547,600 Subtype B viruses, the most intensively studied HIV-1 subtype, are the most prevalent isolates in Europe, North America, and Australia. Subtype C strains are responsible for greater than 50% of infections globally. HIV-1 group O isolates were recovered from individuals living in Cameroon, Gabon, and Equatorial Guinea.691 The overall prevalence of this group has declined over time. Group N and P isolates are extremely rare.

A primate lentivirus designated SIVcpz has a genomic structure very similar to that of HIV-1, including its signature vpu gene. SIVcpz was isolated from two chimpanzee subspecies, Pan troglodytes troglodytes (ptt) and Pan troglodytes schweinfurthii (pts). The prevalence of SIVcpz recovered from ptt animals (SIVcpzPtt) in the wild varies widely compared to the more even and higher distribution of SIVs infecting sooty mangabeys and African green monkeys. Phylogenetically, the SIVcpzPtt strains are related to HIV-1 groups M and N but not to group O or SIVcpzPts (Fig. 17.2B) and geographically separated chimpanzee populations carry distinct genetic lineages of SIVcpzPtt.369 Cross-species transmission of SIVcpz to humans in Central Africa and SIVsm in West Africa gave rise to HIV-1 and HIV-2, respectively.247,248 Based on nucleotide substitutions over time, a common ancestor of HIV-1 group M has been proposed to have emerged in the 1920s,742 whereas 635

progenitors for HIV-2 groups A and B have been dated to the 1940s.415 The SIVcpz strains giving rise to HIV-1 have been introduced into humans at least four times (as reflected in groups M, N, O, and P). It is thought that the source of HIV-1 group M, the cause of the worldwide AIDS epidemic, originated in ptt chimpanzees living in southeast Cameroon.369 SIVcpz is itself a recombinant between two other SIVs, SIVrcm and SIVgsn, which likely coinfected a chimpanzee.650 Because of the high degree of genetic relatedness between chimpanzees and humans, the propagation of SIVcpz in chimpanzees likely increased its ability to replicate in humans when the subsequent interspecies jump occurred.

FIGURE 17.2 HIV-1 genetic subtypes and their worldwide distribution. A: Phylogenetic relationships of HIV-1 groups M, N, O, and P with different SIV (SIVcpz and SIVgpr) isolates. (Reprinted from Hemelaar J. The origin and diversity of the HIV-1 pandemic. Trends Mol Med 2012;18(3):182–192. Copyright © 2011 Elsevier. With permission.) B: The global prevalence of HIV-1 subgroups with the predominant clades or circulating recombinant forms (CRFs) in each geographical region indicated. (Provided by Dr. Sodsai Tovanabutra, U.S. Military HIV Research Program, Henry Jackson Foundation.)

GENOMIC ORGANIZATION OF HIVS Nucleotide sequencing of several of the original HIV-1 isolates revealed that in contrast to the well-characterized and intensively studied prototypical retroviruses, whose genomes often contain only three genes (gag, pol, and env) encoding the structural proteins and enzymes required for productive infection, the HIV-1 genome included several additional and overlapping open reading frames (ORFs) of unknown function (Fig. 17.3). Not only did HIV-1 and HIV-2 contain multiple additional ORFs but also their genomic organizations appeared to be very similar. Further analyses revealed that HIV-1 contained the distinguishing vpu gene141,681 and HIV-2 carried a signature vpx361 gene.

Like all replication–competent orthoretroviruses, the three primary translation products from the gag, pol, and env genes are initially synthesized as polyprotein precursors, which are subsequently processed by viral or cellular proteases into mature, particle-associated proteins (Fig. 17.4). The 55-kDa Gag precursor, Pr55Gag, is cleaved into the matrix (MA), capsid (CA), nucleocapsid (NC), and p6 proteins, and the two spacer peptides SP1 and SP2 during or shortly after the release of progeny virions. Autocatalysis of the 160 kDa Gag–Pol polyprotein, Pr160GagPol, gives rise to the protease (PR), the heterodimeric RT, and integrase (IN) enzymes, whereas proteolytic digestion by a cellular enzyme in the Golgi converts the glycosylated 160-kDa Env precursor, gp160, into the gp120 surface (SU) and gp41 transmembrane (TM) cleavage products. The remaining six HIV-1–encoded proteins (Vif, Vpr, Tat, Rev, Vpu, and Nef) are the primary translation products of spliced mRNAs, as is the Env glycoprotein.

HIV-1 and HIV-2 have incorporated multiple sequence elements into their genomic RNAs that direct the balanced and coordinated production of progeny virions.55,72 The 5′ approximately 400 nucleotide untranslated region of HIV-1 genomic RNA is highly structured and contains multiple elements that mediate transcriptional elongation of viral RNA transcripts, splicing, genomic RNA dimerization, packaging of full-length viral RNA, and reverse transcription (Fig. 17.5A). These include the dimerization initiation signal (DIS), a palindromic sequence that promotes RNA dimerization and packaging into virions (Fig. 17.5A and B).

FIGURE 17.3 Genomic organization of simple and complex retroviruses. The genes of murine leukemia virus (MLV), human T-cell leukemia virus (HTLV), HIV-1, and HIV-2 are depicted as they are arranged in their respective proviral DNA. The sizes of the different proviral DNAs are shown in proportion to the 9.7-kb HIV provirus.

636

FIGURE 17.4 HIV-1–encoded proteins. The location of the HIV genes, the molecular masses (in kDa) of primary translation products (in some cases, polyproteins), and the processed mature viral proteins are indicated. Three regions encompassing the primer binding site (PBS) stem participate in the placement and stabilization of the tRNALys3 primer, which is incorporated into HIV-1 particles and is required for the initiation of reverse transcription. These regions include the PBS itself, which can base pair with the 3′-terminal 18 nucleotides of tRNALys3; the primer activation signal (PAS), which interacts with the thymidine-pseudouridinecytidine (TψC) arm of the tRNA to trigger the reverse transcription reaction; and the A-loop, which is complementary to the anticodon loop of tRNALys3 (Reviewed in Ref.57). The unwinding of both the tRNA primer and the PBS stem within the 5′ untranslated region of the HIV-1 genome, as well as the incorporation of the primer into nascent virions, is facilitated by the NC domain of the Gag polyprotein, which plays the role of a molecular chaperone in this process. This topic, and genomic RNA packaging, will be discussed in more detail below. Lysyl-tRNA synthetase has been reported to bind Gag and facilitate tRNALys3 incorporation into virions.334

FIGURE 17.5 A: Model of the secondary structure at the 5′ terminus of HIV-1 genomic RNA. The positions of the TAR stem-loop, poly(A) stem-loop, primer binding site (PBS), dimerization initiation sequence (DIS), major splice donor (SD) (stem loop 2), ψ stem-loop (SL3), and AUG stem loop (SL4). The AUG serves as translation initiation codon for Gag. B: The self-complementary dimerization initiation sequence (DIS), located at the crown of stem-loop 1, participates in the formation of “kissing loop” intermediates, an initial step in the RNA dimerization reaction. C: RNA stem-loop structure downstream of the UUUUUUA frameshifting sequence. (Penal A adapted with permission of Annual Reviews, Inc. from Bieniasz P, Telesnitsky A. Multiple, switchable protein: RNA interactions regulate human immunodeficiency virus type 1 assembly. Annu Rev Virol 2018;5(1):165–183; permission conveyed through Copyright Clearance Center, Inc.) The viral genomic RNA contains a heptameric UUUUUUA “slippery” sequence within the gag gene where ribosomal frameshifting (FS) can occur during translation of the full-length viral RNA (Fig. 17.5C). This sequence functions in conjunction with a downstream 8-nt spacer and hairpin to mediate (−)1 translational frameshifting at a frequency of about 5%.329 The viral RNA folds into other structures including the transactivation response region (TAR, near the 5′ end of the viral RNA) and Rev-responsive element (RRE, within the env gene), which are involved in RNA synthesis and RNA nuclear-to-cytoplasmic transport, respectively. Multiple inhibitory sequences (INS), associated with the instability or nuclear retention of HIV transcripts, are scattered throughout genes encoding the gag, pol, and env genes.452,604 Finally, although not strictly considered a cis-acting element, HIV-1 genomic RNA contains a significantly higher adenosine (A) content (~39%) than does mammalian DNA; this bias contributes to an unusual codon usage.45

BIOLOGY OF HIV INFECTIONS The main cellular targets of HIV-1 in infected individuals are CD4+ T lymphocytes, which bear a high surface density of CD4. CD4+ cells of the macrophage lineage are efficiently infected by evolutionary variants of HIV-1 selected for the ability to use a low density of surface CD4 for entry. A quintessential property of HIV-1 and the other primate lentiviruses is to sequentially use CD4 and a second receptor (the coreceptor) during entry into susceptible cells.

HIV Replication and Tropism Two coreceptors are predominantly used by primate lentiviruses to infect CD4+ cells: the CC chemokine receptor CCR5 and the CXC chemokine receptor CXCR4. Virus isolates that use CCR5 are classified as “R5-tropic,” viruses that use CXCR4 are designated “X4-tropic,” and strains that can use either coreceptor are referred to as “X4/R5-tropic” or “dual-tropic”. Because X4-tropic viruses evolve from R5-tropic viruses the dualtropic designation is somewhat artificial in that these are viruses that are evolving to use CXCR4 with few primary isolates ever being exclusively X4-tropic. Primary T cells express both CCR5 and CXCR4 and can be infected by all three groups of virus isolates. In contrast to primary CD4+ T cells, many immortalized T-cell lines, which are used extensively to study HIV-1 replication in tissue culture, express CXCR4 but not CCR5. Most T-cell line adapted HIV-1 strains thus use CXCR4 as their coreceptor. Activated PBMCs in culture are highly susceptible to HIV-1 637

infection. In contrast, resting PBMCs are largely refractory to HIV-1 infection676,774 at least in part due to inefficient reverse transcription resulting from low dNTP pools in these cells.

R5-tropic strains are the predominant viruses detected in recently infected individuals and throughout the asymptomatic phase of untreated HIV-1 infection; in addition, the transmitted/asymptomatic virus requires a high density of CD4 for efficient entry and thus can be called R5 T cell-tropic. In approximately 50% of untreated individuals infected with subtype B HIV-1, X4-tropic variants eventually emerge, often coincident with accelerated CD4+ T-cell depletion and more rapid progression to AIDS.

The role that macrophages play in HIV-1 infection remains incompletely understood. Although macrophages express CCR5, they are inefficiently infected by most R5-tropic strains of HIV-1 (i.e., the R5 T cell-tropic form of HIV-1), in part because of low surface expression of CD4 in this cell type. Viruses capable of infecting macrophages are thus adapted to use low CD4 levels during infection and can be referred to as macrophage-tropic (or M-tropic).351 In addition, macrophages express the restriction factor sterile alpha motif and histidine-aspartate domain– containing protein 1 (SAMHD1), which blocks efficient reverse transcription. Members of the HIV-2/SIVsmm/SIVmac lineage of primate lentiviruses express Vpx, which counteracts SAMHD1, but HIV-1 isolates do not encode a vpx gene (see section “Vpx”). In infected individuals, macrophage infection is relatively rare, except in the brain and central nervous system (CNS) and late in disease in untreated individuals when CD4+ T cells have been severely depleted (for review, see Ref.490). HIV-1 can be found in the CNS very soon after transmission initially via the trafficking of infected lymphocytes, which release R5 T cell-tropic virus. HIV-1 does not productively infect neurons but instead targets perivascular macrophages and parenchymal microglial cells, which also express a low density of surface CD4. The establishment of persistent infection within the CNS appears to require the evolution of M-tropic virus given the dearth of CD4+ T cells in the CNS compartment.

Under some circumstances, DCs can be infected with (or at least capture) HIV-1. DCs are antigen-presenting cells that capture, transport, and present antigens to CD4+ and CD8+ T lymphocytes. Three main subtypes of DCs have been identified: plasmacytoid DCs (PDCs) and conventional DCs (cDCs) (previously called myeloid DCs), which are further classified into cDC1 and cDC2 subsets. cDCs reside in multiple tissues including the skin (where they are called Langerhans cells) and the intestinal and genital tract mucosa. PDCs are found in the blood, T-cell zones of lymph nodes, and thymus and can be recruited to sites of inflammation. cDCs present near mucosal surfaces may represent the first line of defense against sexually transmitted HIV-1, transporting virus particles from this portal of entry to draining lymph nodes where they are degraded into antigenic peptides for presentation to CD4+ T lymphocytes. Paradoxically, this critical initial response exacerbates the HIV-1 acute infection; although productive HIV-1 infection of DCs is very inefficient, vigorous virus replication is observed when DCs are first pulsed with virus and then cocultivated with CD4+ T lymphocytes, a process called transinfection.253 During this process, HIV-1 particles bound to the DC surface are internalized and then “presented” to susceptible CD4+ T cells at points of close cell–cell contact, utilizing adhesion molecules involved in the formation of the immunologic synapse.395

HIV Transmission HIV can spread through two general mechanisms: (a) cell-free infection and (b) cell–cell transfer across points of close cell–cell contact, referred to as the virological synapse (VS). Early studies reported that cell-to-cell transmission of HIV-1 is orders of magnitude more efficient than cell-free infection.185 The formation of a VS with T cells, first reported for HTLV-1,321 utilizes some of the components of the immunological synapse machinery involved in the interaction between lymphocytes and antigen-presenting cells.6 The formation of the HIV-1 VS is initiated by the binding of gp120 on the surface of the infected cell to CD4 molecules on the target cell.347 This conjugate is further stabilized by interactions between cellular adhesion molecules, for example, lymphocyte function associated antigen (LFA) and intercellular adhesion molecules (ICAMs), which promote additional recruitment of actin, viral proteins (Gag and Env), CD4, and integrins to cell–cell contact regions. Cytoskeletal remodeling drives virus assembly in the infected cell to the plasma membrane and toward the target cell. Progeny particles released into the synaptic space are then able to directly enter the target cell.625 The copolarization of clustered viral proteins in the donor cell with receptors in multiple adjacent recipients can also lead to the formation of polysynaptic structures.358 Inhibitors of HIV-1 replication, including neutralizing antibodies, are able to block both cell-free and cell-to-cell infection, but transfer across the VS may allow the virus to be less susceptible to inhibition.118,782,783 Consistent with this hypothesis, Env mutations that enhance the efficiency of cell–cell transfer reduce the susceptibility of HIV-1 to antiretroviral drugs, at least in cell culture.714

The genetic diversity of HIV-1 isolates in most newly infected individuals is quite low, indicating that transmission from one individual to another is mediated by a single virus, or a small number of viruses.739 Over time, in the absence of ART, viral diversity increases markedly as a result of high levels of ongoing, error-prone virus replication and evasion of the host immune response. When ART is initiated, viral loads (as measured by the number of viral RNA copies per ml of plasma) rapidly undergo a multi-log decline to levels below the detection limit of standard clinical assays. The development of highly sensitive PCR-based assays to measure viral loads revealed a low level of viremia in the majority of individuals on ART; this low-level viremia is highly stable over many years on therapy.536 The viral genetic diversity in individuals on suppressive ART does not change over time, and drug resistance mutations typically do not arise. These observations suggest that the lowlevel viremia that persists during suppressive ART is not a result of ongoing rounds of virus replication but rather is generated by a population of latently infected cells that produce virion-associated viral RNA that can be measured in the plasma. This latent reservoir is thought to reside primarily in resting memory CD4+ T cells133,217,741; for review see Ref.645 Most latently infected cells harbor defective integrated viral genomes (integrated viral DNA is often referred to as a provirus) but some contain intact proviruses capable of producing replication-competent virus. Latently infected cells have been shown to clonally expand,142,453,723 with every cell in an expanded clone containing a single provirus integrated at an identical site in the human genome. In relatively rare cases, by integrating in host cell genes that control cell proliferation, the integrated provirus contributes to the expansion of the infected cell clone. In the majority of instances, however, clonal expansion is driven not by the integrated provirus but by homeostatic proliferation or antigen stimulation.139,658 Expanded cell clones containing intact proviruses provide a source of replication-competent HIV-1 that can rapidly restore viral loads to pretherapy levels if ART is discontinued. It may be that all proviruses in latently infected cells are maintained by cell replication/expansion, although this is a difficult point to prove with existing technologies.

HIV-1 ANIMAL MODELS Chimpanzees 638

In the search for an HIV-1 animal model during the early phase of the AIDS epidemic, cell suspensions from virus-infected individuals were inoculated into a variety of mammalian species including nonhuman primates, but only chimpanzees consistently became infected. It was subsequently shown that asymptomatic HIV-1 infection could readily be established in chimpanzees, but viremia was not maintained and no long-standing impairment to the immune system occurred.19 This could reflect genetic changes that occurred in the SIVcpz progenitor of HIV-1

following its transmission from chimpanzees and adaptation to humans.626 The failure of chimpanzees to develop disease in a timely fashion following inoculation with tissue culture–adapted or patient-derived HIV-1 isolates, coupled with their endangered-species status, has stopped their use as an animal model of HIV-1–induced immunodeficiency.

SIV Models in Nonhuman Primates When it became apparent that humans and chimpanzees were the only mammalian species that could be infected by HIV-1, attention turned to the SIV/Asian macaque model. Infection of some macaque species with SIV causes a persistent pathogenic infection with similar progression to AIDS as observed in HIV-infected patients and is today widely used for cure-based studies. Rhesus macaques infected with either the reference swarm SIVmac251 or with the SIVmac251-derived infectious molecular clone SIVmac239 reproduce several aspects of human HIV infections, including sustained, high viral loads, progressive depletion of mucosal CD4+ T cells, and chronic immune activation.273,399 But despite these similarities, several limitations exist. The most critical for cure research are the overall higher viral loads, and the natural resistance to nonnucleoside RT inhibitors exhibited by SIV strains.171,592,736 Due to these features, SIV infection is more difficult to control with ART in macaques compared to HIV infection.284,656,686 Other inherent differences between HIV and SIV include Vpx, an accessory protein encoded in SIV, but not HIV. As noted above, Vpx antagonizes the host SAMHD1 protein, which restricts replication of lentiviruses by depleting the deoxynucleotriphosphate pool during reverse transcription.74

In addition to different macaque species, HIV-1 research has also utilized African NHPs that are natural hosts of SIVs, such as African green monkeys, sooty mangabeys, and mandrills. SIV infection of these species with their naturally occurring SIV strains generally does not progress to AIDS, therefore, permitting comparative studies aimed at identifying the correlates of immune protection and the lack of disease progression.106,538,663

Viruses that contain both SIV and HIV gene segments have been used in many studies in NHPs. The earliest versions of these SIV/HIV chimeric viruses (SHIVs) consisted of the genetic backbone of SIVmac239, into which the HIV-1 tat, rev, vpu, env, and, in some instances, portions of vpr and nef genes were inserted. Infection with this virus causes a very aggressive disease phenotype with death from immunodeficiency within 3 to 6 months of inoculation; it was widely used for vaccine studies from 2000 to 2004.322,342,590 Simian-tropic HIV-1 clones have also been developed that are adapted to replicate in monkey cells both in culture and in animals.630

Humanized Mouse Models As mice are inherently nonpermissive to HIV infection due to several blocks in the viral replication cycle including transcription, mice engrafted with human cells and tissue have been utilized as a small-animal model for studies of HIV-1 pathogenesis and vaccine development. In very early studies, severe combined immunodeficiency (SCID) mice, engrafted with human PBMC or fragments of human fetal thymus and fetal liver engrafted under the renal capsule, were shown to support HIV-1 replication and exhibit moderate to profound depletions of CD4+ T lymphocytes but generated no immune responses to the virus.502,518 Relatively low levels of human cell engraftment were achieved in these first-generation humanized mice.

Although the subsequent development of nonobese diabetic (NOD)-SCID animals resulted in improved human PBMC engraftment, the relatively short lifespan of the mice and residual murine NK-cell activity limited their usefulness. Targeted mutations of the mouse interleukin2-receptor-γ chain (γc−/−) and reconstitutions with PBMC, hematopoietic stem cells (HSC), or human cord blood greatly increased engraftment of human tissue.325 Inoculation of these mice with HIV-1 by parenteral and mucosal routes resulted in sustained disseminated infections.26,44

A common engraftment method of human cells is the intravenous or intrahepatic injection of CD34+ HSCs546 into adult or newborn immunodeficient mice, respectively, after myeloablative irradiation or administration of myeloablative doses of drugs.287 The unique engraftment method using surgical implantation of human fetal liver and thymus tissues followed by injection of matched CD34+ HSCs gave rise to the bone marrow liver thymus (BLT) model.73,363,402 The human thymic tissue allows for T-cell education in the context of human cells,660 and the BLT model has become the current gold standard for studying HIV-1 immune responses.250,363 However, limitations include postengraftment graft versus host disease that reduces the lifespan of these mice, the lack of functional B-cell populations, and the technical expertise required to establish and maintain the BLT mice. Humanized mouse models are being continuously improved through genetic manipulations and “further humanization” including knock-in of human HLA alleles, the thymic stromal cell–derived lymphopoietin, and human myeloid–promoting cytokines.254

MOLECULAR BIOLOGY OF HIV-1 REPLICATION Overview The HIV-1 replication cycle (Fig. 17.6) begins with the binding of Env on the surface of virus particles to CD4 molecules on susceptible target cells. Although binding of Env to CD4 is generally essential for HIV infectivity, as noted above the subsequent interaction of Env with a coreceptor—CCR5 or CXCR4—is required for membrane fusion and entry (Reviewed in Ref.113). Unlike some other enveloped viruses that enter cells by receptor-mediated endocytosis, HIV-1 and many other retroviruses fuse directly with the plasma membrane under most conditions. After membrane fusion, the capsid core traffics along the microtubule network where reverse transcription is initiated during movement to the nucleus (Reviewed in Ref.98). The partially double-stranded DNA reverse transcription product is transported through the cytoplasm and to the nucleus as a component of a reverse transcription complex (RTC), which contains the viral enzymes RT and IN and the Gag proteins CA and NC. As noted earlier, lentiviruses are unique among retroviruses in generating RTCs that are actively transported by the nuclear import machinery across nuclear pores in an intact nuclear envelope. Recent evidence has suggested that reverse transcription is completed within the nucleus.420 After the import of the RTC into the nucleus, newly synthesized full-length, linear, double-stranded viral DNA is integrated into the 639

chromosomal DNA of the target cell. The integrated viral DNA—the provirus—remains part of the genetic makeup of the infected cell for the lifetime of that cell. The proviral DNA serves as the template for RNA polymerase II (Pol II)-directed viral RNA synthesis. The coordinated interaction of the HIVencoded Tat protein and the cellular NF-κB and Sp1 transcriptional transactivating proteins with the RNA Pol II transcriptional apparatus ensures the production of high levels of viral RNA. Unspliced or partially spliced HIV transcripts are exported from the nucleus to the cytoplasm by a unique transport mechanism mediated by the virus-encoded Rev protein that allows viral RNAs with introns to exit the nucleus. The subsequent translation of the gp160 Env precursor occurs on the endoplasmic reticulum (ER); gp160 is cleaved by a cellular protease to gp120 and gp41 during its trafficking through the secretory pathway to the plasma membrane. The Gag and Gag–Pol polyprotein precursors are synthesized on free cytoplasmic ribosomes and then move to the cell surface. The Gag and Gag–Pol polyproteins, in association with dimers of genomic RNA, condense at the plasma membrane to form an electron-dense “bud” that gives rise to a spherical, immature particle containing heterotrimeric gp120 and gp41. Proteolytic processing of the Gag and Pol proteins by the viral PR during or immediately after particle release generates the cone-shaped core characteristic of mature HIV virions. Many of the basic replicative steps described above and illustrated in Figure 17.6 are shared with the so-called “simple” retroviruses encoding only the Gag, Pol, and Env proteins. However, during its evolution, HIV acquired additional genes—the regulatory proteins Tat and Rev and the accessory proteins Vif, Vpu, Vif, Vpr, and Nef (and Vpx in the case of HIV-2)—to carry out functions that are either (a) performed by cellular proteins already present in the cells infected by the simple retroviruses or (b) uniquely required for virus replication, transmission, and survival in hematopoietic cells targeted by the primate lentiviruses. In some laboratory cell lines, the HIV-1 accessory proteins are not required for replication—hence the term “accessory”—however, these additional genes are required for replication to high levels and pathogenesis in vivo. HIV-1 encodes only 15 proteins and thus, like all other viruses, must utilize a large number of cellular proteins for successful replication. Over the years, a variety of techniques have been used to identify virus–host protein interactions and proteins required for HIV-1 replication (often referred to as HIV “dependency factors”). A number of cellular factors—in some cases referred to as restriction factors—have also been identified that disrupt virus replication. Many of these inhibitory factors are induced by interferon (IFN) and constitute part of the innate immune defense against invading viral pathogens. A primary function of the HIV accessory proteins is to counteract these cellular inhibitory factors.

FIGURE 17.6 The HIV-1 replication cycle. Productive HIV-1 infection begins with the binding of the viral Env glycoprotein complex to CD4 on the target cell, and subsequent interaction between Env and coreceptor. Conformational changes in Env trigger a membrane fusion reaction between the viral envelope and target cell plasma membrane. The core particle is then released into the cytosol where reverse transcription, mediated by the viral reverse transcriptase (RT), initiates. The core then traffics along microtubules to the nuclear envelope where it docks and is transported across a nuclear pore into the nucleus. Reverse transcription is completed and capsid uncoating occurs. The newly synthesized viral DNA, as part of the preintegration complex (PIC), integrates into cellular chromosomal DNA. Integration is catalyzed by the viral integrase (IN). The integrated viral DNA—the provirus—serves as the template for DNAdependent RNA polymerase (pol II) transcription. Unspliced, singly spliced, and multiply spliced viral RNAs are exported to the cytoplasm, where they are translated into viral proteins or, in the case of a population of unspliced viral RNA, used for packaging. The Env glycoprotein precursor is translated in the ER and the Gag and Gag–Pol polyprotein precursors are translated in the cytosol. Env, Gag, and Gag–Pol are transported to the plasma membrane, where progeny virus particles coassemble with full-length, dimeric viral RNA. The mature Env glycoprotein complex, a heterotrimer of gp120 and gp41, is incorporated into the assembling Gag lattice. The nascent particle buds from the plasma membrane in an immature state. Concomitant with release of the virus particle from the infected cell, the viral protease (PR) cleaves the Gag and Gag–Pol precursors, triggering conformational rearrangements that lead to the formation of a mature, infectious virus particle containing a characteristic conical core.

Virus Binding and Entry: The Env Glycoproteins 640

The HIV Env glycoproteins are translated from the singly spliced, 4.3-kb Vpu/Env bicistronic mRNA on ribosomes associated with the rough ER. The Env glycoprotein precursor, gp160, is an integral membrane protein that is anchored to cell membranes by the gp41 transmembrane domain (TMD) (Fig. 17.7) (for review see Ref.112). gp160 is cotranslationally glycosylated and undergoes trimerization in the lumen of the ER before it is transported to the Golgi, where, like other retroviral Env precursor glycoproteins, it is proteolytically cleaved by cellular furin or furin-like proteases at a polybasic amino acid sequence. Gp160 cleavage results in the generation of the mature gp120 and gp41 glycoprotein subunits, which remain associated via noncovalent interactions (Fig. 17.7). Cleavage of gp160 is strictly required for Env-induced fusion activity and virus infectivity.232,479

The HIV Env glycoprotein complex, in particular the gp120 component, is very heavily glycosylated; approximately half the molecular mass of gp160 is composed of N-linked oligosaccharide side chains. During transport of Env from its site of synthesis in the ER to the plasma membrane, many of the side chains are modified by the addition of complex sugars. The numerous oligosaccharide side chains form a sugar “cloud” obscuring much of the surface of gp120 from host immune recognition. As shown in Figure 17.7, gp120 contains interspersed conserved (C1 to C5) and variable (V1 to V5) domains. The Cys residues present in the gp120 proteins of different isolates are highly conserved and form disulfide bonds that link the first four variable regions in large loops.416

After its arrival at the cell surface, the gp120-gp41 trimer of heterodimers complex is rapidly internalized. A Tyr-X-X-Leu (YxxL) sequence in the membrane-proximal region of the HIV-1 gp41 cytoplasmic tail (see Fig. 17.7) is largely responsible for this rapid internalization.398,608 Analogous motifs are also present in the TM Env glycoproteins of HIV-2, SIV, and several other retroviruses. Tyr-based motifs are known to mediate endocytosis of cellular plasma membrane proteins by binding the μ2 chain of the clathrin-associated adapter protein 2 (AP-2) complex, and these interactions have been observed with gp41 cytoplasmic tails.66,526 A dileucine motif at the C-terminus of the gp41 cytoplasmic tail also participates in Env internalization and trafficking via interactions with the AP-1 complex46,92,751 (Fig. 17.7).

FIGURE 17.7 Linear representation of the HIV-1 Env glycoprotein. The vertical yellow arrow indicates the site of gp160 cleavage to gp120 and gp41; SP denotes the signal peptide. In gp120, variable (V1–V5) and conserved (C1–C5) domains are indicated. The gp41 ectodomain contains the N-terminal fusion peptide (FP), the two heptad repeats (HR1 and HR2), and the membrane-proximal external region (MPER). The transmembrane domain (TMD) is represented by a purple box. In the approximately 150 amino acid gp41 cytoplasmic tail (CT), the Tyr-Ser-Pro-Leu (YSPL) and Leu-Leu (LL) motifs implicated in Env trafficking and internalization are indicated. The putative helical motifs—or lentiviral lytic peptides (LLP-1, LLP-2, and LLP-3)—are shown. (Adapted from Checkley MA, Luttge BG, Freed EO. HIV-1 envelope glycoprotein biosynthesis, trafficking, and incorporation. J Mol Biol 2011;410(4):582 -608. Copyright © 2011 Elsevier. With permission.) CD4 Binding and Coreceptor Interactions As noted above, the first step in HIV/SIV infection is the interaction between gp120 and CD4, the major cell-surface receptor for primate lentiviruses (for review, see Ref.113). CD4 is a 55-kDa member of the immunoglobulin (Ig) superfamily; it is composed of a highly charged cytoplasmic domain, a single hydrophobic membrane-spanning domain, and four distinct extracellular domains, D1 to D4.446 CD4 normally functions to stabilize the interaction between the T-cell receptor on the surface of T lymphocytes and class II major histocompatibility complex (MHC-II) molecules on antigen-presenting cells. The high-affinity binding site for gp120 has been localized to a small segment of the CD4 N-terminal D1 extracellular domain, analogous to the second complementarity-determining region (CDR-2) loop of an Ig light chain variable domain. Important CD4 binding determinants in gp120 map to the C3 and C4 domains of gp120, although a more discontinuous, conformationdependent domain is involved in high-affinity gp120–CD4 binding.

The HIV-1 Env trimer is highly conformationally dynamic, a feature that is required for many of its functions. Distinct conformational states of the trimer have been revealed by structural studies and by other methods such as single-molecule Förster resonance energy transfer (FRET) analyses.510 The crystallization of a gp120 “core” domain—an unglycosylated gp120 derivative lacking the V1/V2 and V3 loops and the N- and C-termini—complexed with fragments of CD4 and a neutralizing antibody contributed substantially to our understanding of the gp120–CD4 interaction.397,748 The core structure revealed two major domains (referred to as the “inner” and “outer” domains) connected by a so-called “bridging sheet” (see Fig. 17.8). The latter is composed of a four-stranded, antiparallel β-sheet derived from sequences in the V1/V2 stem and portions of C4. Comparison of the CD4-bound HIV-1 gp120 structure with that of the non–CD4-bound (unliganded) gp120 from SIVmac and HIV-1 reveals that gp120, the inner domain in particular, undergoes a remarkably extensive conformational change upon CD4 binding.114,534 This shift in conformation leads to a more open gp120 structure, creating the bridging sheet, which is absent in the unliganded structure. Highresolution structures of the Env trimer have been challenging to obtain, but a recent study used cryoelectron tomography of chemically inactivated virus particles to obtain subnanometer structures of unliganded, antibody-bound or CD4-bound intact Env trimers430 (Fig. 17.9).

641

FIGURE 17.8 Ribbon diagram of the gp120 core. In this orientation, the viral membrane would be at the top, the target cell membrane at the bottom. The inner and outer domains are connected by a four-stranded β-bridging sheet. The remnants of variable loops V1/V2, V3, V4, and V5 are shown. (Adapted by permission from Nature: Kwong PD, Wyatt R, Robinson J, et al. Structure of an HIV gp120 envelope glycoprotein in complex with the CD4 receptor and a neutralizing human antibody. Nature 1998;393(6686):648–659. Copyright © 1998 Springer Nature.) Soon after the identification of CD4 as the major HIV/SIV receptor, it was recognized that this protein is not sufficient for HIV-induced membrane fusion and virus entry. In the mid-1990s, a number of studies demonstrated that members of the G protein–coupled receptor superfamily of seven-transmembrane domain proteins provided the long-sought coreceptor function.12,113,126,175 As mentioned above, the two major coreceptors for HIV/SIV infection are CXCR4 and CCR5 (for review see Ref.113) and virus isolates are classified based on their coreceptor usage: X4, R5, and X4/R5 (dual-tropic), with CCR5 being the most commonly used coreceptor and the one used by the vast majority of transmitted viruses.

Several domains within gp120 contribute to Env–coreceptor interactions. As mentioned above, gp120 undergoes extensive conformational changes upon binding to CD4 on the target cell membrane, essentially creating the coreceptor binding site,114 which would be highly susceptible to neutralizing antibodies and thus forms only transiently and close to the host cell membrane.537 The structure of the gp120-CCR5 complex, solved by cryoEM, shows two primary interaction interfaces between the two proteins: the V3 loop of gp120 inserts into the chemokine binding pocket of CCR5 and the bridging sheet of gp120 binds the N-terminus of the coreceptor.646

Certain laboratory-adapted isolates of HIV-1, HIV-2, and SIV use coreceptors in a CD4-independent manner, that is, as primary receptors.206,470,572 The coreceptor-binding surface is highly exposed in variants selected to replicate in a CD4-independent fashion.301,384 Because their coreceptor-binding surface is constitutively exposed, CD4-independent isolates tend to be hypersensitive to neutralization.

It was noted that certain individuals, despite persistent high-risk behavior, remain HIV-1 uninfected. The discovery of CCR5 and CXCR4 as HIV-1 coreceptors raised the possibility that some of these individuals might encode mutant coreceptor alleles. Indeed, a mutant CCR5 allele, referred to as CCR5/∆32, contains a 32-bp deletion that leads to the expression of a truncated protein that is not efficiently expressed at the cell surface and cannot function as an HIV-1 coreceptor.170,315,439,620 Homozygotes for CCR5/∆32 are only rarely infected with HIV-1 (and by X4 viruses), providing strong genetic evidence for the role of CCR5 in HIV-1 infection in vivo.

FIGURE 17.9 Model of the HIV-1 Env trimer. The ectodomain structure is based on fitting cryoEM densities437 into the crystal 642

structure of HIV-1 BG505 SOSIP.664 (PDB ID code 5T3Z).268 The membrane-proximal external region (MPER) is shown in blue, the transmembrane domain (TMD) in green and the cytoplasmic tail (CT) in magenta. (Adapted with permission from Piai A, Fu Q, Sharp AK, et al. NMR model of the entire membrane-interacting region of the HIV-1 fusion protein and its perturbation of membrane morphology. J Am Chem Soc 2021;143(17):6609–6615. Copyright © 2021 American Chemical Society.)

Membrane Fusion The primary function of viral Env glycoproteins is to promote a membrane fusion reaction between viral and target-cell membranes. The fusion process proceeds in a series of steps: gp120 first interacts with CD4, which induces the formation of the coreceptor binding site; a ternary CD4– coreceptor–gp120 complex then forms; and finally, conformational changes take place in gp41 that ultimately trigger membrane fusion (Fig. 17.10). Mutational analyses demonstrated that a hydrophobic region at the N-terminus of gp41, known as the “fusion peptide,” plays a central role in membrane fusion mediated by HIV-1112,233 (see Fig. 17.7). C-terminal to the gp41 fusion peptide are two amphipathic heptad repeat (HR) domains, HR1 and HR2 (see Fig. 17.7), that are also required for membrane fusion. Structures of the ectodomain of HIV-1 and SIV gp41, including HR1 and HR2, were determined by x-ray crystallography and nuclear magnetic resonance (NMR) spectroscopy.93,107,692,731 These studies showed that HR1 and HR2 pack in an antiparallel fashion to generate a six-helix bundle (Fig. 17.10). The three HR1 domains of the Env trimer form a coiled-coil structure in the center of the bundle, with the HR2 domains packing into hydrophobic grooves on the outside of the HR1 trimer. The gp41 ectodomain structure resembles that of the fusion-competent (low-pH induced) form of influenza HA2, indicating that HIV/SIV Env glycoproteins trigger membrane fusion by the same “spring-loaded” mechanism proposed for influenza virus,87 likely an ancient mechanism for fusing membranes. Binding studies using native HIV-1 Env indicate that gp41 interacts with a soluble HR2-derived peptide only after CD4 binding, indicating that HR1 and HR2 undergo conformation changes after CD4 binding and that these rearrangements are required for formation of the six-helix bundle and membrane fusion.242 Located between HR2 and the gp41 TMD is a Trp-rich region, known as the membrane-proximal external region (MPER), which plays a role in regulating Env conformation. Mutations in the MPER disrupt membrane fusion and infectivity,619 and this region is the binding site for one class of broadly neutralizing antibodies. The cytoplasmic tail of gp41 also plays a role in modulating the fusion activity of Env, in part by regulating cell-surface expression levels of the gp120–gp41 complex and by influencing the conformation of gp120 and the ectodomain of gp41115 (for review see Ref.112). The cytoplasmic tail contains several helical motifs often referred to as “lentiviral lytic peptides” (LLP1-LLP3) because of their ability to disrupt membranes (Fig. 17.7). Several NMR structures have been obtained for the cytoplasmic tail of HIV-1 gp41,514,557 including a recent structure of the entire TMD and cytoplasmic tail embedded in a membrane bicell557 (Fig. 17.9). While these structures differ in some important details, they confirm that the LLP regions interact intimately with the membrane and suggest that the cytoplasmic tail forms a trimeric baseplate around the trimeric TMD. The structure of the gp41 cytoplasmic tail in the context of a virus particle (i.e., in the presence of Gag) has not yet been solved.

A number of studies have shown that the HIV-1 membrane envelope is enriched in lipids (e.g., sphingolipids and cholesterol) that are concentrated within the cell plasma membrane in specialized microdomains known as lipid rafts.13,505 This finding is consistent with evidence that HIV-1 assembly takes place in raft-like membrane microdomains (see below in Section on Virus Assembly and Release: The Gag Proteins). It has also been proposed that membrane fusion takes place in lipid rafts in the target cell plasma membrane, and depletion of cholesterol from either the virus particle or the target cell membrane disrupts fusion.531 In addition to promoting cell-free HIV-1 infection, lipid rafts likely play an important role in HIV-1 transmission between T cells, as the VS is enriched in raft-like microdomains.348

Progress in understanding the molecular mechanism of membrane fusion has led to the development of inhibitors that block various aspects of virus fusion and entry (for review, see Ref.752). T-20 (enfuvirtide), approved for clinical use by the US Food and Drug Administration (FDA) in 2003, is a 36-amino acid, gp41 HR2-derived peptide that interacts directly with HR1 during the conformational changes that take place after CD4 binding. The interaction of T-20 with gp41 prevents six-helix bundle formation, thereby blocking fusion. Resistance to T-20 arises both in vitro and in treated individuals, primarily through mutations in HR1.598 T-20 is very costly and not orally bioavailable, reducing its current utility. A number of strategies have been employed to increase the potency of HR1/HR2 peptide-based inhibitors (reviewed in Ref.752). Given that it is straightforward to identify the HR1/HR2 repeats in the ectodomain of similar viral TM proteins, this type of therapeutic strategy could be rapidly developed for other enveloped viruses (such as SARS-CoV-2).

643

FIGURE 17.10 Schematic representation of the steps leading to membrane fusion. In the resting configuration, the Env glycoprotein complex is in its native state (1). CD4 binding (2) induces conformational changes in Env, which include opening up of the gp120 structure, which allow coreceptor binding (3). After the formation of a ternary gp120–CD4–coreceptor complex, gp41 adopts an extended conformation that allows the fusion peptide to insert into the target lipid bilayer (4). The formation of the gp41 six-helix bundle, which involves antiparallel interactions between the gp41 heptad repeats (HR1 and HR2) (5) brings the viral and cellular membranes together and membrane fusion takes place (6). Additional approaches have been explored for inhibiting the fusion process. Ibalizumab, a CD4-targeting, humanized monoclonal antibody, was approved for clinical use in 2018. It binds the D1/D2 junction of CD4 and noncompetitively inhibits gp120 binding to the receptor.205 A CCR5based inhibitor, maraviroc, was approved in 2007 (for review, see Ref.540). Maraviroc functions by binding extracellular domains of CCR5 and inducing an allosteric change in the conformation of CCR5 that prevents gp120–CCR5 interaction.212 Resistance to maraviroc can arise by acquisition of gp120 V3 loop mutations that allow HIV-1 to utilize the drug-bound coreceptor733 or by the outgrowth of CXCR4-tropic strains.494 Inhibitors that block gp120 binding to CD4 are also being developed. The first of these to be approved by the FDA is BMS-663068 (Fostemavir), approved in 2020. Other fusion and entry targets are being pursued, including small molecules that bind the MPER at docking sites for broadly neutralizing antibodies.753

Postentry Trafficking of the HIV Capsid After membrane fusion deposits the viral core in the cytoplasm, it must traffic to the nuclear pore for eventual transit across the nuclear envelope (see Fig. 17.6). During the trafficking of the core particle to the nucleus, the surface of the core—known as the capsid, composed of an assembled lattice of CA protein—interacts with a series of host cell factors that either promote or interfere with core transport and stability. An early report showed that capsids associate with microtubules and use dynein to traffic toward the nucleus.481 The dynein adapter protein bicaudal D2 (BICD2) helps promote trafficking on microtubules.179,180 HIV-1 infection has been reported to stabilize microtubules613 and proteins involved in microtubule stabilization are required for movement of capsids to the nucleus (for review, see Ref.179). The kinesin adapter fasciculation and elongation protein zeta-1 (FEZ1) directly engages incoming capsids454 by binding to the highly basic pore at the center of the capsid hexamer313 (see section below on CA). An additional protein, Sec24C, interacts with a pocket on the capsid that is the binding site for several other proteins involved in capsid nuclear import and viral DNA integration; this interaction involves Phe-Gly (FG) motifs on the host protein. Depletion of Sec24C destabilizes the capsid in the cytosol and thus interferes with the downstream events of nuclear import and DNA integration.589

Reverse Transcription A defining characteristic of retroviruses is the ability to convert their single-stranded RNA genomes into double-stranded DNA during the early stages of infection.28,697 The enzyme that catalyzes this reaction is RT, in conjunction with its associated ribonuclease H (RNase H) activity. Retroviral RTs have two enzymatic centers: (a) a DNA polymerase that can copy either RNA templates (for “minus-strand” DNA synthesis) or DNA templates (for second- or “plus-strand” DNA synthesis) and (b) RNase H (for the degradation of RNA present in DNA–RNA hybrid intermediates). RT was the first viral protein targeted by antiretroviral therapy, and RT inhibitors remain central to the treatment of people living with HIV (for reviews, see Refs.307,659).

The retroviral genome is packaged into the virion as a dimer of single-stranded RNA. As mentioned earlier, the two RNAs are held together in part by the DIS near their 5′ ends. Although each retroviral particle contains two copies of genomic RNA, only one provirus is formed per virion.309 Retroviruses are, therefore, referred to as “pseudodiploid.” As with most other DNA polymerases, RT is dependent on the 3′-OH group of an RNA or DNA primer to initiate polymerization. Retroviruses use specific tRNAs (tRNALys3 in the case of HIV-1) to initiate DNA synthesis. The mechanism of tRNA selection and placement on the template is complex, involving interactions with RT and NC as well as with the 18-nt PBS near the 5′ end of the viral genome.

Reverse transcription of the retroviral RNA genome to a double-stranded DNA copy is initiated after viral entry into the target cell and proceeds 644

via a series of steps that are outlined briefly below (see e-Fig. 17.1). This is also described in Figure 8 of Chapter 15. (a) Minus-strand DNA synthesis is initiated from the 3′-OH of the tRNA bound to the PBS. DNA synthesis then proceeds a short distance to the 5′ end of the genome. (b) RNase H digests the RNA portion of the newly formed RNA–DNA hybrid, freeing the resulting short, single-stranded DNA fragment (known as the minus-strand strong-stop DNA). (c) The minus-strand strong-stop DNA is transferred to the 3′ end of the genome, where it hybridizes by virtue of a short region of homology (the “repeated” or R region) present at both 5′ and 3′ ends of the RNA genome (the first strand transfer). (d) Minus-strand synthesis, accompanied by RNase H-mediated degradation of the RNA in the resulting RNA–DNA hybrid, continues along the length of the genome to the PBS that forms the 5′ end of the genome after the degradation of the early RNA/DNA hybrid of strong-stop DNA. (e) Fragments of RNA that were not removed by RNase H serve as primers for plus-strand synthesis. The major site of plus-strand priming is the polypurine tract (PPT) near the 3′ end of the genome (e-Fig. 17.1); however, residual RNA fragments that remain hybridized to regions outside the PPT can also be used for priming plus-strand synthesis. In the case of HIV-1, one such region, known as the central PPT, appears to be particularly important in this regard. (f) After plus-strand synthesis copies the initial minus-strand DNA product including a portion of the tRNA primer, RNase H removes that portion of tRNA in the RNA/DNA hybrid. This exposes the complement of the PBS (designated in lower case as pbs) at the 3′ end of the initial plus-strand DNA, allowing this short plus-strand DNA to anneal at its 3′ end with the complementary region at the 3′ end of the near full-length minus-strand DNA that paused after copying the PBS (“second-strand transfer”). (g) Plus- and minusstrand syntheses proceed to completion. Plus-strand synthesis terminates at the end of the minus strand and, for HIV-1, at a sequence known as the central termination signal (CTS). The position of the central PPT upstream of the CTS results in the displacement of approximately 100 nucleotides of plus-strand DNA and the formation of a triplex DNA structure. The final product of reverse transcription is a linear, doublestranded DNA molecule capable of serving as the substrate for integration.

e-FIGURE 17.1 Long terminal repeat (LTR) elements mediate steps in the reverse transcription and integration reactions. In addition to providing an essential function in the virus replication cycle, the enzymatic activity of RT is routinely used in the laboratory to quantitatively monitor levels of progeny virions present in the supernatant of infected cultures and to elucidate the molecular details of virus replication. Also, the detection of viral DNA postinfection by real-time PCR provides one of the most reliable methods for monitoring entry and postentry steps in the virus replication cycle. Once inside the nucleus, some double-stranded viral DNA is ligated by nuclear enzymes to generate circular forms; although these circular DNA products do not integrate and are eventually lost, their formation is often used to monitor nuclear import. The mature HIV-1 RT holoenzyme is a heterodimer of 66- and 51-kDa subunits. The 51-kDa subunit (p51) is derived from the 66-kDa (p66) subunit or some larger precursor by proteolytic removal of the C-terminal 15-kDa fragment of p66 by PR, cleaving at an internal site within a presumably partially unfolded RNase H domain (Figs. 17.4 and 17.11). Hundreds of structures of HIV-1 RT have been determined by x-ray crystallography in a number of studies.307 Early studies crystallized RT in several contexts: (a) unliganded,601 (b) bound to a short DNA duplex and the Fab portion of an anti-RT antibody,330 (c) covalently linked to a complex of primer/template and dNTP,310 and (d) bound to an RNA– DNA duplex.621 A number of structures are also available for RT bound to nonnucleoside RT inhibitors (NNRTIs). The crystal structure of HIV-1 RT reveals that the p66 and p51 subunits are folded into similar subdomains, but these subdomains are arranged quite differently in the two subunits of the p66/p51 heterodimer. The p66 subunit can be visualized as a right hand, with the polymerase active site within the palm, and a deep, template-binding cleft formed by the palm, fingers, and thumb subdomains331 (Figs. 17.11 and 17.12). The polymerase domain is linked to RNase H by the connection subdomain. The active site, located in the palm, contains three critical Asp residues (Asp-110, Asp-185, and Asp-186) that are spacially in close proximity, and two Mg2+ ions that are coordinated by the negatively charged Asp side chains. Mutation of these Asp residues abolishes the polymerase activity of RT. The p51 subunit plays a structural role and does not form a polymerizing cleft; Asp-110, Asp-185, and Asp-186 of p51 are buried within the subunit. Approximately 18 bp of the primer/template duplex lie in the nucleic acid–binding cleft, stretching from the polymerase to the RNase H active sites. In the RT-primer/template-dNTP structure,310 the presence of a dideoxynucleotide (ddNTP) at the 3′ end of the primer allows visualization of the catalytic complex blocked just prior to attack on the incoming dNTP. Comparison with previously obtained structures suggested a model whereby the fingers close in to trap the template and dNTP prior to nucleophilic attack of the 3′-OH of the primer on the incoming dNTP. After the addition of the incoming dNTP to the growing chain, the fingers adopt a more open configuration, thereby releasing the pyrophosphate and enabling RT to bind the next dNTP. HIV-1 RNase H, the structure of which has been determined by x-ray crystallography, displays a global folding similar to that of Escherichia coli RNase H163 and other nucleotide metabolizing enzymes, suggesting the ancestral enzyme appeared early in the evolution of life. The structure of human RNase H1 bound to an RNA/DNA hybrid has allowed the modeling of HIV-1 RNase H complexed with its substrate.525 A structure of HIV-1 RNase H engaged in RNA cleavage has also been solved.699

645

FIGURE 17.11 Ribbon diagrams of the p66 (left) and p51 (right) RT subunits. Polymerase active site and fingers, palm, thumb, and connection subdomains are shown. (Modified by Kalyas Das and Eddy Arnold from Jacobo-Molina A, Ding J, Nanni RG, et al. Crystal structure of human immunodeficiency virus type 1 reverse transcriptase complexed with double-stranded DNA at 3.0 Å resolution shows bent DNA. Proc Natl Acad Sci USA 1993;90(13):6320–6324. Copyright © 1993 National Academy of Sciences, USA.) As mentioned above, reverse transcription initiates from a tRNALys3 primer bound to the PBS. Initiation is known to proceed slowly relative to subsequent elongation and is characterized by frequent pausing events. Recent studies on the reverse transcription initiation complex indicate that the structure of the enzyme is regulated by the viral RNA and the primer, providing insights into the slow rate of initiation. The results of these studies also suggest that the initiation complex is particularly vulnerable to inhibition by NNRTIs.407

FIGURE 17.12 HIV-1 RT bound to an RNA/DNA template/primer. (Modified by Karen Kirby and Stefan Sarafianos from Singh K, Marchand B, Kirby KA, et al. Structural aspects of drug resistance and inhibition of HIV-1 reverse transcriptase. Viruses 2010;2(2):606–638. https://creativecommons.org/licenses/by/3.0/.) The high rate of variation among HIV-1 populations poses one of the fundamental challenges to effectively controlling this pathogen. Highly variable virus populations are largely a consequence of rapid virus replication rates, coupled with the error-prone nature of RT (which lacks a proof-reading function), and frequent template switching during reverse transcription138 (for review, see Ref.307). Cell-free error rates for purified RTs have been determined,35,574,599 as have in vivo retrovirus mutation rates.1,192,464,543,544 Errors during retrovirus replication are also likely to be introduced by the cellular DNA-dependent RNA pol II during transcription of the viral RNA. The total HIV-1 in vivo mutation rate (a composite of substitutions, frameshifts, simple deletions, and deletions with insertions) was measured at approximately 1 to 3 × 10−5 per cycle of replication.1,464

As discussed earlier, retroviral particles contain two copies of single-stranded RNA, which are usually identical. During reverse transcription, RT frequently switches from one RNA template to the other in the dimer. RT also undergoes intramolecular jumps, that is, within the same RNA template.542 Intramolecular jumps lead to mutations (e.g., deletions, insertions, and duplications), whereas intermolecular jumps generate recombinants if the two packaged RNAs are not genetically identical.308 The intermolecular template switches that lead to recombination can occur when RT encounters a break in the RNA during polymerization—the so-called “forced copy choice” model.137 Jumps during minus-strand synthesis can also occur in the absence of strand breaks,355 and the stability of RT association with template is influenced by the balance between the enzyme’s RNase H and polymerase activities—the “dynamic copy-choice” model.319 Intermolecular strand transfers that lead to recombination are required for successful completion of reverse transcription, as blocking these events leads to large deletions in the viral genome and severe reductions in particle infectivity.585 The impact of high rates of recombination and high replication rates on HIV-1 biology is substantial. As noted earlier, recombinants constitute the predominant strains currently circulating in certain parts of the world. Recombination also provides a mechanism for the rapid generation of multidrug-resistant HIV-1 variants by combining multiple drug-resistance mutations in one genome.

RT has long been a target for antiviral compounds, with AZT being the first antiretroviral approved for treatment of people living with HIV (in 1987). RT inhibitors are routinely prescribed, typically in combination with PR or IN inhibitors. There are two major classes of drugs that block reverse transcription: nucleoside RT inhibitors (NRTIs) and nonnucleoside RT inhibitors (NNRTIs). NRTIs are nucleotide mimics that lack the 3′-OH and thus act as chain terminators upon incorporation into DNA, whereas the NNRTIs inhibit DNA polymerization by binding a small hydrophobic pocket near the RT active site and inducing a change in the structure of RT that blocks DNA synthesis.383 Resistance to RT inhibitors can develop in treated individuals; escape from NRTIs generally takes place by one of two mechanisms: (a) resistant RTs acquire the ability to selectively incorporate natural dNTPs but (at least partially) exclude the NRTI or (b) mutant RTs incorporate the NRTI but subsequently excise it from the terminated primer with the evolution of a rudimentary 3′-exonuclease activity. Resistance to NNRTIs occurs when mutations in RT interfere with the binding of the drug to the enzyme by disrupting key drug–enzyme interactions, changing the shape of the NNRTI binding pocket, or preventing entry of the drug into the binding pocket.622 Finally, a significant effort has been directed toward developing inhibitors that target the RNaseH activity of RT, as this remains the only HIV enzymatic activity against which no drug is available.

646

Nuclear Import Most retroviral genomes gain access to host cell chromosomal DNA during progression of the target cell through the cell cycle when the nuclear membrane dissolves. These retroviruses are, therefore, unable to efficiently infect noncycling cells. In contrast, as noted earlier, lentiviruses are able to productively infect nondividing cells (e.g., macrophages) and their genomes are competent to access the interphase nucleus of dividing cells. Although early studies proposed that the HIV-1 capsid uncoats (i.e., the CA subunits dissociate from the capsid) rapidly after membrane fusion, subsequent work demonstrated that at least some (and maybe most) CA subunits remain associated with the core particle, or RTC, as it traffics to the nucleus.209 Analysis of chimeras between MLV (which is unable to infect nondividing cells) and HIV-1 revealed that the ability to infect nondividing cells maps to CA.759 A C-terminally truncated, cytosolic form of cleavage and polyadenylation factor 6 (CPSF6, which is normally localized in the nucleus), referred to as CPSF6-358, was shown to block HIV-1 nuclear import but not to affect MLV infectivity.413 Virus selection experiments in the presence of CPSF6-358 gave rise to a resistant mutant that contained a single amino acid change in CA (N74D). This CA-N74D mutant was not only resistant to inhibition by CPSF6-358 but also displayed an altered requirement for karyopherins and nuclear pore factors. These data supported a role for CA in nuclear import of the RTC and demonstrated that CA regulates HIV’s utilization of nuclear transport and pore components.

Additional evidence accumulated in support of the model that not only is CA an integral part of the RTC but also that the capsid lattice must remain at least partially intact during trafficking to the nuclear envelope: (a) restriction factors such as tripartite motif-containing 5α (TRIM5α) and myxovirus resistance protein 2 (Mx2, also known as MxB), which block postentry and/or nuclear import events in the virus replication cycle (see below), recognize the assembled hexameric CA lattice on the capsid235,245; (b) nuclear pore components required for nuclear import (e.g., Nup153 and Nup358) bind to the capsid with much higher affinity than they bind nonassembled CA54; and (c) an intact capsid is likely required to shield the viral nucleic acid from detection by host innate immune sensors in the cytosol.194 Recent studies have indeed provided compelling evidence that the HIV-1 capsid remains intact, or largely intact, until after import into the nucleus. Live-cell imaging of CA-labeled capsids revealed the import of intact capsids into the nucleus, with full uncoating taking place near the site of viral DNA integration.89,420 This analysis used a system in which GFP, packaged into virions as an internal Gag–GFP fusion protein, becomes trapped within the capsid during Gag proteolysis and maturation and thus serves as a content marker for measuring the intactness of the capsid. GFP loss from the capsid took place in the nucleus almost simultaneously with loss of CA near the site of DNA integration.420 Studies that examined the kinetics with which a nuclear pore-blocking agent inhibited HIV-1 infection178 and sensitivity to the capsid-targeting small-molecule inhibitor PF-3450074 (PF74)89,178 concluded that reverse transcription is likely completed in the nucleus and that intact CA hexamers survive nuclear import. One of the key arguments originally supporting the hypothesis that capsid uncoating must occur before nuclear entry was that the intact capsid is too large to traffic through the nuclear pore. However, a study that used a combination of light microscopy and cryoEM observed that whereas capsids in the nucleus appeared to be disrupted, apparently intact conical capsids could be visualized inside the central channel of the nuclear pore complex (Fig. 17.13). These observations indicated that the nuclear pore is large enough to accommodate an intact HIV-1 capsid.789 The findings that reverse transcription is likely completed in the nucleus,420 and that endogenous reverse transcription in vitro requires an intact capsid,130 also support the hypothesis that uncoating takes place predominantly in the nucleus. Despite the evidence cited above, the topic of where uncoating occurs remains controversial, as some data support the hypothesis that uncoating occurs in the cytosol461 or at the nuclear envelope.226,227

FIGURE 17.13 Cryoelectron tomographic reconstruction of an HIV-1 capsid entering a nuclear pore. Nuclear envelope (NE) is highlighted in yellow, HIV-1 capsid in purple, nuclear pore complex (NPC) in light blue, microtubules (MTs) in red. (Image provided by Vojtech Zila based on Zila V, Margiotta E, Turonova B, et al. Cone-shaped HIV-1 capsids are transported through intact nuclear pores. Cell 2021;184(4):1032–1046 e18.)

Postentry Blocks to Lentiviral Infection A number of host cell restriction factors have been described that interfere with postentry steps in the HIV replication cycle. Several of these factors have likely imposed barriers to lentiviral transmission across primate species, and viruses that have crossed species have had to adapt to these factors in the new host. The best characterized of the factors that affect viral replication prior to integration include the cytidine deaminase apolipoprotein B mRNA-editing enzyme, catalytic polypeptide-like 3G (APOBEC3G) protein, and family members; TRIM5α; Mx2; and SAMHD1. APOBEC3G is counteracted by Vif, and SAMHD1 in antagonized by the HIV-2 Vpx protein; these restriction factors will, therefore, be discussed later in the context of the HIV-1 and HIV-2 accessory proteins. TRIM5α and TRIMCyp Decades ago, it was discovered that cells from mice of specific genetic backgrounds express dominant factors that block infection by certain subtypes of MLV. For example, the Friend virus susceptibility-1 (Fv1) allele431,558 encodes resistance to distinct strains of MLV; Fv1n confers resistance to B-tropic MLV, whereas Fv1b cells cannot be efficiently infected by N-tropic MLV (N-MLV). The viral determinant of N- and B-tropism maps to a specific amino acid (residue 110) in the CA domain of Gag.385 The Fv1 block occurs early postentry, after reverse transcription but before integration.345 It was demonstrated in the mid-1990s that Fv1 encodes an endogenous Gag-like protein.51

Nonmurine cells do not harbor an Fv1 gene.51 However, seemingly analogous postentry restrictions have also been observed in a variety of 647

mammalian cells. N-tropic MLV poorly infects cells from a number of mammalian species.703 Similarly, HIV-1 infection is inefficient in cells derived from Old World (e.g., African green and rhesus) monkeys, whereas New World (e.g., owl and squirrel) monkey cells are poorly infected by SIVmac.298,302,653 The host factor responsible for postentry restriction of HIV-1 in rhesus macaque cells was demonstrated to be the IFN-

inducible protein TRIM5α.682 Expression of rhesus TRIM5α in human cells potently inhibits HIV-1 infection but has no effect on Moloney MLV infectivity. Conversely, knock-down of TRIM5α expression using small interfering RNA (siRNA) in rhesus cells markedly increases HIV-1 infectivity.682 Subsequent studies indicated that, when expressed in human cells, TRIM5α derived from several New World monkey species restricts infection by SIV from African green monkeys (SIVagm) and SIV adapted to replicate in macaques (SIVmac).669 Rhesus, African green monkey, and human TRIM5α can also diminish infection by the nonprimate lentivirus equine infectious anemia virus (EIAV).286

As their name implies, TRIM proteins contain three major domains: a RING domain that possesses ubiquitin ligase activity, a B-box 2 domain, and a coiled-coil domain (Fig. 17.14). The α isoform of TRIM5 (TRIM5α) also contains a C-terminal B30.2 or SPRY domain that is absent in other TRIM5 isoforms. This C-terminal B30.2/SPRY domain harbors the determinants responsible for the species specificity of TRIM5αmediated restriction to retroviral infection517,550,668,684,766; a single amino acid change in the B30.2/SPRY domain of human TRIM5α converts it from a weak inhibitor of HIV-1 infection to one that behaves like rhesus TRIM5α in potently restricting HIV-1 infection.766 This specificity is mediated through direct binding between the B30.2/SPRY domain and the CA protein on the incoming capsid.405,427,642 Binding of TRIM5α to CA requires both TRIM5α dimerization and higher-order multimerization.405,427 The higher-order multimers have been reported to be hexameric,245,521 an observation that is relevant to the mechanism of TRIM5 restriction (see below). The importance of TRIM5α in combating retroviral infections is supported by the finding that the CA-binding B30.2/SPRY domain in TRIM5α has undergone positive selection during primate evolution.344,627,668

In owl monkey cells, HIV-1 infection is highly inefficient but is significantly enhanced by disrupting the interaction between CA and the cellular protein cyclophilin A (CypA) (see section on capsid). This observation suggested a link between CA–CypA binding and postentry restriction. The basis for this link was revealed by the discovery that owl monkey cells express a form of TRIM5α in which the C-terminal B30.2/SPRY domain has been replaced by CypA524,628 (Fig. 17.14). This unusual TRIM5 variant, designated TRIMCyp, restricts HIV-1 infection through a direct binding between the CypA portion of TRIMCyp and CA of the incoming particle. Thus, the ability of TRIMCyp to restrict HIV-1 infectivity in owl monkey cells is eliminated by mutations in the CypA-binding domain of CA or by treating the infected cells with drugs like cyclosporine A that block the CA–CypA interaction.524,628 Given the short period of time HIV-1 has circulated in the human population, TRIM evolution in primates was not driven by selective pressure by HIV-1 but rather by SIVs. Nevertheless, TRIM-mediated restriction of HIV-1 provides useful information in understanding the antiviral activity of this family of restriction factors. In addition, human TRIM5α has been reported to potently block HIV-1 infection in primary T cells when the CA–CypA interaction is disrupted, suggesting that CypA binding to the incoming capsid protects it from TRIM5α restriction in human cells.375

FIGURE 17.14 Domain organization of TRIM5α and TRIMCyp. The major domains—RING, B-box 2, coiled-coil, and B30.2 (SPRY), and cyclophilin A (CypA)—are shown. Gag-binding determinants of TRIM5α and TRIMCyp reside in the B30.2 (SPRY) and CypA domains, respectively. TRIM family proteins, when overexpressed in cells, assemble into cytosolic complexes referred to as “cytoplasmic bodies”.594 Although preexisting TRIM5α or TRIMCyp cytoplasmic bodies are not required for restriction of retroviral infection,550,667 microscopic examination of cells overexpressing rhesus TRIM5α early after HIV-1 infection revealed a colocalization between TRIM5α-induced cytoplasmic bodies and viral capsids.99 These cytoplasmic bodies display features that suggest that they are associated with macroautophagy.101

Several lines of evidence support a model whereby binding of TRIM5α or TRIMCyp to CA on the incoming capsid contributes to a destabilization of the capsid and, consequently, a block to reverse transcription: (a) TRIM5α appears to accelerate CA uncoating early postinfection in the context of either HIV-1 or N-MLV.683 (b) Incubation of in vitro-assembled capsid-like complexes with cell lysates containing TRIM5α or TRIMCyp disrupts the assembled structures.61 (c) Treatment of infected cells with proteasome inhibitors reverses the ability of TRIM5α to prevent viral DNA synthesis, suggesting that TRIM5α induces the degradation of capsid in the proteasome.745 It is important to note, however, that the mechanism of TRIM restriction in cells is still not fully understood, and although proteasome inhibitors rescue the TRIM-induced defect in DNA synthesis they do not rescue virus infectivity, suggesting that restriction is a multistep process.745

Structural studies have shed light on the mechanism of action of TRIM5α and TRIMCyp restriction.246 In vitro, TRIM5α forms dimers, which assemble into a hexagonal lattice,245 a feature of both TRIM5α and TRIMCyp.428 Although TRIM lattice assembly occurs spontaneously, it is greatly facilitated by the presence of hexagonal arrays of in vitro-assembled CA. These findings suggest that restricting TRIM proteins assemble on top of incoming CA hexamers, thereby contributing to capsid disassembly and degradation (Fig. 17.15). Consistent with this model, a fragment of rhesus TRIM5α comprising the coiled-coil and B30.2/SPRY domain was shown to bind HIV-1 CA assemblies or virus-derived capsids in vitro and induce their disassembly from an extended CA hexameric lattice to individual hexamers.780 Although binding of TRIM5α or TRIMCyp to the viral capsid induces capsid destabilization in vitro, the E3 ubiquitin ligase activity of the TRIM protein, which mediates TRIM autoubiquitylation, appears to contribute to restriction, presumably through recruitment of the proteasome. The immunoproteasome, a type of proteasome that is up-regulated by IFN, may promote capsid degradation.341 TRIM5α binding to the CA lattice not only contributes to CA lattice disassembly and direct antiviral restriction but also triggers the innate immune response against retroviral infection. CA lattice recognition by TRIM5α reportedly leads to the generation of free polyubiquitin chains, which could in turn activate innate immune signaling.551 In addition, the hexameric nature of the TRIM5α assemblies on the viral capsid leads to the formation of K63-linked polyubiquitin chains at the N-terminus of TRIM5α through RING domain–mediated autoubiquitylation; trivalent RING domain interactions at the vertices of the TRIM5α hexamer (Fig. 17.15) have been shown to be required for extension of K63-linked polyubiquitin chains. These chains are proposed to promote 648

capsid disassembly and trigger immune signaling.221

Mx2 The Mx proteins are highly conserved across vertebrates and are IFN-inducible, dynamin-like GTPases. Humans encode Mx1 and Mx2 (also referred to as MxA and MxB, respectively). Mx1 interferes with the replication of a large number of viruses, but does not inhibit HIV. In 2013, it was reported independently by several groups that Mx2 exhibits anti-HIV activity.265,359,440 Interestingly, the GTPase activity of Mx2 is not required for its antiviral activity. Mx2 is a dimeric protein that bears an N-terminal nuclear localization signal, and Mx1/Mx2 chimeras that retain this region of Mx2 exhibit anti–HIV-1 activity.265 This observation, coupled with the finding that Mx2 restriction is elicited not at the level of reverse transcription but at a nuclear import step,265,359,440 suggests that Mx2 disrupts some aspect of capsid import into the nucleus. Indeed, Mx2 has been shown to interact with several nuclear pore components, potentially positioning the protein to disrupt the nuclear import of incoming HIV-1 capsids.182

FIGURE 17.15 Capsid recognition by TRIM5α. A: Two SPRY domains bind the CA-NTD on the assembled capsid. B: TRIM5α (in purple) assembles on top of the capsid. CA hexamers are shown in dark green, pentamers in light green/yellow. The location of a trivalent RING domain interaction is indicated. (Adapted by permission from Nature: Ganser-Pornillos BK, Pornillos O. Restriction of HIV-1 and other retroviruses by TRIM5. Nat Rev Microbiol 2019;17(9):546–556. Copyright © 2019 Springer Nature.) Primate Mx2 proteins exhibit variable abilities to inhibit lentiviral infection; this species specificity maps to a single residue near the N-terminus of the protein that is under positive selection.90 While the precise mechanism of Mx2 restriction is still uncertain, a number of studies have shown that mutations in CA can alleviate the antiviral activity of this host factor. Dimerization but not higher-order oligomerization of Mx2 is required for its antiviral activity and direct Mx2 binding to assembled CA has been demonstrated.53,235

Integration Mutations that block retroviral DNA integration were first reported in avian and murine retroviral systems,191,539,580,641 leading to the discovery of the essential viral enzyme IN, generated by PR-mediated cleavage of the C-terminal portion of the Gag–Pol polyprotein (see Fig. 17.6). The steps in the integration process were originally elucidated in studies using MLV,81,241 but these findings apply to HIV as well. In all retroviral systems, integration proceeds in the same series of steps (Fig. 17.16): (a) 3′ processing: After its assembly with the viral DNA, IN cleaves immediately downstream from the invariant nucleotide sequence CA, typically removing two nucleotides from the 3′ termini of both strands of full-length, linear viral DNA, generating a preintegration substrate with 3′-recessed ends. (b) DNA strand transfer: In the nucleus, IN catalyzes a staggered cleavage in the cellular target DNA. The 3′-recessed ends of viral DNA are joined to the 5′ “overhanging” termini of the cleaved cellular DNA. (c) Gap repair: The cellular repair machinery fills the gap, thereby completing the integration process.81,241,606 All integrated proviruses terminate with dinucleotides 5′-TG and CA-3′, whereas different genera yield different sized flanking duplications based on the spacing of the staggered chromosomal DNA cut; the lentiviral duplication of cellular DNA is 5 bp.

Our understanding of the chemistry of the retroviral integration reaction was greatly assisted by the development of in vitro integration assays. Purified IN can carry out 3′ processing and strand transfer reactions when combined with short synthetic oligonucleotides that mimic the viral DNA ends and a divalent metal ion (Mg2+ or Mn2+).152,365 Initial studies observed that the predominant product in in vitro assays using purified HIV-1 IN was a single end joined to one strand of the target, rather than the more physiologically relevant product in which both ends are integrated into the target (“concerted” or “full-site” integration).

Retroviral IN proteins are composed of three structurally and functionally distinct domains: an N-terminal, zinc-binding domain (NTD); a catalytic core domain (CCD); and a C-terminal domain (CTD) (Fig. 17.17). Problems with low solubility and propensity to aggregate initially hindered progress in defining the structure of the IN holoenzyme in its active state. However, the structures of all three domains independently and in various combinations were solved by x-ray crystallography or NMR methods. The NMR structure of the NTD95 revealed three helical bundles with a zinc coordinated by invariant amino acids His-12, His-16, Cys-40, and Cys-43. Attempts at crystallography of the CCD were successful when a mutation was identified that greatly increased solubility without disrupting in vitro catalytic activity.198 Each monomer of the core domain is composed of a five-stranded β-sheet flanked by helices; this structure bears striking resemblance to that of other polynucleotidyl transferases including RNase H and the bacteriophage MuA transposase.198,596 Three highly conserved residues are found in analogous positions at the catalytic center of these enzymes; in HIV-1 IN, these residues are Asp-64, Asp-116, and Glu-152—often referred to as the D,D-35-E motif (Fig. 17.17). Mutations in this catalytic triad block HIV-1 IN function both in vivo and in vitro. The IN CTD, whose structure was initially solved by NMR,201,442 adopts a five-stranded β-barrel folding topology reminiscent of Src homology 3 (SH3) domains. The crystal 649

structure of an HIV-1 NTD-CCD IN fragment726 revealed a dimer of dimers, with two inner monomers and two outer monomers, with an extensive intermolecular NTD–CTD interface that provided a preview of the interface evident in the structure of the full retroviral tetramer.

FIGURE 17.16 Schematic depiction of the HIV-1 integration process. During 3′ processing, IN cleaves immediately downstream from the CA dinucleotide, removing two nucleotides from the 3′ termini of both strands of full-length, linear viral DNA, generating a preintegration substrate with 3′-recessed ends. IN then catalyzes a staggered cleavage in the cellular target DNA. The 3′-recessed ends of viral DNA are joined to the 5′ “overhanging” termini of the cleaved cellular DNA (strand transfer). The cellular repair machinery fills the gap, thereby completing the integration process (gap repair). A breakthrough in IN structural biology was made possible by the realization that the IN of prototype foamy virus (PFV) is highly soluble and capable of concerted integration in vitro with short DNA substrates.710 Although divergent at the overall amino acid sequence level, HIV-1 and PFV IN are structurally conserved. The IN tetramer bound to DNA oligonucleotides that mimic the viral DNA ends—which together form the active nucleoprotein complex referred to as the “intasome”—was crystallized and the structure solved.277 This structure revealed that two monomers in the tetramer interact at a dimer interface and are responsible for all contacts with the DNA substrate; the other two monomers are on the outside of the complex and do not interact with each other or with DNA. The residues of the catalytic triad (D,D-35-E) of the inner monomers are oriented close to the 3′-OH of the viral DNA ends. Two metal ions are coordinated at the active site by the residues of the catalytic triad. The addition of model target DNA to the intasome allowed for crystallization of a complex containing both viral and target DNAs.450 In the presence of Mg2+, the strand transfer reaction took place during crystallization, whereas in the absence of divalent metal ions, strand transfer was blocked. This allowed capture of postcatalytic strand transfer and prestrand transfer complexes. These structures revealed that the target DNA, which binds in a cleft between the two halves of the intasome complex, adopts a highly bent conformation allowing the wellseparated intasome active sites to access the scissile phosphodiester bonds in the target DNA. The viral (donor) DNA duplex is unpaired at the active site. Conformational changes that occur during catalysis are predicted to render the strand transfer reaction irreversible.450 Following the original publication of the structures described above,277,450 a number of additional PFV structures that contained potent inhibitors were solved (for review, see Ref.426).

FIGURE 17.17 Schematic representation of an IN monomer. The three major structural domains—the N-terminal domain (NTD), catalytic core domain (CCD), and C-terminal domain (CTD)—are shown. The catalytic triad Asp-64, Asp-116, and Glu-152 is represented as D-D-E. Despite the utility of the PFV intasome structure, as mentioned, the sequences of the HIV-1 and PFV enzymes are highly divergent, sharing only approximately 20% amino acid sequence identity in the catalytic domain. Many of the mutations in HIV-1 IN that confer resistance to drugs targeting IN (see below) map to residues that are not conserved between the HIV-1 and PFV enzymes. These considerations led to efforts to solve the structure of the HIV-1 intasome by cryoEM methods.541 These studies made use of the finding that fusion of the DNA-binding protein Sso7D to the N-terminus of HIV-1 IN increased enzyme activity and solubility, overcoming the propensity of HIV-1 IN to aggregate. Using this fusion protein, a cryoEM structure was solved for the HIV-1 IN tetramer in complex with joined viral and target DNA ends, referred to as the strand transfer complex (STC) (Fig. 17.18). The core tetrameric unit was observed to assemble into higher-order structures containing 12 or 16 IN protomers.541 The biological relevance of these higher-order complexes remains to be fully explored, but it appears that the active complex contains multiple IN tetramers.

650

FIGURE 17.18 Structure of the HIV-1 strand transfer complex (STC). A: Cryo-EM reconstruction of the STC, with IN protomers indicated in red, green, yellow, and blue. Viral and target DNA are highlighted in dark and light gray, respectively. B: Atomic model derived from the cryo-EM density, colored as in (A). C: Segmented cryo-EM density and (D) asymmetric subunit of the atomic model, with IN N-terminal domain (NTD) in green, IN catalytic core domain (CCD) in beige, NTD-CCD linker in blue, and C-terminal domain (CTD) in purple. A number of host factors have been reported to bind directly to HIV-1 IN, including a 75-kDa, predominantly nuclear, chromatin-associated protein known as lens epithelium–derived growth factor (LEDGF/p75).123,204 This protein was found to stimulate the strand transfer step of the integration reaction in vitro123 and increase binding of IN to DNA.91 Near-complete knock-down of LEDGF/p75 levels,441 or use of a murine LEDGF/p75 knock-out cell line,655 revealed markedly reduced HIV-1 infectivity in the absence of detectable LEDGF/p75 expression. Infectivity was also severely compromised by overexpression of the IN-binding domain (IBD) of LEDGF/p75.168,441 In the case of both LEDGF/p75 depletion and LEDGF/p75-IBD overexpression, infectivity was disrupted at the integration step, and the effect was specific to lentiviruses, establishing LEDGF/p75 as a cellular cofactor for lentiviral integration. Studies in which siRNA-resistant LEDGF/p75 mutants were added back to LEDGF/p75-depleted cells identified domains of LEDGF/p75 required for its activity as an integration cofactor.441,655,717 The observation that the LEDGF/p75-IBD and the chromatin-binding regions (the PWWP domain and AT-hook) (Fig. 17.19) are required for function supported the hypothesis that LEDGF/p75 functions to tether the PIC to target chromosomal DNA. Structural studies have provided further details of the IN–LEDGF/p75 interaction.4,122,200

Although the basic mechanism by which retroviruses integrate their DNAs into the host cell genome is highly conserved, different retroviruses differ in their target site selection.488 For example, HIV-1 preferentially integrates within the bodies of actively transcribed genes in gene-rich regions; MLV favors transcription start sites but only weakly selects active genes. ASLV displays a strong preference for neither active genes nor transcription start sites. Target site selection, which impacts the development of retroviruses as vehicles for gene therapy, is influenced by interactions between components of the incoming viral complex—specifically CA and IN—and chromatin-associated host factors. CA–CPSF6 and IN–LEDGF/p75 interactions strongly influence, by different mechanisms, HIV-1 integration into genes. The interaction between the incoming capsid and CPSF6 directs the viral complex to nuclear speckles in speckle-associated domains and the incoming capsid causes CPSF6 to cluster in these speckles.644 Depletion of CPSF6, or mutation of the CPSF6 binding site in CA, in most cases does not substantially reduce the overall efficiency of virus infection but causes integration targeting to shift from gene-rich regions of the chromatin to gene-sparse regions.2,225 Following CPSF6-mediated delivery of the PIC to gene-dense regions, IN–LEDGF/p75 binding tethers the viral DNA to chromatin (Fig. 17.19).

FIGURE 17.19 Model for the tethering of the preintegration complex to chromosomal DNA by LEDGF/p75. The preintegration complex (PIC), containing multimeric IN (red) and viral DNA (blue), is bound by the IN binding domain (IBD) of LEDGF. The AT hook (AT) domain of LEDGF is shown in complex with DNA; the PWWP domain is depicted bound to DNA and histone proteins in the nucleosome. The function of LEDGF/p75 in promoting HIV-1 integration raises the possibility that the IN–LEDGF/p75 interaction could serve as a target for developing novel antiviral therapeutics. Small molecule “LEDGINs” (LEDGF/p75-IN inhibitors) were identified that block the IN–LEDGF/p75

651

interaction in vitro and inhibit HIV-1 infection at the integration step.129 Although these molecules were originally identified based on their ability to block the IN–LEDGF/p75 interaction, it has become clear that their antiviral mechanism of action primarily results from their ability to induce IN aggregation and thus disrupt the role of IN in RNA condensation into the capsid during maturation (for review, see Ref.379) (also see section on Maturation). These compounds are now most often referred to as allosteric IN inhibitors (ALLINIs).

The IN protein is subject to extensive posttranscriptional modifications in the form of acetylation, SUMOylation, and phosphorylation. While the roles of these modifications in IN function are still under investigation, recent work has shown that mutation of Lys residues that are modified by acetylation, in particular IN residue Lys-258, has no effect on integration but markedly reduces viral gene expression from the integrated provirus.735 Reduced viral RNA transcription is a result of misdirected integration into centromeric repeats. Polymorphisms in residue Lys-258 could thus lead to the formation of latent proviruses, suggesting a possible connection between altered integration site selection and latency.734

Diketo acid inhibitors have been developed that specifically and potently block the IN-mediated strand transfer reaction.288 These IN strand transfer inhibitors (INSTIs) bind IN only in the presence of divalent metal and viral DNA and prevent the complex from associating with cellular (target) DNA. Although structurally diverse, INSTIs appear to have common features, including (a) a hydrophobic moiety composed of halobenzyl groups and (b) coplanar oxygen atoms that chelate the Mg+2 ions in the active site.662 Work in the early 2000s showed that naphthyridine carboxamides inhibit the replication of a SHIV in rhesus macaques, setting the stage for in vivo use of INSTIs.289 Further structure–activity relationship studies led to the development of raltegravir687 and elvitegravir.624 Raltegravir and elvitegravir were approved for clinical use in people living with HIV by the U.S. FDA in 2007 and 2014, respectively, and three other INSTIs, dolutegravir, bictegravir, and cabotegravir, are now approved for clinical use. The long-acting injectable Cabenuva combines cabotegravir with the NNRTI rilpivirine.

The observation that INSTIs are active against a wide range of retroviruses,382 including PFV,710 allowed structures to be determined for raltegravir and elvitegravir bound to the PFV IN active site.277 These structural analyses showed that INSTIs chelate the metals in the enzyme active site and demonstrated that the halobenzyl moieties of the INSTI stack against the cytidine of the invariant CA dinucleotide and in doing so supplant the adenine ring and eject its associated DNA strand transfer 3′-OH nucleophile from the active site.277,278,389 As with other HIV inhibitors, resistance develops both in vitro and in vivo. Several major pathways of resistance have been observed for raltegravir, and secondary mutations arise at a number of additional positions. The more recently developed INSTIs are more difficult for the virus to escape. The advances noted above in solving the structure of HIV-1 IN have provided a significant amount of structural information on the binding of INSTIs to HIV-1 IN.354,662 This information will be useful in designing modified inhibitors that are active against INSTI-resistant HIV-1 mutants.

In addition to its primary role in viral DNA integration, a variety of studies have demonstrated a role for IN in RNA condensation in the viral core during maturation (see Maturation section).

Viral Gene Expression The regulation of viral gene expression is rate-limiting in the replication cycle. It occurs at the transcriptional and posttranscriptional levels and uses common cellular and unique viral mechanisms. Two viral proteins play a role in HIV gene expression, Tat and Rev, and both are RNAbinding proteins. After viral integration, the HIV promoter, located within the 5′ LTR, is packaged into host chromatin and behaves in many ways like a highly signal-dependent human promoter. Viral transcription is executed by the cellular RNA polymerase II (Pol II), which forms a nonprocessive “paused” complex downstream of the start of transcription, generating short viral RNAs. RNA Pol II transitions to a processive state in elongation, allowing synthesis of full-length viral RNA, after cellular activation or in the presence of Tat. The activity of the viral promoter is tightly connected to extracellular stimuli that induce cellular transcription factors (TFs) to translocate into the nuclear compartment and bind to various DNA elements that make up the promoter (Fig. 17.20). Distinct TF combinations drive transcription in different cell types (e.g., T cells and monocyte-derived macrophages), activated states (e.g., naive, effector and memory CD4+ T lymphocytes), and tissues (e.g., lymph nodes, gastrointestinal tract, and CNS) targeted by the virus. Posttranscriptionally, viral RNAs undergo distinct splicing steps that facilitate viral RNA export and translation; unspliced RNAs are effectively exported by the action of Rev.

Here we summarize the steps of basal and Tat-induced HIV transcription, and the roles of RNA splicing, nuclear export, and the Rev protein. Basal HIV Transcription After Integration Transcription of HIV, similar to eukaryotic genes comprises four key phases: initiation, promoter-proximal pausing, elongation, and termination.647 Each phase involves dynamic, reversible, posttranslational modifications of Pol II.280 Pol II contains 12 subunits, and the catalytic activity is encoded in the largest subunit, RPB1. RBP1 has a unique region called the C-terminal domain (CTD), which contains a heptad sequence (consensus Tyr1-Ser2-Pro3-Thr4-Ser5-Pro6-Ser7) that is repeated 52 times, making the CTD a highly repetitive, unstructured region of low sequence diversity.570 This region is modified extensively posttranslationally; for example, serine residues, specifically Ser2 and Ser5, of the CTD are dynamically and reversibly phosphorylated.88,199,335,556,773 Transcription also involves other features of gene regulation, such as nucleosome position, DNA sequence, DNA structure, and cotranscriptional processes.15,42,349,569,616

652

FIGURE 17.20 HIV-1 and HIV-2 long terminal repeat (LTR). The HIV-1 LTR is a duplicated 630+ base-pair (bp) element located at the termini of integrated proviral DNA. A blow-up of the 5′ LTR and adjacent cellular and Gag coding sequences is presented at the top. The HIV-1 LTR has been subdivided into three domains: the R (repeat) region is defined as a 96-nt repeat present at the 3′ and 5′ termini of HIV-1 genomic RNA; U5 is an 84-nt segment located immediately 3′ to the R region; and U3 is a 454-nt segment situated immediately 5′ to R. The 5′ LTR and adjacent gag leader sequence (GLS) are aligned with the subdivided LTR (middle) to indicate functionally important binding sites for transcriptional regulatory proteins. The positions of nucleosomes nuc 0, nuc 1, and nuc 2 are indicated. For HIV-2 (bottom), the unique binding sites for the peri-ets (pets) and E26 transforming specific (Ets) transcriptional factors are indicated. HIV was one of the first systems in which promoter-proximal pausing of Pol II was described and intensely studied.360 Multiple TF binding sites on the promoter recruit the Pol II preinitiation complex, a process observed even in latently infected cells. However, progression from promoterproximal pausing into the elongation phase is prevented by the presence of negative elongation factors, suppressive placement of the first nucleosome (Nuc-1), and an absence of positive elongation factors, including the virally encoded Tat protein (Fig. 17.21). Without Tat, randomly and prematurely terminated HIV transcripts are produced. This Tat-deficient phenotype can be studied after transfection of LTRdriven reporter gene constructs in the absence of Tat and in cells harboring integrated HIV-1 proviruses with mutated Tat genes.214,408,702 The presence or absence of Tat is also critical for the latent state of the virus.

LTR The two LTRs flanking the HIV genome are assembled from different regions of the RNA genome representing the U3, R, and U5 regions (Fig. 17.20). The HIV-1 transcriptional promoter and multiple regulatory elements are located within the U3 region and function in the context of the 5′ LTR (Fig. 17.20). Their principal function is to recruit the Pol II holoenzyme to the start site (+1) of viral RNA synthesis (transcription start site/TSS), also defined as the first nucleotide in the LTR R region. In the presence of a functional 5′ LTR, transcription from the 3′ LTR is usually suppressed.378

The HIV-1 promoter can be divided into four main functional regions: a modulatory region (−455 to −104), the enhancer (−109 to −79), the core promoter (−78 to +1), and the leader region (+1 to +188) (Fig. 17.20). Both constitutive and inducible TFs recognize the cis-regulatory elements contained within these functional regions to enhance or repress HIV-1 transcription. Three TFs (nuclear factor-kappa B [NF-κB], activator protein 1 [AP-1], and nuclear factor of activated T cells [NFAT]) are important in regulating HIV-1 RNA synthesis in activated T cells after antigen-specific/MHC-restricted signaling through the T-cell receptor complex, whereas specificity protein 1 (Sp1) regulates basal transcription. These four TFs are further discussed below. In addition, there are binding sites for other TFs (e.g., COUP-TF, c-MyB, C/EBP, Ets-1, LEF-1, RBF-2, ATF4, and AP-4) that are involved in HIV transcription.

FIGURE 17.21 Regulation of HIV transcription. Tat-promoted phosphorylation of the C-terminal domain (CTD) of RNA polymerase II (Pol II) results in processive synthesis of human immunodeficiency virus 1 (HIV-1) messenger RNA (mRNA). In the absence of Tat binding to transactivation response region (TAR), the processivity of the RNA Pol II complex is impaired. When the activation domain of Tat interacts with the cyclin T (cycT1)–CDK9 complex, the conformation of Tat changes and its affinity and specificity for TAR RNA increase. By recruiting cycT1–CDK9 and the Super Elongation Complex (SEC, composed of AFF, ELL2, ENL/AF9) to a promoterproximal location, Tat mediates the hyperphosphorylation of the CTD and promoter clearance (elongation) of the transcriptional complex. (Modified from Ott M, Geyer M, Zhou Q. The control of HIV transcription: keeping RNA polymerase II on track. Cell Host Microbe 2011;10(5):426–435. Copyright © 2011 Elsevier. With permission.)

653

Sp1 The LTRs of HIV and other primate lentiviruses contain three tandemly arranged binding sites for the constitutively expressed Sp1; these DNA elements are situated immediately upstream of a canonical Pol II TATA box (Fig. 17.20). The Sp1 and TATA elements constitute the HIV-1 core promoter and must all be present for basal levels of LTR-directed RNA synthesis.281 Mutations that functionally inactivate all three Sp1 motifs eliminate detectable replication in Jurkat T cells and delay progeny virus production in CEM and H9 cells but have little effect on replication in activated PBMC.417,605

NF-κB The two binding sites for the NF-κB/Rel family of TFs constitute the principal activatable enhancer elements in the HIV-1 LTR. The NF-κB sites (consensus sequence: GGGRNNYYCC) are located upstream and adjacent to the Sp1 binding motifs in all HIV-1 isolates.516 In clade E strains, a single inactivating nucleotide deletion in the upstream NF-κB element converts it to a GA-binding protein (GABP) site, and clade C isolates carry three or four NF-κB sites.25 In vivo, NF-κB is intimately involved in regulating inflammation, cell proliferation, and apoptosis. Because the various members of the NFκB family are present intracellularly as dimeric molecules, a heterogeneous population of homo- and heterodimers, with different functional specificities, bind to the two NF-κB sites in the HIV-1 LTR. For HIV-1 gene expression, the two most important forms of NF-κB are the p50/p50 homodimer, which functions as a repressor, and the p50/p65 heterodimer, which functions as an activator. In addition to its role in basal transcription, the Rel family member RelB interacts with Tat to further enhance HIV transcription.727 Importantly, NF-κB synergizes with Sp1 and AP-1 family TFs in the activation of the HIV promoter. The LTRs of HIV-2 and SIV, while retaining the triplicated Sp1 motifs, each contain only a single NF-κB binding site (Fig. 17.20).

NFAT Like NF-κB, members of the NFAT family of proteins are sequestered in the cytoplasm and transported into the nucleus after an increase in the levels of intracellular calcium.700 Those increases activate the calcineurin serine/threonine phosphatase and result in the phosphorylation of NFAT, exposure of the NFAT nuclear localization sequence (NLS), and nuclear translocation of NFAT. This process is suppressed in resting T cells by the master regulator of cell quiescence, FOXO1. This protein acts by enhancing autophagy and reducing ER stress signals, which would otherwise activate HIV transcription by mobilizing NFAT and ATF4 TFs.340,607,711

AP-1 The rate of HIV-1 viral RNA synthesis can also be stimulated by AP1, which consists of Jun homodimers or Jun/Fos heterodimers. Depending on the HIV-1 subtype, AP1 binds to one or two DNA elements located immediately upstream of the two NF-κB sites in the HIV-1 LTR, or the three sites downstream of the TSS in the leader region. AP1 also cooperates with NF-κB and activates the viral promoter via the two NF-κB binding motifs. AP1 is activated by the c-Jun N-terminal kinase (JNK) and the extracellular signal-related kinase (ERK).362

Epigenetic Regulation of HIV Eukaryotic gene expression is closely associated with nucleosome positioning.631 Nucleosomes comprise 147 bp of DNA wrapped around an octamer of pairs of the four core histone proteins (i.e., H2A, H2B, H3, and H4). Promoter TF binding site accessibility is affected by the position of nucleosomes, and transcription machinery accesses DNA through the process of nucleosome remodeling. The positioning and stability of nucleosomes are regulated by their movement, deposition, or ejection by chromatin-remodeling complexes or posttranslational modification of the histone N-terminal tails.71 These PTMs serve as a “histone code,” allowing proteins with cognate recognition domains to interact with histones and then affect gene expression. The HIV epigenome is being targeted experimentally and clinically to manipulate the activity of the HIV provirus during latency.

When HIV-1 DNA becomes stably integrated into the host cell chromosomal DNA, it acquires precise nucleosomal positioning, independent from the integration site. Studies using DNA footprinting and restriction enzyme accessibility of integrated viral DNA revealed nucleosomes both upstream and downstream of the HIV-1 promoter (designated nuc-0, nuc-1, and nuc-2 in Fig. 17.20).143 Nucleosome positioning defines two open chromatin regions encompassing (a) the promoter/enhancer sequences (−250 to +11 [relative to the transcription start site]) and (b) a downstream segment that begins within the 3′ portion of U5 and extends downstream of the PBS (+150 to +250). TFs are thought to bind to their cognate recognition motifs to maintain these open chromatin regions (Fig. 17.20). The effects of TFs on HIV-1 promoter activity depend on the function of coactivators and corepressors, which often induce chromatin remodeling by posttranslationally modifying histone tails on assembled nucleosomes.

Acetylation is one of the best studied histone modifications, and recruitment of histone acetyl transferases (HATs) and histone deacetyl transferases (HDACs) to the HIV LTR is mediated by TFs and other key transcriptional regulatory factors, such as BRD4, YY1, and the HUSH complex. Inhibitors of HDACs were the first epigenetic interventions experimentally and clinically tried to reverse HIV latency.79,582,716 BRD4, a member of the bromodomain and ET domain (BET) protein family, binds acetylated histones through its double bromodomains. BRD4 exists in long and short isoforms; the long isoform recruits a critical cellular host factor, the positive transcription elongation factor (P-TEFb) to cellular promoters and is a transcriptional co-activator.764 The short isoform, BRD4S, acts as a corepressor of HIV-1 transcription by binding to the catalytic subunit BRG1 of the SWI/SNF BAF complex, which promotes remodeling of nuc-1 in a repressive conformation downstream of the TSS.60,146 Interestingly, other members of SWI/SNF activate HIV transcription in conjunction with the acetylated form of Tat (see below). Other bromodomain-containing proteins act on the HIV promoter: for example, the cellular factor P300/CBP-associated factor (PCAF) also interacts with the acetylated form of Tat through its bromodomain and coactivates the HIV LTR.533,716

The HIV promoter is also regulated by reversible methylation of lysines, arginines, and DNA. Proteins can be mono-, di-, or trimethylated, depending on the specificity of the methyltransferase. The histone lysine methyltransferase, SMYD2, represses HIV reactivation from latency by monomethylating lysine 20 of histone H4 at the HIV LTR. In addition, the reader protein L3MBTL1, which acts as a chromatin compactor, is recruited to the LTR in an SMYD2-dependent manner. The TF YY1 also represses HIV promoter activity by recruiting HDAC1.47,151 Enhancer of Zeste 2 polycomb repressive complex 2 subunit (EZH2) associates with the promoter/enhancer region of HIV-1 proviruses in latently infected Jurkat T-cell lines and is found with the corresponding trimethylation mark at lysine27 in histone H3 (H3K27me3).236 Knockdown of EZH2 with shRNA or treatment with an EZH2 inhibitor reactivates a significant portion of silenced proviruses.522 Euchromatin histone 654

methyltransferases (EHMTs) mono- and dimethylate histone 3 on lysine 9 (H3K9me1 and H3K9me2) in euchromatic regions of the genome386 and both mammalian EHMTs, G9a and GLP, are responsible for transcriptional repression of the HIV-LTR by promoting repressive dimethylation at H3K9.186,323 SUV39H1 specifically catalyzes trimethylation on histone H3 lysine 9 (H3K9me3) using monomethylated H3K9 as substrate.588 H3K9me3 is a hallmark of facultative and constitutive heterochromatin and recruits heterochromatic protein 1α (HP1α), which binds H3K9me2/3 through its chromodomain.237 Several studies link SUV39H1 to HIV-1 latency in microglial cells465 and peripheral blood mononucleated cells (PBMCs) isolated from infected individuals.196

The human silencing hub (HUSH) complex is a key regulator of silencing endogenous retroviruses and transposable elements in mammalian genomes. The complex comprises three core proteins, TASOR, MPP8, and PPHLN1. Their exact functions are unknown, but by recruiting SETDB1, the complex is involved in reading and writing histone H3 lysine 9 trimethyl marks (H3K9me3) and spreads repressive chromatin marks. In several HIV latency models, inhibiting the HUSH complex reactivated the virus.694

DNA methylation involves the transfer of a methyl group onto cytosines to form 5-methylcytosine. DNA methylation of CpG (cytosine-proximal guanine) islands (CPGI) influences the chromatin environment and plays a role in epigenetic regulation of mammalian gene expression,149 generally by recruiting repressors or blocking the binding of TFs.495 The HIV-1 genome contains five CpGIs111: two surround the promoter region and flank the HIV-1 TSS and several TF binding sites at the 5′ LTR, two other CpGIs are located in the env gene, surrounding the HIV-1 antisense open reading frame, and the fifth CpGI is located in the 3′ LTR, where the antisense transcription start site is located.104 In cultured HIV-1–infected cells, promoter DNA hypermethylation stabilizes HIV-1 latency and demethylating agents induce activation of HIV-1 transcription.64,69,111,366,705 However, studies performed in infected individuals yielded controversial results.63,300

Tat Tat is an indispensable viral protein that increases steady-state levels of viral RNA in a positive feedback loop by directing formation of a more processive Pol II transcription complex to produce itself and viral RNAs in virus-infected cells. When the tat gene is mutagenized, no detectable progeny virions are produced.164 Tat is a small, pleiotropic protein that interacts with a set of critical host factors and links to TFs, the Pol II complex and the epigenetic machinery controlling HIV transcription. Non–LTR-related functions of Tat include regulation of T-cell activity, apoptosis, oxidative stress, and inflammation.7 Tat exerts its transactivator function by binding to the TAR RNA element. HIV-1 TAR encompasses the 5′-terminal 59 nt of all viral RNAs and folds into a stable stem-loop structure (Fig. 17.22). The minimal TAR element (mapping between bases +19 and +42) has three critical components: a base-paired stem, a trinucleotide bulge (with the sequence UCU at positions +23 to +25), and a hexanucleotide G-rich loop.216 Sequences located in both the hexanucleotide loop and the bulge of HIV-1 TAR are required for Tat function. HIV-1 Tat binds to wild-type (WT) or “loop” mutants of TAR but not to TAR elements with alterations that affect the bulge region.188,609 Interestingly, HIV-2 TAR forms a double stem-loop structure, each arm of which possesses a dinucleotide bulge and the hexanucleotide G-rich loop (Fig. 17.22).202 The interaction between Tat and TAR is being explored as a drug target to block the interaction either to suppress HIV replication202 or to enhance HIV latency by inducing a state of “deep latency.”421,485,503,504

P-TEFb The Tat/TAR complex activates HIV transcription both at the initiation and elongation steps. After initiation, Pol II pauses proximal to the promoter, at an average of 25 to 50 bp downstream of the TSS.148 Establishment and maintenance of promoter-proximal pausing is performed by the interaction of the Pol II CTD with negative elongation factors NELF and DSIF.121,758 This state is critical for HIV transcriptional regulation. Recruitment of P-TEFb catalyzes the transition of Pol II from a paused state to an actively elongating complex, called pause release.148

HIV research was critical in the discovery of P-TEFb. Reports describing the specific interaction of HIV-2 Tat with a cellular kinase provided the first mechanistic clues about Tat function.294,295 The Tat-related kinase component in human P-TEFb was subsequently identified as CDK9, a 42-kDa CDC2-related kinase. However, the CDK9 catalytic subunit alone failed to bind to Tat in in vitro assays.763 These unresolved issues were clarified with the isolation of an 87-kDa protein from nuclear extracts bound to Tat.730 The new protein, named cyclin T (because it bound to Tat), is one of many cyclin regulatory subunits interacting with the CDK9 family of kinases (i.e., T1, T2a, T2b, or K).548 Of these P-TEFb complexes, only those containing cyclin T1 directly bind to the HIV-1 Tat activation domain.730

P-TEFb is incorporated into a larger complex containing additional transcription elongation factors termed the superelongation complex (SEC). Besides P-TEFb, the SEC consists of the Pol II elongation factors eleven-nineteen Lys-rich leukemia (ELL) proteins and several frequent mixed lineage leukemia (MLL) translocation partners. ELL2 and P-TEFb act cooperatively to promote efficient Tat-mediated elongation.290 AFF4, ENL, AF9, and elongation factor ELL2 are components of the Tat-P-TEFb complex, and Tat stabilizes ELL2.290 P-TEFb phosphorylates Ser2 residues of the CTD to enhance polymerase processivity, but in the presence of Tat, it also phosphorylates serine 5.469,552,575,636,785 P-TEFb also phosphorylates DSIF and NELF; phosphorylation of NELF leads to its dissociation from the Pol II complex, whereas phosphorylation of DSIF turns it into a positive elongation factor.148

655

FIGURE 17.22 Structure of Tat and its transactivation response region (TAR) element. A: Schematic representation of the human immunodeficiency virus 1 (HIV-1) Tat protein with the cysteine-rich activation, core, and TAR-binding domains indicated. B: Structure of the stem-bulge-loop configurations of HIV-1 and HIV-2 TAR elements. C: Crystal structure of Tat/TAR binding in the presence of the super elongation complex. In proliferating cells, P-TEFb is maintained in an inactive form in the cytoplasm through its reversible association with the 7SK small nuclear ribonucleoprotein (snRNP). The 7SK small nuclear RNA folds into a 3D scaffold that is stabilized by interactions with several proteins (Fig. 17.23). Of particular importance to HIV is the homodimer of the kinase inhibitor hexamethylene bisacetamide-inducible protein 1/2 (HEXIM). Several phosphorylation events, including especially Thr-186 on the CDK9 subunit, and acetylation of CDK9 and Cyclin T1, regulate activity and availability outside the 7SK snRNP.125 In resting CD4+ T cells, P-TEFb regulation occurs through miRNA-mediated downregulation of CycT1 and sequestration of inactive CDK9 in the cytoplasm.124

Tat expression during the early phase of productive infection results in the dissociation of 7SK snRNA from P-TEFb and formation of the Tat/PTEFb complex.643 This is facilitated, in part, by the fact that the Tat RNA-binding domain is highly homologous to the 7SK snRNP-interacting domain in the HEXIM1 inhibitor.555,768 The interaction of Tat and the 7SKsnRNA releases HEXIM-1 from the 7SKsnRNA and relieves P-TEFb inhibition (Fig. 17.23). Moreover, subsequent P-TEFb inactivation is prevented by the interaction of Tat and the 7SKsnRNA, which prevents reassembly of the 7SKsnRNP complex.29,509,555

Structure HIV-1 Tat contains 86 to 101 amino acid residues encoded by two exons (Fig. 17.22). A shorter, 72-amino acid “one-exon” HIV-1 Tat protein possesses all the transcriptional activating properties of full-length Tat, as measured in tissue culture infections or in LTR-driven reporter gene experiments. The termination codon after the first Tat exon is highly conserved among diverse HIV-1 isolates, suggesting that the one-exon and two-exon Tat proteins mediate different functions during productive viral infections in vivo. Immune modulatory effects of the second exon of Tat have been described.532

FIGURE 17.23 Differential Recruitment of P-TEFb from the 7SK snRNP during HIV Infection. The 7SK snRNP is a cellular reservoir of inactive P-TEFb that contains HEXIM1, an inhibitor of the CDK9 kinase activity. P-TEFb is recruited from the 7SK snRNP by Tat for transactivation of HIV-1 transcription or is released upon exposure of cells to hypertrophic or stress signals to associate with bromodomain-containing protein 4 (Brd4) to induce basal HIV transcription in the absence of Tat. (Modified from Ott M, Geyer M, Zhou Q. The control of HIV transcription: keeping RNA polymerase II on track. Cell Host Microbe 2011;10(5):426–435. Copyright © 2011 Elsevier. With permission. Ref.5)

The functional organization of the HIV-1 Tat protein was deduced from TAR binding and transcriptional activation experiments with WT and mutagenized derivatives of Tat. The tripartite Tat activation (or effector) domain encompasses the N-terminal 48 residues, which include (a) a string of highly acidic amino acids (residues 1 to 21); (b) a Cys-rich region (seven invariant, six of which are required for function) at positions 22 to 37; and (c) a hydrophobic core segment (amino acids 38 to 48) that is highly conserved among HIV isolates (Fig. 17.22). The activation domain is critical for recruiting P-TEFb to TAR; mutations affecting this region drastically reduce or eliminate transactivation activity.610 The TAR RNA binding domain of Tat was mapped to a Lys/Arg-rich region at residues 48 to 57 and is highly posttranslationally modified (Fig. 17.22). Peptides from this Tat segment bind to the TAR bulge region and a few base pairs surrounding the bulge, but with somewhat less affinity and specificity than does full-length Tat.147

While many structural studies have investigated Tat binding to TAR and P-TEFb binding to Tat, Tat binding to TAR is highly cooperative and involves formation of a trimolecular complex composed of Tat, TAR, and P-TEFb (including the SEC).56,249,730,762,785 Upon binding of Tat/SEC, the TAR loop is stabilized by cross-loop hydrogen bonds. It makes structure-specific contacts with the side chains of the Tat–TAR recognition motif (TRM) of cyclin T1 and the zinc-coordinating region of Tat.637 Despite limited resolution, it is clear that the TAR loop evolved 656

to make high-affinity interactions with the cyclin T1 TRM, whereas Tat has three roles in the trimolecular complex: scaffolding and stabilizing the TRM, making specific interactions through its zinc-coordinating loop, and engaging in electrostatic interactions through its ARM (Fig. 17.22). Besides P-TEFb, Tat has numerous other interacting partners in host cells as seen in proteomic screens252,332 and described in functional studies.56,249,730,785,788 These studies identified the SEC as a Tat/P-TEFb cofactor. Additional cofactors regulate the Tat/TAR/P-TEFb interaction, enhance HIV transcription elongation, link Tat to the cellular epigenetic machinery, or modify the host response to HIV infection.120,290,535

Posttranslational Modifications Tat function is regulated by a series of PTMs, mainly in its RNA-binding domain. These modifications fine-tune Tat’s function in transcription. For example, this function is inhibited by methylation of Arg52 and Arg53 by the arginine methyltransferase PRMT6 and methylation of Lys50 and Lys51 by the lysine methyltransferase SETDB1 as both inhibit the interaction with TAR and P-TEFb.70,713

Activating Tat modifications include phosphorylation, polyubiquitination, acetylation, and monomethylation. For example, Tat is acetylated on its lysine 28 residue by PCAF to mediate P-TEFb recruitment to the Tat–TAR complex.156,374 Lys28 acetylation can be reversed by HDAC6.316 Tat is also acetylated on Lys50 by the cellular proteins hGCN5 and p300/CBP to release the Tat–P-TEFb complex from TAR.356,507 In addition, Tat is monomethylated by the lysine methyltransferase Set7/9 on lysine 51 and 71—both are needed for full Tat transactivation ability11,535— and Lys71 is also polyubiquinated.76 Demethylation of Tat on Lys51 by the demethylase LSD1 is important for Tat-mediated viral reactivation.617 LSD1 also represses HIV transcription in microglial cells through its normal function as a histone demethylase.410

Function in Mice Tat transactivation of HIV-1 LTR-driven gene expression is quite low in rodent cells but can be greatly augmented in rodent somatic cell hybrids containing human chromosome 12.283 This defect can also be corrected by overexpressing human cyclin T in rodent cells, which results in enhanced Tat transactivation of the HIV-1 LTR to levels measured in human cells.730 The murine homolog of human cyclin T binds to HIV-1 Tat, but the resulting Tat/murine cyclin T heterodimer is not efficiently recruited to TAR.240,249 Substitution of Tyr260 in murine cyclin T with the Cys residue at that position in the human homolog also restores Tat activity in rodent cells.56,249 Thus, the extremely weak activity of Tat in rodent cells is not due to a failure to form the Tat/cyclin T heterodimer; rather, the cross-species heterodimer that does form is unable to bind to TAR and deliver P-TEFb to a poorly processive transcription complex.

Latency In addition to initiating productive virus infections, HIV-1 establishes latent infections in memory CD4+ T cells and possibly other reservoir cells (e.g., macrophages) that persist in patients receiving ART.132,217,741 The long half-life of memory T cells and their expansion after periodic restimulation prevent natural elimination of infected cells during the lifetime of infected individuals. Although most current latency studies are performed in the blood, the majority of reservoir cells reside in tissues, especially the gut and lymph nodes. Latency is maintained through homeostatic proliferation, antigenic stimulation and clonal expansion. In individuals on ART, lymph node CD4+ T cells composed of about 65% T follicular helper cells, as defined by the expression of the cell-surface receptors CXCR5 and PD-1, were identified as the major sources of replication-competent HIV-1.155

Unlike other viruses such as herpes viruses, HIV lacks a dedicated latency program and encodes no proteins to actively regulate the establishment and maintenance of latency. In many ways, HIV latency can be defined as a lack of Tat; the HIV promoter becomes inactive and dependent on regulation through cellular TFs and extracellular stimuli. As such, HIV latency is a labile state of HIV with periodic low-level reactivation and homeostatic expansion of latent clones.477 Importantly, latently infected lymphocytes carry replication-competent proviruses versus lymphocytes that contain defective proviruses that are not considered latent. As no or very few viral proteins are produced in latently infected cells, these latently infected cells are invisible to the immune system and persist during ART.

The size of the viral reservoir varies between individuals and depends on the onset of treatment, the viral set point, clade, and genetic diversity.24,473,529 Approximately 1 in 106 CD4+ T cells is latently infected in most patients.657 The half-life of latently infected T cells was estimated to be 44 months, but predictions about whether the reservoir decays or increases over time are at odds.24,473,529,549 The mechanisms responsible for establishing and maintaining latently infected CD4+ T cells have only been partially elucidated. A prevailing model posits that latently infected cells arise from reversion of activated effector T cells to a quiescent memory T lymphocyte phenotype,33,67,468,587 but they may also be the product of direct infection of resting CD4+ T cells97 or may arise in activated infected T cells before conversion to cellular quiescence.110,587

It is important to distinguish latent cells from the large number of cells with defective proviruses. The latter may outnumber truly latent cells by greater than a factor of greater than 1083 and obscure accurate detection of the active reservoir. While culturing T cells from ART-suppressed individuals in an in vitro viral outgrowth assay [such as the quantitative viral outgrowth assay (QVOA)] underestimates the size of the reservoir, analysis of integrated viral DNA overestimates it.83 Recent advances in intact proviral DNA measurements either by droplet digital PCR or next generation sequencing aid in more accurately assessing the number of cells containing nondefective proviruses.83,84,401

There has been debate as to whether the very low levels of viremia ( 200 x 106/l. AIDS 2001;15:1509–1515. 270. Koot M, Keet IPM, Vos AHV, et al. Prognostic value of HIV-1 syncytium-inducing phenotype for rate of CD4+ cell depletion and progression to AIDS. Ann Intern Med 1993;118:681–688. 271. Kopp JB, Smith MW, Nelson GW, et al. MYH9 is a major-effect risk gene for focal segmental glomerulosclerosis. Nat Genet 2008;40:1175–1184. 272. Korber B, Muldoon M, Theiler J, et al. Timing the ancestor of the HIV-1 pandemic strains. Science 2000;288:1789–1796. 273. Kostrikis LG, Huang Y, Moore JP, et al. A chemokine receptor CCR2 allele delays HIV-1 disease progression and is associated with a CCR5 promoter mutation. Nat Med 1998;4:350–353. 274. Kotler DP, Grunfeld C. Pathophysiology and treatment of the AIDS wasting syndrome. In: Volberding P, Jacobson MA, eds. AIDS Clinical Review 1995/1996. New York: Marcel Dekker, Inc.; 1995:229–275. 725

275. Koup RA, Safrit JT, Cao Y, et al. Temporal association of cellular immune responses with the initial control of viremia in primary human immunodeficiency virus type 1 syndrome. J Virol 1994;68:4650–4655. 276. Kozal M, Aberg J, Pialoux G, et al. Fostemsavir in adults with multidrug-resistant HIV-1 infection. N Engl J Med 2020;382:1232–1243. 277. Kozal MJ, Kroodsma K, Winters MA, et al. Didanosine resistance in HIV-infected patients switched from zidovudine to didanosine monotherapy. Ann Intern Med 1994;121:263–268. 278. Kuller LH, Tracy R, Belloso W, et al. Inflammatory and coagulation biomarkers and mortality in patients with HIV infection. PLoS Med 2008;5:e203. 279. Kuritzkes DR, Bell S, Bakhtiari M. Rapid CD4+ cell decline after sexual transmission of a zidovudine-resistant syncytium-forming isolate of HIV-1. AIDS 1994;8:1017–1019. 280. Kuritzkes DR, Jacobson J, Powderly WG, et al. Antiretroviral activity of the anti-CD4 monoclonal antibody TNX-355 in patients infected with human immunodeficiency virus type 1. J Infect Dis 2004;189:286–291. 281. Lalezari JP, Henry K, O’Hearn M, et al. Enfuvirtide, an HIV-1 fusion inhibitor, for drug-resistant HIV infection in North and South America. N Engl J Med 2003;348:2175–2185. 282. Landovitz RJ, Donnell D, Clement ME, et al. Cabotegravir for HIV prevention in cisgender men and transgender women. N Engl J Med 2021;385:595–608. 283. Lavreys L, Baeten JM, Chohan V, et al. Higher set point plasma viral load and more-severe acute HIV type 1 (HIV-1) illness predict mortality among high-risk HIV-1-infected African women. Clin Infect Dis 2006;42:1333–1339. 284. Le Gall S, Erdtmann L, Benichou S, et al. Nef interacts with the mu subunit of clathrin adaptor complexes and reveals a cryptic sorting signal in MHC I molecules. Immunity 1998;8:483–495. 285. Ledergerber B, Lundgren JD, Walker AS, et al. Predictors of trend in CD4-positive T-cell count and mortality among HIV-1-infected individuals with virological failure to all three antiretroviral-drug classes. Lancet 2004;364:51–62. 286. Lederman M, Connick E, Landay A, et al. Immunologic responses associated with 12 weeks of combination antiretroviral therapy consisting of zidovudine, lamivudine, and ritonavir: results of AIDS clinical trials group protocol 315. J Infect Dis 1998;178:70–79. 287. Lennox JL, DeJesus E, Lazzarin A, et al. Safety and efficacy of raltegravir-based versus efavirenz-based combination therapy in treatmentnaive patients with HIV-1 infection: a multicentre, double-blind randomised controlled trial. Lancet 2009;374:796–806. 288. Levi J, Raymond A, Pozniak A, et al. Can the UNAIDS 90-90-90 target be achieved? A systematic analysis of national HIV treatment cascades. BMJ Glob Health 2016;1:e000010. 289. Li Q, Duan L, Estes JD, et al. Peak SIV replication in resting memory CD4+ T cells depletes gut lamina propria CD4+ T cells. Nature 2005;434:1148–1152. 290. Li JZ, Paredes R, Ribaudo HJ, et al. Low-frequency HIV-1 drug resistance mutations and risk of NNRTI-based antiretroviral treatment failure: a systematic review and pooled analysis. JAMA 2011;305:1327–1335. 291. Li Y, Svehla K, Louder MK, et al. Analysis of neutralization specificities in polyclonal sera derived from human immunodeficiency virus type 1-infected individuals. J Virol 2009;83:1045–1059. 292. Lindback S, Karlsson AC, Mittler J, et al. Viral dynamics in primary HIV-1 infection. Karolinska Institutet Primary HIV Infection Study Group. AIDS. 2000;14:2283–2291. 293. Lingappa JR, Thomas KK, Hughes JP, et al. Partner characteristics predicting HIV-1 set point in sexually acquired HIV-1 among African seroconverters. AIDS Res Hum Retroviruses 2013;29:164–171. 294. Link JO, Rhee MS, Tse WC, et al. Clinical targeting of HIV capsid protein with a long-acting small molecule. Nature 2020;584:614–618. 295. Little SJ, Holte S, Routy JP, et al. Antiretroviral-drug resistance among patients recently infected with HIV. N Engl J Med 2002;347:385–394. 296. Liu Z, Cumberland WG, Hultin LE, et al. Elevated CD38 antigen expression on CD8+ T cells is a stronger marker for the risk of chronic HIV disease progression to AIDS and death in the Multicenter AIDS Cohort Study than CD4+ cell count, soluble immune activation markers, or combinations of HLA-DR and CD38 expression. J Acquir Immune Defic Syndr Hum Retrovirol 1997;16:83–92. 297. Liu R, Paxton WA, Choe S, et al. Homozygous defect in HIV-1 coreceptor accounts for resistance of some multiply-exposed individuals to HIV-1 infection. Cell 1996;86:367–377. 298. Liu L, Zhang Q, Chen P, et al. Foxp3(+)Helios(+) regulatory T cells are associated with monocyte subsets and their PD-1 expression during acute HIV-1 infection. BMC Immunol 2019;20:38. 299. Lok JJ, Bosch RJ, Benson CA, et al. Long-term increase in CD4+ T-cell counts during combination antiretroviral therapy for HIV-1 infection. AIDS 2010;24:1867–1876. 300. Lonergan JT, Behling C, Pfander H, et al. Hyperlactatemia and hepatic abnormalities in 10 human immunodeficiency virus-infected patients receiving nucleoside analogue combination regimens. Clin Infect Dis 2000;31:162–166. 301. Lopez-Vazquez A, Mina-Blanco A, Martinez-Borra J, et al. Interaction between KIR3DL1 and HLA-B*57 supertype alleles influences the progression of HIV-1 infection in a Zambian population. Hum Immunol 2005;66:285–289. 302. Lore K, Sonnerborg A, Brostrom C, et al. Accumulation of DC-SIGN+CD40+ dendritic cells with reduced CD80 and CD86 expression in lymphoid tissue during acute HIV-1 infection. AIDS 2002;16:683–692. 303. Lorenzi JC, Cohen YZ, Cohn LB, et al. Paired quantitative and qualitative assessment of the replication-competent HIV-1 reservoir and comparison with integrated proviral DNA. Proc Natl Acad Sci U S A 2016;113:E7908–E7916. 304. Lu W, Ma F, Churbanov A, et al. Virus-host mucosal interactions during early SIV rectal transmission. Virology 2014;464-465:406–414. 305. Lyles RH, Munoz A, Yamashita TE, et al. Natural history of human immunodeficiency virus type 1 viremia after seroconversion and proximal to AIDS in a large cohort of homosexual men. J Infect Dis 2000;181:872–880. 306. Lynch RM, Boritz E, Coates EE, et al. Virologic effects of broadly neutralizing antibody VRC01 administration during chronic HIV-1 infection. Sci Transl Med 2015;7:319ra206. 307. Macdonald JC, Torriani FJ, Morse LS, et al. Lack of reactivation of cytomegalovirus (CMV) retinitis after stopping CMV maintenance therapy in AIDS patients with sustained elevations in CD4 T cells in response to highly active antiretroviral therapy. J Infect Dis 1998;177:1182–1187. 308. Maldarelli F, Wu X, Su L, et al. HIV latency. Specific HIV integration sites are linked to clonal expansion and persistence of infected cells. Science 2014;345:179–183. 309. Mallal S, Nolan D, Witt C, et al. Association between presence of HLA-B*5701, HLA-DR7, and HLA-DQ3 and hypersensitivity to HIV-1reverse-transcriptase inhibitor abacavir. Lancet 2002;359:727–732. 310. Mallon PW, Brunet L, Hsu RK, et al. Weight gain before and after switch from TDF to TAF in a U.S. cohort study. J Int AIDS Soc 2021;24:e25702. 311. Manches O, Frleta D, Bhardwaj N. Dendritic cells in progression and pathology of HIV infection. Trends Immunol 2014;35:114–122. 312. Mangili A, Murman DH, Zampini AM, et al. Nutrition and HIV infection: review of weight loss and wasting in the era of highly active antiretroviral therapy from the nutrition for healthy living cohort. Clin Infect Dis 2006;42:836–842. 313. Margot NA, Lu B, Cheng A, et al. Resistance development over 144 weeks in treatment-naive patients receiving tenofovir disoproxil fumarate or stavudine with lamivudine and efavirenz in Study 903. HIV Med 2006;7:442–450. 726

314. Marin B, Thiebaut R, Bucher HC, et al. Non-AIDS-defining deaths and immunodeficiency in the era of combination antiretroviral therapy. AIDS 2009;23:1743–1753. 315. Markowitz M, Grobler JA. Islatravir for the treatment and prevention of infection with the human immunodeficiency virus type 1. Curr Opin HIV AIDS 2020;15:27–32. 316. Marrazzo JM, Ramjee G, Richardson BA, et al. Tenofovir-based preexposure prophylaxis for HIV infection among African women. N Engl J Med 2015;372:509–518. 317. Martin MP, Dean M, Smith MW, et al. Genetic acceleration of AIDS progression by a promoter variant of CCR5. Science 1998;282:1907–1911. 318. Martin MP, Gao X, Lee JH, et al. Epistatic interaction between KIR3DS1 and HLA-B delays the progression to AIDS. Nat Genet 2002;31:429–434. 319. Martin AM, Nolan D, Gaudieri S, et al. Predisposition to abacavir hypersensitivity conferred by HLA-B*5701 and a haplotypic Hsp70Hom variant. Proc Natl Acad Sci U S A 2004;101:4180–4185. 320. Martin-Gayo E, Buzon MJ, Ouyang Z, et al. Potent cell-intrinsic immune responses in dendritic cells facilitate HIV-1-specific T cell immunity in HIV-1 elite controllers. PLoS Pathog 2015;11:e1004930. 321. Masur H, Ognibene FP, Yarchoan R, et al. CD4 counts as predictors of opprotunistic pneumonia in human immunodeficiency virus (HIV) infection. Ann Intern Med 1989;111:223–231. 322. Matheron S, Pueyo S, Damond F, et al. Factors associated with clinical progression in HIV-2 infected-patients: the French ANRS cohort. AIDS 2003;17:2593–2601. 323. Mattapallil JJ, Douek DC, Hill B, et al. Massive infection and loss of memory CD4+ T cells in multiple tissues during acute SIV infection. Nature 2005;434:1093–1097. 324. Mayer KH, Mimiaga MJ, Gelman M, et al. Raltegravir, tenofovir DF, and emtricitabine for postexposure prophylaxis to prevent the sexual transmission of HIV: safety, tolerability, and adherence. J Acquir Immune Defic Syndr 2012;59:354–359. 325. Mayer KH, Molina JM, Thompson MA, et al. Emtricitabine and tenofovir alafenamide vs emtricitabine and tenofovir disoproxil fumarate for HIV pre-exposure prophylaxis (DISCOVER): primary results from a randomised, double-blind, multicentre, active-controlled, phase 3, non-inferiority trial. Lancet 2020;396:239–254. 326. McCarthy KR, Kirmaier A, Autissier P, et al. Evolutionary and functional analysis of old world primate TRIM5 reveals the ancient emergence of primate lentiviruses and convergent evolution targeting a conserved capsid interface. PLoS Pathog 2015;11:e1005085. 327. McComsey GA, Kitch D, Daar ES, et al. Bone mineral density and fractures in antiretroviral-naive persons randomized to receive abacavir-lamivudine or tenofovir disoproxil fumarate-emtricitabine along with efavirenz or atazanavir-ritonavir: AIDS Clinical Trials Group A5224s, a substudy of ACTG A5202. J Infect Dis 2011;203:1791–1801. 328. McCormack S, Dunn DT, Desai M, et al. Pre-exposure prophylaxis to prevent the acquisition of HIV-1 infection (PROUD): effectiveness results from the pilot phase of a pragmatic open-label randomised trial. Lancet 2016;387:53–60. 329. McElrath MJ, De Rosa SC, Moodie Z, et al. HIV-1 vaccine-induced immunity in the test-of-concept Step Study: a case-cohort analysis. Lancet 2008;372:1894–1905. 330. McMahon D, Jones J, Wiegand A, et al. Short-course raltegravir intensification does not reduce persistent low-level viremia in patients with HIV-1 suppression during receipt of combination antiretroviral therapy. Clin Infect Dis. 2010;50:912–919. 331. McMichael AJ, Carrington M. Topological perspective on HIV escape. Science 2019;364:438–439. 332. Mehandru S, Poles MA, Tenner-Racz K, et al. Primary HIV-1 infection is associated with preferential depletion of CD4+ T lymphocytes from effector sites in the gastrointestinal tract. J Exp Med 2004;200:761–770. 333. Mellors JW, Kingsley LA, Rinaldo CR, et al. Quantitation of HIV-1 RNA in plasma predicts outcome after seroconversion. Ann Intern Med 1995;122:573–579. 334. Mellors JW, Munoz A, Giorgi JV, et al. Plasma viral load and CD4 + lymphocytes as prognostic markers of HIV-1 infection. Ann Intern Med 1997;126:946–954. 335. Mellors JW, Rinaldo CR, Gupta P, et al. Prognosis of HIV-1 infection predicted by quantity of virus in plasma. Science 1996;272:1167–1170. 336. Meng TC, Fischl MA, Boota AM, et al. Combination therapy with zidovudine and dideoxycytidine in patients with advanced human immunodeficiency virus infection. A phase I/II study. Ann Intern Med 1992;116:13–20. 337. Michael NL, Louie LG, Rohrbaugh AL, et al. The role of CCR5 and CCR2 polymorphisms in HIV-1 transmission and disease progression. Nat Med 1997;3:1160–1162. 338. Migueles SA, Connors M. Long-term nonprogressive disease among untreated HIV-infected individuals: clinical implications of understanding immune control of HIV. JAMA 2010;304:194–201. 339. Migueles SA, Osborne CM, Royce C, et al. Lytic granule loading of CD8+ T cells is required for HIV-infected cell elimination associated with immune control. Immunity 2008;29:1009–1021. 340. Mitsuki YY, Tuen M, Hioe CE. Differential effects of HIV transmission from monocyte-derived dendritic cells vs. monocytes to IL-17+CD4+ T cells. J Leukoc Biol 2017;101:339–350. 341. Mocroft A, Gill MJ, Davidson W, et al. Are there gender differences in starting protease inhibitors, HAART, and disease progression despite equal access to care? J Acquir Immune Defic Syndr 2000;24:475–482. 342. Molina JM, Andrade-Villanueva J, Echevarria J, et al. Once-daily atazanavir/ritonavir versus twice-daily lopinavir/ritonavir, each in combination with tenofovir and emtricitabine, for management of antiretroviral-naive HIV-1-infected patients: 48 week efficacy and safety results of the CASTLE study. Lancet 2008;372:646–655. 343. Molina JM, Capitant C, Spire B, et al. On-demand preexposure prophylaxis in men at high risk for HIV-1 infection. N Engl J Med 2015;373:2237–2246. 344. Molla A, Korneyeva M, Gao Q, et al. Ordered accumulation of mutations in HIV protease confers resistance to ritonavir. Nat Med 1996;2:760–766. 345. Moore RD, Wong WME, Keruly JC, et al. Incidence of neuropathy in HIV-infected patients on monotherapy versus those on combination therapy with didanosine, stavudine and hydroxyurea. AIDS 2000;14:273–278. 346. Moran CA, Weitzmann MN, Ofotokun I. The protease inhibitors and HIV-associated bone loss. Curr Opin HIV AIDS 2016;11:333–342. 347. Morou A, Brunet-Ratnasingham E, Dube M, et al. Altered differentiation is central to HIV-specific CD4(+) T cell dysfunction in progressive disease. Nat Immunol 2019;20:1059–1070. 348. Moyle GJ, Wildfire A, Mandalia S, et al. Epidemiology and predictive factors for chemokine receptor use in HIV-1 infection. J Infect Dis 2005;191:866–872. 349. Muema DM, Akilimali NA, Ndumnego OC, et al. Association between the cytokine storm, immune cell dynamics, and viral replicative capacity in hyperacute HIV infection. BMC Med 2020;18:81. 350. Muller M, Wandel S, Colebunders R, et al. Immune reconstitution inflammatory syndrome in patients starting antiretroviral therapy for HIV infection: a systematic review and meta-analysis. Lancet Infect Dis 2010;10:251–261. 351. Munoz A, Sabin CA, Phillips AN. The incubation period of AIDS. AIDS 1997;11(Suppl A):S69–S76. 727

352. Murata H, Hruz PW, Mueckler M. Indinavir inhibits the glucose transporter isoform Glut4 at physiologic concentrations. AIDS 2002;16:859–863. 353. Murphy RL, Sommadossi JP, Lamson M, et al. Antiviral effect and pharmacokinetic interaction between nevirapine and indinavir in persons infected with HIV-1. J Infect Dis 1999;179:1116–1123. 354. Napravnik S, Edwards D, Stewart P, et al. HIV-1 drug resistance evolution among patients on potent combination antiretroviral therapy with detectable viremia. J Acquir Immune Defic Syndr 2005;40:34–40. 355. National Institutes of Health. News Releases. https://www.nih.gov/news-events/news-releases/hiv-vaccine-candidate-does-notsufficiently-protect-women-against-hiv-infection. Accessed November 9, 2021. 356. Ndhlovu ZM, Kamya P, Mewalal N, et al. Magnitude and kinetics of CD8+ T cell activation during hyperacute HIV infection impact viral set point. Immunity 2015;43:591–604. 357. Nettles RE, Kieffer TL. Update on HIV-1 viral load blips. Curr Opin HIV AIDS 2006;1:157–161. 358. Nettles RE, Kieffer TL, Kwon P, et al. Intermittent HIV-1 viremia (blips) and drug resistance in patients receiving HAART. JAMA 2005;293:817–829. 359. Neuhaus J, Angus B, Kowalska JD, et al. Risk of all-cause mortality associated with nonfatal AIDS and serious non-AIDS events among adults infected with HIV. AIDS 2010;24:697–706. 360. Nielsen C, Pedersen C, Lundgren JD, et al. Biological properties of HIV isolates in primary HIV infection: consequences for the subsequent course of infection. AIDS 1993;7:1035–1040. 361. Nishijima T, Gatanaga H, Teruya K, et al. Skin rash induced by ritonavir-boosted darunavir is common, but generally tolerable in an observational setting. J Infect Chemother 2014;20:285–287. 362. Nishimura Y, Brown CR, Mattapallil JJ, et al. Resting naive CD4+ T cells are massively infected and eliminated by X4-tropic simianhuman immunodeficiency viruses in macaques. Proc Natl Acad Sci U S A 2005;102:8000–8005. 363. Notermans DW, Pakker NG, Hamann D, et al. Immune reconstitution after 2 years of successful potent antiretroviral therapy in previously untreated human immunodeficiency virus type 1-infected adults. J Infect Dis 1999;180:1050–1056. 364. O’Connell RJ, Merritt TM, Malia JA, et al. Performance of the OraQuick rapid antibody test for diagnosis of human immunodeficiency virus type 1 infection in patients with various levels of exposure to highly active antiretroviral therapy. J Clin Microbiol 2003;41:2153–2155. 365. Orkin C, Arasteh K, Gorgolas Hernandez-Mora M, et al. Long-acting cabotegravir and rilpivirine after oral induction for HIV-1 infection. N Engl J Med 2020;382:1124–1135. 366. Orkin C, DeJesus E, Sax PE, et al. Fixed-dose combination bictegravir, emtricitabine, and tenofovir alafenamide versus dolutegravircontaining regimens for initial treatment of HIV-1 infection: week 144 results from two randomised, double-blind, multicentre, phase 3, non-inferiority trials. Lancet HIV 2020;7:e389–e400. 367. Overton ET, Richmond G, Rizzardini G, et al. Long-acting cabotegravir and rilpivirine dosed every 2 months in adults with HIV-1 infection (ATLAS-2M), 48-week results: a randomised, multicentre, open-label, phase 3b, non-inferiority study. Lancet 2021;396:1994–2005. 368. Pacheco D, Ballesteros F, Gonzalez M, et al. [Acute polyarthritis in a patient with AIDS. Clinical report]. Rev Med Chil 1989;117:910–913. 369. Pakker NG, Notermans DW, de Boer RJ, et al. Biphasic kinetics of peripheral blood T cells after triple combination therapy in HIV-1 infection: a composite of redistribution and proliferation. Nat Med 1998;4:208–214. 370. Palella FJ, Delaney KM, Moorman AC, et al. Declining morbidity and mortality among patients with advanced human immunodeficiency virus infection. HIV outpatient study investigators. N Engl J Med 1998;338:853–860. 371. Palmer S, Maldarelli F, Wiegand A, et al. Low-level viremia persists for at least 7 years in patients on suppressive antiretroviral therapy. Proc Natl Acad Sci U S A 2008;105:3879–3884. 372. Pancera M, McLellan JS, Wu X, et al. Crystal structure of PG16 and chimeric dissection with somatically related PG9: structure-function analysis of two quaternary-specific antibodies that effectively neutralize HIV-1. J Virol 2010;84:8098–8110. 373. Pandori MW, Hackett J Jr, Louie B, et al. Assessment of the ability of a fourth-generation immunoassay for human immunodeficiency virus (HIV) antibody and p24 antigen to detect both acute and recent HIV infections in a high-risk setting. J Clin Microbiol 2009;47:2639–2642. 374. Panel on Antiretroviral Guidelines for Adults and Adolescents. Guidelines for the use of antiretroviral agents in adults and adolescents with HIV. Department of Health and Human Services. http://www.aidsinfo.nih.gov/ContentFiles/AdultandAdolescentGL.pdf. Accessed May 3, 2021. 375. Panel on Treatment of Pregnant Women with HIV Infection and Prevention of Perinatal Transmission. Recommendations for use of antiretroviral drugs in pregnant HIV-1-infected women for maternal health and interventions to reduce perinatal HIV transmission in the United States. 2020:1–470. https://clinicalinfo.hiv.gov/en/guidelines/perinatal/. Accessed April 30, 2021. 376. Parker WB, White EL, Shaddix SC, et al. Mechanism of inhibition of human immunodeficiency virus type 1 reverse transcriptase and human DNA polymerases alpha, beta, and gamma by the 5′-triphosphates of carbovir, 3′-azido-3′-deoxythymidine, 2′,3′-dideoxyguanosine and 3′-deoxythymidine. A novel RNA template for the evaluation of antiretroviral drugs. J Biol Chem 1991;266:1754–1762. 377. Patel P, Borkowf CB, Brooks JT, et al. Estimating per-act HIV transmission risk: a systematic review. AIDS 2014;28:1509–1519. 378. Patick AK, Potts KE. Protease inhibitors as antiviral agents. Clin Microbiol Rev 1998;11:614–627. 379. Peeters M, Gueye A, Mboup S, et al. Geographical distribution of HIV-1 group O viruses in Africa. AIDS 1997;11:493–498. 380. Percus JK, Percus OE, Markowitz M, et al. The distribution of viral blips observed in HIV-1 infected patients treated with combination antiretroviral therapy. Bull Math Biol 2003;65:263–277. 381. Pereira GFM, Kim A, Jalil EM, et al. Dolutegravir and pregnancy outcomes in women on antiretroviral therapy in Brazil: a retrospective national cohort study. Lancet HIV 2021;8:e33–e41. 382. Perelson AS, Neumann AU, Markowitz M, et al. HIV-1 dynamics in vivo: virion clearance rate, infected cell life-span, and viral generation time. Science 1996;271:1582–1586. 383. Pereyra F, Jia X, McLaren PJ, et al. The major genetic determinants of HIV-1 control affect HLA class I peptide presentation. Science 2010;330:1551–1557. 384. Petrovas C, Casazza JP, Brenchley JM, et al. PD-1 is a regulator of virus-specific CD8+ T cell survival in HIV infection. J Exp Med 2006;203:2281–2292. 385. Phair J, Jacobson L, Detels R, et al. Acquired immune deficiency syndrome occurring within 5 years of infection with human immunodeficiency virus type-1: the Multicenter AIDS Cohort Study. J Acquir Immune Defic Syndr 1992;5:490–496. 386. Phillips AN. Reduction of HIV concentration during acute infection: independence from a specific immune response. Science 1996;271:497–499. 387. Phillips A. Short-term risk of AIDS according to current CD4 cell count and viral load in antiretroviral drug-naive individuals and those treated in the monotherapy era. AIDS 2004;18:51–58. 728

388. Phillips AN, Carr A, Neuhaus J, et al. Interruption of antiretroviral therapy and risk of cardiovascular disease in persons with HIV-1 infection: exploratory analyses from the SMART trial. Antivir Ther 2008;13:177–187. 389. Phillips AN, Staszewski S, Weber R, et al. HIV viral load response to antiretroviral therapy according to the baseline CD4 cell count and viral load. JAMA 2001;286:2560–2567. 390. Phillips EJ, Wong GA, Kaul R, et al. Clinical and immunogenetic correlates of abacavir hypersensitivity. AIDS 2005;19:979–981. 391. Pitisuttithum P, Gilbert P, Gurwith M, et al. Randomized, double-blind, placebo-controlled efficacy trial of a bivalent recombinant glycoprotein 120 HIV-1 vaccine among injection drug users in Bangkok, Thailand. J Infect Dis 2006;194:1661–1671. 392. Plantier JC, Leoz M, Dickerson JE, et al. A new human immunodeficiency virus derived from gorillas. Nat Med 2009;15:871–872. 393. Poignard P, Sabbe R, Picchio GR, et al. Neutralizing antibodies have limited effects on the control of established HIV-1 infection in vivo. Immunity 1999;10:431–438. 394. Pollard RB, Robinson P, Dransfield K. Safety profile of nevirapine, a nonnucleoside reverse transcriptase inhibitor for the treatment of human immunodeficiency virus infection. Clin Ther 1998;20:1071–1092. 395. Pontesilli O, Kerkhof-Garde S, Noterman DW, et al. Functional T cell Reconstitution and HIV-1-specific cell-mediated immunity during highly active antiretroviral therapy. J Infect Dis 1999;180:76–86. 396. Popovic M, Sarngadharan MG, Read E, et al. Detection, isolation, and continuous production of cytopathic retroviruses (HTLV-III) from patients with AIDS and pre-AIDS. Science 1984;224:497–500. 397. Quinn TC, Wawer MJ, Sewankambo N, et al. Viral load and heterosexual transmission of human immunodeficiency virus type 1. N Engl J Med 2000;342:921–928. 398. Rabezanahary H, Moukambi F, Palesch D, et al. Despite early antiretroviral therapy effector memory and follicular helper CD4 T cells are major reservoirs in visceral lymphoid tissues of SIV-infected macaques. Mucosal Immunol 2020;13:149–160. 399. Raffi F, Rachlis A, Stellbrink HJ, et al. Once-daily dolutegravir versus raltegravir in antiretroviral-naive adults with HIV-1 infection: 48 week results from the randomised, double-blind, non-inferiority SPRING-2 study. Lancet 2013;381:735–743. 400. Ramduth D, Chetty P, Mngquandaniso NC, et al. Differential immunogenicity of HIV-1 clade C proteins in eliciting CD8+ and CD4+ cell responses. J Infect Dis 2005;192:1588–1596. 401. Ramsuran V, Naranbhai V, Horowitz A, et al. Elevated HLA-A expression impairs HIV control through inhibition of NKG2A-expressing cells. Science 2018;359:86–90. 402. Ranasinghe S, Flanders M, Cutler S, et al. HIV-specific CD4 T cell responses to different viral proteins have discordant associations with viral load and clinical outcome. J Virol 2012;86:277–283. 403. Ratnam I, Chiu C, Kandala NB, et al. Incidence and risk factors for immune reconstitution inflammatory syndrome in an ethnically diverse HIV type 1-infected cohort. Clin Infect Dis 2006;42:418–427. 404. Ren J, Nichols C, Bird L, et al. Structural mechanisms of drug resistance for mutations at codons 181 and 188 in HIV-1 reverse transcriptase and the improved resilience of second generation non-nucleoside inhibitors. J Mol Biol 2001;312:795–805. 405. Rerks-Ngarm S, Pitisuttithum P, Nitayaphan S, et al. Vaccination with ALVAC and AIDSVAX to prevent HIV-1 infection in Thailand. N Engl J Med 2009;361:2209–2220. 406. Reynolds SJ, Makumbi F, Nakigozi G, et al. HIV-1 transmission among HIV-1 discordant couples before and after the introduction of antiretroviral therapy. AIDS 2011;25:473–477. 407. Ribaudo HJ, Benson CA, Zheng Y, et al. No risk of myocardial infarction associated with initial antiretroviral treatment containing abacavir: short and long-term results from ACTG A5001/ALLRT. Clin Infect Dis 2011;52:929–940. 408. Ribaudo HJ, Daar ES, Tierney C, et al. Impact of UGT1A1 Gilbert variant on discontinuation of ritonavir-boosted atazanavir in AIDS Clinical Trials Group Study A5202. J Infect Dis 2013;207:420–425. 409. Richman DD, Bozzette SA. The impact of the syncytium-inducing phenotype of human immunodeficiency virus on disease progression. J Infect Dis 1994;169:968–974. 410. Richman DD, Fischl MA, Grieco MH, et al. The toxicitiy of azidothymidine (AZT) in the treatment of patients with AIDS and AIDS-related complex. A double-blind, placebo-controlled trial. N Engl J Med 1987;317:192–197. 411. Richman DD, Wrin T, Little SJ, et al. Rapid evolution of the neutralizing antibody response to HIV type 1 infection. Proc Natl Acad Sci U S A 2003;100:4144–4149. 412. Robbins GK, Addo MM, Troung H, et al. Augmentation of HIV-1-specific T helper cell responses in chronic HIV-1 infection by therapeutic immunization. AIDS 2003;17:1121–1126. 413. Rockstroh JK. Influence of viral hepatitis on HIV infection. J Hepatol 2006;44(1 Suppl):S25–S27. 414. Rockwood N, Mandalia S, Bower M, et al. Ritonavir-boosted atazanavir exposure is associated with an increased rate of renal stones compared with efavirenz, ritonavir-boosted lopinavir and ritonavir-boosted darunavir. AIDS 2011;25:1671–1673. 415. Rodger AJ, Cambiano V, Bruun T, et al. Sexual activity without condoms and risk of HIV transmission in serodifferent couples when the HIV-positive partner is using suppressive antiretroviral therapy. JAMA 2016;316:171–181. 416. Rodger AJ, Fox Z, Lundgren JD, et al. Activation and coagulation biomarkers are independent predictors of the development of opportunistic disease in patients with HIV infection. J Infect Dis 2009;200:973–983. 417. Rodriguez B, Sethi AK, Cheruvu VK, et al. Predictive value of plasma HIV RNA level on rate of CD4 T-cell decline in untreated HIV infection. JAMA 2006;296:1498–1506. 418. Rosenberg E, Altfeld M, Poon SH, et al. Immune control of HIV-1 after early treatment of acute infection. Nature 2000;407:523–526. 419. Rosenberg ES, Graham BS, Chan ES, et al. Safety and immunogenicity of therapeutic DNA vaccination in individuals treated with antiretroviral therapy during acute/early HIV-1 infection. PLoS One 2010;5:e10555. 420. Ross AC, Rizk N, O’Riordan MA, et al. Relationship between inflammatory markers, endothelial activation markers, and carotid intimamedia thickness in HIV-infected patients receiving antiretroviral therapy. Clin Infect Dis 2009;49:1119–1127. 421. Royce RA, Sena A, Cates W, et al. Sexual transmission of HIV. N Engl J Med 1997;336:1072–1078. 422. Ryan ES, Micci L, Fromentin R, et al. Loss of function of intestinal IL-17 and IL-22 producing cells contributes to inflammation and viral persistence in SIV-infected rhesus macaques. PLoS Pathog 2016;12:e1005412. 423. Saag MS, Gandhi RT, Hoy JF, et al. Antiretroviral drugs for treatment and prevention of HIV infection in adults: 2020 recommendations of the international antiviral society-USA panel. JAMA 2020;324:1651–1669. 424. Sabin CA, Telfer P, Phillips AN, et al. The association between hepatitis C virus genotype and human immunodeficiency virus disease progression in a cohort of hemophilic men. J Infect Dis 1997;175:164–168. 425. Sabin CA, Worm SW, Weber R, et al. Use of nucleoside reverse transcriptase inhibitors and risk of myocardial infarction in HIV-infected patients enrolled in the D:A:D study: a multi-cohort collaboration. Lancet 2008;371:1417–1426. 426. Salemi M, Strimmer K, Hall WW, et al. Dating the common ancestor of SIVcpz and HIV-1 group M and the origin of HIV-1 subtypes using a new method to uncover clock-like molecular evolution. FASEB J 2001;15:276–278. 427. Salie ZL, Kirby KA, Michailidis E, et al. Structural basis of HIV inhibition by translocation-defective RT inhibitor 4′-ethynyl-2-fluoro2′-deoxyadenosine (EFdA). Proc Natl Acad Sci U S A 2016;113:9274–9279. 428. Samson M, Libert F, Doranz BJ, et al. Resistance to HIV-1 infection in caucasian individuals bearing mutant alleles of th eCCR-5 729

chemokine receptor gene. Nature 1996;382:722–725. 429. Sanne I, Mommeja-Marin H, Hinkle J, et al. Severe hepatotoxicity associated with nevirapine use in HIV-infected subjects. J Infect Dis 2005;191:825–829. 430. Santiago ML, Range F, Keele BF, et al. Simian immunodeficiency virus infection in free-ranging sooty mangabeys (Cercocebus atys atys) from the Tai Forest, Cote d’Ivoire: implications for the origin of epidemic human immunodeficiency virus type 2. J Virol 2005;79:12515–12527. 431. Sarngadharan MG, Popovic M, Bruch L, et al. Antibodies reactive with human T-lymphotropic retroviruses (HTLV-III) in the serum of patients with AIDS. Science 1984;224:506–508. 432. Sauter D, Kirchhoff F. Key viral adaptations preceding the AIDS pandemic. Cell Host Microbe 2019;25:27–38. 433. Sax PE, Erlandson KM, Lake JE, et al. Weight gain following initiation of antiretroviral therapy: risk factors in randomized comparative clinical trials. Clin Infect Dis 2020;71:1379–1389. 434. Sax PE, Pozniak A, Montes ML, et al. Coformulated bictegravir, emtricitabine, and tenofovir alafenamide versus dolutegravir with emtricitabine and tenofovir alafenamide, for initial treatment of HIV-1 infection (GS-US-380-1490): a randomised, double-blind, multicentre, phase 3, non-inferiority trial. Lancet 2017;390:2073–2082. 435. Schmitt N, Nugeyre MT, Scott-Algara D, et al. Differential susceptibility of human thymic dendritic cell subsets to X4 and R5 HIV-1 infection. AIDS 2006;20:533–542. 436. Schmitz JE, Kuroda MJ, Santra S, et al. Control of viremia in simian immunodeficiency virus infection by CD8+ lymphocytes. Science 1999;283:857–860. 437. Schooley RT, Ramirez-Ronda C, Lange JMA, et al. Virologic and immunologic benefits of initial combination therapy with zidovudine and zalcitabine or didanosine compared to zidovudine monotherapy. J Infect Dis 1996;173:1354–1366. 438. Schreiber GB, Busch MP, Kleinman SH, et al. The risk of transfusion-transmitted viral infections. The Retrovirus Epidemiology Donor Study. N Engl J Med 1996;334:1685–1690. 439. Schuitemaker H, Koot M, Kootstra NA, et al. Biological phenotype of human immunodeficiency virus type 1 clones at different stages of infection: progression of disease is associated with a shift from monocytotropic to T-cell-tropic virus populations. J Virol 1992;66:1354–1360. 440. Schuurman R, Nijhuis M, van Leeuwen R, et al. Rapid changes in human immunodeficiency virus type 1 RNA load and appearance of drug-resistant virus populations in persons treated with lamivudine (3TC). J Infect Dis 1995;171:1411–1419. 441. Schwartz O, Marechal V, Le Gall S, et al. Endocytosis of major histocompatibility complex class I molecules is induced by the HIV-1 Nef protein. Nat Med 1996;2:338–342. 442. Segal-Maurer S, Castagna A, Berhe M, et al. Potent antiviral activity of lenacapavir in phase 2/3 in heavily ART-experienced PWH. Virtual CROI 2021. (held virtually): International Antiviral Society-USA; 2021 [Abstract 127]. 443. Shapiro RL, Hughes MD, Ogwu A, et al. Antiretroviral regimens in pregnancy and breast-feeding in Botswana. N Engl J Med 2010;362:2282–2294. 444. Sharp PM, Hahn BH. Origins of HIV and the AIDS pandemic. Cold Spring Harb Perspect Med 2011;1:a006841. 445. Sharp PM, Shaw GM, Hahn BH. Simian immunodeficiency virus infection of chimpanzees. J Virol 2005;79:3891–3902. 446. Shelburne SA III, Darcourt J, White AC Jr, et al. The role of immune reconstitution inflammatory syndrome in AIDS-related Cryptococcus neoformans disease in the era of highly active antiretroviral therapy. Clin Infect Dis 2005;40:1049–1052. 447. Shelburne SA, Visnegarwala F, Darcourt J, et al. Incidence and risk factors for immune reconstitution inflammatory syndrome during highly active antiretroviral therapy. AIDS 2005;19:399–406. 448. Shiels MS, Pfeiffer RM, Engels EA. Age at cancer diagnosis among persons with AIDS in the United States. Ann Intern Med 2010;153:452–460. 449. Shiels MS, Pfeiffer RM, Hall HI, et al. Proportions of Kaposi sarcoma, selected non-Hodgkin lymphomas, and cervical cancer in the United States occurring in persons with AIDS, 1980-2007. JAMA 2011;305:1450–1459. 450. Siliciano JD, Kajdas J, Finzi D, et al. Long-term follow-up studies confirm the stability of the latent reservoir for HIV-1 in resting CD4+ T cells. Nat Med 2003;9:727–728. 451. Simek MD, Rida W, Priddy FH, et al. Human immunodeficiency virus type 1 elite neutralizers: individuals with broad and potent neutralizing activity identified by using a high-throughput neutralization assay together with an analytical selection algorithm. J Virol 2009;83:7337–7348. 452. Simmons RP, Scully EP, Groden EE, et al. HIV-1 infection induces strong production of IP-10 through TLR7/9-dependent pathways. AIDS 2013;27:2505–2517. 453. Simon F, Mauclere P, Roques P, et al. Identification of a new human immunodeficiency virus type 1 distinct from group M and group O. Nat Med 1998;4:1032–1037. 454. Simonetti FR, White JA, Tumiotto C, et al. Intact proviral DNA assay analysis of large cohorts of people with HIV provides a benchmark for the frequency and composition of persistent proviral DNA. Proc Natl Acad Sci U S A 2020;117:18692–18700. 455. Simonetti FR, Zhang H, Soroosh GP, et al. Antigen-driven clonal selection shapes the persistence of HIV-1-infected CD4+ T cells in vivo. J Clin Invest 2021;131:e145254. 456. Simpson DM, Tagliati M. Neurologic manifestations of HIV infection. Ann Intern Med 1994;121:769–785. 457. Sirivichayakul S, Phanuphak P, Tanprasert S, et al. Evaluation of a 2-minute anti-human immunodeficiency virus (HIV) test using the autologous erythrocyte agglutination technique with populations differing in HIV prevalence. J Clin Microbiol 1993;31:1373–1375. 458. Smith K, Aga E, Bosch RJ, et al. Long-term changes in circulating CD4 T lymphocytes in virologically suppressed patients after 6 years of highly active antiretroviral therapy. AIDS 2004;18:1953–1956. 459. Smith MW, Dean M, Carrington M, et al. Contrasting genetic influence of CCR2 and CCR5 variants on HIV-1 infection and disease progression. Science 1997;277:959–965. 460. Smith PF, Ogundele A, Forrest A, et al. Phase I and II study of the safety, virologic effect, and pharmacokinetics/pharmacodynamics of single-dose 3-o-(3′,3′-dimethylsuccinyl)betulinic acid (bevirimat) against human immunodeficiency virus infection. Antimicrob Agents Chemother 2007;51:3574–3581. 461. So YT, Holtzman DM, Abrams DI, et al. Peripheral neuropathy associated with acquired immunodeficiency syndrome. Prevalence and clinical features from a population-based survey. Arch Neurol 1988;45:945–948. 462. Soghoian DZ, Jessen H, Flanders M, et al. HIV-specific cytolytic CD4 T cell responses during acute HIV infection predict disease outcome. Sci Transl Med 2012;4:123ra25. 463. Solomon SS, Mehta SH, McFall AM, et al. Community viral load, antiretroviral therapy coverage, and HIV incidence in India: a crosssectional, comparative study. Lancet HIV 2016;3:e183–e190. 464. Spector SA, Wong R, Hsia K, et al. Plasma cytomegalovirus (CMV) DNA load predicts CMV disease and survival in AIDS patients. J Clin Invest 1998;101:497–502. 465. Spence RA, Kati WM, Anderson KS, et al. Mechanism of inhibition of HIV-1 reverse transcriptase nonnucleoside inhibitors. Science 1995;267:988–993. 730

466. Sperling RS, Shapiro DE, Coombs RW, et al. Maternal viral load, zidovudine treatment, and the risk of transmission of human immunodeficiency virus type 1 from mother to infant. N Engl J Med 1996;335:1621–1629. 467. Spinner C, Felizarta FB, Rizzardini G, et al. Phase IIA proof-of-concept trial of next-generation maturation inhibitor GSK3630254. Virtual CROI 2021. (held virtually): International Antiviral Society-USA; 2021 [Abstract 126]. 468. Stamatatos L, Morris L, Burton DR, et al. Neutralizing antibodies generated during natural HIV-1 infection: good news for an HIV-1 vaccine? Nat Med. 2009;15:866–870. 469. Stellbrink HJ, Orkin C, Arribas JR, et al. Comparison of changes in bone density and turnover with abacavir-lamivudine versus tenofoviremtricitabine in HIV-infected adults: 48-week results from the ASSERT study. Clin Infect Dis 2010;51:963–972. 470. Stephenson KE, Wagh K, Korber B, et al. Vaccines and broadly neutralizing antibodies for HIV-1 prevention. Annu Rev Immunol 2020;38:673–703. 471. Sterling TR, Vlahov D, Astemborski J, et al. Initial plasma HIV-1 RNA levels and progression to AIDS in women and men. N Engl J Med 2001;344:720–725. 472. Stevens WS, Noble L, Berrie L, et al. Ultra-high-throughput, automated nucleic acid detection of human immunodeficiency virus (HIV) for infant infection diagnosis using the Gen-Probe Aptima HIV-1 screening assay. J Clin Microbiol 2009;47:2465–2469. 473. Sulkowski MS, Mehta SH, Chaisson RE, et al. Hepatotoxicity associated with protease inhibitor-based antiretroviral regimens with or without concurrent ritonavir. AIDS 2004;18:2277–2284. 474. Sullivan PS, Hanson DL, Teshale EH, et al. Effect of hepatitis C infection on progression of HIV disease and early response to initial antiretroviral therapy. AIDS 2006;20:1171–1179. 475. Swindells S, Andrade-Villanueva JF, Richmond GJ, et al. Long-acting cabotegravir and rilpivirine for maintenance of HIV-1 suppression. N Engl J Med 2020;382:1112–1123. 476. Szczech LA, Gupta SK, Habash R, et al. The clinical epidemiology and course of the spectrum of renal diseases associated with HIV infection. Kidney Int 2004;66:1145–1152. 477. Takata H, Buranapraditkun S, Kessing C, et al. Delayed differentiation of potent effector CD8(+) T cells reducing viremia and reservoir seeding in acute HIV infection. Sci Transl Med 2017;9:eaag1809. 478. Tanser F, de Oliveira T, Maheu-Giroux M, et al. Concentrated HIV subepidemics in generalized epidemic settings. Curr Opin HIV AIDS 2014;9:115–125. 479. TEMPRANO ANRS Study Group. A trial of early antiretrovirals and isoniazid preventive therapy in Africa. N Engl J Med 2015;373:808–822. 480. Tenorio A, Smith KY, Kuritzkes DR, et al. HIV-1-infected antiretroviral-treated patients with prolonged partial viral suppression: clinical, virologic, and immunologic course. J Acquir Immune Defic Syndr 2003;34:491–496. 481. Tenorio AR, Zheng Y, Bosch RJ, et al. Soluble markers of inflammation and coagulation but not T-cell activation predict non-AIDSdefining morbid events during suppressive antiretroviral treatment. J Infect Dis 2014;210:1248–1259. 482. Thein HH, Yi Q, Dore GJ, et al. Natural history of hepatitis C virus infection in HIV-infected individuals and the impact of HIV in the era of highly active antiretroviral therapy: a meta-analysis. AIDS 2008;22:1979–1991. 483. Tindall B, Barker S, Donovan B, et al. Characterization of the acute clinical illness associated with human immunodeficiency virus infection. Arch Intern Med 1988;148:945–949. 484. Trautmann L, Janbazian L, Chomont N, et al. Upregulation of PD-1 expression on HIV-specific CD8+ T cells leads to reversible immune dysfunction. Nat Med 2006;12:1198–1202. 485. Triant VA, Brown TT, Lee H, et al. Fracture prevalence among human immunodeficiency virus (HIV)-infected versus non-HIV-infected patients in a large U.S. healthcare system. J Clin Endocrinol Metab 2008;93:3499–3504. 486. Triant VA, Lee H, Hadigan C, et al. Increased acute myocardial infarction rates and cardiovascular risk factors among patients with human immunodeficiency virus disease. J Clin Endocrinol Metab 2007;92:2506–2512. 487. Tsai CC, Follis KE, Sabo A, et al. Prevention of SIV infection in macaques by (R)-9-(2-phosphonylmethoxypropyl)adenine. Science 1995;270:1197–1199. 488. Tural C, Romeu J, Sirera G, et al. Long-lasting remission of cytomegalovirus retinitis without maintenance therapy in human immunodeficiency virus-infected patients. J Infect Dis 1998;177:1080–1083. 489. Van Damme L, Corneli A, Ahmed K, et al. Preexposure prophylaxis for HIV infection among African women. N Engl J Med 2012;367:411–422. 490. van Rij RP, de Roda Husman AM, Brouwer M, et al. Role of CCR2 genotype in the clinical course of syncytium-inducing (SI) or non-SI HIV-1 infection and in the time to conversion to SI virus variants. J Infect Dis 1998;178:1806–1811. 491. van Sighem A, Gras L, Reiss P. Life expectancy of recently diagnosed asymptomatic HIV-infected patients approaches that of uninfected individuals. AIDS 2010;24:1527–1535. 492. Van de Perre P, Nzaramba D, Allen S, et al. Comparison of six serological assays for human immunodeficiency virus antibody detection in developing countries. J Clin Microbiol 1988;26:552–556. 493. Veazey RS. Intestinal CD4 depletion in HIV/SIV infection. Curr Immunol Rev 2019;15:76–91. 494. Veazey RS, DeMaria M, Chalifoux LV, et al. Gastrointestinal tract as a major site of CD4+ T cell depletion and viral replication in SIV infection. Science 1998;280:427–431. 495. Venter WDF, Moorhouse M, Sokhela S, et al. Dolutegravir plus two different prodrugs of tenofovir to treat HIV. N Engl J Med 2019;381:803–815. 496. Venter WDF, Sokhela S, Simmons B, et al. Dolutegravir with emtricitabine and tenofovir alafenamide or tenofovir disoproxil fumarate versus efavirenz, emtricitabine, and tenofovir disoproxil fumarate for initial treatment of HIV-1 infection (ADVANCE): week 96 results from a randomised, phase 3, non-inferiority trial. Lancet HIV 2020;7:e666–e676. 497. Verhoeven D, Sankaran S, Silvey M, et al. Antiviral therapy during primary simian immunodeficiency virus infection fails to prevent acute loss of CD4+ T cells in gut mucosa but enhances their rapid restoration through central memory T cells. J Virol 2008;82:4016–4027. 498. Vilches C, Parham P. KIR: diverse, rapidly evolving receptors of innate and adaptive immunity. Annu Rev Immunol 2002;20:217–251. 499. Walensky RP, Arbelaez C, Reichmann WM, et al. Revising expectations from rapid HIV tests in the emergency department. Ann Intern Med 2008;149:153–160. 500. Walker LM, Phogat SK, Chan-Hui PY, et al. Broad and potent neutralizing antibodies from an African donor reveal a new HIV-1 vaccine target. Science 2009;326:285–289. 501. Walker LM, Simek MD, Priddy F, et al. A limited number of antibody specificities mediate broad and potent serum neutralization in selected HIV-1 infected individuals. PLoS Pathog 2010;6:e1001028. 502. Walmsley SL, Antela A, Clumeck N, et al. Dolutegravir plus abacavir-lamivudine for the treatment of HIV-1 infection. N Engl J Med 2013;369:1807–1818. 503. Walmsley S, Bernstein B, King M, et al. Lopinavir-Ritonavir versus nelfinavir for the initial treatment of HIV infection. N Engl J Med 2002;346:2039–2046. 504. Wei X, Decker JM, Wang S, et al. Antibody neutralization and escape by HIV-1. Nature 2003;422:307–312. 731

505. Wei X, Ghosh SK, Taylor ME, et al. Viral dynamics in human immunodeficiency virus type 1 infection. Nature 1995;373:117–122. 506. Wensing AM, Calvez V, Ceccherini-Silberstein F, et al. 2019 update of the drug resistance mutations in HIV-1. Top Antivir Med 2019;27:111–121. 507. Wherry EJ, Blattman JN, Murali-Krishna K, et al. Viral persistence alters CD8 T-cell immunodominance and tissue distribution and results in distinct stages of functional impairment. J Virol 2003;77:4911–4927. 508. Wherry EJ, Teichgraber V, Becker TC, et al. Lineage relationship and protective immunity of memory CD8 T cell subsets. Nat Immunol 2003;4:225–234. 509. Whitney JB, Hill AL, Sanisetty S, et al. Rapid seeding of the viral reservoir prior to SIV viraemia in rhesus monkeys. Nature 2014;512:74–77. 510. Whitney JB, Lim SY, Osuna CE, et al. Prevention of SIVmac251 reservoir seeding in rhesus monkeys by early antiretroviral therapy. Nat Commun 2018;9:5429. 511. Wild C, Greenwell T, Shugars D, et al. The inhibitory activity of an HIV type 1 peptide correlates with its ability to interact with a leucine zipper structure. AIDS Res Hum Retroviruses 1995;11:323–325. 512. Wild CT, Shugars DC, Greenwell TK, et al. Peptides corresponding to a predictive alpha-helical domain of human immunodeficiency virus type 1 gp41 are potent inhibitors of virus infection. Proc Natl Acad Sci U S A 1994;91:9770–9774. 513. Winkler C, Modi W, Smith MW, et al. Genetic restriction of AIDS pathogenesis by an SDF-1 chemokine gene variant. ALIVE Study, Hemophilia Growth and Development Study (HGDS), Multicenter AIDS Cohort Study (MACS), Multicenter Hemophilia Cohort Study (MHCS), San Francisco City Cohort (SFCC). Science 1998;279:389–393. 514. Winston JA, Bruggeman LA, Ross MD, et al. Nephropathy and establishment of a renal reservoir of HIV type 1 during primary infection. N Engl J Med 2001;344:1979–1984. 515. Winston A, De Francesco D, Post F, et al. Comorbidity indices in people with HIV and considerations for coronavirus disease 2019 outcomes. AIDS 2020;34:1795–1800. 516. Wong JK, Hezareh M, Gunthard HF, et al. Recovery of replication-competent HIV despite prolonged suppression of plasma viremia. Science 1997;278:1291–1295. 517. Wood E, Hogg RS, Yip B, et al. Effect of medication adherence on survival of HIV-infected adults who start highly active antiretroviral therapy when the CD4+ cell count is 0.200 to 0.350 x 109 cells/L. Ann Intern Med 2003;139:810–816. 518. World Health Organization. WHO Case Definitions of HIV for Surveillance and Revised Clinical Staging and Immunological Classification of HIV-Related Disease in Adults and Children. Geneva: WHO Press; 2007. 519. World Health Organization. Programmatic Update: Antiretroviral Treatment as Prevention (TASP) of HIV and TB. Geneva: World Health Organization; 2012. 520. World Health Organization. Update of Recommendations on First- and Second-line Antrietroviral Regimens. Geneva, Switzerland: World Health Organization; 2019:1–16. 521. Worm SW, Sabin C, Weber R, et al. Risk of myocardial infarction in patients with HIV infection exposed to specific individual antiretroviral drugs from the 3 major drug classes: the data collection on adverse events of anti-HIV drugs (D:A:D) study. J Infect Dis 2010;201:318–330. 522. Worobey M, Gemmel M, Teuwen DE, et al. Direct evidence of extensive diversity of HIV-1 in Kinshasa by 1960. Nature 2008;455:661–664. 523. Wu X, Yang ZY, Li Y, et al. Rational design of envelope identifies broadly neutralizing human monoclonal antibodies to HIV-1. Science 2010;329:856–861. 524. Xia H, Jiang W, Zhang X, et al. Elevated level of CD4(+) T Cell immune activation in acutely HIV-1-infected stage associates with increased IL-2 production and cycling expression, and subsequent CD4(+) T Cell preservation. Front Immunol 2018;9:616. 525. Xu H, Wang X, Liu DX, et al. IL-17-producing innate lymphoid cells are restricted to mucosal tissues and are depleted in SIV-infected macaques. Mucosal Immunol 2012;5:658–669. 526. Yamamoto T, Price DA, Casazza JP, et al. Surface expression patterns of negative regulatory molecules identify determinants of virusspecific CD8+ T-cell exhaustion in HIV infection. Blood 2011;117:4805–4815. 527. Youngblood B, Noto A, Porichis F, et al. Cutting edge: prolonged exposure to HIV reinforces a poised epigenetic program for PD-1 expression in virus-specific CD8 T cells. J Immunol 2013;191:540–544. 528. Yue FY, Merchant A, Kovacs CM, et al. Virus-specific interleukin-17-producing CD4+ T cells are detectable in early human immunodeficiency virus type 1 infection. J Virol 2008;82:6767–6771. 529. Yue L, Prentice HA, Farmer P, et al. Cumulative impact of host and viral factors on HIV-1 viral-load control during early infection. J Virol 2013;87:708–715. 530. Yusim K, Peeters M, Pybus OG, et al. Using human immunodeficiency virus type 1 sequences to infer historical features of the acquired immune deficiency syndrome epidemic and human immunodeficiency virus evolution. Philos Trans R Soc Lond B Biol Sci 2001;356:855–866. 531. Zash R, Holmes L, Diseko M, et al. Update on neural tube defects with antiretroviral exposure in the Tsepamo study, Botswana. 23rd International AIDS Conference (conducted virtually), 6-10 July, 2020. International AIDS Society. Abstract OAXLB0102. 532. Zash R, Makhema J, Shapiro RL. Neural-tube defects with dolutegravir treatment from the time of conception. N Engl J Med 2018;379:979–981. 533. Zhang ZQ, Notermans DW, Sedgewick G, et al. Kinetics of CD4+ T cell repopulation of lymphoid tissues after treatment of HIV-1 infection. Proc Natl Acad Sci U S A 1998;95:1154–1159. 534. Zhang ZQ, Schuler T, Cavert W, et al. Reversibility of the pathological changes in the follicular dendritic cell network with treatment of HIV-1 infection. Proc Natl Acad Sci U S A 1999;96:5169–5172. 535. Zuniga R, Lucchetti A, Galvan P, et al. Relative dominance of Gag p24-specific cytotoxic T lymphocytes is associated with human immunodeficiency virus control. J Virol 2006;80:3122–3125.

732

CHAPTER 19 Nonhuman Lentiviruses Ronald C. Desrosiers • David T. Evans History Infectious agent Genome organization and composition Propagation Propagation and cell culture Host range Restriction Receptor use Germline integration Pathogenesis and pathology Portals of entry Cell and tissue tropism Immune responses and persistence Virulence Clinical and pathologic features Contributions of individual genes and genetic elements Genetic resistance Diagnosis Prevention and control Research on vaccine development Research on therapeutic regimens Perspective

HISTORY Use of the term “slow virus infections” and identification of the first lentivirus is generally credited to Sigurdsson et al.321–323 Twenty karakul sheep imported into Iceland from Germany in 1933 resulted in the transmission of a chronic disease that led to the death of more than 100,000 sheep in the decades that followed. Sigurdsson et al. not only described the disease but also demonstrated that it was due to a transmissible agent321–323 and used the term “slow virus infections” to refer to this disease.320 In 1960, Sigurdsson et al. described the cultivation of the transmissible agent in tissue culture,324 and Gudnadottir and Palsson were able to reproduce the disease with the culture-grown virus.126 The diseases in the sheep were called maedi (Icelandic for dyspnea; i.e., a lung disease resulting in difficulty breathing) and visna (Icelandic for a state of progressive apathy, a “fading away” resulting from brain disease). Both disease states result from the same virus, now referred to as maedi/visna virus (MVV). MVV and related viruses are called lentiviruses, which is derived from the Latin lentus for slow. Approximately 600,000 sheep were slaughtered in Iceland in 1965 to eradicate MVV from the island.

The maedi/visna disease in Icelandic sheep, although the first specifically shown to be caused by a lentivirus, is probably not, however, the first description of a lentiviral disease. Vallée and Carré described in 1904 the infectious nature of a chronic disease in horses,354 which is now known to be caused by the lentivirus equine infectious anemia virus (EIAV). A chronic, progressive interstitial pneumonia had also been described in South African sheep in 1915 and in Montana sheep in 1923 before the identification of MVV.222 Work with EIAV was largely on a parallel track with that of MVV, and the first description of EIAV cultivation appeared in 1961.180 Only subsequently were EIAV and MVV shown to belong to the lentivirus genus on the basis of morphologic criteria.118,258,375

The scenario of disease outbreak leading to the identification of a new lentivirus has repeated itself dramatically on several occasions in more recent history. In 1964, an emerging infectious disease was first detected in Bali cattle in the Jembrana district of Bali.282,328 Bali cattle are the domesticated form of the wild banteng (Bos javanicus). Within 12 months, 26,000 of the 300,000 cattle on the island died of the disease. The cause of the disease was subsequently traced to a bovine lentivirus, now called Jembrana disease virus (JDV), which is a distinct variant of 733

bovine immunodeficiency virus (BIV).43,170 Bali cattle are particularly sensitive to disease caused by this virus.329,330,380 Outbreaks of immunodeficiency disease and lymphoma in captive colonies of macaques were subsequently traced to the introduction of a simian lentivirus from African monkeys.23,68,214,219 Simian immunodeficiency viruses (SIVs) naturally infect African apes and Old World monkeys but are not found in macaques or other Asian nonhuman primate species. The origins of the human immunodeficiency viruses have followed a similar pattern. HIV-2 in western Africa quite clearly originated from SIVsmm in sooty mangabey monkeys.49,79,104,142,197,225 Sooty mangabeys are naturally infected with SIVsmm at high frequency, and the habitat of sooty mangabeys coincides with the same geographic region of western Africa where HIV-2 is most prevalent. SIVsmm also groups phylogenetically with HIV-2. The origins of HIV-1 have similarly been traced to SIVs of great apes in West Central Africa.313

The earliest descriptions of the isolation of HIV and its association with acquired immunodeficiency syndrome (AIDS) did not appreciate that the virus was a lentivirus.18,275 Only subsequently, through more careful examination of electron micrographs and sequence analysis, did this become clear.119,235 At the time, study of lentiviruses was an obscure discipline with which many scientists were not familiar.

From a historical perspective, MVV and EIAV were discovered, isolated, and characterized long before the discovery of HIV.180,324 Lentiviruses identified after the discovery of HIV-1 have used a nomenclature similar to that for HIV-1 (i.e., “immunodeficiency virus”). BIV, originally isolated from a cow with a chronic disease by Van Der Maaten et al. in 1972,356 received little attention until after the discovery of HIV. Subsequent to the discovery of HIV-1, lentiviruses were isolated from monkeys and cats (Table 19.1). Although the discovery of MVV, EIAV, and BIV predates that of HIV-1, HIV-1 has received such intense scrutiny that much more is known about it than any of the other lentiviruses. New information about the nonhuman lentiviruses is thus usually compared with what is known for HIV-1.

INFECTIOUS AGENT Lentiviruses have been isolated from sheep, goats, horses, cattle, cats, monkeys, and humans (Table 19.1). Genetic analysis of a lentivirus from goats, caprine arthritis–encephalitis virus (CAEV) revealed that it clusters closely with MVV353 and belongs in a single group with MVV. Therefore, based on host species and viral genetics, five distinct groups of lentiviruses are recognized (Fig. 19.1). It is important to note that even within a single group, discrete subgroups can be defined based on host species, geography, and genetic distance. For example, among the nonhuman primate lentiviruses, different subgroups exist for the SIVs from African green monkeys, sooty mangabeys, Sykes’ monkeys, and L’Hoest’s monkeys (Fig. 19.2 and Table 19.2).

FIGURE 19.1 Phylogeny of the lentiviruses. Five distinct phylogenetic groups of lentiviruses. The unrooted tree depicts the phylogenetic relationships among the five recognized groups of lentiviruses. The tree is based on an amino acid alignment of 470 residues of reverse transcriptase from representative members of each group, including bovine immunodeficiency virus (BIV), equine infectious anemia virus (EIAV), feline immunodeficiency virus (FIV), maedi/visna virus, caprine arthritis–encephalitis virus (MVV/CAEV), and the simian and human immunodeficiency viruses (SIV/HIV). Maximum parsimony (shown) and neighbor-joining (not shown) analyses give trees of nearly identical topology. Branch lengths are proportional to the number of amino acid substitutions. TABLE 19.1 Known lentiviruses Species

Virus

Year That Cultivation Was First Published

Sheep/goats

Maedi/visna virus; caprine arthritis encephalitis 1960324 virus

Horses

Equine infectious anemia virus

734

1961180

Cattle

Bovine immunodeficiency virus; Jembrana disease virus

1972356

Humans

Human immunodeficiency virus

198318

Monkeys

Simian immunodeficiency virus

1985356

Cats

Feline immunodeficiency virus

1987260

FIGURE 19.2 Phylogeny of primate lentiviruses. The tree depicts the phylogenetic relationships among the human and simian immunodeficiency viruses and is based on neighbor-joining analysis of a 294 bp sequence in pol. Branch lengths that are drawn to scale indicate the number of nucleotide substitutions per site. (Courtesy of Martine Peeters. Redrawn from Peeters M, Ma D, Liegeois F, et al. Simian immunodeficiency virus infections in the wild. In: Ansari AA, Silvestri G, eds. Natural Hosts of SIV: Implication in AIDS. Amsterdam, The Netherlands: Elsevier; 2014:37–67. Copyright © 2014 Elsevier. With permission.) All lentiviruses have a common morphology that distinguishes them from other retroviruses (see Chapter 17). Lentiviruses bud from the plasma membrane without a preformed nucleoid, and mature particles typically have a conical or rod-shaped nucleoid core (Fig. 19.3). Classification into the lentivirus subgroup by morphologic criteria alone is entirely consistent with phylogenetic analysis of pol gene sequences. Viruses classified as lentiviruses have pol gene sequences more closely related to one another than to other retroviruses, and all have the same characteristic morphology. All lentiviruses have a propensity to replicate in macrophages, all exhibit long-term persistent viral replication in their natural host, and all are associated with a chronic progressive disease course in susceptible hosts. The primate lentiviruses have acquired use of CD4 as one of two receptors used sequentially by virus for entry into cells (Table 19.3). FIV has also been shown to use a similar two receptor mechanism for entry. However, nonprimate lentiviruses, including FIV, do not use CD4 as an entry receptor. The chronic disease induced by primate lentiviruses has an immunodeficiency component because of the infection of CD4+ lymphocytes. All lentiviruses have a number of auxiliary (or accessory) genes in addition to the gag, pol, and env genes found in all retroviruses.

GENOME ORGANIZATION AND COMPOSITION Widespread use of DNA sequencing has greatly enhanced our understanding of the phylogenetic relationships and the gene products of lentiviruses. Based on sequence analysis, five discrete groups of lentiviruses are now recognized (Fig. 19.1). Extensive diversity exists even within a group. For example, among the SIVs, distinct subgroups have been defined from the African green monkey, sooty mangabey monkey, L’Hoest’s monkey, Sykes’ monkey, and other genera (Table 19.2 and Fig. 19.2). Of the 69 nonhuman primate species known to inhabit subSaharan Africa, SIV infection has been demonstrated in at least 45 of them, and partial or complete viral sequence information is available for 39 of them.262 Since some species have not yet been surveyed, additional SIV lineages may still be discovered. Extensive diversity has also been observed for FIV in wild and captive cat species.42,346 Among the four subspecies of African green monkeys (vervet, grivet, tantalus, and sabeus), discrete sub-subgroups of SIVagm have been defined (Table 19.2). These four subspecies naturally inhabit distinct or sometimes partially overlapping habitats that cover almost all of sub-Saharan Africa. The JDV also represents a distinct subgroup relative to the original BIV isolate.43

TABLE 19.2 Partial listing of primate lentivirusesa

735

The pol gene generally exhibits the greatest degree of sequence conservation. Pol sequences are therefore often used for the comparison of lentiviruses from different groups, subgroups, or sub-subgroups. Sequences from one subgroup of SIV (e.g., SIVsmm) will typically exhibit only 55% to 60% amino acid identity in Pol when compared with sequences from another SIV subgroup (e.g., SIVagm). Diverse members within a subgroup may exhibit as little as 75% to 80% amino acid identity in Pol, but the number is typically higher. When different groups of lentiviruses are compared, for example, MVV with SIV, amino acid identity in Pol is typically 35% or less. Similarity in other genes is even lower. Nonetheless, lentiviral sequences are clearly more closely related to one another than to other retroviruses. All known lentiviruses have at least three genes in addition to gag, pol, and env that are present in all replication-competent retroviruses (Fig. 19.4 and Tables 19.4 and 19.5). These additional genes contribute to the more complex biology of lentiviruses, which includes persistent virus replication and immune evasion as discussed in more detail later in this chapter. A rev gene that encodes a protein responsible for the nuclear export of unspliced viral RNA transcripts through interactions with a region of RNA secondary structure, referred to as the Rev response element (RRE), is present in all lentiviruses.274 Interestingly, although neither the sequence nor the location of the RRE is conserved, its function appears to be the same in all lentiviruses.274 A vif gene, whose primary function is to counteract restriction by members of the APOBEC3 family of cytosine deaminases, is consistently present in five of the six lentivirus groups; EIAV stands alone in lacking a vif gene. A nef gene is found at the 3′ end of all primate lentiviruses (Table 19.5). The nonprimate lentiviruses do not have a nef gene. However, cells infected with EIAV and BIV make spliced mRNAs that encode the Ttm and Tmx proteins, which correspond to the carboxy-terminal portion of Env.22,117 It is possible that these viral proteins represent evolutionary precursors of Nef. All lentiviruses except FIV contain an unusually long (>120 amino acids) Env cytoplasmic domain.277 In the intergenic region between vif and env, a number of other genes are present, particularly in the primate lentiviruses. A vpr gene is present in all known primate lentiviruses. The HIV-2/SIVsmm, SIVmnd-2, and SIVrcm lineages also have a vpx gene in this region that is a paralog of vpr. The nonprimate lentiviruses have genes between vif and env with varied roles. BIV encodes a Tat protein from this location that acts on a trans-activation response element similar to the primate lentiviruses.210 In other ruminant lentiviruses, a small protein encoded from this region was originally thought to be transcriptional trans-activator but may actually have activity more similar to Vpr.365 Likewise, a small protein encoded in a similar region of FIV, termed OrfA, down-regulates the CD134 receptor for this virus from the surface of infected cells, similar to CD4 down-regulation by primate lentiviral Nef proteins.144 Only two groups of primate lentiviruses have consistently been found to have a vpu gene: HIV-1/SIVcpz/gor and SIVgsn/mon/mus (Table 19.5). SIVden from a pet Dent’s Mona monkey (Cercopithecus mona denti) was also found to contain a vpu gene, although this virus clusters phylogenetically with SIVdeb from DeBrazza monkeys.70 736

FIGURE 19.3 Lentivirus morphogenesis and morphology. This electron micrograph of a cell infected with simian immunodeficiency virus shows virus budding from the cell in the absence of preformed particles (lower arrow) and a mature particle with a cylindrical or rod-shaped nucleoid (upper arrow). (Courtesy of John MacKey.) TABLE 19.3 Properties of lentiviruses Property

HIV-1

SIV

MVV, EIAV, BIV, FIV

Morphogenesis/morphology

Lenti

Lenti

Lenti

Macrophage tropism

Yes

Yes

Yes

CD4 lymphocyte tropism

Yes

Yes

No

Use of CD4 as receptor for virus entry

Yes

Yes

No

Use of chemokine receptors as receptor for virus entry

Yes

Yes

Yes (FIV)

Natural modes of transmission

Sex, blood, vertical

Sex, blood, vertical

Insects, saliva/aerosols, blood, sex, vertical

6

5 or 6

3 or more

Yes

Yes

Yes

dUTPase

No

No

Yes

Persistent viral replication

Yes

Yes

Yes

Chronic, debilitating diseasea

Yes

Yes

Yes

Immunodeficiencya

Yes

Yes

No (MVV, EIAV, BIV), Yes (FIV)

Genes in addition to gag, pol, env, tat, and rev activities

aIn susceptible hosts.

Not all hosts are susceptible to disease.

737

FIGURE 19.4 Genome organization of representative lentiviruses. TABLE 19.4 Accessory genes in nonprimate lentiviruses MVVa

EIAVb

vif

tat (S1) vif

vif

tat

rev (S3) tat

rev

rev

S2

rev

tat (orfA)

ttm

tmx

BIVc

FIVd

vpw? vpy? aValas S,

Benoit C, Guionaud C, et al. North American and French caprine arthritis-encephalitis viruses emerge from ovine maedi-visna viruses. Virology 1997;237:307–318. Ref.353

bMaury W. Regulation of equine infectious anemia virus

cGonda M. The lentiviruses

expression. J Biomed Sci 1998:11–23. Ref.228

of cattle. In: Levy JA, ed. Retroviridae. New York: Plenum Press; 1994. Ref.116

dOlmsted RA, Barnes AK,

Yamamoto JK, et al. Molecular cloning of feline immunodeficiency virus. Proc Natl Acad Sci U S A 1989;86:2448–2452. Ref.252

738

All nonprimate lentiviruses except BIV encode deoxyuridine triphosphatase (dUTPase) from a distinct open reading frame within the pol gene.89 dUTPase converts dUTP to a precursor of dTTP, which reduces dUTP concentrations in infected cells. By doing so, dUTPase prevents mis-incorporation of uracil into viral DNA, thereby reducing the accumulation of G-to-A transitions in the viral genome. dUTPase is particularly important for lentiviral replication in nondividing cells such as macrophages where cellular dUTPase levels are especially low. Curiously, dUTPase coding sequences are absent from the primate lentiviruses. The explanation for this is not entirely clear, but may be related to the evolution of Vpr and Vpx to facilitate virus replication in nondividing cells. This is consistent with the transitional genomic structure of an endogenous lentivirus found in a prosimian species, the gray mouse lemur (Microcebus murinus), which encodes dUTPase but lacks a vpr gene.111 Thus, dUTPase may have been lost during the course of primate lentiviral evolution as a result of the acquisition of Vpr.

TABLE 19.5 Accessory genes of primate lentiviruses

PROPAGATION Propagation and Cell Culture Primary isolates from all lentivirus groups can be grown in macrophage cultures of their respective host species. Other types of cells may also be used depending on the virus (Table 19.6), and virus isolates may be adapted to replicate in particular cell types. Examples include FIV replication in Crandell feline kidney cells and the replication of SIV and HIV-1 in immortalized T-cell lines (Table 19.6). However, it is important to note that lentiviral replication in cell culture can select for phenotypic changes that do not represent natural replication in infected hosts.

Host Range SIVs naturally infect a variety of African apes and Old World monkeys, but natural infection of Asian nonhuman primates has never been documented. The SIVs from sooty mangabeys and African green monkeys when introduced into macaque species (Asian monkeys) can cause an AIDS-like disease.100,140,242 In fact, SIVsmm from sooty mangabeys was accidentally introduced into captive macaque colonies in the United States and disseminated unknowingly for more than a decade before it was discovered and eliminated.7,66,219 At least one case of laboratoryacquired human infection with SIVmac has been documented.173,333

Baboons do not appear to naturally harbor their own SIV. However, cross-species transmission of SIV to baboons in the wild has been documented. Of 279 baboon sera taken from native habitats in Tanzania and Ethiopia, 2 animals had strong reactivity to SIVagm antigens and 1 of these animals was later shown to harbor SIVagm in plasma.181 The virus sequences from this baboon clustered with the vervet subtype of SIVagm, consistent with the overlapping habitat of vervet monkeys in the region where the baboon samples were taken.158 An SIVagm variant has similarly been detected in a chacma baboon of southern Africa.359 There is additional evidence for the cross-species transmission of SIVagm from West African green monkeys (Cercopithecus aethiops sabeus) to patas monkeys (Erythrocebus patas).28 Baboons can also be infected experimentally with HIV-2.211

TABLE 19.6 Cell substrates for growing lentiviruses Lentivirus Group

Growth on Macrophages

Other Cell Types

Adaptation to Growth

HIV-1

Yes

Stimulated, primary, human CD4+ lymphocytes

Human tumor T-cell lines

SIV

Yes

Stimulated, primary, monkey CD4+ lymphocytes

Human tumor T-cell lines

MVV/CAEV

Yes

Choroid plexus cells; primary synovial cells; endothelial cells

—

FIV

Yes

Primary, stimulated feline lymphocytes

Crandell feline kidney cells

EIAV

Yes

—

Fetal equine kidney; equine dermal

739

BIV

Yes

—

Bovine, rabbit, canine fibroblasts

It is generally accepted that HIV-2 arose from the zoonotic transmission of SIVsmm from sooty mangabeys to humans. HIV-2 is primarily found in western Africa and has only slowly made its way to other parts of the world. The native habitat of the sooty mangabey is the coastal forest region of western Africa, which corresponds to regions where HIV-2 is most prevalent. SIVsmm also has the same genomic organization and groups phylogenetically with HIV-2 (Figs. 19.2 and 19.4).104,142 Of the nine distinct groups of HIV-2 thought to represent independent crossspecies transmission events, most are limited to a few cases in West Africa.313 The two groups of HIV-2 that account for the majority of spread (groups A and B) are closely related to SIVsmm isolates that have been identified in wild sooty mangabeys of the Taï forest in Côte d’Ivoire.294

Noninvasive fecal sampling techniques for great apes in the wild have provided a wealth of data on the origins of HIV-1. Molecular epidemiological studies of thousands of samples collected from wild chimpanzee and gorilla populations across Africa have identified SIV strains related to each of the four groups of HIV-1 (groups M, N, O, and P). SIVcpz has been found in only two of the four subspecies of the common chimpanzee; Pan troglodytes troglodytes in West Central Africa and P. t. schweinfurthii in East Africa.168,357 These subspecies harbor their own lineages (SIVcpzPtt and SIVcpzPts), and while SIVcpzPtt sequences are closely related to HIV-1, viruses similar to SIVcpzPts have not been found in humans.262 Detailed phylogenetic analyses indicate that HIV-1 group M, which is responsible for the global AIDS pandemic, and HIV-1 group N, which has only been found in a few individuals, originated from SIVcpzPtt in geographically distinct populations of chimpanzees in southern Cameroon.168,203,291,357 Extensive fecal sampling and sequencing have similarly shown that western lowland gorillas (Gorilla gorilla gorilla) are naturally infected with their own SIV and that SIVgor probably originated from the transmission of SIVcpzPtt from chimpanzees to gorillas.358 HIV-1 groups O and P are more closely related to SIVgor than to SIVcpzPtt, and phylogeographic analyses have traced the ancestors of these viruses to distinct populations of gorillas in southern Cameroon.69,262 Whereas HIV-1 group O has been found in approximately 100,000 individuals in West Central Africa, HIV-1 group P has only been identified in a handful of patients.

The absence of SIVcpz in two of the four chimpanzee subspecies (P. t. ellioti and P. t. vellerosus) and the genetic composition of the virus in the subspecies that do harbor SIVcpz suggest that chimpanzees may have acquired SIV relatively recently. A recent origin for SIVcpz is also consistent with the finding that at least some chimpanzees develop an AIDS-like disease from SIVcpz infection.92,290,342 SIVcpz is more closely related to SIVrcm in Pol and to SIVgsn in Env than it is to the corresponding sequences of other SIVs.14 Furthermore, the range of the chimpanzees (P. t. troglodytes) in West Central Africa overlaps with the ranges of both red-capped mangabeys and greater spot-nosed monkeys, and chimpanzees have been known to prey on monkeys for food (Fig. 19.5). Thus, SIVcpz may have recently emerged from an ancestral recombination between viruses similar to SIVrcm and SIVgsn.

Attempts to infect species other than great apes with HIV-1 have failed. HIV-1 is infectious for chimpanzees and chimpanzee-passaged HIV-1 has shown pathogenic potential249,253; however, HIV-1 does not replicate in Old World monkeys. The two strongest blocks to HIV-1 replication in monkeys are the restriction factors TRIM5 and APOBEC3.315,337 Simian-tropic strains of HIV-1 have now been engineered with defined sequence changes that overcome each of these restriction factors to allow persistent virus replication in macaques.134,305

The nonprimate lentiviruses are also restricted in their host range, as reflected by their replication only in the same or in closely related genera. BIV is a notable exception in that it has been reported to infect New Zealand white rabbits.270 However, cross-species transmission does occur. The transmission of CAEV from goats to sheep has been observed,271 although CAEV is genetically similar enough to MVV to qualify as a quasispecies of this lentivirus. Studies have also shown that domestic cats can be infected by puma FIV (FIVpco) and lion FIV (FIVple).360,361 Although no overt disease symptoms have been documented for the latter, long-term studies have not been performed.

Restriction Mammals express a number of gene products termed “restriction factors” that interfere with specific stages of virus replication at the cellular level. The best characterized of these are the APOBEC3, TRIM5, tetherin, SAMHD1, MX2, SERINC5, and ZAP proteins.123,148,165,189,243,315,337,355 To replicate efficiently in their respective hosts, lentiviruses have evolved mechanisms to overcome each of these factors. Resistance to restriction can occur by the expression of a viral protein (usually an accessory protein) that directly antagonizes the restriction factor or by changes in the viral target sequence that reduce sensitivity to restriction. Viral pathogens have in turn caused selection for sequence variation in restriction factors among different species and in some cases within the same species. As a consequence, a few of these species-specific differences represent important barriers to the cross-species transmission of lentiviruses.232,234,299,301 Key properties of each of the restriction factors discussed below are summarized in Table 19.7.

740

FIGURE 19.5 Chimpanzee “Sagu” eating a spine and rib cage of a red colobus monkey. Photo by Cristina Gomes, Max-PlanckInstitute for Evolutionary Anthropology. (Photo by Cristina Gomes, Max-Plank Institute for Evolutionary Anthropology. Reprinted from Leendertz SA, Locatelli S, Boesch C, et al. No evidence for transmission of SIVwrc from western red colobus monkeys (Piliocolobus badius badius) to wild West African chimpanzees (Photo by Cristina Gomes, Max-Planck-Institute for Evolutionary Anthropology; Tropical Conservation Institute, Florida International University. Reprinted from Leendertz SA, Locatelli S, Boesch C, et al. No evidence for transmission of SIVwrc from western red colobus monkeys (Piliocolobus badius badius) to wild West African chimpanzees (Pan troglodytes verus) despite high exposure through hunting. BMC Microbiol 2011;11(1):24. Ref.196)

TABLE 19.7 Lentiviral restriction factors

APOBEC3 The apolipoprotein B mRNA editing enzyme, catalytic polypeptide-like 3 (APOBEC3 or A3) proteins are a family of mammalian cytosine deaminases that inhibit the replication of lentiviruses as well as other retroviruses and retroelements. Humans and other primate species express seven distinct A3 genes (A3A, A3B, A3C, A3D, A3F, A3G, and A3H),190 of which A3D, A3F, A3G, and A3H are able to restrict HIV-1 and SIV.151 If unopposed, A3 proteins become incorporated into virions where they catalyze the conversion of cytosines into uracils during the minus-strand cDNA synthesis step of reverse transcription. These uracils then template the insertion of adenines during plus-strand cDNA synthesis, resulting in guanine-to-adenine (G-to-A) transitions in the proviral DNA.132,396 Lentiviruses overcome this restriction by expressing Vif, which mediates A3 degradation in productively infected cells thereby preventing A3 incorporation into virions. In the absence of antagonism by Vif, the accumulation of extensive G-to-A substitutions results in catastrophic hypermutation of the viral genome, which prevents further virus replication.133,194

A3 antagonism by Vif is often species-dependent. HIV-1 Vif can degrade the A3G proteins of humans and chimpanzees but not those of African green monkeys or rhesus macaques.325 Conversely, SIVagm Vif can degrade African green monkey A3G but not human A3G.325 This specificity is governed by a single amino acid difference between human and monkey A3G that is critical for binding to Vif.30,307 The failure of HIV-1 Vif to counteract the A3 proteins of Old World monkeys represents a major barrier to HIV-1 replication in these species. Likewise, the inability of SIVagm Vif to counteract human APOBEC3G is a significant barrier to the zoonotic transmission of this group of viruses to humans. As an illustration of the evolutionary dynamics driving A3G variation and viral adaptation, amino acid differences were identified in the A3G proteins of four different subspecies of African green monkeys that confer resistance to antagonism by the Vif proteins of SIVagm strains from other subspecies but not from the same subspecies.56

TRIM5 Isoforms of the tripartite motif-containing protein 5 (TRIM5), most notably the alpha isoform (TRIM5α), impose a postentry block to virus infection that represents a major host range determinant for lentiviruses and other types of retroviruses.337 Evolutionary analyses indicate that TRIM5 has been coevolving with retroviral pathogens for tens of millions of years, perhaps since the radiation of eutherian mammals.300 Homologs of the TRIM5 gene have been found in the genomes of primates, cows, pigs, dogs, rabbits, rats, and mice,300 and antiviral activity has been demonstrated for the TRIM5 proteins of various species of Old and New World primates136,292,331 as well as for the TRIM5 proteins of rabbits and cows.303,395 TRIM5 typically does not block infection by retroviruses adapted to a particular host but exhibits variable patterns of restriction against the retroviruses of other species.136 Differences in sensitivity to restriction reflect sequence variation in the viral capsid protein and in the domains of TRIM5 predicted to interact with capsid.246,383,394 The mechanism of restriction remains to be fully defined but appears to involve TRIM5 recognition of the hexagonal capsid symmetry of intact retroviral core particles and the formation of a higher-order lattice around the core that destabilizes it before the completion of reverse transcription.103

One of the more peculiar twists in TRIM5 evolution is the independent emergence of a TRIM5-cyclophilin A (TRIM5-Cyp) isoform in at least two different primate lineages. Owl monkey cells exhibit a potent postentry block to HIV-1 infection due to a TRIM5-Cyp fusion resulting from the retrotransposition of an open reading frame for CypA into an intron of TRIM5.302 A similar TRIM5-Cyp fusion occurred in macaques as a result the insertion of CypA into the 3′ UTR of TRIM5.366,384 In this case, macaque TRIM5-Cyp poorly restricts HIV-1 but does block infection by other lentiviruses such as HIV-2, SIVagm, and FIV.366,384

TRIM5 polymorphisms may also result in variable courses of infection within a single species. In contrast to SIV strains such as SIVmac239 that are well-adapted to rhesus macaques and consistently result in high viral loads, rhesus macaques infected with SIVsmE543-3 exhibit considerable animal-to-animal variation in plasma viremia. Kirmaier et al. found that this variation reflects differences in TRIM5 genotype and that the adaptation of SIVsmE543-3 for efficient replication in rhesus macaques is associated with amino acid changes in the viral capsid that confer resistance to restrictive variants of TRIM5α.176 The susceptibility of SIVsmE543-3 to certain allotypes of rhesus macaque TRIM5α 741

appears to be a result of the incomplete adaptation of the virus to this species, since SIVsmE543-3 was derived after the sequential passage of SIVsmm in only two rhesus macaques.135,138

Tetherin Tetherin (BST-2 or CD317) is an interferon-inducible transmembrane protein that inhibits the detachment of enveloped viruses from infected cells. Although initially identified as the cellular factor that accounts for a defect in the release of vpu-deleted HIV-1 from restrictive cells,243,355 tetherin has since been shown to have broad antiviral activity against diverse families of enveloped viruses.244 The antiviral activity of tetherin is a function of its unique topology, which includes an N-terminal cytoplasmic domain, a membrane-spanning domain, an extracellular coiled-coil domain, and a C-terminal glycosyl-phosphatidylinositol anchor.184,265 These features allow opposite ends of tetherin dimers to become incorporated into viral and cellular membranes, physically linking budding virions to the surface of infected cells.95,127,265 Tetherin may also amplify other immune responses. Virion-induced cross-linking of tetherin stimulates NK-κB signaling and the release of proinflammatory cytokines,102 and the accumulation of captured virions on the cell surface increases the susceptibility of infected cells to antibody-dependent cellular cytotoxicity.10 Thus, tetherin may serve as a link between innate and adaptive immunity to enhance the elimination of virus-infected cells by antibodies.

Among the primate lentiviruses, at least three different viral proteins have evolved to counteract tetherin. Most SIVs, including phylogenetically diverse viruses such as SIVcpz, SIVagm, and SIVsmm, use Nef to counteract the tetherin proteins of their simian hosts.156,298,402 However, the SIVs of Old World monkeys that have a vpu gene (SIVgsn, SIVmon, and SIVmus) use Vpu as a tetherin antagonist.298 HIV-1 group M and HIV-2 evolved to use Vpu and Env, respectively, to counteract human tetherin because of the loss of a five amino acid sequence from the cytoplasmic domain of human tetherin that confers susceptibility to Nef.156,192,243,355 Thus, even though the SIVs that gave rise to HIV-1 and HIV-2 (SIVcpz and SIVsmm) use Nef to antagonize tetherin in their natural hosts, the absence of sequences from the N-terminus of human tetherin that confer sensitivity to Nef explains why HIV-1 group M evolved to use Vpu and why HIV-2, which lacks a vpu gene, evolved to use Env to counteract human tetherin. The resistance of human tetherin to Nef does not appear to be absolute, however, since the Nef proteins of HIV-1 group O and certain HIV-1 group M isolates are able to counteract human tetherin.9,178

Species-specific differences in tetherin represent a potential barrier to cross-species transmission. HIV-1 Vpu counteracts human, chimpanzee, and gorilla tetherin but is ineffective against the tetherin orthologs of Old World monkeys.156,232,298 Conversely, the Nef proteins of SIVcpz, SIVsmm, and SIVagm counteract the tetherin proteins of their respective hosts but are generally ineffective against human tetherin.156,298,402 Differences in the susceptibility of tetherin to antagonism by Vpu and Nef map to amino acid residues at sites of contact with these viral proteins. Whereas variation in the membrane-spanning domain accounts for differences in susceptibility to Vpu, it is the cytoplasmic domain of tetherin that accounts for differences in susceptibility to Nef. This is consistent with the rapid evolution of sequences coding for the cytoplasmic and transmembrane domains of tetherin in response to the selective pressure of viral pathogens.232

Several instances of lentiviral adaptation to tetherin in primates have been documented. These include compensatory changes in the Env cytoplasmic domain of a nef-deleted strain of SIV that regained a pathogenic phenotype in rhesus macaques,311 changes in Nef that restore the ability to counteract tetherin in HIV-1–infected chimpanzees,122 adaptation of the Vpu protein of a simian-tropic HIV-1 to antagonize macaque tetherin,134 and compensatory changes in Nef that restore tetherin antagonism during the replication of a tetherin-sensitive SIV nef mutant in rhesus macaques.340 Together, these observations indicate that the primate lentiviruses are under strong selective pressure to overcome restriction by tetherin.

SAMHD1 Sterile alpha motif and histidine–aspartic acid domain–containing protein 1 (SAMHD1) is a GTP/dGTP-activated triphosphohydrolase that cleaves deoxynucleotide triphosphates (dNTPs) into their deoxynucleoside and inorganic triphosphate components.115 In nondividing cells, SAMHD1 forms catalytically active tetramers that maintain intracellular dNTP pools below concentrations needed to complete reverse transcription.11,115 The Vpx proteins of HIV-2 and SIVsmm/mac alleviate this barrier to infection of differentiated myeloid cells (macrophages and dendritic cells) and resting CD4+ T cells by targeting SAMHD1 for proteasomal degradation.15,148,189 However, Vpx is only expressed by the HIV-2/SIVsmm, SIVmnd-2, and SIVrcm lineages of primate lentiviruses (Table 19.5).

An evolutionary analysis of SAMHD1 antagonism revealed that the Vpr proteins of some SIVs, including SIVagm, SIVdeb, and SIVmus, acquired the ability to degrade their host’s SAMHD1 proteins.208 Vpr and Vpx are related by gene duplication; however, unlike Vpx, Vpr is expressed by all primate lentiviruses. Phylogenetic analyses suggest that the functionalization of Vpr to counteract SAMHD1 occurred prior to the gene duplication that gave rise to Vpx in SIVs of the Cercopithecinae subfamily of Old World monkeys.208 Comparisons of the SAMHD1-coding sequences among Old and New World primates also revealed evidence of positive selection among Cercopithecinae species, including at sites essential for interactions with Vpx.208 Lentiviral antagonism may therefore have driven SAMHD1 evolution in these species, and speciesspecific differences in SAMHD1 represent a potential barrier to cross-species transmission.

MX2 The myxovirus resistance (MX) proteins are a family of dynamin-like GTPases found in vertebrates that are strongly inducible by type I interferons. Mammals typically express two paralogous MX proteins, MX1 and MX2 (also known as MXA and MXB), which differ in their spectrum of antiviral activity. Whereas MX1 has broad activity against diverse families of RNA viruses, including orthomyxoviruses, paramyxoviruses, and rhabdoviruses (but not retroviruses), MX2 was thought to lack antiviral activity until it was identified as an interferonstimulated gene product responsible for differences in resistance to HIV-1 infection.123,165 Although its mechanism of antiviral activity is not fully understood, MX2 is thought to impede nuclear import of pre-integration complexes. This is consistent with the inhibition of HIV-1 replication at a step after reverse transcription but before proviral integration, the localization of MX2 to nuclear pores, and the physical interactions of MX2 with multiple components of the nuclear import machinery.84,123,165

Similar to TRIM5, MX2 binds to surfaces of the HIV-1 capsid, and it is differences in capsid protein sequences that differentially affect virus 742

restriction by MX2.25,39,326 However, MX2 does not impose as stringent of a block to HIV-1 infection as TRIM5, and sensitivity to MX2 is dependent on cyclophilin A as well as cell type and cell cycle variation in the expression of nucleoporins.164 Sequence variation and signatures of positive selection among the MX2 orthologs of different species indicate evolutionary conflict with viral pathogens.39 Serial passage of HIV-1 in cell lines that overexpress MX239 and the adaptation of simian-tropic HIV-1 to macaques305 can also select for amino acid changes in capsid that confer resistance to MX2. However, the sensitivity of primary HIV-1 isolates to human MX2, including interferon-resistant transmitted/founder viruses, as well as the general sensitivity of other primate lentiviruses to the MX2 proteins of their simian hosts,39 suggests that MX2 is not a significant barrier to the cross-species transmission of the primate lentiviruses.

SERINC5 Serine incorporator 3 and 5 (SERINC3 and SERINC5) were identified as the cellular factors that account for producer cell-dependent impairment of the infectivity of nef-deleted HIV-1.287,352 In the absence of Nef, these multipass transmembrane proteins become incorporated into budding virions and potently inhibit HIV-1 infectivity.287,352 SERINC homologs are found in all eukaryotes, including five paralogous gene products in humans (SERINC1-5), that are distinguished by a unique bipartite domain structure composed of 10 membrane-spanning alpha helices.279 Although SERINC3 and SERINC5 both impair HIV-1 infectivity and are expressed in primary lymphocytes, the majority of antiretroviral activity can be attributed to SERINC5.287,352

There is a general consensus that SERINC5 inhibits the formation and expansion of fusion pores between viral and cellular membranes,287,352,372 thereby impairing the delivery of the viral nucleocapsid to the cytoplasm. How this occurs is less clear. SERINC5 does not appear to alter the lipid composition of viral membranes as its name might suggest.345 Instead, differences in the sensitivity of HIV-1 to SERINC5 map to Env.352 The ability of SERINC5 to increase the sensitivity of HIV-1 to certain neutralizing antibodies332 suggests that SERINC5 directly modifies Env conformation. There is also evidence that SERINC5 prevents the clustering of Env trimers on the surface of virions,47 which may inhibit fusion by reducing the number of Env–receptor contacts that can mediate entry.

SERINC5 antagonism is broadly conserved among the Nef proteins of phylogenetically diverse primate lentiviruses.137 Accessory proteins of other retroviruses have also evolved to counteract SERINC5, including the glycoGag protein of murine leukemia virus (MLV) and the S2 protein of EIAV.45,287,352 Thus, the SERINC proteins appear to have broad antiviral activity that has impacted the course of retroviral evolution. However, unlike other restriction factors discussed thus far, the SERINCs do not appear to be evolving under positive selection. SERINC proteins are generally well conserved with few amino acid differences between species (e.g., human and macaque SERINC5 share 99% amino acid identity). As a consequence, the Nef proteins of HIV-1, HIV-2, and diverse SIVs are able to counteract human, simian, and even murine orthologs of SERINC3 and SERINC5.137 Likewise, EIAV S2 and MLV glycoGag counteract human, rodent, and lagomorph SERINC3/5.75 Thus, the SERINC proteins are unlikely to represent a significant barrier to lentiviral transmission between species.

ZAP The zinc finger antiviral protein (ZAP, also known as PARP13) was initially identified as a cellular gene product that specifically depletes viral mRNAs in the cytoplasm of MLV-infected cells.105 This factor was subsequently shown to have broad antiviral activity against diverse families of RNA viruses.121,404 The specificity of ZAP is dependent on the CG dinucleotide content of viral RNAs; increasing the frequency of GCs in HIV-1 increases the depletion of viral transcripts in the cytoplasm.339 Because vertebrate genomes contain a lower than expected frequency of CG dinucleotides, ZAP can differentiate viral RNAs from host cellular mRNAs based on their CG content.339

The antiviral activity of ZAP is not fully understood but is dependent on binding to RNA GC dinucleotides233 and the formation of a complex with at least two cofactors, TRIM25 and KHNYN.94,202 TRIM25 regulates ZAP by a poorly understood mechanism,403 and KHNYN contains an endonuclease domain that is essential for the cytoplasmic depletion of HIV-1 RNAs.94 ZAP and TRIM25 are both interferon-inducible,314 and the ZAP gene encodes long and short protein isoforms (ZAP-L and ZAP-S) that are products of alternative splicing.169 Although both isoforms have antiviral activity, ZAP-L is reported to have greater activity against retroviruses. Signatures of recurrent positive selection are also localized to the PARP-like domain that is present in ZAP-L but not ZAP-S.169 Lentiviruses are unlikely, however, to account for this selective pressure; since like many RNA viruses, they have evolved to evade ZAP by maintaining low CG frequencies. Thus, ZAP is unlikely to represent a significant barrier to the cross-species transmission of lentiviruses.

Receptor Use HIV-1 uses a sequential two-receptor system that includes CD4 as a primary receptor and a seven transmembrane chemokine receptor as a secondary receptor for entry into target cells (see Chapter 17). The overwhelming majority of naturally transmitted HIV-1 field isolates use CCR5 as a second receptor, although variants that are able to use CXCR4 as an alternative co-receptor may arise at later stages of infection.162 A 32 base pair deletion in the CCR5 gene (CCR5δ32) occurs with an allelic frequency of approximately 10% in people of European descent. CCR5δ32 homozygotes are extremely resistant to HIV-1 infection. However, among the few documented cases of HIV-1 infection of CCR5δ32 homozygotes, CCR5 inactivation does not protect against high viral loads and disease progression because of the utilization of CXCR4 by HIV-1 in these individuals.327 The SIVs that have been examined to date are similar to HIV-1 in their use of CD4 and CCR5 as primary and secondary receptors, but rarely use CXCR4. However, many SIVs are also able to use additional chemokine receptors that are not utilized by HIV-1. Depending on the SIV strain, these receptors may include CXCR6 (STRL33 or Bonzo), GPR15 (BOB), CCR2b, CCR3, and GPR1.

The use of alternative co-receptors may help to explain high levels of persistent SIV replication without progressive CD4+ T-cell depletion in natural hosts. Early studies revealed that SIVrcm can use CXCR6 and CCR2b rather than CCR5 for entry,21,48 which probably accounts for the replication of this virus in red-capped mangabeys (Cercocebus torquatus torquatus) with homozygous deletions in CCR5. More recently, SIVsmm and SIVagm were also found to use CXCR6 for infection of primary CD4+ T cells from their natural hosts, the sooty mangabey (Cercocebus atys) and the African green monkey (Chlorocebus spp.).90,286,377 This is consistent with exceedingly low or absent CCR5 expression on CD4+ lymphocytes of these species,256 and possibly also with the downmodulation of CD4 on a subset of memory T cells in African green monkeys.19 On the other hand, SIVcpz and SIVmac, which are pathogenic in chimpanzees and rhesus macaques, respectively, are 743

unable to use their hosts’ CXCR6 for entry—which appears to be a property of Env in the case of SIVcpz and an amino acid difference in rhesus macaque CXCR6 in the case of SIVmac.378 Hence, these viruses depend on high levels of CCR5 for efficient replication in their respective hosts. The observation that CXCR6 is expressed on a subset of effector memory CD4+ T cells in sooty mangabeys distinct from the central memory population that expresses CCR5 raises the intriguing possibility that some natural hosts may be able to sustain high levels of SIV replication without CD4+ T-cell depletion because the virus is replicating in a cell population that is more easily replaced.378

FIV also uses a two-receptor mechanism and shares with HIV the use of CXCR4 as a co-receptor for cell entry.272,381 FIV infection is modulated by soluble stromal cell–derived factor-1 (SDF-1), the natural ligand for CXCR4,145 and is inhibited by the CXCR4 antagonist AMD3100. However, FIV uses the T-cell activation marker CD134 as the initial binding receptor rather than CD4.72,318 CD134 is a member of the TNF receptor superfamily and has the typical four-domain structure. The outermost domain is responsible for receptor activity and as few as five amino acid changes can make human CD134 a viable receptor for FIV.71 Certain lion FIVs also use CD134 and CXCR4 sequentially for entry.230 This two-receptor mechanism used by the primate lentiviruses and by FIV conceals critical surfaces of the envelope glycoprotein from host antibody responses prior to conformational changes that mediate entry.

EIAV also uses a member of the TNF receptor superfamily for entry, a protein called ELR1, which is expressed on macrophages that are the primary target cells for this virus.399,400 Little information is currently available about the receptors used by other nonprimate lentiviruses, leaving a number of fundamental questions central to lentiviral evolution and receptor biology unanswered. Do all lentiviruses use a tworeceptor system for entry into target cells? Is the second receptor always a seven membrane-spanning chemokine receptor? Did evolutionary changes result in a switch from a one-receptor system to a two-receptor system or a switch from one two-receptor system to another tworeceptor system?

Germline Integration Many retroviruses not only infect their hosts exogenously but may also be inherited in a mendelian fashion, either as highly defective genomes, solo long terminal repeats (LTRs), or in the case of the murine and feline gammaretroviruses as inducible infectious agents. However, endogenous lentiviruses have not been found in the vast majority of species examined, suggesting that germline integration for lentiviruses is much rarer than for other types of retroviruses. The absence of endogenous lentiviruses was thought to reflect the recent evolutionary origin of lentiviruses, the lack of lentiviral receptors on germ cells, and the cytopathic nature of lentiviral infection. Nevertheless, there are now four examples of germline transmission of nonreplicating lentiviral elements. Since entering the germline, these elements have been expanded and shuffled by host genetic mechanisms and are now inherited as multiple copies of various defective forms by all individuals of their respective host species. These elements no longer express functional viral proteins due to the accumulation of numerous in-frame stop codons and frameshift mutations in coding regions. The first endogenous lentivirus to be identified was in the European rabbit (Oryctolagus cuniculus). Rabbit endogenous lentivirus type K (RELIK) as it was termed has unmistakable lentiviral sequences, including gag, pol, env, tat, and rev genes.166 The pol gene of RELIK encodes dUTPase similar to the nonprimate lentiviruses and like EIAV lacks an open reading frame for vif.166 Subsequent to the identification of RELIK, endogenous lentiviruses were also identified in two different genera of Malagasy lemurs (Microcebus and Cheirogaleus),111,112 mustelids (weasels and ferrets),65,128 and a colugo (Galeopterus variegatus).129 These lentiviral elements are phylogenetically distinct from one another in accordance with independent introductions into the germlines of these species. Estimates of the time since these lentiviral sequences entered the germline vary between 1.9 and 14.3 million years (Table 19.8). These findings therefore indicate that lentiviruses are much older than previously appreciated.

Inspection of the endogenous lentiviruses found in the gray mouse lemur (M. murinus) and the fat-tailed dwarf lemur (C. medius) revealed that they represent distinct germline insertions and are phylogenetically basal to all extant lineages of exogenous primate lentiviruses.111,112 In addition to gag, pol, and env, these prosimian immunodeficiency viruses (pSIVgml and pSIVfdl) contain a vif gene and a putative open reading frame in a similar position as nef, albeit with no discernible similarity to the nef genes of known SIVs.111 However, similar to the nonprimate lentiviruses, pSIVgml and pSIVfdl have a dUTPase coding region in pol and lack orthologs of the primate lentiviral vpr and vpx genes. Hence, the endogenous lentiviruses of lemurs exhibit transitional features intermediate to the nonprimate and primate lentiviruses.

TABLE 19.8 Summary of known endogenous lentiviruses Host

Geographic Location

Estimated Time of Endogenization Genes Present

Rabbit

Europe

5.7–7 M years

gag, pol, env, tat, rev

Lemur

Madagascar

1.9–4.2 M years

gag, pol, env, tat, rev, vif

Weasel (Ferret)

Widespread

8.8–11.8 M years

gag, pol, env, tat, rev, vif

Colugo

Southeast Asia

~14.3 M years

gag, pol, env, tat, orf1a, orf2a

aThe proteins encoded by orf1 and orf2 do

not share any significant similarity with other known lentiviral accessory elements.

PATHOGENESIS AND PATHOLOGY Portals of Entry

744

The primary mode of natural transmission of the nonhuman lentiviruses varies considerably with the virus. EIAV may be the most interesting because it is the only lentivirus for which there is good evidence for vector-borne transmission. During disease episodes, levels of infectious virus in the plasma of horses can exceed 104/mL. The horse fly appears to be more efficient than mosquitoes, fleas, or other insects for being able to transmit EIAV. Transmission has been documented by following a single horse fly that took a blood meal on an infected pony during an acute clinical period.155 Transmission via blood can also be mediated by inappropriate veterinary practices involving needles or scalpels.

Work on natural modes of infection has demonstrated the capability of both vertical and horizontal transmission of FIV, MVV/CAEV, and EIAV.40 Vertical transmission can occur in utero, during parturition, and postnatally via milk. Vertical transmission by these viruses parallels that observed with HIV in humans. Contact transmission is also common, particularly for MVV/CAEV and EIAV, when animals are herded closely together in barns or stables. These viruses can be found in semen, lung excretions, and saliva. For cats, bite wounds are believed to be the most important route of transmission in adult animals.390 Transmission via grooming/licking has also been observed,74 and experimental infection of cats can be achieved via oronasal administration. FIV infection is much more prevalent in free-roaming males than in females, consistent with increased fighting and biting among male cats. Cats allowed to roam free in areas with high cat density are at the greatest risk of becoming infected.

Information about natural modes of SIV transmission has been harder to come by. One study of wild grivet monkeys in Awash National Park in Ethiopia analyzed SIVagm serologic status with age, sex, and risk.267 Infection was nearly universal in females of reproductive age and nearly absent in younger females. In males, infection was observed in only those that were fully adult. The findings support a predominantly sexual mode of SIV transmission among grivets. Male-to-male transmission by aggressive contacts may also be a prominent mode by which SIV is spread.245 Maternal–infant transmission of SIV has been documented in captive animals.179 Experimental SIV infection of laboratory animals is routinely achieved by intravenous injection or by mucosal inoculation.

Cell and Tissue Tropism Despite the varied modes of transmission, all lentiviruses are disseminated to an assortment of tissue sites by the blood. Dissemination can occur as cell-free virus or by virus-infected monocytes or lymphocytes. Differences in the cellular tropism of primate versus nonprimate lentiviruses relate largely to receptor use as discussed earlier. Infection by the primate lentiviruses SIV and HIV is seen in CD4+ lymphocytes and macrophages, with CD4+ lymphocytes vastly predominating in terms of numbers of infected cells. For the nonprimate lentiviruses, infection of macrophages typically predominates, but infection can also be seen in other cell types. Replication in tissue macrophages is a unifying feature of all lentiviruses.

The principal anatomical sites of MVV and CAEV infection have varied with the study and with the strain of virus used. Because many studies have used experimental infection, the origin of the virus and the cell type on which it was grown must be considered when interpreting the results. Various reports have localized MVV principally to the lungs, mammary glands, joints, lymph nodes, the spleen, and the brain. Tissue macrophages are the principal target cell in which MVV and CAEV sequences are consistently found. However, in keeping with the broad range of cell types that can support replication in culture, evidence has been presented for MVV/CAEV infection of other cell types, including epithelial and fibroblast cells of the choroid plexus, intestine, and kidneys. An important observation first made with MVV is that although the virus may reside in relatively undifferentiated monocytes in peripheral blood, MVV expression is greatest in differentiated tissue macrophages.109,263 This led to the “Trojan horse” concept for MVV and for other lentiviruses whereby undifferentiated monocytes in peripheral blood may carry the proviral genome with little or no viral protein expression until they reach tissues where they differentiate into macrophages and begin to produce virus.

A variety of studies have also shown tissue macrophages to be the predominant cell type for productive infection with EIAV.231,250,310 Virus replication has been noted in the spleen, liver, kidney, lymph nodes, lung, heart, brain, stomach, bone marrow, thymus, adrenals, and intestine. Although EIAV can be adapted to replicate on fibroblasts, the virus that comes out of horses during viremic episodes is derived principally from macrophages. The anemia caused by EIAV is hemolytic and results from the formation of antigen–antibody complexes that can associate with the surface of erythrocytes. The kidneys are also affected by antigen–antibody complex formation.

FIV appears to be unusual among the nonprimate lentiviruses in that it has been found more consistently in a broader range of cell types, particularly lymphocytes, in addition to macrophages. Analyses of tissue and cellular tropism have primarily depended on experimentally infected cats and FIV grown in activated peripheral blood mononuclear cell (PBMC) cultures. However, studies with naturally infected cats have yielded similar findings. FIV has been found in a variety of cell types including CD4+ T lymphocytes, B cells, CD8+ T cells, macrophages, bone marrow–derived cells, and cells of the central nervous system (CNS). Macrophages, microglia, and astrocytes, but not neurons, in the brains of infected cats have been identified as targets of FIV infection.86 The propensity of FIV to replicate in and deplete CD4+ T lymphocytes is consistent with use of the activated T-cell marker CD134 (OX40) as the initial receptor and CXCR4 as a co-receptor for cell entry.71,72,318

The major sites of SIV replication in macaques at early stage of infection are the gastrointestinal tract, thymus, spleen, and other lymphoid tissues.187,362 SIV is found at early time points within periarteriolar lymphoid sheaths in the spleen, paracortex of lymph nodes, and the medulla of the thymus. The gastrointestinal tract contains most of the lymphoid tissue in the body, and activated T lymphocytes of the gastrointestinal mucosa express CCR5 in great abundance. SIV infection of rhesus monkeys results in profound and selective depletion of CD4+ T cells in the intestine within days of infection, before such changes are evident in peripheral lymphoid tissues. Thus, the gastrointestinal tract appears to be a major site of SIV replication and CD4+ T-cell depletion.362,363 Subsequent to these pioneering studies in SIV-infected macaques, the gut-associated lymphoid tissues were shown to be the major site of HIV-1 replication during acute infection.35 The predominate replication of HIV-1 in the human gut is again associated with a preference for activated CD4+CCR5+ memory T cells.227 Within the thymus, marked depletion of thymic progenitors occurs by 21 days after infection of macaques with pathogenic SIV; this depletion is followed temporally by increased cell proliferation in the thymus and a rebound in thymocyte progenitors.389 The distribution of virus within lymphoid organs varies with the inoculum. In some animals, SIV can be found in the CNS early after infection. The infected cells in the brain, whether at early or later stages of SIV-induced encephalitis (SIVE), are primarily cells of the monocyte/macrophage lineage296 and may include perivascular macrophages that have migrated from the blood or resident microglial cells.382 A neurovirulent infectious molecular clone of SIV that results in SIVE in approximately 50% of rhesus macaques after about a year of infection was recently described as a more physiological model for HIVassociated neuropathology than earlier systems that depend on immunomodulation of SIV infection to induce rapid progression to SIVE.195,226

745

Immune Responses and Persistence Monkeys infected with SIV and animals infected with other lentiviruses typically develop strong antibody and CD8+ T-cell responses to the virus. These immune responses persist at high levels for the lifetime of the infected host, regardless of whether infection occurs by natural or experimental means. Approximately 20% to 30% of macaques infected with some AIDS-causing strains of SIV develop a more rapid disease course, make few or no antibodies to the virus, and develop AIDS within 3 to 7 months. The magnitude of the SIV-specific antibody response varies with the extent of virus replication. SIV deletion mutants that are progressively more attenuated and result in lower viral loads elicit weaker antibody responses (Fig. 19.6).80

FIGURE 19.6 Attenuation by accessory gene deletion lowers viral load and reduces the strength of the antiviral antibody response. SIVmac239δ4 has inactivating deletions in vpr, vpx, and two in nef. CD8+ T cells recognize and kill virus-infected cells presenting viral peptides on their cell surface in complex with MHC class I molecules. The importance of CD8+ lymphocytes in limiting the extent of SIV replication was demonstrated by depleting CD8+ T cells with anti-CD8 antibodies.306 When CD8+ cells are depleted during primary infection, virus replication proceeds unabated after peak viremia; this contrasts with nondepleted animals, in which immune responses typically reduce viral loads after 14 days of infection.306 Depletion of CD8+ lymphocytes during chronic SIV infection results in an immediate increase in viral loads that is suppressed again by the reappearance of virus-specific CD8+ T cells.306

In macaques infected with wild-type, pathogenic strains of SIV, CD4+ T-cell responses to viral antigens are typically weak or absent. However, infection with nef-deleted, live attenuated strains of SIV results in strong CD4+ helper T-cell responses.107 This situation may be analogous to HIV-1 infection in which virus-specific CD4+ T-cell responses are weak or absent in most individuals but are often strong in nonprogressors that are able to control virus replication.288 These observations suggest a struggle during early stages of infection, where CD4+ helper T cells respond to sites of virus replication only to become targets of virus infection. In most cases, the virus wins resulting in progressive CD4+ T-cell depletion, immunodeficiency, and ultimately progression to AIDS. However, under certain circumstances virus-specific CD4+ T cells are preserved and are able to support the development of antibody and CD8+ T-cell responses needed to contain virus replication.

Lentiviral persistence is achieved primarily through continuous virus replication. In HIV-1–infected individuals and SIV-infected animals, millions of CD4+ T lymphocytes are infected and turned over every day throughout months and years of ongoing virus replication.143,374 This does not mean that latent infection does not occur. Latent infection of quiescent cells with transcriptionally silent proviruses is a key component of the viral reservoir, which persists indefinitely even under conditions of complete suppression of virus replication with antiretroviral drugs. However, in the absence of antiretroviral therapy, virus replication continues unabated in most HIV-infected individuals and SIV-infected animals. This is also true of nonpathogenic SIV infection of natural hosts, which exhibit ongoing virus replication, high viral loads in plasma, CD4+ T-cell turnover, and sustained antibody and CD8+ T-cell responses.44

EIAV is unusual among the lentiviruses for the episodic nature of its persistent viral replication. Infection of horses with EIAV is associated with recurring episodes of fever, anemia, and thrombocytopenia. These episodes of clinical disease coincide with bursts of virus replication, which may be weeks or months apart. While the virus replicating in an infected horse during these periods is resistant to neutralization by contemporaneous serum, EIAV variants from earlier episodes can be neutralized by the same serum.236

Lentiviruses have evolved multiple mechanisms of immune evasion that allow them to replicate persistently in the face of vigorous host immune responses. In addition to overcoming innate immunity as discussed in the context of restriction factors, these include mechanisms of resistance to adaptive immunity. A few of the features that confer resistance to antibody and T-cell responses are summarized below and depicted in Figure 19.7. Resistance to Antibodies Structural studies have confirmed that the envelope glycoproteins of HIV-1 and SIV form trimers consisting of three noncovalently bound gp120 and gp41 subunits.163,379 Outer domain surfaces of Env trimers are covered by extensive N-linked glycosylation that creates a “glycan shield” to prevent antibody binding to underlying polypeptides.283 SIV strains lacking N-linked glycosylation sites in Env are more sensitive to antibodymediated neutralization and are able to elicit higher neutralizing antibody titers in infected macaques.283 Surfaces of Env that must be conserved to interact with cellular receptors are concealed by “conformational masking.”186 The CD4-binding site in gp120 is a deeply recessed pocket that is not accessible to most antibodies, and the co-receptor binding site is not even formed prior to CD4 engagement. Some of the most antigenic surfaces of Env (those that elicit the strongest binding antibody responses) are occluded by oligomerization via gp120-gp120 and gp120-gp41 contacts. Other Env surfaces that are readily accessible to antibodies, such as the variable loops, rapidly acquire amino acid changes to escape antibody binding.38,373,387 On average, there are also only 8 to 14 Env trimers per virion,50,398 which prevents antibodies from crosslinking Env trimers on virions, thereby preventing neutralization by all but the highest affinity antibodies that are capable of binding to Env using a single Fab arm.

Many of the structural features of Env that provide resistance to neutralization also protect virus-infected cells from elimination by antibodies. With a few notable exceptions,96,367 antibodies that mediate antibody-dependent cellular cytotoxicity (ADCC) also neutralize virus 746

infectivity,368 which is consistent with the idea that antibodies that can bind to functional Env trimers on virions can also bind to Env trimers on the surface of virus-infected cells. Additional mechanisms of resistance to ADCC include CD4 downmodulation by Vpu and Nef,364 which prevents exposure of CD4-inducible epitopes on the inner domain of gp120 that are normally occluded in “closed” Env trimers, and tetherin downmodulation by Vpu,10 which prevents the accumulation of captured virions on the plasma membrane.

FIGURE 19.7 Mechanisms of immune evasion by human and simian immunodeficiency viruses. MHC Class I Down-regulation The Nef proteins of HIV-1 and SIV downmodulate HLA A and B molecules from the surface of virus-infected cells, reducing their sensitivity to lysis by cytotoxic CD8+ T cells.55,309,338 More recently, the Vpu proteins of certain primary HIV-1 isolates were also found to downmodulate HLA C,8 providing additional resistance to HLA C–restricted CD8+ T cells.

Destruction of CD4+ Helper T Cells

The primate lentiviruses preferentially replicate in virus-specific CD4+ lymphocytes,85 which are activated by ongoing virus replication. As a result, CD4+ helper T cells needed to support antibody and CD8+ T-cell responses are depleted. A similar mechanism may apply to FIV, which uses CD134 to infect CD4+ lymphocytes in cats.

Escape Variants The error-prone nature of reverse transcription coupled with high levels of persistent virus replication generates an enormous pool of genetic diversity within a single infected host for the selection of immunological escape variants. The emergence of lentiviral variants that escape neutralizing antibodies was first described for EIAV, MVV, and CAEV. Subsequent work has provided exquisite detail on the emergence of SIV and HIV-1 variants that escape both antibody and CD8+ T-cell responses.32,38,93,285,373

Virulence Not all lentiviral infections are uniformly pathogenic. Although a few instances of AIDS-like disease have been reported in African monkeys naturally infected with SIV,209,229,257 the primate lentiviruses are generally not associated with disease in their natural hosts. Lifelong subclinical infections have also been documented for EIAV and MVV. Breeds of horses and sheep vary in their susceptibility to EIAV and MVV and may even vary in the degree of susceptibility to specific disease manifestations. For example, the classic CNS form of MVV infection originally observed in Iceland is only rarely seen elsewhere. Some strains of SIV are also much less pathogenic than others in susceptible macaque monkeys, and certain strains attenuated by deletion mutations appear to be nonpathogenic.

The diseases associated with lentiviral infections are typically chronic and manifest over a prolonged period. However, there are some prominent exceptions. SIVsmPbj14 is acutely lethal in rhesus monkeys. Monkeys infected with SIVsmPBj14 typically die within 14 days with very high viral loads, severe gastrointestinal disease, cytokine dysregulation, lymphoproliferative disease, and organ system failure.82,99,101 The unusual properties of this strain have been attributed in large part to the presence of a tyrosine residue at position 17 of the Nef protein.87 This tyrosine creates an immunoreceptor tyrosine-based activation motif (ITAM) that imparts on the virus the ability to cause lymphocyte activation and to replicate to high titers in unstimulated PBMC cultures.87 The BIV variant that is the cause of Jembrana disease in Bali cattle also can be acutely pathogenic. About 17% of Bali cattle infected either naturally or experimentally with JDV die with an acute disease within the first few weeks.83,380 During acute disease, infectious titers reach 108/mL of plasma. JDV has remarkable similarities to the disease induced by 747

SIVsmPBj14. There is marked enlargement of lymph nodes and spleen, which feature proliferating lymphoblastoid cells. Proliferating lymphoid infiltrates are also found in many other tissues. The disease in horses induced by EIAV is often considered more acute than that occurring with other lentiviruses. The first episode of anemia usually occurs 2 to 6 weeks after EIAV infection. Subsequent disease cycles are irregular, appearing weeks to months apart, and usually last 3 to 5 days. The frequency and severity of disease episodes usually declines with time, and usually ends within the first year after an average of six to eight episodes. Everything that we know about the pathogenesis of SIV in macaques and HIV-1 in humans points to the importance of viral loads. Whereas high viral loads bode poorly, low viral loads are indicative of a better prognosis. Because sooty mangabeys and African green monkeys do not get sick from their SIVs, most scientists assumed that the viral loads in these species would be low. Quite unexpectedly, they are not.36,114,284 Naturally infected sooty mangabeys and African green monkeys live normal lifespans with plasma viral loads of 105 to 106 RNA copies/mL, levels at which disease progression usually occurs with HIV-1 in humans and SIV in macaques.36,114,167,284 The SIVs infecting these species are fully capable of inducing AIDS when passaged in macaques. Distinguishing features of SIV infection of natural hosts include the absence of chronic immune activation, preservation of mucosal integrity, lack of microbial translocation, maintenance of healthy CD4+ T-cell counts, and preservation of normal lymph node architecture.44 A strong case has been made that chronic immune activation as a consequence of the loss of mucosal integrity and translocation of microbial antigens (e.g., bacterial lipopolysaccharide, peptidoglycans, and nucleic acids) is a major factor contributing to pathogenic HIV-1 and SIV infection.34

Clinical and Pathologic Features Good reviews are available on the clinical and pathologic features of nonhuman lentiviral infections.26,40,161,218,254,264 Some of the most prominent findings are highlighted here.

EIAV Only equine species are susceptible to natural or experimental infection with EIAV. In contrast to infections with HIV, SIV, and FIV, immunosuppression is not a feature of EIAV. Clinical disease is usually divided into acute, subacute, and chronic phases. Acute disease typically results in a fever as high as 108°F 1 to 4 weeks after infection. Anemia is not a prominent feature at the outset. Excessive thirst, loss of appetite, weakness, depression, and hemorrhage are seen in the acute phase. Acute disease may result in death. The subacute form is characterized by relapsing fever and recurrence of other signs. Recurrent episodes may be brought on by hard work or malnutrition. In its chronic form, animals may remain thin despite adequate availability of food, and red cell counts are typically well below normal. Clinical signs in late disease appear to result principally from hemolytic anemia. Erythrocytes of infected horses are coated with antibodies and complement factor 3, and destruction of red cells is immunologically mediated. Osmotic fragility, shortened half-life, and phagocytosis contribute to erythrocyte destruction. Bone marrow may also be depressed, but this seems less important than immune-mediated destruction. Hemorrhage, jaundice, and edema are commonly found at necropsy. The nature and severity of lesions vary with disease course and duration of illness. MVV and CAEV Although maedi (pneumonia) and visna (wasting, depression, paralysis) were once thought to be separate diseases, it is now clear that they are both caused by the same virus. In contrast to infections with HIV, SIV, and FIV, immunosuppression is not a feature of maedi/visna. Polyarthritis and mastitis are also seen as a result of viral infection. Disease is usually seen only in adult sheep because of the lengthy incubation period, typically 3 to 8 years. The lungs of affected sheep may be two to five times their normal weight and exhibit a rubbery loss of elasticity.161 These abnormalities result from a gross thickening of the alveolar walls caused by infiltration and proliferation of reticuloendothelial or mesenchymal cells that invade the septa (Fig. 19.8A and B). Lymph nodules occur along the bronchi and bronchioles. There is progressive weight loss. Dyspnea is initially apparent only after exercise, but it progresses. Severely dyspneic sheep spend much time lying down. Lesions in the brain consist of demyelination and lymphocytic infiltration (Fig. 19.8C and D). Trembling of facial muscles and lips may occur. Clinical signs of visna usually begin with weakness of the hind legs, and this eventually leads to paraplegia. Diseases caused by CAEV in goats are similar to those of MVV in sheep, except arthritis is usually most prominent and pneumonia is usually of lesser severity. Joints are swollen and painful, and this is exacerbated by cold weather. The basic lesion is a proliferative synovitis of joints, tendon sheaths, and bursae.

FIV Primary infection by FIV may lead to low-grade fever, generalized lymphadenopathy, and sometimes diarrhea. During the ensuing months and years, progressing disease is associated with lymphopenia, recurrent fever, lymphadenopathy, anemia, diarrhea, and weight loss of protracted duration. CD4 T-cell counts and other cell subsets may be depressed. The final stages of disease are associated with chronic secondary infections, particularly gingivitis, dermatitis, and infections of the upper respiratory tract. Opportunistic infections that have been observed include calicivirus, herpesviruses, toxoplasma, and cryptococcus. Neurologic abnormalities also have been noted, including dementia, twitching tremors, and convulsions. Pathologic lesions primarily reflect those of opportunistic infection.

748

FIGURE 19.8 Pathology of maedi/visna virus (MVV) in sheep. A and B: Pneumonia. C and D: Encephalitis. (Reprinted with permission from Jones TC, Hunt RD, King NW. Diseases caused by viruses. In: Cann C, ed. Veterinary Pathology. 6th ed. Baltimore, MD: Lippincott Williams & Wilkins; 1997:197–370.) SIV SIV infection of rhesus monkeys is generally considered the closest model of HIV-1 infection of humans. SIV infection of natural hosts (e.g., SIVagm in African green monkeys or SIVsmm in sooty mangabeys) is in general not associated with disease, although there may be occasional exceptions.209,229,257 When SIVsmm was inadvertently introduced into macaque species (Asian Old World primates) in captivity, AIDS-like disease and lymphomas ensued.7,68,219 AIDS-like disease is generally induced experimentally in macaque monkeys with SIVmac, SIVsmm, or less frequently with SIVagm. As with other lentiviruses, SIV establishes a chronic active infection with a prodromal period of months to years before clinical signs appear. Immunodeficiency is usually, but not always, associated with marked declines in CD4+ lymphocyte counts. Macrophages are also infected. Generalized lymphadenopathy typically occurs and is characterized by hyperplasia followed by lymphoid depletion in terminal stages (Fig. 19.9A and B). The gastrointestinal tract, where activated T lymphocytes predominate, appears to be the major site of early viral replication and loss of CD4+ T cells (Fig. 19.9C).362 However, CXCR4-using viruses may cause a profound loss of CD4+ T cells in the periphery that is not paralleled in the intestine.269 Marked depletion of progenitor cells occurs in the thymus by 21 days postinfection; although a rebound occurs subsequently, thymic dysinvolution is typically seen at terminal stages (Fig. 19.9D and E).389 Nodular lymphocytic infiltrates in a variety of tissues (Fig. 19.9C), interstitial pneumonia with syncytial cells (“giant cell pneumonia”) (Fig. 19.9H and I), and granulomatous encephalitis (Fig. 19.9F and G) are variably present. Opportunistic infections usually occur, and these can influence the specific nature of the clinical signs. Common opportunistic infections include Pneumocystis carinii pneumonia, generalized cytomegalovirus infection, cryptosporidiosis, and Mycobacterium avium.

A significant number of AIDS research studies in macaques now utilize recombinant forms of SIVmac with HIV-1 env, tat, and rev genes, referred to as simian-human immunodeficiency viruses (SHIVs). First-generation SHIVs, such as SHIV89.6P, were X4-tropic (use CXCR4 as coreceptor) and acutely pathogenic. However, these SHIVs proved to be paradoxically easy to protect against by vaccination and are no longer considered rigorous challenge viruses for vaccine studies.135 Several pathogenic SHIVs that use CCR5 as a co-receptor have now been developed. A few of these SHIVs result in high viral loads during chronic infection and progressive CD4+ T-cell turnover that resembles the pathogenesis of the SIVmac strains from which they were derived.201,319 These SHIVs have become especially useful for the preclinical evaluation of HIV-1 Env-specific antibodies in macaque models.

Contributions of Individual Genes and Genetic Elements Lentiviruses, like other retroviruses, replicate through a proviral DNA intermediate. Proviral DNA clones that contain a full-length lentiviral genome are therefore sufficient to initiate a spreading infection after physical or chemical delivery into permissive cells. Infectious molecular clones (IMCs) of lentiviruses have been used to study the functions of open reading frames not found in other retroviruses and to gauge their relative importance in the context of experimental animal infection. The first molecular clone of a lentivirus that was not only shown to be infectious but also pathogenic was for SIVmac.171 Pathogenic IMCs were subsequently obtained for EIAV, CAEV, FIV, and other SIV isolates.58,74,138,259 749

Deoxyuridine Triphosphatase A dUTPase reading frame is located within the pol gene of the nonprimate lentiviruses and in the transition region between gag and pol of type D retroviruses. dUTPase catalyzes the conversion of dUTP to dUMP and inorganic pyrophosphate. dUMP is a key precursor of dTTP, which is required for cDNA synthesis during reverse transcription. dUTPase activity also minimizes the mutagenic effects of misincorporation of dUTP into viral DNA. The impact of eliminating the dUTPase open reading frame (DU) has been studied for EIAV, FIV, CAEV, and MVV. In all cases, DU− viruses still replicate in cultured cells. However, the loss of DU dramatically impairs virus replication in nondividing macrophages with low dNTP pools.343,369 Compared to wild-type EIAV, DU− EIAV exhibited 5- to 10-fold lower peak viral loads in plasma, and the pathogenicity of both DU− EIAV and DU− CAEV was attenuated.204,369 Moreover, DU− FIV and DU− CAEV were found to accumulate increased levels of mutations, particularly G-to-A transitions.198,350 Visna viruses lacking DU also showed decreased viral loads in experimentally infected sheep but still produced neuropathogenic effects upon direct intracerebral inoculation.266

Tat Lentiviral Tat proteins can be divided into two groups depending on whether their transactivating activities involve binding to an RNA secondary structure at the 5′ end of nascent viral transcripts known as the transactivation response (TAR) element.31,73,237 Whereas the efficiency of transcription is strongly enhanced for HIV-1, SIV, EIAV, and BIV by their respective Tat proteins in a cyclin T1– and viral RNA– dependent manner, transcription is fully active for MVV and FIV in the absence of any viral proteins. The products of MVV tat and FIV orfA may therefore have functions more similar to those of other accessory proteins.46,108,365 Similar to CD4 downmodulation by Nef, FIV OrfA downmodulates the CD134 receptor of the virus from the surface of infected cells.144 BIV Tat is unusual in that it is able to bind to an RNA sequence element independent of cyclin T1, but transactivation is nonetheless dependent on cyclin T1.31

Rev The lentiviral Rev protein mediates the nuclear export of unspliced viral genomic RNA (gRNA) and a subset of partially spliced, introncontaining mRNAs that encode the Vif, Vpr, Vpx, Vpu, and Env proteins.274 Rev multimerizes on an RNA secondary structure at the 3′ end of the viral genome known as the Rev-responsive element (RRE).274 Interactions between Rev and the chromosomal region maintenance 1 (CRM1) protein (also known as exportin-1) shuttle viral transcripts from the nucleus to the cytoplasm, where Rev and CRM1 disengage to allow protein translation or packaging of gRNA.274 Importantly, Rev and Tat are translated from multiply spliced transcripts and therefore are not dependent on Rev for nuclear export. Although Rev and RRE sequences vary, this mechanism is used by all lentiviruses and is essential for replication.274

S2 The S2 gene is unique to EIAV and encodes a 7-kDa protein that bears no homology to other known retroviral proteins. The S2 protein was found to counteract the effects of SERINC3 and SERINC5 on viral infectivity by excluding these factors from virions.45 Similar to Nef, this activity is dependent on N-terminal myristoylation and a dileucine motif in S2 required for the recruitment of AP-2 for clathrin-mediated endocytosis of SERINC3/5.45 The phenotype of S2-deleted EIAV also resembles the phenotype of nef-deleted SIV. An EIAV derivative lacking S2 replicated normally in fetal equine kidney cells, monocyte-derived macrophages, and differentiated macrophages200 but exhibited lower viral loads and decreased pathogenesis compared to wild-type EIAV in infected horses.199

750

FIGURE 19.9 Acquired immunodeficiency syndrome in monkeys: histopathologic lesions and simian immunodeficiency virus localization. A–C: In situ hybridization for SIV RNA in lymph nodes (A, B) and intestine (C). In (A), numerous individual positive cells can be seen in the paracortex of the lymph node of an animal 8 weeks after infection. In addition, diffuse staining of the germinal center (GC) of a lymphoid follicle consistent with trapping of virus on follicular dendritic cells can be seen. Note that the lymphoid architecture is relatively intact at this early time point, in contrast to a lymph node from an animal with terminal AIDS in (B). Although the lymph node in (B) shows severe lymphoid depletion with no evidence of lymphoid follicles, numerous SIV-infected cells can be seen. In (C), numerous infected cells in the intestine can be seen in a submucosal lymphoid nodule with a rare positive cell (arrow) in the overlying lamina propria of this animal 2 months after infection. D and E: Thymic atrophy. Infection with SIV is associated with profound dysinvolution of the thymus (D), as opposed to a normal thymus (E) with discrete cortex and medulla. F and G: SIV encephalitis. Infection with SIV results in inflammation of the brain in 25% to 50% of the animals that are infected. The brain lesions are characterized by aggregates of mononuclear cells and multinucleated giant cells (F). Cells in the lesions contain abundant SIV nucleic acid by in situ hybridization (G). H and I: Giant cell pneumonia. Multinucleated giant cells in this pneumonia (H) as well as numerous mononuclear cells contain abundant SIV nucleic acid by in situ hybridization (I). (Figure and legend courtesy of Andrew Lackner.) Vif With the exception of EIAV, all lentiviruses have a vif gene (Fig. 19.4 and Tables 19.4 and 19.5). Vif is required to overcome a producer-cell block to virus infectivity imposed by members of the APOBEC3 family of cytidine deaminases.29,205,315 Accordingly, Vif is critical for virus replication in vivo and in cells expressing A3G, A3F, and A3H.80,312 In the absence of Vif, A3G, A3F, and in some cases, A3H become incorporated into virus particles and catalyze cytidine deamination of minus strand DNA during reverse transcription.132,194 This leads to the accumulation of G-to-A transitions in the plus strand that inactivate the viral genome (see “Restriction” section of this chapter). Vif counteracts this restriction by recruiting the cullin-5-elongin B/C-Rbx ubiquitin ligase complex, which mediates the polyubiquitylation and proteasomal degradation of A3 proteins.220,316 By depleting intracellular A3 levels, Vif prevents the incorporation of these proteins into virions.

Nef A nef gene is present in all of the primate lentiviruses but is not found in the nonprimate lentiviruses (Fig. 19.4 and Tables 19.4 and 19.5). The Nef sequences of HIV-1 and SIVsmm/mac correspond in most regions. However, SIVsmm/mac Nef is approximately 40 to 50 amino acids longer due to additional N-terminal sequences that are not found in HIV-1 Nef. A multitude of functional activities have been attributed to Nef. Most of these functions, including CD4 and MHC class I downmodulation, lymphocyte activation, and infectivity enhancement, are shared by the Nef proteins of HIV-1 and SIV. CD4 downmodulation prevents CD4 binding to Env and the exposure of CD4-inducible epitopes that render virus-infected cells susceptible to elimination by antibodies.364 Selective downmodulation of certain MHC class I molecules, but not others, enables HIV- and SIV-infected cells to evade recognition and destruction by virus-specific CD8+ T cells, while simultaneously inhibiting NK cell responses.54,77,338 Infectivity enhancement reflects the downmodulation of SERINC5, and to a lesser extent SERINC3, which prevents the incorporation of these factors into virions.287,352

Additional functions of the Nef proteins of SIV and HIV-2 have been identified that are not shared by HIV-1 Nef. The Nef proteins of HIV-2 and diverse SIVs down-regulate the T-cell receptor (TCR) through interactions with the CD3ζ chain.146,304 TCR downmodulation reduces the sensitivity of infected CD4+ T cells to activation and has been proposed to contribute to the nonpathogenic nature of SIV infection in natural 751

hosts.304 The Nef proteins of most SIVs, with the exception of SIVgsn/mon/mus, also counteract restriction by the tetherin proteins of their simian hosts by downmodulating tetherin from the surface of infected cells.156,298,402 This activity is dependent on a five-amino acid sequence that is present in the cytoplasmic domain of the tetherin orthologs of apes and Old World monkeys but is missing from human tetherin.156,402 Recent structural studies have revealed that Nef acts as a cargo adaptor for clathrin-mediated endocytosis of cellular transmembrane proteins.37,157,185 By using different surfaces and conformations to form multimeric complexes with the cytoplasmic domains of specific cargo proteins and AP-1 or AP-2 subunits,37,157,185 Nef is able to selectively downmodulate several different transmembrane proteins from the cell surface, which helps to explain how Nef is able to serve so many functions.

Nef also plays a role in lymphocyte activation. The Nef proteins of HIV-1 and SIV potentiate NF-κB activation, which enhances virus replication by stimulating viral gene transcription from the LTR promoter.297 Nef additionally increases IL-2 production from infected cells, and nefdeleted viruses are accordingly more dependent on exogenous IL-2 for replication.1,113 Polymorphisms in Nef can also impact cellular activation, as illustrated by the identification of an ITAM in SIVpbj14 Nef that accounts for the unusual ability of SIVpbj14 to replicate in unstimulated CD4+ T cells and for the highly pathogenic phenotype of this virus in macaques.87

Nef is not essential for virus replication in vitro but is important for efficient virus replication and pathogenesis in infected hosts. SIV strains with deletions in nef are highly attenuated compared with wild-type virus.172 Peak viral loads are typically two to three logs lower during acute infection, and set-point viral loads are frequently undetectable during chronic infection with nef-deleted SIV. Most monkeys infected with SIVmac239∆nef have survived without disease and with undetectable viral loads for as long as they have been studied. However, despite marked attenuation, SIVmac239∆nef infection is clearly persistent. Approximately 10% to 20% of monkeys infected with SIVmac239∆nef develop moderate viral loads and eventually progress to AIDS as a result of the accumulation of genetic changes by the virus that restore a pathogenic phenotype.2 Among these changes, substitutions in the gp41 cytoplasmic domain of serially passaged, nef-deleted SIV have been shown to restore the ability to counteract tetherin.311 The phenotype of nef-deleted HIV-1 in humans appears to be similar to nef-deleted SIV in monkeys. Although infection with nef-deleted HIV-1 is clearly attenuated,76,174 some individuals have developed signs of disease progression.125,193

Vpr A vpr gene is present in all primate lentiviruses and encodes a protein of approximately 14-kDa that is specifically incorporated into virions through binding interactions with the p6 domain of Gag. Vpr directs a number of cellular proteins for proteasomal degradation by recruiting the cullin-4A–containing E3 ubiquitin ligase complex (CRL4) through binding interactions with VprBP (DCAF1).147,191,308 Several DNA repair enzymes have been identified as targets of Vpr-mediated degradation, including the nuclear isoform of uracil–DNA glycosylase (UNG2),33,149 helicase-like transcription factor (HTLF),149 exonuclease 1 (EXO1),391 the structure-specific endonuclease regulator complex (SLX4),188 and the methylcytosine dioxygenase TET2.216 Degradation of the base excision repair enzymes UNG2 and TET2 may prevent cDNA products of reverse transcription from triggering DNA repair mechanisms as a consequence of high levels of uracil incorporation owing to elevated dUTP pools in macrophages.149 Vpr-mediated degradation of HTLF and EXO1 also affords an advantage to HIV-1 replication in T cells,392 consistent with the idea that the turnover of these enzymes prevents the processing of branched cDNA intermediates of reverse transcription.

Vpr additionally targets the coiled-coil domain–containing-137 (CCDC137) protein for proteasomal degradation.401 Although the cellular function of CCDC137 is poorly understood, the degradation of this chromosome periphery protein is sufficient to account for the long-standing observation that Vpr induces G2/M cell cycle arrest.401 Whereas CCDC137 depletion increases HIV-1 gene expression by approximately 10-fold in macrophages, it only increases viral gene expression by approximately twofold in CD4+ lymphocytes.401 These observations are consistent with a greater effect of vpr deletion on virus replication in macrophages than in CD4+ lymphocytes16,57 and the modest attenuation of SIVmac239∆vpr replication in rhesus macaques.110 Interestingly, HIV-2 Vpr does not degrade human HTLF or UNG2,149 only partially depletes CCDC137,401 and SIVmac Vpr does not degrade human CCDC137,401 which suggests species-specific differences in these activities.

Vpx Vpx and vpr are related by gene duplication. However, in contrast to vpr, which is present in all of the primate lentiviruses, the vpx gene is only found in a subset of primate lentiviruses (Table 19.5). Like Vpr, Vpx is packaged into virions through interactions with the p6 domain of Gag and mediates the proteasomal degradation of cellular proteins by binding to DCAF1 and recruiting CRL4.335 Vpx is required for efficient SIVsmm/mac replication in terminally differentiated, nondividing myeloid cells (macrophages and dendritic cells) and in resting CD4+ lymphocytes. Vpx can also enhance HIV-1 infection of these cell types. This phenotype reflects the degradation of SAMHD1,15,148,189 a triphosphohydrolase that maintains dNTP concentrations in nondividing cells below the threshold required to complete reverse transcription.115 Accordingly, deletion of vpx has a greater effect than deletion of vpr on the attenuation of SIV replication in macaques. Nevertheless, more than half of the animals infected with SIVmac239∆vpx progress to AIDS.110 An analysis of necropsy tissue from animals infected with vpx-deleted SIV revealed little or no infected macrophages at the time of death indicating that AIDS can occur without significant macrophage infection.376

In addition to SAMHD1, the Vpx proteins of SIVsmm/mac and HIV-2 degrade the transgene activation suppressor (TASOR) subunit of the human silencing hub (HUSH).51 The HUSH complex recruits an H3K9me3 methyltransferase implicated in the epigenetic repression of retroelements. Vpx-mediated depletion of HUSH reduces H3K9me3 chromosomal marks and reactivates transcription from latent HIV-1 proviruses.51 The effects of Vpx are stronger for SIVmac LTR–driven gene expression, suggesting that this activity is especially important for the replication of SIVsmm/mac and HIV-2.51 However, whereas the Vpx proteins of SIVrcm and SIVmnd2 are unable to degrade human TASOR, the Vpr proteins of two phylogenetically distinct SIVs (SIVagm and SIVlst) can degrade human TASOR and activate HIV-1 transcription.51 These observations suggest species specificity in the antagonism of HUSH and that this function also preceded evolution the of vpx gene.

Env Env encodes an extensively glycosylated viral protein that is cleaved into surface (SU) and transmembrane (TM) subunits by furin proteases in the Golgi. Three SU and three TM subunits (which correspond to gp120 and gp41 for the primate lentiviruses) associate noncovalently to form 752

Env trimers on the surface of virions. Env binding to receptor(s) on the surface of susceptible cells induces conformational changes that mediate membrane fusion and virus entry. As the only viral protein exposed on the surface of infected cells and virions, it is the only viral-encoded target of antibodies that can neutralize viral infectivity and eliminate virus-infected cells by Fc-mediated mechanisms. A distinguishing feature of lentiviral envelope glycoproteins is the unusual length of their cytoplasmic tails. These sequences range in length from approximately 50 amino acids for FIV to over 200 amino acids for EIAV and are typically 150 and 164 amino acids for HIV-1 and SIVsmm/mac, respectively. The cytoplasmic domains (CDs) of the primate lentiviral Env proteins have been studied most extensively and are known to participate in the trafficking and incorporation of Env into virions.277 Both HIV-1 and SIV Env contain a highly conserved, membrane-proximal, tyrosine-based motif in their CDs that serves as an AP-2–binding site for clathrin-mediated endocytosis.24,398 The HIV-1 CD also contains a motif implicated in Rab11-family interacting protein 1C (FIP1C)- and Rab14-dependent sorting of Env to sites of virus assembly.280,281 The incorporation of SIV Env into virions is not dependent on FIP1C, however,177 and SIV replication in human CD4+ T cells selects for CD truncations,182 suggesting species-specific differences in Env trafficking. Although a direct physical interaction between the Env CD and the viral matrix (MA) protein has been difficult to confirm biochemically, genetic evidence supports an interaction between the HIV-1 Env CD and MA.240,341 The CDs of HIV-1 and SIV Env may also enhance virus replication in minimally activated CD4+ T cells by stimulating NF-κB activation.276

U3 The U3 regions of the primate lentiviruses are unusually long compared with their counterparts in the nonprimate lentiviruses.153 Most of this length can be accounted for by the overlap of nef coding sequences with the U3 region of the LTR. This is supported by several observations. In monkeys infected with nef-deleted SIV missing 182 bp in the region that is unique to nef, the virus consistently loses approximately 300 bp of sequence in the nef-U3 overlap.175 However, the 12 terminal nucleotides of U3 required for integration and an approximately 50 bp sequence at the 3′ end of nef immediately upstream of the NF-κB binding site are retained.152,221,273 In one case of human infection with a nef-deleted HIV-1 variant, sequences in the region of nef-U3 overlap were also progressively lost over time, while terminal U3 nucleotides and an approximately 50 bp sequence upstream of the NF-κB site were retained.174

NF-κB and Sp1 Sequence elements in the U3 region of the LTR for binding NF-κB and Sp1 transcription factors have been defined as major promoter elements for HIV-1, HIV-2, and SIV. Nevertheless, the removal of NF-κB or Sp1 binding sites from both LTRs surprisingly did not detectably attenuate SIVmac239 replication in rhesus monkeys.152 The removal of both NF-κB and Sp1 binding sites did attenuate SIVmac239 but not markedly so.154 This is likely to be predominantly or exclusively due to an enhancer element present within the 50 bp sequence immediately upstream of the NF-κB binding region that coincides with the nef coding region.152,221,273 Although HIV-1 replication appears more heavily dependent on the presence of the NF-κB and Sp1 sites,289 there is also evidence for an enhancer element upstream of the NF-κB binding sequence of this virus.88,174

Genetic Resistance Host genetic determinants that influence resistance to disease occur for all lentiviruses. The best documented examples come from studies of HIV-1 in humans because of the availability of large, well-characterized patient cohorts. Differences in susceptibility to HIV-1 infection and progression to disease are associated with polymorphisms in many genes, but particularly in the CCR5, HLA, and KIR genes (see Chapter 17). Among the nonhuman lentiviruses, genetic resistance is best characterized for SIV infection of the rhesus macaque because of the extensive use of this species as an animal model. Certain MHC class I alleles have been associated with the control of SIV replication in rhesus macaques. Mamu (Macaca mulatta)-A*01, a common MHC class I allele present in 22% of Indian origin rhesus macaques, is associated with a fivefold reduction in chronic phase viral loads.238 Mamu-B*08 and Mamu-B*17, which are present in 6% and 11% of Indian origin rhesus macaques, respectively, are significantly overrepresented among elite controllers (animals that contain plasma viremia below 1,000 copies/mL).212,393 The protective effect of MamuB*08 is particularly strong and is associated on average with more than a sevenfold reduction in chronic phase viral loads.212 The protective effect of Mamu-B*17 is less consistent, since viral loads in Mamu-B*17–positive animals vary considerably.385 Interestingly, Mamu-B*08 binds a similar set of peptides as HLA-B*2705, which is associated with the control of HIV-1 replication in humans.213 CD8 depletion studies and studies of CD8+ T-cell escape suggest that these MHC class I associations primarily reflect virus-specific CD8+ T-cell responses.

TRIM5 polymorphisms are also associated with differences in the ability of macaques to contain the replication of some SIV strains. SIVsmE543-3 and SIVsmE660 were derived by experimental infection of rhesus macaques with SIVsmm isolated from sooty mangabeys, and as a consequence of the incomplete adaptation to rhesus macaques, they exhibit considerable animal-to-animal variation in both acute and chronic phase viral loads. This variation in viral loads reflects differences in susceptibility to restriction by TRIM5.176 Thus, the sensitivity of SIVsmE543-3 and SIVsmE660 to restrictive variants of rhesus macaque TRIM5 is an important consideration for the interpretation and design of animal studies using these viruses. SIVsmm replication in rhesus macaques expressing restrictive variants of TRIM5 nevertheless selects for amino acid changes in capsid that confer resistance to these variants at later stages of infection. These capsid changes can be engineered back into SIVsmm to generate infectious molecular clones that bypass the confounding effects of TRIM5 genotype in animal studies.386

DIAGNOSIS Because lentiviral infections are persistent, antiviral antibodies are present throughout the lifetime of the infected host. Detection of antibodies to viral antigens is thus the most widely used method for diagnosing viral infection. A variety of methods are commonly used for antibody detection. These include enzyme-linked immunosorbent assay (ELISA), Western blot, gel diffusion, indirect immunofluorescence, hemagglutination, complement fixation, and neutralization assays. The commonly used Coggins test for the detection of EIAV infection is a gel diffusion assay for the detection of antiviral antibodies. ELISA methods for the detection of antibodies to FIV are available. It is estimated that 2% to 3% of cats in the United States are FIV positive. ELISA and Western blots are most commonly used for the detection of antibodies to SIV; 753

detection by ELISA is usually routine but can be complicated by the history of the monkey, whether the antibodies are to the same or a different type of SIV (i.e., cross-reactive), and in an experimental setting, the presence of antibodies at low levels due to attenuation or intervention. Positivity can be confirmed by virus isolation or by identification of viral antigens or viral RNA in plasma or cells. The presence of specific clinical signs and clear demonstration of the presence of virus-specific antibodies is usually sufficient for a definitive diagnosis.

PREVENTION AND CONTROL MVV was eradicated from Iceland by a drastic slaughter policy before the availability of diagnostic tests. Test and removal programs, either voluntary or mandated, continue to be used as an effective means of control. Buyers of horses have increasingly sought negative test certification for EIAV, and negative test certification is required as a condition for entry to many racetracks, sales yards, and shows. Horses imported into the United States and some other countries are required to have a negative test certificate. Testing within a state is not always compulsory, nor is it compulsory for an owner to destroy a positive horse. For FIV, testing is routinely available for cats under veterinary care, and animals in shelters are also routinely screened. However, test and removal programs and certification at the point of sale have been sparingly applied to FIV. Two vaccines have been used in the field for the prevention of lentivirus infections. A live attenuated EIAV strain was developed by researchers by repeated passage of the virus in donkey cells.317 This EIAV vaccine has been extensively used in China and in Cuba, where it has been administered to millions of horses with apparent safety and efficacy. An accumulation of nucleotide substitutions rather than deletions appears to be the basis for the attenuation of EIAV.370 An EIAV vaccine virus derived from a single proviral clone did not fare as well in vaccine challenge experiments as the actual vaccine, which contains extensive sequence diversity.217 The live attenuated EIAV vaccine was administered to 61 million horses and mules in China from 1975 to the 1990s, and this nationwide vaccination program ended the incidence of equine infectious anemia in the country.371 A vaccine against FIV consisting of two inactivated virus subtypes was available in the United States and Canada from 2002 to 2017278 but has been discontinued because of questionable efficacy.336

RESEARCH ON VACCINE DEVELOPMENT One important application of nonhuman lentiviruses is in the area of vaccine research. The development of a safe and effective AIDS vaccine is certainly one of the greatest challenges of our time. Yet, after more than three decades of research, we still lack the basic scientific knowledge needed to achieve this goal. While vaccines designed to stimulate virus-specific T-cell responses can reduce viral loads in nonhuman primate models, at least under conditions where protection is assessed against viruses closely matched to the vaccine and the time of challenge is only a few weeks after the last vaccine dose, conventional T-cell responses alone are not sufficient to prevent infection. On the other hand, passive antibody transfer experiments have shown that potent broadly neutralizing antibodies that are able to bind with high affinity to conserved, conformational features of the HIV-1 envelope glycoprotein can afford complete protection. However, it is presently unclear how to elicit these types of antibodies by vaccination. A myriad of vaccine approaches have been evaluated in macaque models, including many prime and boost vaccine regimens with different combinations of recombinant DNA, poxviral, or adenoviral vectors. While pathogenic strains of SIV have been used most commonly as challenge viruses, SHIVs have also been extensively used to assess protection by vaccines designed to elicit antibodies to the HIV-1 envelope glycoprotein. Rather than attempting to cover the multitude of vectors and prime-boost vaccine regimens that have been tested over the years, we will focus on a few approaches that highlight key concepts or promising advances. Early expectations were raised when inactivated whole SIV was found to provide protection against pathogenic strains of SIV.81,241 Hope was quickly dashed, however, when it was found that protection occurred only when the vaccine and challenge stocks were grown in human cells.63,64 When the vaccine was prepared in human cells and the challenge virus was grown in monkey cells, protection was not observed. Human cellular antigens present in virus preparations appear to have conferred protection when the challenge virus was grown in the same human cells. MHC class II antigens were present in greater abundance in virus particles than the viral envelope glycoprotein and may have been among the xenoantigens that contributed to protection.12,13

There are good reasons for believing that development of an effective vaccine against HIV is going to be a very difficult task. First and foremost, despite enormous antibody and cellular responses to HIV-1, these immune responses do not stop continuous virus replication and CD4+ T-cell destruction. Not only do these immune responses not control virus replication in an already infected individual, they are routinely unable to protect against superinfection by a different strain of the virus.268 HIV-1 field isolates have extraordinary genetic variability, and neutralizing antibody responses are typically strain specific285; methods by which a vaccine could deal with this diversity have yet to be devised. These difficulties have been borne out through seven large-scale efficacy trials in humans, the most recent of which was another disappointing failure.124

Vaccine studies in animal models can provide useful information in several ways. Head-to-head comparisons of different vaccine approaches can shed light on which approaches perform more effectively, at least under defined experimental conditions. In-depth analyses of individual vaccine approaches also may provide fundamental insights into immunologic control and what is needed for protective immunity. With these goals in mind, vaccine approaches that have been tried in monkey models include a variety of Env subunit approaches, replication-competent and replication-defective recombinant viruses, single-cycle SIV, recombinant DNA and RNA, and prime-boost protocols that use combinations of these approaches. A few themes arising from these studies are worth noting. The particular virus that is used for challenge is one of the most important factors that determine whether or not a vaccine/challenge study will be successful. Easy-to-neutralize, nonpathogenic challenge viruses have proven relatively easy to protect against. Difficult-to-neutralize, pathogenic viruses, which are considered representative of naturally transmitted HIV-1 field isolates, have proven very difficult to protect against. A homologous cloned virus challenge is less stringent than a closely matched, uncloned virus challenge, which in turn is much less stringent than a challenge with a virus with natural levels of sequence divergence. The timing of the challenge is also a factor. Many of the vaccine approaches mentioned above induce transient immune responses that decline dramatically after peaking a few weeks after the last vaccine boost. Other important considerations are the dose and route of challenge. High doses of challenge virus are more difficult to protect against than low challenge doses and it is more difficult to protect against intravenous challenge than mucosal challenge. For these reasons, the use of repeated, low-dose mucosal inoculation, usually via the rectal mucosa, is now widely used as a challenge model to better approximate sexual transmission of HIV-1. 754

Live attenuated strains of SIV have performed most impressively as vaccines in experimental monkey studies.67,98 Single and combinations of mutations in nef, vpr, vpx, and LTR sequences have been used. In general, the ability to achieve protection has varied inversely with the degree of attenuation.160 While protection against homologous challenge with SIVmac239 has been impressive, only minimal protection has been observed against challenge with a heterologous strain of SIV.388 Nevertheless, because of the potential for live attenuated viruses to regain a pathogenic phenotype as a result of the accumulation of genetic changes,2 this approach is not under serious consideration for clinical use in people. Interesting results have also been obtained with recombinant herpesviruses. Herpesviruses persist for life, and immune responses to their antigens are maintained in an active state. Herpesviruses have large genomes and can accommodate large amounts of genetic information. A lab-adapted strain of rhesus macaque cytomegalovirus (rhCMV) called 68-1 has been used as a vaccine vector to express SIV antigens in monkeys. These recombinants stimulate unconventional CD8+ T-cell responses with extraordinary breadth and unusual restriction characteristics. More than half of the monkeys immunized with such rhCMV recombinants have exhibited early and complete virologic control following SIVmac239 challenge.130,131 A gamma-2 herpesvirus of rhesus monkeys, rhesus rhadinovirus (RRV), has also been used to deliver a near-full-length SIV genome that expresses all nine SIV gene products and assembles non-infectious virus particles. Protection against SIVmac239 acquisition has been observed with this vaccine strain following challenge by both mucosal and intravenous routes.120,224 The use of adeno-associated virus (AAV) as a vector to achieve long-term delivery of potent broadly neutralizing monoclonal antibodies is another promising approach being intensively studied in rhesus monkey models. When delivered to muscle, the muscle cells can become factories for long-term antibody production. Protection against SIV and SHIV challenges has been observed with this approach.96,106,159 However, antidrug antibody (ADA) responses to vectored antibodies or antibody-like molecules can severely impair antibody delivery. ADA responses remain a major challenge to realizing the potential of this approach.223

RESEARCH ON THERAPEUTIC REGIMENS Antiviral drugs used against HIV-1 in people have been developed with little or no input from animal models of lentiviral infection. However, animal models can provide valuable information for investigation of certain types of therapeutic intervention. For example, there is currently great interest in the idea of using so-called latency reversing agents (LRAs) in combination with antiretroviral therapy (ART) to reactivate viral gene expression in latently infected cells so that these cells can be targeted and eliminated by antiviral responses. LRAs are highly exploratory, and monkey models are quite reasonably being used to examine their feasibility.247 A number of companies are also exploring approaches to long-acting ART formulations in an attempt to reduce the frequency with which these drugs must be taken, and monkey models are being used to address their feasibility.6 Many of the antiretroviral drugs developed for use against HIV-1 in humans are less potent against SIV, and the bioavailability of some of these drugs can also differ between humans and macaques.5 After extensive testing, the current “gold standard” ART regimen for suppressing SIV replication in monkeys is a daily subcutaneous injection of tenofovir, emtricitabine, and dolutegravir.78 Reverse transcriptase (RT)-SHIVs, in which the RT reading frame of SIV has been replaced with that of HIV-1, have been developed.4,351 Macaques infected with these RT-SHIVs have been used to study the emergence of drug resistance mutations.5,351 A minimally modified simian-tropic HIV-1 that replicates in pigtail macaques may also be useful for therapeutic studies in primate models.305 Early studies showed that treatment of SIV-infected monkeys with tenofovir (PMPA; R-9-2-phosphonylmethoxypropyl adenine) for 4 weeks starting at 24 hours after experimental SIV infection resulted in impressive virological control after discontinuing drug treatment.206,348 Delaying the initiation of therapy to 48 or 72 hours, or shortening the duration of treatment, significantly reduced efficacy.347 Interestingly, when three such animals off therapy for 6 weeks were subsequently challenged intravenously with SIVsmE660, one animal was completely protected, and the other two showed dramatic reductions in viral load.207 What is most startling and difficult to understand is that protection was achieved in these animals in the absence of readily measurable SIV-specific antibody or cellular immune responses at the time of challenge. Continuation of this line of investigation could provide important insights into immune-mediated control outside the bounds of our current level of understanding. A variety of gene therapy strategies for inhibiting lentiviral replication and depleting viral reservoirs are also being tested in nonhuman primate models. Recent approaches include editing of the CCR5 gene in hematopoietic stem cells (HSCs) and transduction of HSCs with chimeric antigen receptors (CARs).17,41,397 These approaches rely on either the differentiation of HSCs into CCR5-deleted T cells that are resistant to HIV-1 and SHIV infection or into cytotoxic T lymphocytes expressing CARs that allow them to recognize and kill HIV-infected cells. Another promising gene therapy approach that is currently under investigation in macaque models is the use of AAV vectors for long-term delivery of potent broadly neutralizing antibodies.223

PERSPECTIVE The global AIDS crisis brought on by HIV-1 has focused attention on nonhuman lentiviruses as a source of information that will shed light on the human condition. Lentiviral infections of domesticated animals are economically important in their own right and have been studied historically in this context. The lack of a practical animal model for infection with HIV-1 that recapitulates features of human infection has led to the extensive investigation of nonhuman lentiviruses in naturally or experimentally infected hosts as analog model systems. Critical areas for future progress include a better understanding of lentiviral pathogenesis, improvements in therapy, and perhaps most importantly, the development of a safe, effective, long-acting, and affordable vaccine or other preventative measure. The most remarkable feature of lentiviruses, and perhaps the most important for the eventual control of HIV-1, is their ability to replicate continuously and unrelentingly in the presence of strong immune responses. The advancement of therapeutic regimens and the development of effective vaccines against HIV-1 will ultimately need to deal with the propensity of this group of viruses to generate and tolerate genetic changes that enable the evasion of host immune responses.

755

References 1. Alexander L, Du Z, Rosenzweig M, et al. A role for natural simian immunodeficiency virus and human immunodeficiency virus type 1 nef alleles in lymphocyte activation. J Virol 1997;71:6094–6099. 2. Alexander L, Illyinskii PO, Lang SM, et al. Determinants of increased replicative capacity of serially passaged simian immunodeficiency virus with nef deleted in rhesus monkeys. J Virol 2003;77:6823–6835. 3. Allan JS, Kanda P, Kennedy RC, et al. Isolation and characterization of simian immunodeficiency viruses from two subspecies of African green monkeys. AIDS Res Hum Retroviruses 1990;6:275–285. 4. Ambrose Z, Boltz V, Palmer S, et al. In vitro characterization of a simian immunodeficiency virus-human immunodeficiency virus (HIV) chimera expressing HIV type 1 reverse transcriptase to study antiviral resistance in pigtail macaques. J Virol 2004;78:13553–13561. 5. Ambrose Z, Palmer S, Boltz VF, et al. Suppression of viremia and evolution of human immunodeficiency virus type 1 drug resistance in a macaque model for antiretroviral therapy. J Virol 2007;81:12145–12155. 6. Andrews CD, Bernard LS, Poon AY, et al. Cabotegravir long acting injection protects macaques against intravenous challenge with SIVmac251. AIDS 2017;31:461–467. 7. Apetrei C, Kaur A, Lerche NW, et al. Molecular epidemiology of simian immunodeficiency virus SIVsm in U.S. primate centers unravels the origin of SIVmac and SIVstm. J Virol 2005;79:8991–9005. 8. Apps R, Del Prete GQ, Chatterjee P, et al. HIV-1 Vpu mediates HLA-C downregulation. Cell Host Microbe 2016;19:686–695. 9. Arias JF, Colomer-Lluch M, von Bredow B, et al. Tetherin Antagonism by HIV-1 Group M Nef Proteins. J Virol 2016;90:10701–10714. 10. Arias JF, Heyer LN, von Bredow B, et al. Tetherin antagonism by Vpu protects HIV-infected cells from antibody-dependent cell-mediated cytotoxicity. Proc Natl Acad Sci U S A 2014;111:6425–6430. 11. Arnold LH, Groom HC, Kunzelmann S, et al. Phospho-dependent Regulation of SAMHD1 Oligomerisation Couples Catalysis and Restriction. PLoS Pathog 2015;11:e1005194. 12. Arthur LO, Bess JW Jr, Sowder RC II, et al. Cellular proteins bound to immunodeficiency viruses: implications for pathogenesis and vaccines. Science 1992;258:1935–1938. 13. Arthur LO, Bess JW Jr, Urban RG, et al. Macaques immunized with HLA-DR are protected from SIV challenge. J Virol 1995;69:3117–3124. 14. Bailes E, Gao F, Bibollet-Ruche F, et al. Hybrid origin of SIV in chimpanzees. Science 2003;300:1713. 15. Baldauf HM, Pan X, Erikson E, et al. SAMHD1 restricts HIV-1 infection in resting CD4(+) T cells. Nat Med 2012;18:1682–1687. 16. Balliet JW, Kolson DL, Eiger G, et al. Distinct effects in primary macrophages and lymphocytes of the human immunodeficiency virus type 1 accessory genes vpr, vpu, and nef: mutational analysis of a primary HIV-1 isolate. Virology 1994;200:623–631. 17. Barber-Axthelm IM, Barber-Axthelm V, Sze KY, et al. Stem cell-derived CAR T cells traffic to HIV reservoirs in macaques. JCI Insight 2021;6(1):e141502. 18. Barre-Sinoussi F, Chermann JC, Rey F, et al. Isolation of a T-lymphotropic retrovirus from a patient at risk for acquired immune deficiency syndrome (AIDS). Science 1983;220:868–871. 19. Beaumier CM, Harris LD, Goldstein S, et al. CD4 downregulation by memory CD4+ T cells in vivo renders African green monkeys resistant to progressive SIVagm infection. Nat Med 2009;15:879–885. 20. Beer BE, Bailes E, Goeken R, et al. Simian immunodeficiency virus (SIV) from sun-tailed monkeys (Cercopithecus solatus): evidence for host-dependent evolution of SIV within the C. lhoesti superspecies. J Virol 1999;73:7734–7744. 21. Beer BE, Foley BT, Kuiken CL, et al. Characterization of novel simian immunodeficiency viruses from red-capped mangabeys from Nigeria (SIVrcmNG409 and -NG411). J Virol 2001;75:12014–12027. 22. Beisel CE, Edwards JF, Dunn LL, et al. Analysis of multiple mRNAs from pathogenic equine infectious anemia virus (EIAV) in an acutely infected horse reveals a novel protein, Ttm, derived from the carboxy terminus of the EIAV transmembrane protein. J Virol 1993;67:832–842. 23. Benveniste RE, Arthur LO, Tsai C-C, et al. Isolation of a lentivirus from a macaque with lymphoma: comparison with HTLV-III/LAV and other lentiviruses. J Virol 1986;60:483–490. 24. Berlioz-Torrent C, Shacklett BL, Erdtmann L, et al. Interactions of the cytoplasmic domains of human and simian retroviral transmembrane proteins with components of the clathrin adaptor complexes modulate intracellular and cell surface expression of envelope glycoproteins. J Virol 1999;73:1350–1361. 25. Betancor G, Dicks MDJ, Jimenez-Guardeno JM, et al. The GTPase domain of MX2 interacts with the HIV-1 capsid, enabling its short isoform to moderate antiviral restriction. Cell Rep 2019;29:1923–1933 e3. 26. Bhatia S, Sood R. Bovine immunodeficiency virus. In: Bayry J, ed. Emerging and Re-emerging Infectious Diseases of Livestock. Switzerland, Cham: Springer; 2017:301–308. 27. Bibollet-Ruche F, Bailes E, Gao F, et al. New simian immunodeficiency virus infecting De Brazza's monkeys (Cercopithecus neglectus): evidence for a cercopithecus monkey virus clade. J Virol 2004;78:7748–7762. 28. Bibollet-Ruche F, Galat-Luong A, Cuny G, et al. Simian immunodeficiency virus infection in a patas monkey (Erythrocebus patas): evidence for cross-species transmission from African green monkeys (Cercopithecus aethiops sabaeus) in the wild. J Gen Virol 1996;77:773–781. 29. Bishop KN, Holmes RK, Sheehy AM, et al. Cytidine deamination of retroviral DNA by diverse APOBEC proteins. Curr Biol 2004;14:1392–1396. 30. Bogerd HP, Doehle BP, Wiegand HL, et al. A single amino acid difference in the host APOBEC3G protein controls the primate species specificity of HIV type 1 virion infectivity factor. Proc Natl Acad Sci U S A 2004;101:3770–3774. 31. Bogerd HP, Wiegand HL, Bieniasz PD, et al. Functional differences between human and bovine immunodeficiency virus tat transcription factors. J Virol 2000;74:4666–4671. 32. Borrow P, Lewicki H, Wei X, et al. Antiviral pressure exerted by HIV-1-specific cytotoxic T lymphocytes (CTLs) during primary infection demonstrated by rapid selection of CTL escape virus. Nat Med 1997;3:205–211. 33. Bouhamdan M, Benichou S, Rey F, et al. Human immunodeficiency virus type 1 Vpr protein binds to the uracil DNA glycosylase DNA repair enzyme. J Virol 1996;70:697–704. 34. Brenchley JM, Price DA, Schacker TW, et al. Microbial translocation is a cause of systemic immune activation in chronic HIV infection. Nat Med 2006;12:1365–1371. 35. Brenchley JM, Schacker TW, Ruff LE, et al. CD4+ T cell depletion during all stages of HIV disease occurs predominantly in the gastrointestinal tract. J Exp Med 2004;200:749–759. 36. Broussard SR, Staprans SI, White R, et al. Simian immunodeficiency virus replicates to high levels in naturally infected African green monkeys without inducing immunologic or neurologic disease. J Virol 2001;75:2262–2275. 37. Buffalo CZ, Sturzel CM, Heusinger E, et al. Structural basis for tetherin antagonism as a barrier to zoonotic lentiviral transmission. Cell Host Microbe 2019;26:359–368 e8. 756

38. Burns D, Collignon, C, Desrosiers, RC. Simian immunodeficiency virus mutants resistant to serum neutralization arise during persistent infection of rhesus monkeys. J Virol 1993;67:4104–4113. 39. Busnadiego I, Kane M, Rihn SJ, et al. Host and viral determinants of Mx2 antiretroviral activity. J Virol 2014;88:7738–7752. 40. Campbell R, Robinson, WF. The comparative pathology of the lentiviruses. J Comp Pathol 1998;119:333–395. 41. Cardozo-Ojeda EF, Duke ER, Peterson CW, et al. Thresholds for post-rebound SHIV control after CCR5 gene-edited autologous hematopoietic cell transplantation. Elife 2021;10:e57646. 42. Carpenter MA, Brown EW, Culver M, et al. Genetic and phylogenetic divergence of feline immunodeficiency virus in the puma (Puma concolor). J Virol 1996;70:6682–6693. 43. Chadwick BJ, Coelen RJ, Wilcox GE, et al. Nucleotide sequence analysis of Jembrana disease virus: a bovine lentivirus associated with an acute disease syndrome. J Gen Virol 1995;76:1637–1650. 44. Chahroudi A, Bosinger SE, Vanderford TH, et al. Natural SIV hosts: showing AIDS the door. Science 2012;335:1188–1193. 45. Chande A, Cuccurullo EC, Rosa A, et al. S2 from equine infectious anemia virus is an infectivity factor which counteracts the retroviral inhibitors SERINC5 and SERINC3. Proc Natl Acad Sci U S A 2016;113:13197–13202. 46. Chatterji U, de Parseval A, Elder JH. Feline immunodeficiency virus OrfA is distinct from other lentivirus transactivators. J Virol 2002;76:9624–9634. 47. Chen YC, Sood C, Marin M, et al. Super-resolution fluorescence imaging reveals that serine incorporator protein 5 inhibits human immunodeficiency virus fusion by disrupting envelope glycoprotein clusters. ACS Nano 2020;14:10929–10943. 48. Chen Z, Kwon D, Jin Z, et al. Natural infection of a homozygous delta24 CCR5 red-capped mangabey with an R2b-tropic simian immunodeficiency virus. J Exp Med 1998;188:2057–2065. 49. Chen Z, Luckay A, Sodora DL, et al. Human immunodeficiency virus type 2 (HIV-2) seroprevalence and characterization of a distinct HIV-2 genetic subtype from the natural range of simian immunodeficiency virus-infected sooty mangabeys. J Virol 1997;71:3953–3960. 50. Chertova E, Bess JW Jr, Crise BJ, et al. Envelope glycoprotein incorporation, not shedding of surface envelope glycoprotein (gp120/SU), is the primary determinant of SU content of purified human immunodeficiency virus type 1 and simian immunodeficiency virus. J Virol 2002;76:5315–5325. 51. Chougui G, Munir-Matloob S, Matkovic R, et al. HIV-2/SIV viral protein X counteracts HUSH repressor complex. Nat Microbiol 2018;3:891–897. 52. Clavel F, Guetaro D, Brun-Vezinet F, et al. Isolation of a new human retrovirus from West African patients with AIDS. Science 1986;233:343–346. 53. Clewley JP, Lewis JC, Brown DW, et al. A novel simian immunodeficiency virus (SIVdrl) pol sequence from the drill monkey, Mandrillus leucophaeus. J Virol 1998;72:10305–10309. 54. Cohen GB, Gandhi RT, Davis DM, et al. The selective downregulation of class I major histocompatibility complex proteins by HIV-1 protects HIV-infected cells from NK cells. Immunity 1999;10:661–671. 55. Collins KL, Chen BK, Kalams SA, et al. HIV-1 Nef protein protects infected primary cells against killing by cytotoxic T lymphocytes. Nature 1998;391:397–401. 56. Compton AA, Hirsch VM, Emerman M. The host restriction factor APOBEC3G and retroviral Vif protein coevolve due to ongoing genetic conflict. Cell Host Microbe 2012;11:91–98. 57. Connor RI, Chen BK, Choe S, et al. Vpr is required for efficient replication of human immunodeficiency virus type-1 in mononuclear phagocytes. Virology 1995;206:935–944. 58. Cook RF, Leroux C, Cook SJ, et al. Development and characterization of an in vivo pathogenic molecular clone of equine infectious anemia virus. J Virol 1998;72:1383–1393. 59. Courgnaud V, Abela B, Pourrut X, et al. Identification of a new simian immunodeficiency virus lineage with a vpu gene present among different cercopithecus monkeys (C. mona, C. cephus, and C. nictitans) from Cameroon. J Virol 2003;77:12523–12534. 60. Courgnaud V, Formenty P, Akoua-Koffi C, et al. Partial molecular characterization of two simian immunodeficiency viruses (SIV) from African colobids: SIVwrc from Western red colobus (Piliocolobus badius) and SIVolc from olive colobus (Procolobus verus). J Virol 2003;77:744–748. 61. Courgnaud V, Pourrut X, Bibollet-Ruche F, et al. Characterization of a novel simian immunodeficiency virus from guereza colobus monkeys (Colobus guereza) in Cameroon: a new lineage in the nonhuman primate lentivirus family. J Virol 2001;75:857–866. 62. Courgnaud V, Salemi M, Pourrut X, et al. Characterization of a novel simian immunodeficiency virus with a vpu gene from greater spotnosed monkeys (Cercopithecus nictitans) provides new insights into simian/human immunodeficiency virus phylogeny. J Virol 2002;76:8298–8309. 63. Cranage MP, Ashworth LAE, Greenaway PJ, et al. AIDS vaccine developments. Nature 1992;355:684–686. 64. Cranage MP, Polyanskaya N, McBride B, et al. Studies on the specificity of the vaccine effect elicited by inactivated simian immunodeficiency virus. AIDS Res Hum Retroviruses 1993;9:13–22. 65. Cui J, Holmes EC. Endogenous lentiviruses in the ferret genome. J Virol 2012;86:3383–3385. 66. Daniel M, Letvin N, Sehgal P, et al. Prevalence of antibodies to 3 retroviruses in a captive colony of macaque monkeys. Int J Cancer 1988;41:601–608. 67. Daniel MD, Kirchhoff F, Czajak SC, et al. Protective effects of a live-attenuated SIV vaccine with a deletion in the nef gene. Science 1992;258:1938–1941. 68. Daniel MD, Letvin NL, King NW, et al. Isolation of T-cell tropic HTLV-III-like retrovirus from macaques. Science 1985;228:1201–1204. 69. D'Arc M, Ayouba A, Esteban A, et al. Origin of the HIV-1 group O epidemic in western lowland gorillas. Proc Natl Acad Sci U S A 2015;112:E1343–E1352. 70. Dazza MC, Ekwalanga M, Nende M, et al. Characterization of a novel vpu-harboring simian immunodeficiency virus from a Dent's Mona monkey (Cercopithecus mona denti). J Virol 2005;79:8560–8571. 71. de Parseval A, Chatterji U, Morris G, et al. Structural mapping of CD134 residues critical for interaction with feline immunodeficiency virus. Nat Struct Mol Biol 2005;12:60–66. 72. de Parseval A, Chatterji U, Sun P, et al. Feline immunodeficiency virus targets activated CD4+ T cells by using CD134 as a binding receptor. Proc Natl Acad Sci U S A 2004;101:13044–13049. 73. de Parseval A, Elder, JH. Demonstration that orf2 encodes the feline immunodeficiency virus transactivating (Tat) protein and characterization of a unique gene product with partial rev activity. J Virol 1999;73:608–617. 74. de Rozieres S, Mathiason CK, Rolston MR, et al. Characterization of a highly pathogenic molecular clone of feline immunodeficiency virus clade C. J Virol 2004;78:8971–8982. 75. de Sousa-Pereira P, Abrantes J, Bauernfried S, et al. The antiviral activity of rodent and lagomorph SERINC3 and SERINC5 is counteracted by known viral antagonists. J Gen Virol 2019;100:278–288. 76. Deacon NJ, Tsykin A, Solomon A, et al. Genomic structure of an attenuated quasi species of HIV-1 from a blood transfusion donor and recipients. Science 1995;270:988–991. 77. DeGottardi MQ, Specht A, Metcalf B, et al. Selective downregulation of rhesus macaque and sooty mangabey major histocompatibility 757

complex class I molecules by Nef alleles of simian immunodeficiency virus and human immunodeficiency virus type 2. J Virol 2008;82:3139–3146. 78. Del Prete GQ, Smedley J, Macallister R, et al. Short communication: comparative evaluation of coformulated injectable combination antiretroviral therapy regimens in simian immunodeficiency virus-infected rhesus Macaques. AIDS Res Hum Retroviruses 2016;32:163–168. 79. Desrosiers RC. HIV-1 origins: A finger on the missing link. Nature 1990;345:288–289. 80. Desrosiers RC, Lifson JD, Gibbs JS, et al. Identification of highly attenuated mutants of simian immunodeficiency virus. J Virol 1998;72:1431–1437. 81. Desrosiers RC, Wyand MS, Kodama T, et al. Vaccine protection against simian immunodeficiency virus infection. Proc Natl Acad Sci U S A 1989;86:6353–6357. 82. Dewhurst S, Embretson JE, Anderson DC, et al. Sequence analysis and acute pathogenicity of molecularly cloned SIVsmmPBj14. Nature 1990;345:636–640. 83. Dharma DM, Budiantono A, Campbell RS, et al. Studies on experimental Jembrana disease in Bali cattle. III. Pathology. J Comp Pathol 1991;105:397–414. 84. Dicks MDJ, Betancor G, Jimenez-Guardeno JM, et al. Multiple components of the nuclear pore complex interact with the amino-terminus of MX2 to facilitate HIV-1 restriction. PLoS Pathog 2018;14:e1007408. 85. Douek DC, Brenchley JM, Betts MR, et al. HIV preferentially infects HIV-specific CD4+ T cells. Nature 2002;417:95–98. 86. Dow SW, Poss ML, Hoover EA. Feline immunodeficiency virus: a neurotropic lentivirus. J Acquir Immune Defic Syndr 1990;3:658–668. 87. Du Z, Lang SM, Sasseville VG, et al. Identification of a nef allele that causes lymphocyte activation and acute disease in macaque monkeys. Cell 1995;82:665–674. 88. Duverger A, Wolschendorf F, Zhang M, et al. An AP-1 binding site in the enhancer/core element of the HIV-1 promoter controls the ability of HIV-1 to establish latent infection. J Virol 2013;87:2264–2277. 89. Elder JH, Lerner DL, Hasselkus-Light CS, et al. Distinct subsets of retroviruses encode dUTPase. J Virol 1992;66:1791–1794. 90. Elliott ST, Wetzel KS, Francella N, et al. Dualtropic CXCR6/CCR5 simian immunodeficiency virus (SIV) infection of sooty mangabey primary lymphocytes: distinct coreceptor use in natural versus pathogenic hosts of SIV. J Virol 2015;89:9252–9261. 91. Emau P, McClure HM, Isahakia M, et al. Isolation from African Sykes’ monkeys (Cercopithecus mitis) of a lentivirus related to human and simian immunodeficiency viruses. J Virol 1991;65:2135–2140. 92. Etienne L, Nerrienet E, LeBreton M, et al. Characterization of a new simian immunodeficiency virus strain in a naturally infected Pan troglodytes troglodytes chimpanzee with AIDS related symptoms. Retrovirology 2011;8:4. 93. Evans DT, O'Connor DH, Jing P, et al. Virus-specific cytotoxic T-lymphocyte responses select for amino-acid variation in simian immunodeficiency virus Env and Nef. Nat Med 1999;5:1270–1276. 94. Ficarelli M, Wilson H, Pedro Galao R, et al. KHNYN is essential for the zinc finger antiviral protein (ZAP) to restrict HIV-1 containing clustered CpG dinucleotides. Elife 2019;8:e46767. 95. Fitzpatrick K, Skasko M, Deerinck TJ, et al. Direct restriction of virus release and incorporation of the interferon-induced protein BST-2 into HIV-1 particles. PLoS Pathog 2010;6:e1000701. 96. Fuchs SP, Martinez-Navio JM, Piatak M Jr, et al. AAV-delivered antibody mediates significant protective effects against SIVmac239 challenge in the absence of neutralizing activity. PLoS Pathog 2015;11:e1005090. 97. Fukasawa M, Muira T, Hasegawa A, et al. Sequence of simian immunodeficiency virus from African green monkey, a new member of the HIV/SIV group. Nature 1988;333:457–461. 98. Fukazawa Y, Park H, Cameron MJ, et al. Lymph node T cell responses predict the efficacy of live attenuated SIV vaccines. Nat Med 2012;18:1673–1681. 99. Fultz PN. Replication of an acutely lethal simian immunodeficiency virus activates and induces proliferation of lymphocytes. J Virol 1991;65:4902–4909. 100. Fultz PN, McClure HM, Anderson DC, et al. Isolation of a T-lymphotropic retrovirus from naturally infected sooty mangabey monkeys (Cercocebus atys). Proc Natl Acad Sci U S A 1986;83:5286–5290. 101. Fultz PN, McClure HM, Anderson DC, et al. Identification and biologic characterization of an acutely lethal variant of simian immunodeficiency virus from sooty mangabeys (SIV/SMM). AIDS Res Hum Retroviruses 1989;5:397–409. 102. Galao RP, Pickering S, Curnock R, et al. Retroviral retention activates a Syk-dependent HemITAM in human tetherin. Cell Host Microbe 2014;16:291–303. 103. Ganser-Pornillos BK, Pornillos O. Restriction of HIV-1 and other retroviruses by TRIM5. Nat Rev Microbiol 2019;17:546–556. 104. Gao F, Yue L, White AT, et al. Human infection by genetically diverse SIVsm-related HIV-2 in west Africa. Nature 1992;358:495–499. 105. Gao G, Guo X, Goff SP. Inhibition of retroviral RNA production by ZAP, a CCCH-type zinc finger protein. Science 2002;297:1703–1706. 106. Gardner MR, Kattenhorn LM, Kondur HR, et al. AAV-expressed eCD4-Ig provides durable protection from multiple SHIV challenges. Nature 2015;519:87–91. 107. Gauduin MC, Yu Y, Barabasz A, et al. Induction of a virus-specific effector-memory CD4+ T cell response by attenuated SIV infection. J Exp Med 2006;203:2661–2672. 108. Gemeniano MC, Sawai ET, Sparger EE. Feline immunodeficiency virus Orf-A localizes to the nucleus and induces cell cycle arrest. Virology 2004;325:167–174. 109. Gendelman HE, Narayan O, Kennedy-Stoskopf S, et al. Tropism of sheep lentiviruses for monocytes: susceptibility to infection and virus gene expression increase during maturation of monocytes to macrophages. J Virol 1986;58:67–74. 110. Gibbs JS, Lackner AA, Lang SM, et al. Progression to AIDS in the absence of genes for vpr or vpx. J Virol 1995;69:2378–2383. 111. Gifford RJ, Katzourakis A, Tristem M, et al. A transitional endogenous lentivirus from the genome of a basal primate and implications for lentivirus evolution. Proc Natl Acad Sci U S A 2008;105:20362–20367. 112. Gilbert C, Maxfield DG, Goodman SM, et al. Parallel germline infiltration of a lentivirus in two Malagasy lemurs. PLoS Genet 2009;5:e1000425. 113. Glushakova S, Grivel JC, Suryanarayana K, et al. Nef enhances human immunodeficiency virus replication and responsiveness to interleukin-2 in human lymphoid tissue ex vivo. J Virol 1999;73:3968–3974. 114. Goldstein S, Ourmanov I, Brown CR, et al. Wide range of viral load in healthy African green monkeys naturally infected with simian immunodeficiency virus. J Virol 2000;74:11744–11753. 115. Goldstone DC, Ennis-Adeniran V, Hedden JJ, et al. HIV-1 restriction factor SAMHD1 is a deoxynucleoside triphosphate triphosphohydrolase. Nature 2011;480:379–382. 116. Gonda M. The lentiviruses of cattle. In: Levy JA, ed. Retroviridae. New York: Plenum Press; 1994. 117. Gonda MA, Braun MJ, Carter SG, et al. Characterization and molecular cloning of a bovine lentivirus related to human immunodeficiency virus. Nature 1987;330:388–391. 118. Gonda MA, Charman HP, Walker JL, et al. Scanning and transmission electron microscopic study of equine infectious anemia virus. Am J Vet Res 1978;39:731–740. 758

119. Gonda MA, Wong-Staal F, Gallo RC, et al. Sequence homology and morphologic similarity of HTLV-III and visna virus, a pathogenic lentivirus. Science 1985;227:173–177. 120. Gonzalez-Nieto L, Castro IM, Bischof GF, et al. Vaccine protection against rectal acquisition of SIVmac239 in rhesus macaques. PLoS Pathog 2019;15:e1008015. 121. Goodier JL, Pereira GC, Cheung LE, et al. The broad-spectrum antiviral protein ZAP restricts human retrotransposition. PLoS Genet 2015;11:e1005252. 122. Gotz N, Sauter D, Usmani SM, et al. Reacquisition of Nef-mediated tetherin antagonism in a single in vivo passage of HIV-1 through its original chimpanzee host. Cell Host Microbe 2012;12:373–380. 123. Goujon C, Moncorge O, Bauby H, et al. Human MX2 is an interferon-induced post-entry inhibitor of HIV-1 infection. Nature 2013;502:559–562. 124. Gray GE, Bekker LG, Laher F, et al. Vaccine efficacy of ALVAC-HIV and bivalent subtype C gp120-MF59 in adults. N Engl J Med 2021;384:1089–1100. 125. Greenough T, Sullivan JL, Desrosiers RC. Declining CD4 T-cell counts in a person infected with nef-deleted HIV-1. N Engl J Med 1999;340:236–237. 126. Gudnadottir M, Palsson PA. Transmission of maedi by inoculation of a virus grown in tissue culture from maedi-affected lungs. J Infect Dis 1967;117:1–6. 127. Hammonds J, Wang JJ, Yi H, et al. Immunoelectron microscopic evidence for Tetherin/BST2 as the physical bridge between HIV-1 virions and the plasma membrane. PLoS Pathog 2010;6:e1000749. 128. Han GZ, Worobey M. Endogenous lentiviral elements in the weasel family (Mustelidae). Mol Biol Evol 2012;29:2905–2908. 129. Han GZ, Worobey M. A primitive endogenous lentivirus in a colugo: insights into the early evolution of lentiviruses. Mol Biol Evol 2015;32:211–215. 130. Hansen SG, Ford JC, Lewis MS, et al. Profound early control of highly pathogenic SIV by an effector memory T-cell vaccine. Nature 2011;473:523–527. 131. Hansen SG, Marshall EE, Malouli D, et al. A live-attenuated RhCMV/SIV vaccine shows long-term efficacy against heterologous SIV challenge. Sci Transl Med 2019;11(501):eaaw2607. 132. Harris RS, Bishop KN, Sheehy AM, et al. DNA deamination mediates innate immunity to retroviral infection. Cell 2003;113:803–809. 133. Harris RS, Sheehy AM, Craig HM, et al. DNA deamination: not just a trigger for antibody diversification but also a mechanism for defense against retroviruses. Nat Immunol 2003;4:641–643. 134. Hatziioannou T, Del Prete GQ, Keele BF, et al. HIV-1-induced AIDS in monkeys. Science 2014;344:1401–1405. 135. Hatziioannou T, Evans DT. Animal models for HIV/AIDS research. Nat Rev Microbiol 2012;10:852–867. 136. Hatziioannou T, Perez-Caballero D, Yang A, et al. Retrovirus resistance factors Ref1 and Lv1 are species-specific variants of TRIM5alpha. Proc Natl Acad Sci U S A 2004;101:10774–10779. 137. Heigele A, Kmiec D, Regensburger K, et al. The potency of Nef-mediated SERINC5 antagonism correlates with the prevalence of primate lentiviruses in the wild. Cell Host Microbe 2016;20:381–391. 138. Hirsch V, Adger-Johnson D, Campbell B, et al. A molecularly cloned, pathogenic, neutralization-resistant simian immunodeficiency virus, SIVsmE543-3. J Virol 1997;71:1608–1620. 139. Hirsch VM, Campbell BJ, Bailes E, et al. Characterization of a novel simian immunodeficiency virus (SIV) from L'Hoest monkeys (Cercopithecus l'hoesti): implications for the origins of SIVmnd and other primate lentiviruses. J Virol 1999;73:1036–1045. 140. Hirsch VM, Dapolito G, Johnson PR, et al. Induction of AIDS by simian immunodeficiency virus from an African green monkey: speciesspecific variation in pathogenicity correlates with the extent of in vivo replication. J Virol 1995;69:955–967. 141. Hirsch VM, Dapolito GA, Goldstein S, et al. A distinct African lentivirus from Sykes' monkeys. J Virol 1993;67:1517–1528. 142. Hirsch VM, Olmsted RA, Murphey-Corb M, et al. An African primate lentivirus (SIVsm) closely related to HIV-2. Nature 1989;339:389–392. 143. Ho DD, Neuman AU, Perelson AS, et al. Rapid turnover of plasma virions and CD4 lymphocytes in HIV-1 infection. Nature 1995;373:123–126. 144. Hong Y, Fink E, Hu QY, et al. OrfA downregulates feline immunodeficiency virus primary receptor CD134 on the host cell surface and is important in viral infection. J Virol 2010;84:7225–7232. 145. Hosie MJ, Broere N, Hesselgesser J, et al. Modulation of feline immunodeficiency virus infection by stromal cell-derived factor. J Virol 1998;72:2097–2104. 146. Howe AY, Jung JU, Desrosiers RC. Zeta chain of the T-cell receptor interacts with nef of simian immunodeficiency virus and human Immunodeficiency virus type 2. J Virol 1998;72:9827–9834. 147. Hrecka K, Gierszewska M, Srivastava S, et al. Lentiviral Vpr usurps Cul4-DDB1[VprBP] E3 ubiquitin ligase to modulate cell cycle. Proc Natl Acad Sci U S A 2007;104:11778–11783. 148. Hrecka K, Hao C, Gierszewska M, et al. Vpx relieves inhibition of HIV-1 infection of macrophages mediated by the SAMHD1 protein. Nature 2011;474:658–661. 149. Hrecka K, Hao C, Shun MC, et al. HIV-1 and HIV-2 exhibit divergent interactions with HLTF and UNG2 DNA repair proteins. Proc Natl Acad Sci U S A 2016;113:E3921–E3930. 150. Hu J, Switzer WM, Foley BT, et al. Characterization and comparison of recombinant simian immunodeficiency virus from drill (Mandrillus leucophaeus) and mandrill (Mandrillus sphinx) isolates. J Virol 2003;77:4867–4880. 151. Hultquist JF, Lengyel JA, Refsland EW, et al. Human and rhesus APOBEC3D, APOBEC3F, APOBEC3G, and APOBEC3H demonstrate a conserved capacity to restrict Vif-deficient HIV-1. J Virol 2011;85:11220–11234. 152. Ilyinskii PO, Desrosiers RC. Efficient transcription and replication of simian immunodeficiency virus in the absence of NF-kB and Sp1 binding elements. J Virol 1996;70:3118–3126. 153. Ilyinskii PO, Daniel MD, Simon MA, et al. The role of upstream U3 sequences in the pathogenesis of simian immunodeficiency virusinduced AIDS in rhesus monkeys. J Virol 1994;68:5933–5944. 154. Ilyinskii PO, Simon MA, Czajak SC, et al. Induction of AIDS by simian immunodeficiency virus lacking NF-kB and SP1 binding elements. J Virol 1997;71:1880–1887. 155. Issel C, Foil, LD. Studies on equine infectious anemia virus transmission by insects. J Am Vet Med Assoc 1984;184:293–297. 156. Jia B, Serra-Moreno R, Neidermyer W, et al. Species-specific activity of SIV Nef and HIV-1 Vpu in overcoming restriction by tetherin/BST2. PLoS Pathog 2009;5:e1000429. 157. Jia X, Singh R, Homann S, et al. Structural basis of evasion of cellular adaptive immunity by HIV-1 Nef. Nat Struct Mol Biol 2012;19:701–706. 158. Jin MJ, Rogers J, Phillips-Conroy JE, et al. Infection of a yellow baboon with simian immunodeficiency virus from African green monkeys: evidence for cross-species transmission in the wild. J Virol 1994;68:8454–8460. 159. Johnson PR, Schnepp BC, Zhang J, et al. Vector-mediated gene transfer engenders long-lived neutralizing activity and protection against SIV infection in monkeys. Nat Med 2009;15:901–906. 759

160. Johnson RP, Lifson JD, Czajak SC, et al. Highly attenuated vaccine strains of simian immunodeficiency virus protect against vaginal challenge: inverse relation of degree of protection with level of attenuation. J Virol 1999;73:4952–4961. 161. Jones T, Hunt RD, King NW. Diseases caused by viruses. In: Cann C, ed. Veterinary Pathology. Baltimore, MD: Lippincott Williams & Wilkins; 1997:197–370. 162. Joseph SB, Swanstrom R. The evolution of HIV-1 entry phenotypes as a guide to changing target cells. J Leukoc Biol 2018;103:421–431. 163. Julien JP, Cupo A, Sok D, et al. Crystal structure of a soluble cleaved HIV-1 envelope trimer. Science 2013;342:1477–1483. 164. Kane M, Rebensburg SV, Takata MA, et al. Nuclear pore heterogeneity influences HIV-1 infection and the antiviral activity of MX2. Elife 2018;7:e35738. 165. Kane M, Yadav SS, Bitzegeio J, et al. MX2 is an interferon-induced inhibitor of HIV-1 infection. Nature 2013;502:563–566. 166. Katzourakis A, Tristem M, Pybus OG, et al. Discovery and analysis of the first endogenous lentivirus. Proc Natl Acad Sci U S A 2007;104:6261–6265. 167. Kaur A, Grant RM, Means RE, et al. Diverse host responses and outcomes following simian immunodeficiency virus SIVmac239 infection in sooty mangabeys and rhesus macaques. J Virol 1998;72:9597–9611. 168. Keele BF, Van Heuverswyn F, Li Y, et al. Chimpanzee reservoirs of pandemic and nonpandemic HIV-1. Science 2006;313:523–526. 169. Kerns JA, Emerman M, Malik HS. Positive selection and increased antiviral activity associated with the PARP-containing isoform of human zinc-finger antiviral protein. PLoS Genet 2008;4:e21. 170. Kertayadnya G, Wilcox GE, Soeharsono S, et al. Characteristics of a retrovirus associated with Jembrana disease in Bali cattle. J Gen Virol 1993;74:1765–1778. 171. Kestler H, Kodama T, Ringler D, et al. Induction of AIDS in rhesus monkeys by molecularly cloned simian immunodeficiency virus. Science 1990;248:1109–1112. 172. Kestler HW III, Ringler DJ, Mori K, et al. Importance of the nef gene for maintenance of high virus loads and for the development of AIDS. Cell 1991;65:651–662. 173. Khabbaz RF, Heneine W, George JR, et al. Brief report: infection of a laboratory worker with simian immunodeficiency virus. N Engl J Med 1994;330:172–177. 174. Kirchhoff F, Greenough TC, Brettler DB, et al. Absence of intact nef sequences in a long-term survivor with nonprogressive HIV-1 infection. N Engl J Med 1995;332:228–232. 175. Kirchhoff F, Kestler HW III, Desrosiers RC. Upstream U3 sequences in simian immunodeficiency virus are selectively deleted in vivo in the absence of an intact nef gene. J Virol 1994;68:2031–2037. 176. Kirmaier A, Wu F, Newman RM, et al. TRIM5 suppresses cross-species transmission of a primate immunodeficiency virus and selects for emergence of resistant variants in the new species. PLoS Biol 2010;8(8):e1000462. 177. Kirschman J, Qi M, Ding L, et al. HIV-1 envelope glycoprotein trafficking through the endosomal recycling compartment is required for particle incorporation. J Virol 2018;92(5):e01893-17. 178. Kluge SF, Mack K, Iyer SS, et al. Nef proteins of epidemic HIV-1 group O strains antagonize human tetherin. Cell Host Microbe 2014;16:639–650. 179. Klumpp SA, Novembre FJ, Anderson DC, et al. Clinical and pathologic findings in infant rhesus macaques infected with SIVsmm by maternal transmission. J Med Primatol 1993;22:169–176. 180. Kobayashi K. Studies on the cultivation of equine infectious anemia virus in vitro. I. Serial cultivation of the virus in the culture of various horse tissues. Virus 1961;11:177–189. 181. Kodama T, Silva DP, Daniel MD, et al. Prevalence of antibodies to SIV in baboons in their native habitat. AIDS Res Hum Retroviruses 1989;5(3):337–343. 182. Kodama T, Wooley DP, Naidu YM, et al. Significance of premature stop codons in env of simian immunodeficiency virus. J Virol 1989;63:4709–4714. 183. Kraus G, Werner A, Baier M, et al. Isolation of human immunodeficiency virus-related simian immunodeficiency viruses from African green monkeys. Proc Natl Acad Sci U S A 1989;86:2892–2896. 184. Kupzig S, Korolchuk V, Rollason R, et al. Bst-2/HM1.24 is a raft-associated apical membrane protein with an unusual topology. Traffic 2003;4:694–709. 185. Kwon Y, Kaake RM, Echeverria I, et al. Structural basis of CD4 downregulation by HIV-1 Nef. Nat Struct Mol Biol 2020;27:822–828. 186. Kwong PD, Doyle ML, Casper DJ, et al. HIV-1 evades antibody-mediated neutralization through conformational masking of receptorbinding sites. Nature 2002;420:678–682. 187. Lackner A, Vogel P, Ramos R, et al. Early events in tissues during infection with pathogenic (SIVmac239) and nonpathogenic (SIVmac1A11) molecular clones of SIV. Am J Pathol 1994;145:428–439. 188. Laguette N, Bregnard C, Hue P, et al. Premature activation of the SLX4 complex by Vpr promotes G2/M arrest and escape from innate immune sensing. Cell 2014;156:134–145. 189. Laguette N, Sobhian B, Casartelli N, et al. SAMHD1 is the dendritic- and myeloid-cell-specific HIV-1 restriction factor counteracted by Vpx. Nature 2011;474:654–657. 190. LaRue RS, Jonsson SR, Silverstein KA, et al. The artiodactyl APOBEC3 innate immune repertoire shows evidence for a multi-functional domain organization that existed in the ancestor of placental mammals. BMC Mol Biol 2008;9:104. 191. Le Rouzic E, Belaidouni N, Estrabaud E, et al. HIV1 Vpr arrests the cell cycle by recruiting DCAF1/VprBP, a receptor of the Cul4-DDB1 ubiquitin ligase. Cell Cycle 2007;6:182–188. 192. Le Tortorec A, Neil SJ. Antagonism to and intracellular sequestration of human tetherin by the human immunodeficiency virus type 2 envelope glycoprotein. J Virol 2009;83:11966–11978. 193. Learmont JC, Geczy AF, Mills J, et al. Immunologic and virologic status after 14 to 18 years of infection with an attenuated strain of HIV-1. A report from the Sydney Blood Bank Cohort. N Engl J Med 1999;340:1715–1722. 194. Lecossier D, Bouchonnet F, Clavel F, et al. Hypermutation of HIV-1 DNA in the absence of the Vif protein. Science 2003;300:1112. 195. Lee CA, Beasley E, Sundar K, et al. Simian immunodeficiency virus-infected memory CD4(+) T cells infiltrate to the site of infected macrophages in the neuroparenchyma of a chronic macaque model of neurological complications of AIDS. mBio 2020;11:e00602-20. 196. Leendertz SA, Locatelli S, Boesch C, et al. No evidence for transmission of SIVwrc from western red colobus monkeys (Piliocolobus badius badius) to wild West African chimpanzees (Pan troglodytes verus) despite high exposure through hunting. BMC Microbiol 2011;11:24. 197. Lemey P, Pybus OG, Wang B, et al. Tracing the origin and history of the HIV-2 epidemic. Proc Natl Acad Sci U S A 2003;100:6588–6592. 198. Lerner DL, Wagaman PC, Phillips TR, et al. Increased mutation frequency of feline immunodeficiency virus lacking functional deoxyuridine-triphosphatase. Proc Natl Acad Sci U S A 1995;92:7480–7484. 199. Li F, Leroux C, Craigo JK, et al. The S2 gene of equine infectious anemia virus is a highly conserved determinant of viral replication and virulence properties in experimentally infected ponies. J Virol 2000;74:573–579. 200. Li F, Puffer BA, Montelaro RC. The S2 gene of equine infectious anemia virus is dispensable for viral replication in vitro. J Virol 1998;72:8344–8348. 201. Li H, Wang S, Kong R, et al. Envelope residue 375 substitutions in simian-human immunodeficiency viruses enhance CD4 binding and 760

replication in rhesus macaques. Proc Natl Acad Sci U S A 2016;113:E3413–E3422. 202. Li MM, Lau Z, Cheung P, et al. TRIM25 enhances the antiviral action of Zinc-Finger Antiviral Protein (ZAP). PLoS Pathog 2017;13:e1006145. 203. Li Y, Ndjango JB, Learn GH, et al. Eastern chimpanzees, but not bonobos, represent a simian immunodeficiency virus reservoir. J Virol 2012;86:10776–10791. 204. Lichtenstein DL, Rushlow KE, Cook RF, et al. Replication in vitro and in vivo of an equine infectious anemia virus mutant deficient in dUTPase activity. J Virol 1995;69:2881–2888. 205. Liddament MT, Brown WL, Schumacher AJ, et al. APOBEC3F properties and hypermutation preferences indicate activity against HIV-1 in vivo. Curr Biol 2004;14:1385–1391. 206. Lifson JD, Rossio JL, Arnaout R, et al. Containment of simian immunodeficiency virus infection: cellular immune responses and protection from rechallenge following transient postinoculation antiretroviral treatment. J Virol 2000;74:2584–2593. 207. Lifson JD, Rossio JL, Piatak M Jr, et al. Role of CD8(+) lymphocytes in control of simian immunodeficiency virus infection and resistance to rechallenge after transient early antiretroviral treatment. J Virol 2001;75:10187–10199. 208. Lim ES, Fregoso OI, McCoy CO, et al. The ability of primate lentiviruses to degrade the monocyte restriction factor SAMHD1 preceded the birth of the viral accessory protein Vpx. Cell Host Microbe 2012;11:194–204. 209. Ling B, Apetrei C, Pandrea I, et al. Classic AIDS in a sooty mangabey after an 18-year natural infection. J Virol 2004;78:8902–8908. 210. Liu ZQ, Sheridan D, Wood C. Identification and characterization of the bovine immunodeficiency-like virus tat gene. J Virol 1992;66:5137–5140. 211. Locher CP, Barnett SW, Herndier BG, et al. Human immunodeficiency virus-2 infection in baboons is an animal model for human immunodeficiency virus pathogenesis in humans. Arch Pathol Lab Med 1998;122:523–533. 212. Loffredo JT, Maxwell J, Qi Y, et al. Mamu-b*08-positive macaques control simian immunodeficiency virus replication. J Virol 2007;81:8827–8832. 213. Loffredo JT, Sidney J, Bean AT, et al. Two MHC class I molecules associated with elite control of immunodeficiency virus replication, Mamu-B*08 and HLA-B*2705, bind peptides with sequence similarity. J Immunol 2009;182:7763–7775. 214. Lowenstine LJ, Lerche NW, Yee JL, et al. Evidence for a lentiviral etiology in an epizootic of immune deficiency and lymphoma in stumptailed macaques (Macaca arctoides). J Med Primatol 1992;21:1–14. 215. Lowenstine LJ, Pedersen NC, Higgins J, et al. Seroepidemiologic survey of captive Old-World primates for antibodies to human and simian retroviruses, and isolation of a lentivirus from sooty mangabeys (Cercocebus atys). Int J Cancer 1986;38:563–574. 216. Lv L, Wang Q, Xu Y, et al. Vpr targets TET2 for degradation by CRL4(VprBP) E3 ligase to sustain IL-6 expression and enhance HIV-1 replication. Mol Cell 2018;70:961–970 e5. 217. Ma J, Shi N, Jiang CG, et al. A proviral derivative from a reference attenuated EIAV vaccine strain failed to elicit protective immunity. Virology 2011;410:96–106. 218. Malik P, Singha H, Sarkar S. Equine infectious anemia. In: Bayry J, ed. Emerging and Re-emerging Infectious Diseases of Livestock. 2017:215–336. 219. Mansfield KG, Lerch NW, Gardner MB, et al. Origins of simian immunodeficiency virus infection in macaques at the New England Regional Primate Research Center. J Med Primatol 1995;24:116–122. 220. Marin M, Rose KM, Kozak SL, et al. HIV-1 Vif protein binds the editing enzyme APOBEC3G and induces its degradation. Nat Med 2003;9:1398–1403. 221. Markovitz DM, Smith MJ, Hilfinger J, et al. Activation of the human immunodeficiency virus type 2 enhancer is dependent on purine box and kB regulatory elements. J Virol 1992;66:5479–5484. 222. Marsh H. Progressive pneumonia in sheep. J Am Vet Med Assoc 1923;62:458–473. 223. Martinez-Navio JM, Fuchs SP, Pantry SN, et al. Adeno-associated virus delivery of anti-HIV monoclonal antibodies can drive long-term virologic suppression. Immunity 2019;50:567–575 e5. 224. Martins MA, Bischof GF, Shin YC, et al. Vaccine protection against SIVmac239 acquisition. Proc Natl Acad Sci U S A 2019;116:1739–1744. 225. Marx P. Unsolved questions over the origin of HIV and AIDS. ASM News 2005;71:15–17. 226. Matsuda K, Riddick NE, Lee CA, et al. A SIV molecular clone that targets the CNS and induces neuroAIDS in rhesus macaques. PLoS Pathog 2017;13:e1006538. 227. Mattapallil JJ, Douek DC, Hill B, et al. Massive infection and loss of memory CD4+ T cells in multiple tissues during acute SIV infection. Nature 2005;434:1093–1097. 228. Maury W. Regulation of equine infectious anemia virus expression. J Biomed Sci 1998;5(1):11–23. 229. McClure HM, Anderson DC, Gordon TP, et al. Natural simian immunodeficiency virus infection in nonhuman primates. Top Primatol 1992;3:425–438. 230. McEwan WA, McMonagle EL, Logan N, et al. Genetically divergent strains of feline immunodeficiency virus from the domestic cat (Felis catus) and the African lion (Panthera leo) share usage of CD134 and CXCR4 as entry receptors. J Virol 2008;82:10953–10958. 231. McGuire TC, Crawford TB, Henson JB. Immunofluorescent localization of equine infectious anemia virus in tissue. Am J Pathol 1971;62:283–294. 232. McNatt MW, Zang T, Hatziioannou T, et al. Species-specific activity of HIV-1 Vpu and positive selection of tetherin transmembrane domain variants. PLoS Pathog 2009;5:e1000300. 233. Meagher JL, Takata M, Goncalves-Carneiro D, et al. Structure of the zinc-finger antiviral protein in complex with RNA reveals a mechanism for selective targeting of CG-rich viral sequences. Proc Natl Acad Sci U S A 2019;116:24303–24309. 234. Monit C, Morris ER, Ruis C, et al. Positive selection in dNTPase SAMHD1 throughout mammalian evolution. Proc Natl Acad Sci U S A 2019;116:18647–18654. 235. Montagnier L, Dauguet C, Axler C, et al. A new type of retrovirus isolated from patients presenting with lymphadenopathy and acquired immune deficiency syndrome: structural and antigenic relatedness with equine infectious anemia virus. Ann Virol 1984;135 E:119–134. 236. Montelaro RC, Ball JM, Rushlow KE. Equine retroviruses. In: Levy JA, ed. The Retroviridae. New York: Plenum Press; 1993:257. 237. Morse BA, Carruth LM, Clements JE. Targeting of the visna virus tat protein to AP-1 sites: interactions with the bZIP domains of fos and jun in vitro and in vivo. J Virol 1999;73:37–45. 238. Mothe BR, Weinfurter J, Wang C, et al. Expression of the major histocompatibility complex class I molecule Mamu-A*01 is associated with control of simian immunodeficiency virus SIVmac239 replication. J Virol 2003;77:2736–2740. 239. Muller MC, Saksena NK, Nerrienet E, et al. Simian immunodeficiency viruses from central and western Africa: evidence for a new species-specific lentivirus in tantalus monkeys. J Virol 1993;67:1227–1235. 240. Murakami T, Freed EO. Genetic evidence for an interaction between human immunodeficiency virus type 1 matrix and alpha-helix 2 of the gp41 cytoplasmic tail. J Virol 2000;74:3548–3554. 241. Murphey-Corb M, Martin LN, Davison-Fairburn B, et al. A formalin-inactivated whole SIV vaccine confers protection in macaques. Science 1989;246:1293–1297. 761

242. Murphey-Corb M, Martin LN, Rangan SRS, et al. Isolation of an HTLV-III-related retrovirus from macaques with simian AIDS and its possible origin in asymptomatic mangabeys. Nature 1986;321:435–437. 243. Neil SJ, Zang T, Bieniasz PD. Tetherin inhibits retrovirus release and is antagonized by HIV-1 Vpu. Nature 2008;451:425–430. 244. Neil SJ. The antiviral activities of tetherin. Curr Top Microbiol Immunol 2013;371:67–104. 245. Nerrienet E, Amouretti X, Muller-Trutwin MC, et al. Phylogenetic analysis of SIV and STLV type I in mandrills (Mandrillus sphinx): indications that intracolony transmissions are predominantly the result of male-to-male aggressive contacts. AIDS Res Hum Retroviruses 1998;14:785–796. 246. Newman RM, Hall L, Connole M, et al. Balancing selection and the evolution of functional polymorphism in Old World monkey TRIM5alpha. Proc Natl Acad Sci U S A 2006;103:19134–19139. 247. Nixon CC, Mavigner M, Sampey GC, et al. Systemic HIV and SIV latency reversal via non-canonical NF-kappaB signalling in vivo. Nature 2020;578:160–165. 248. Novembre FJ, Hirsch VM, McClure HM, et al. SIV from stump-tailed macaques: molecular characterization of a highly transmissible primate lentivirus. Virology 1992;186:783–787. 249. Novembre FJ, Saucier M, Anderson DC, et al. Development of AIDS in a chimpanzees infected with human immunodeficiency virus type. J Virol 1997;71:4086–4091. 250. Oaks JL, McGuire TC, Ulibarri C, et al. Equine infectious anemia virus is found in tissue macrophages during subclinical infection. J Virol 1998;72:7263–7269. 251. Ohta Y, Masuda T, Tsujimoto H, et al. Isolation of simian immunodeficiency virus from African green monkeys and seroepidemiologic survey of the virus in various non-human primates. Int J Cancer 1988;41:115–122. 252. Olmsted RA, Barnes AK, Yamamoto JK, et al. Molecular cloning of feline immunodeficiency virus. Proc Natl Acad Sci U S A 1989;86:2448–2452. 253. O'Neil SP, Novembre FJ, Hill AB, et al. Progressive infection in a subset of HIV-1-positive chimpanzees. J Infect Dis 2000;182:1051–1062. 254. Orandle MS, Baldwin CJ. Clinical and pathological features associated with feline immunodeficiency virus infection in cats. Iowa State Univ Vet. 1995;7(2):61–65. 255. Osterhaus AD, Pedersen N, van Amerongen G, et al. Isolation and partial characterization of a lentivirus from talapoin monkeys (Myopithecus talapoin). Virology 1999;260:116–124. 256. Pandrea I, Apetrei C, Gordon S, et al. Paucity of CD4+CCR5+ T cells is a typical feature of natural SIV hosts. Blood 2007;109:1069–1076. 257. Pandrea I, Onanga R, Rouquet P, et al. Chronic SIV infection ultimately causes immunodeficiency in African non-human primates. AIDS 2001;15:2461–2462. 258. Parekh B, Issel CJ, Montelaro RC. Equine infectious anemia virus, a putative lentivirus, contains polypeptides analogous to prototype-C oncornaviruses. Virology 1980;107:520–525. 259. Payne SL, Qi XM, Shao H, et al. Disease induction by virus derived from molecular clones of equine infectious anemia virus. J Virol 1998;72:483–487. 260. Pedersen NC, Ho EW, Brown ML, et al. Isolation of a T-lymphotropic virus from domestic cats with an immunodeficiency-like syndrome. Science 1987;235:790–793. 261. Peeters M, Courgnaud V, Abela B, et al. Risk to human health from a plethora of simian immunodeficiency viruses in primate bushmeat. Emerg Infect Dis 2002;8:451–457. 262. Peeters M, Ma D, Liegeois F, et al. Simian immunodeficiency virus infections in the wild. In: Ansari AA, Silvestri G, eds. Natural Hosts of SIV. Elsevier; 2014:37–67. 263. Peluso R, Haase A, Stowring L, et al. A Trojan horse mechanism for the spread of visna virus in monocytes. Virology 1985;147:231–236. 264. Pepin M, Vitu C, Russo P, et al. Maedi-visna virus infection in sheep: a review. Vet Res 1998;29:341–367. 265. Perez-Caballero D, Zang T, Ebrahimi A, et al. Tetherin inhibits HIV-1 release by directly tethering virions to cells. Cell 2009;139:499–511. 266. Petursson G, Turelli P, Matthiasdottir S, et al. Visna virus dUTPase is dispensable for neuropathogenicity. J Virol 1998;72:1657–1661. 267. Phillips-Conroy JE, Jolly CJ, Petros B, et al. Sexual transmission of SIVagm in wild grivet monkeys. J Med Primatol 1994;23:1–7. 268. Piantadosi A, Chohan B, Chohan V, et al. Chronic HIV-1 infection frequently fails to protect against superinfection. PLoS Pathog 2007;3:e177. 269. Picker LJ, Hagen SI, Lum R, et al. Insufficient production and tissue delivery of CD4+ memory T cells in rapidly progressive simian immunodeficiency virus infection. J Exp Med 2004;200:1299–1314. 270. Pifat DY, Ennis WH, Ward JM, et al. Persistent infection of rabbits with bovine immunodeficiency-like virus. J Virol 1992;66:4518–4524. 271. Pisoni G, Quasso A, Moroni P. Phylogenetic analysis of small-ruminant lentivirus subtype B1 in mixed flocks: evidence for natural transmission from goats to sheep. Virology 2005;339:147–152. 272. Poeschla EM, Looney DJ. CXCR4 is required by a nonprimate lentivirus: heterologous expression of feline immunodeficiency virus in human, rodent, and feline cells. J Virol 1998;72:6858–6866. 273. Pohlmann S, Floos S, Ilyinskii PO, et al. Sequences just upstream of the simian immunodeficiency virus core enhancer allow efficient replication in the absence of NF-kappaB and Sp1 binding elements. J Virol 1998;72:5589–5598. 274. Pollard VW, Malim MH. The HIV-1 Rev protein. Annu Rev Microbiol 1998;52:491–532. 275. Popovic M, Sarngadharan MG, Read E, et al. Detection, isolation, and continuous production of cytopathic retroviruses (HTLV-III) from patients with AIDS and pre-AIDS. Science 1984;224:497–500. 276. Postler TS, Desrosiers RC. The cytoplasmic domain of the HIV-1 glycoprotein gp41 induces NF-kappaB activation through TGF-betaactivated kinase 1. Cell Host Microbe 2012;11:181–193. 277. Postler TS, Desrosiers RC. The tale of the long tail: the cytoplasmic domain of HIV-1 gp41. J Virol 2013;87:2–15. 278. Pu R, Coleman J, Omori M, et al. Dual-subtype FIV vaccine protects cats against in vivo swarms of both homologous and heterologous subtype FIV isolates. AIDS 2001;15:1225–1237. 279. Pye VE, Rosa A, Bertelli C, et al. A bipartite structural organization defines the SERINC family of HIV-1 restriction factors. Nat Struct Mol Biol 2020;27:78–83. 280. Qi M, Chu H, Chen X, et al. A tyrosine-based motif in the HIV-1 envelope glycoprotein tail mediates cell-type- and Rab11-FIP1Cdependent incorporation into virions. Proc Natl Acad Sci U S A 2015;112:7575–7580. 281. Qi M, Williams JA, Chu H, et al. Rab11-FIP1C and Rab14 direct plasma membrane sorting and particle incorporation of the HIV-1 envelope glycoprotein complex. PLoS Pathog 2013;9:e1003278. 282. Ramachandran S. Early observations and research of Jembrana disease in Bali and other Indonesian islands. In: Jembrana Disease and The Bovine Lentiviruses. Canberra, Australia: ACIAR Proceedings; 1997. 283. Reitter J, Means RE, Desrosiers RC. A role for carbohydrates in immune evasion in AIDS. Nat Med 1998;4:679–684. 284. Rey-Cuille MA, Berthier JL, Bomsel-Demontoy MC, et al. Simian immunodeficiency virus replicates to high levels in sooty mangabeys without inducing disease. J Virol 1998;72:3872–3886. 762

285. Richman DD, Wrin T, Little SJ, et al. Rapid evolution of the neutralizing antibody response to HIV type 1 infection. Proc Natl Acad Sci U S A 2003;100:4144–4149. 286. Riddick NE, Wu F, Matsuda K, et al. Simian immunodeficiency virus SIVagm efficiently utilizes non-CCR5 entry pathways in African Green Monkey lymphocytes: potential role for GPR15 and CXCR6 as viral coreceptors. J Virol 2015;90:2316–2331. 287. Rosa A, Chande A, Ziglio S, et al. HIV-1 Nef promotes infection by excluding SERINC5 from virion incorporation. Nature 2015;526:212–217. 288. Rosenberg ES, Billingsley JM, Caliendo AM, et al. Vigorous HIV-1-specific CD4+ T cell responses associated with control of viremia. Science 1997;278:1447–1450. 289. Ross EK, Buckler-White AJ, Rabson AB, et al. Contribution of NF-kB and Sp1 binding motifs to the replicative capacity of human immunodeficiency virus type 1: distinct patterns of viral growth are determined by T-cell types. J Virol 1991;65:4350–4358. 290. Rudicell RS, Holland Jones J, Wroblewski EE, et al. Impact of simian immunodeficiency virus infection on chimpanzee population dynamics. PLoS Pathog 2010;6:e1001116. 291. Rudicell RS, Piel AK, Stewart F, et al. High prevalence of simian immunodeficiency virus infection in a community of savanna chimpanzees. J Virol 2011;85:9918–9928. 292. Saenz DT, Teo W, Olsen JC, et al. Restriction of feline immunodeficiency virus by Ref1, Lv1, and primate TRIM5alpha proteins. J Virol 2005;79:15175–15188. 293. Santiago ML, Lukasik M, Kamenya S, et al. Foci of endemic simian immunodeficiency virus infection in wild-living eastern chimpanzees (Pan troglodytes schweinfurthii). J Virol 2003;77:7545–7562. 294. Santiago ML, Range F, Keele BF, et al. Simian immunodeficiency virus infection in free-ranging sooty mangabeys (Cercocebus atys atys) from the Tai Forest, Cote d'Ivoire: implications for the origin of epidemic human immunodeficiency virus type 2. J Virol 2005;79:12515–12527. 295. Santiago ML, Rodenburg CM, Kamenya S, et al. SIVcpz in wild chimpanzees. Science 2002;295:465. 296. Sasseville V, Lackner A A. Neuropathogenesis of simian immunodeficiency virus infection in macaque monkeys. J Neurovirol 1997;3:1–9. 297. Sauter D, Hotter D, Van Driessche B, et al. Differential regulation of NF-kappaB-mediated proviral and antiviral host gene expression by primate lentiviral Nef and Vpu proteins. Cell Rep 2015;10:586–599. 298. Sauter D, Schindler M, Specht A, et al. Tetherin-driven adaptation of Vpu and Nef function and the evolution of pandemic and nonpandemic HIV-1 strains. Cell Host Microbe 2009;6:409–421. 299. Sawyer SL, Emerman M, Malik HS. Ancient adaptive evolution of the primate antiviral DNA-editing enzyme APOBEC3G. PLoS Biol 2004;2:E275. 300. Sawyer SL, Emerman M, Malik HS. Discordant evolution of the adjacent antiretroviral genes TRIM22 and TRIM5 in mammals. PLoS Pathog 2007;3:e197. 301. Sawyer SL, Wu LI, Emerman M, et al. Positive selection of primate TRIM5alpha identifies a critical species-specific retroviral restriction domain. Proc Natl Acad Sci U S A 2005;102:2832–2837. 302. Sayah DM, Sokolskaja E, Berthoux L, et al. Cyclophilin A retrotransposition into TRIM5 explains owl monkey resistance to HIV-1. Nature 2004;430:569–573. 303. Schaller T, Hue S, Towers GJ. An active TRIM5 protein in rabbits indicates a common antiviral ancestor for mammalian TRIM5 proteins. J Virol 2007;81:11713–11721. 304. Schindler M, Munch J, Kutsch O, et al. Nef-mediated suppression of T cell activation was lost in a lentiviral lineage that gave rise to HIV-1. Cell 2006;125:1055–1067. 305. Schmidt F, Keele BF, Del Prete GQ, et al. Derivation of simian tropic HIV-1 infectious clone reveals virus adaptation to a new host. Proc Natl Acad Sci U S A 2019;116:10504–10509. 306. Schmitz JE, Kuroda MJ, Santra S, et al. Control of viremia in simian immunodeficiency virus infection by CD8+ lymphocytes. Science 1999;283:857–860. 307. Schrofelbauer B, Chen D, Landau NR. A single amino acid of APOBEC3G controls its species-specific interaction with virion infectivity factor (Vif). Proc Natl Acad Sci U S A 2004;101:3927–3932. 308. Schrofelbauer B, Hakata Y, Landau NR. HIV-1 Vpr function is mediated by interaction with the damage-specific DNA-binding protein DDB1. Proc Natl Acad Sci U S A 2007;104:4130–4135. 309. Schwartz O, Marechal V, LeGall S, et al. Endocytosis of major histocompatibility complex class I molecules is induced by the HIV-1 nef protein. Nat Med 1996;2:338–342. 310. Sellon DC, Perry ST, Coggins L, et al. Wild-type equine infectious anemia virus replicates in vivo predominantly in tissue macrophages, not in peripheral blood monocytes. J Virol 1992;66:5906–5913. 311. Serra-Moreno R, Jia B, Breed M, et al. Compensatory changes in the cytoplasmic tail of gp41 confer resistance to tetherin/BST-2 in a pathogenic nef-deleted SIV. Cell Host Microbe 2011;9:46–57. 312. Shacklett BL, Luciw PA. Analysis of the vif gene of feline immunodeficiency virus. Virology 1994;204:860–867. 313. Sharp PM, Hahn BH. Origins of HIV and the AIDS pandemic. Cold Spring Harb Perspect Med 2011;1:a006841. 314. Shaw AE, Hughes J, Gu Q, et al. Fundamental properties of the mammalian innate immune system revealed by multispecies comparison of type I interferon responses. PLoS Biol 2017;15:e2004086. 315. Sheehy AM, Gaddis NC, Choi JD, et al. Isolation of a human gene that inhibits HIV-1 infection and is suppressed by the viral Vif protein. Nature 2002;418:646–650. 316. Sheehy AM, Gaddis NC, Malim MH. The antiretroviral enzyme APOBEC3G is degraded by the proteasome in response to HIV-1 Vif. Nat Med 2003;9:1404–1407. 317. Shen R, Wang Z. Development and use of an equine infectious anemia donkey leukocyte attenuated vaccine. In: Tashjian R, ed. Equine Infectious Anemia: A National Review of Policies, Programs, and Future Objections. Amarillo, TX: American Quarter Horse Association; 1985:135–248. 318. Shimojima M, Miyazawa T, Ikeda Y, et al. Use of CD134 as a primary receptor by the feline immunodeficiency virus. Science 2004;303:1192–1195. 319. Shingai M, Donau OK, Schmidt SD, et al. Most rhesus macaques infected with the CCR5-tropic SHIV(AD8) generate cross-reactive antibodies that neutralize multiple HIV-1 strains. Proc Natl Acad Sci U S A 2012;109:19769–19774. 320. Sigurdsson B. RIDA; a chronic encephalitis of sheep. With general remarks on infections which develop slowly and some of their special characteristics. Br Vet J 1954;110:354–358. 321. Sigurdsson B, Grimsson H, Palsson PA. Maedi, a chronic progressive infection of sheep's lungs. J Infect Dis 1952;90:23–241. 322. Sigurdsson B, Pálsson PA. Visna of sheep. A slow, demyelinating infection. Br J Exp Pathol 1958;39:519. 323. Sigurdsson B, Palsson PA, Grimsson H. Visna, a demyelinating transmissible disease of sheep. J Neuropathol Exp Neurol 1957;16:389–403. 324. Sigurdsson B, Thormar H, Pálsson PA. Cultivation of visna virus in tissue culture. Arch Gesamte Virusforsch 1960;10:368. 763

325. Simon JH, Miller DL, Fouchier RA, et al. The regulation of primate immunodeficiency virus infectivity by Vif is cell species restricted: a role for Vif in determining virus host range and cross-species transmission. EMBO J 1998;17:1259–1267. 326. Smaga SS, Xu C, Summers BJ, et al. MxB restricts HIV-1 by targeting the Tri-hexamer interface of the viral capsid. Structure 2019;27:1234–1245 e5. 327. Smolen-Dzirba J, Rosinska M, Janiec J, et al. HIV-1 infection in persons homozygous for CCR5-Delta32 allele: the next case and the review. AIDS Rev 2017;19:219–230. 328. Soeharsono S. Current information on Jembrana disease distribution in Indonesia. In: Jembrana Disease and the Bovine Lentiviruses. ACIAR Proceedings. Canberra, Australia: ACIAR; 1997. 329. Soeharsono S, Wilcox GE, Dharma DM, et al. Species differences in the reaction of cattle to Jembrana disease virus infection. J Comp Pathol 1995;112:391–402. 330. Soeharsono S, Wilcox GE, Putra AA, et al. The transmission of Jembrana disease, a lentivirus disease of Bos javanicus cattle. Epidemiol Infect 1995;115:367–374. 331. Song B, Javanbakht H, Perron M, et al. Retrovirus restriction by TRIM5alpha variants from Old World and New World primates. J Virol 2005;79:3930–3937. 332. Sood C, Marin M, Chande A, et al. SERINC5 protein inhibits HIV-1 fusion pore formation by promoting functional inactivation of envelope glycoproteins. J Biol Chem 2017;292:6014–6026. 333. Sotir M, Switzer W, Schable C, et al. Risk of occupational exposure to potentially infectious nonhuman primate materials and to simian immunodeficiency virus. J Med Primatol 1997;26:233–240. 334. Souquiere S, Bibollet-Ruche F, Robertson DL, et al. Wild Mandrillus sphinx are carriers of two types of lentivirus. J Virol 2001;75:7086–7096. 335. Srivastava S, Swanson SK, Manel N, et al. Lentiviral Vpx accessory factor targets VprBP/DCAF1 substrate adaptor for cullin 4 E3 ubiquitin ligase to enable macrophage infection. PLoS Pathog 2008;4:e1000059. 336. Stickney A, Ghosh S, Cave NJ, et al. Lack of protection against feline immunodeficiency virus infection among domestic cats in New Zealand vaccinated with the Fel-O-Vax(R) FIV vaccine. Vet Microbiol 2020;250:108865. 337. Stremlau M, Owens CM, Perron MJ, et al. The cytoplasmic body component TRIM5alpha restricts HIV-1 infection in Old World monkeys. Nature 2004;427:848–853. 338. Swigut T, Alexander L, Morgan J, et al. Impact of Nef-mediated downregulation of major histocompatibility complex class I on immune response to simian immunodeficiency virus. J Virol 2004;78:13335–13344. 339. Takata MA, Goncalves-Carneiro D, Zang TM, et al. CG dinucleotide suppression enables antiviral defence targeting non-self RNA. Nature 2017;550:124–127. 340. Tavakoli-Tameh A, Janaka SK, Zarbock K, et al. Loss of tetherin antagonism by Nef impairs SIV replication during acute infection of rhesus macaques. PLoS Pathog 2020;16:e1008487. 341. Tedbury PR, Ablan SD, Freed EO. Global rescue of defects in HIV-1 envelope glycoprotein incorporation: implications for matrix structure. PLoS Pathog 2013;9:e1003739. 342. Terio KA, Kinsel MJ, Raphael J, et al. Pathologic lesions in chimpanzees (Pan trogylodytes schweinfurthii) from Gombe National Park, Tanzania, 2004-2010. J Zoo Wildl Med 2011;42:597–607. 343. Threadgill DS, Steagall WK, Flaherty MT, et al. Characterization of equine infectious anemia virus dUTPase: growth properties of a dUTPase-deficient mutant. J Virol 1993;67:2592–2600. 344. Tomonaga K, Katahira J, Fukasawa M, et al. Isolation and characterization of simian immunodeficiency virus from African white-crowned mangabey monkeys (Cercocebus torquatus lunulatus). Arch Virol 1993;129:77–92. 345. Trautz B, Wiedemann H, Luchtenborg C, et al. The host-cell restriction factor SERINC5 restricts HIV-1 infectivity without altering the lipid composition and organization of viral particles. J Biol Chem 2017;292:13702–13713. 346. Troyer JL, Pecon-Slattery J, Roelke ME, et al. Patterns of feline immunodeficiency virus multiple infection and genome divergence in a free-ranging population of African lions. J Virol 2004;78:3777–3791. 347. Tsai CC, Emau P, Follis KE, et al. Effectiveness of postinoculation (R)-9-(2-phosphonylmethoxypropyl) adenine treatment for prevention of persistent simian immunodeficiency virus SIVmne infection depends critically on timing of initiation and duration of treatment. J Virol 1998;72:4265–4273. 348. Tsai CC, Follis KE, Sabo A, et al. Prevention of SIV infection in macaques by (R)-9-(2-phosphonylmethoxypropyl)adenine. Science 1995;270:1197–1199. 349. Tsujimoto H, Hasegawa A, Maki N, et al. Sequence of a novel simian immunodeficiency virus from a wild-caught African mandrill. Nature 1989;341:539–541. 350. Turelli P, Guiguen F, Mornex JF, et al. dUTPase-minus caprine arthritis-encephalitis virus is attenuated for pathogenesis and accumulates G-to-A substitutions. J Virol 1997;71:4522–4530. 351. Überla K, Stahl Hennig C, Böttiger D, et al. Animal model for the therapy of acquired immunodeficiency syndrome with reverse transcriptase inhibitors. Proc Natl Acad Sci U S A 1995;92:8210–8214. 352. Usami Y, Wu Y, Gottlinger HG. SERINC3 and SERINC5 restrict HIV-1 infectivity and are counteracted by Nef. Nature 2015;526:218–223. 353. Valas S, Benoit C, Guionaud C, et al. North American and French caprine arthritis-encephalitis viruses emerge from ovine maedi-visna viruses. Virology 1997;237:307–318. 354. Vallée H, Carré H. Sur la nature infectieuse de l'anaemie du cheval. C R Hebd Seances Acad Sci 1904;139:331–333. 355. Van Damme N, Goff D, Katsura C, et al. The interferon-induced protein BST-2 restricts HIV-1 release and is downregulated from the cell surface by the viral Vpu protein. Cell Host Microbe 2008;3:245–252. 356. Van Der Maaten MJ, Boothe AD, Seger CL. Isolation of a virus from cattle with persistent lymphocytosis. J Natl Cancer Inst 1972;49:1649. 357. Van Heuverswyn F, Li Y, Bailes E, et al. Genetic diversity and phylogeographic clustering of SIVcpzPtt in wild chimpanzees in Cameroon. Virology 2007;368:155–171. 358. Van Heuverswyn F, Li Y, Neel C, et al. Human immunodeficiency viruses: SIV infection in wild gorillas. Nature 2006;444:164. 359. van Rensburg EJ, Engelbrecht S, Mwenda J, et al. Simian immunodeficiency viruses (SIVs) from eastern and southern Africa: detection of a SIVagm variant from a chacma baboon. J Gen Virol 1998;79:1809–1814. 360. VandeWoude S, O'Brien SJ, Hoover EA. Infectivity of lion and puma lentiviruses for domestic cats. J Gen Virol 1997;78(Pt 4):795–800. 361. VandeWoude S, Troyer J, Poss M. Restrictions to cross-species transmission of lentiviral infection gleaned from studies of FIV. Vet Immunol Immunopathol 2010;134:25–32. 362. Veazey RS, DeMaria M, Chalifoux LV, et al. Gastrointestinal tract as a major site of CD4+ T cell depletion and viral replication in SIV infection. Science 1998;280:427–431. 363. Veazey RS, Tham IC, Mansfield KG, et al. Identifying the target cell in primary simian immunodeficiency virus (SIV) infection: highly activated memory CD4(+) T cells are rapidly eliminated in early SIV infection in vivo. J Virol 2000;74:57–64. 764

364. Veillette M, Desormeaux A, Medjahed H, et al. Interaction with cellular CD4 exposes HIV-1 envelope epitopes targeted by antibodydependent cell-mediated cytotoxicity. J Virol 2014;88:2633–2644. 365. Villet S, Bouzar BA, Morin T, et al. Maedi-visna virus and caprine arthritis encephalitis virus genomes encode a Vpr-like but no Tat protein. J Virol 2003;77:9632–9638. 366. Virgen CA, Kratovac Z, Bieniasz PD, et al. Independent genesis of chimeric TRIM5-cyclophilin proteins in two primate species. Proc Natl Acad Sci U S A 2008;105:3563–3568. 367. von Bredow B, Andrabi R, Grunst M, et al. Differences in the binding affinity of an HIV-1 V2 apex-specific antibody for the SIVsmm/mac envelope glycoprotein uncouple antibody-dependent cellular cytotoxicity from neutralization. MBio 2019;10(4):e01255-19. 368. von Bredow B, Arias JF, Heyer LN, et al. Comparison of antibody-dependent cell-mediated cytotoxicity and virus neutralization by HIV-1 Env-specific monoclonal antibodies. J Virol 2016;90:6127–6139. 369. Wagaman PC, Hasselkus-Light CS, Henson M, et al. Molecular cloning and characterization of deoxyuridine triphosphatase from feline immunodeficiency virus (FIV). Virology 1993;196:451–457. 370. Wang X, Wang S, Lin Y, et al. Genomic comparison between attenuated Chinese equine infectious anemia virus vaccine strains and their parental virulent strains. Arch Virol 2011;156:353–357. 371. Wang XF, Lin YZ, Li Q, et al. Genetic evolution during the development of an attenuated EIAV vaccine. Retrovirology 2016;13:9. 372. Ward AE, Kiessling V, Pornillos O, et al. HIV-cell membrane fusion intermediates are restricted by Serincs as revealed by cryo-electron and TIRF microscopy. J Biol Chem 2020;295:15183–15195. 373. Wei X, Decker JM, Wang S, et al. Antibody neutralization and escape by HIV-1. Nature 2003;422:307–312. 374. Wei X, Ghosh SK, Taylor ME, et al. Viral dynamics in human immunodeficiency virus type 1 infection. Nature 1995;373:117–122. 375. Weiland F, Matheka HD, Coggins L, et al. Electron microscopic studies on equine infectious anemia virus (EIAV). Brief report. Arch Virol 1977;55:335–340. 376. Westmoreland SV, Converse AP, Hrecka K, et al. SIV vpx is essential for macrophage infection but not for development of AIDS. PLoS One 2014;9:e84463. 377. Wetzel KS, Yi Y, Elliott STC, et al. CXCR6-mediated simian immunodeficiency virus SIVagmSab entry into Sabaeus African Green Monkey lymphocytes implicates widespread use of non-CCR5 pathways in natural host infections. J Virol 2017;91(4):e01626-16. 378. Wetzel KS, Yi Y, Yadav A, et al. Loss of CXCR6 coreceptor usage characterizes pathogenic lentiviruses. PLoS Pathog 2018;14:e1007003. 379. White TA, Bartesaghi A, Borgnia MJ, et al. Three-dimensional structures of soluble CD4-bound states of trimeric simian immunodeficiency virus envelope glycoproteins determined by using cryo-electron tomography. J Virol 2011;85:12114–12123. 380. Wilcox GE. Jembrana disease. Aust Vet J 1997;75:492–493. 381. Willett BJ, Hosie MJ, Neil JC, et al. Common mechanisms of infection by lentiviruses. Nature 1997;385:587. 382. Williams KC, Corey S, Westmoreland SV, et al. Perivascular macrophages are the primary cell type productively infected by simian immunodeficiency virus in the brains of macaques: implications for the neuropathogenesis of AIDS. J Exp Med 2001;193:905–915. 383. Wilson SJ, Webb BL, Maplanka C, et al. Rhesus macaque TRIM5 alleles have divergent antiretroviral specificities. J Virol 2008;82:7243–7247. 384. Wilson SJ, Webb BL, Ylinen LM, et al. Independent evolution of an antiviral TRIMCyp in rhesus macaques. Proc Natl Acad Sci U S A 2008;105:3557–3562. 385. Wojcechowskyj JA, Yant LJ, Wiseman RW, et al. Control of simian immunodeficiency virus SIVmac239 is not predicted by inheritance of Mamu-B*17-containing haplotypes. J Virol 2007;81:406–410. 386. Wu F, Kirmaier A, Goeken R, et al. TRIM5 alpha drives SIVsmm evolution in rhesus macaques. PLoS Pathog 2013;9:e1003577. 387. Wu F, Ourmanov I, Kuwata T, et al. Sequential evolution and escape from neutralization of simian immunodeficiency virus SIVsmE660 clones in rhesus macaques. J Virol 2012;86:8835–8847. 388. Wyand MS, Manson K, Montefiori DC, et al. Protection by live, attenuated simian immunodeficiency virus against heterologous challenge. J Virol 1999;73:8356–8363. 389. Wykrzykowska JJ, Rosenzweig M, Veazey RS, et al. Early regeneration of thymic progenitors in rhesus macaques infected with simian immunodeficiency virus. J Exp Med 1998;187:1767–1778. 390. Yamamoto JK, Sparger E, Ho EW, et al. Epidemiologic and clinical aspects of feline immunodeficiency virus infection in cats from the continental United States and Canada and possible mode of transmission. J Am Vet Med Assoc 1989;194:213–220. 391. Yan J, Shun MC, Hao C, et al. HIV-1 Vpr reprograms CLR4(DCAF1) E3 ubiquitin ligase to antagonize exonuclease 1-mediated restriction of HIV-1 infection. mBio 2018;9(5):e01732-18. 392. Yan J, Shun MC, Zhang Y, et al. HIV-1 Vpr counteracts HLTF-mediated restriction of HIV-1 infection in T cells. Proc Natl Acad Sci U S A 2019;116:9568–9577. 393. Yant LJ, Friedrich TC, Johnson RC, et al. The high-frequency major histocompatibility complex class I allele Mamu-B*17 is associated with control of simian immunodeficiency virus SIVmac239 replication. J Virol 2006;80:5074–5077. 394. Yap MW, Nisole S, Stoye JP. A single amino acid change in the SPRY domain of human Trim5alpha leads to HIV-1 restriction. Curr Biol 2005;15:73–78. 395. Ylinen LM, Keckesova Z, Webb BL, et al. Isolation of an active Lv1 gene from cattle indicates that tripartite motif protein-mediated innate immunity to retroviral infection is widespread among mammals. J Virol 2006;80:7332–7338. 396. Yu Q, Konig R, Pillai S, et al. Single-strand specificity of APOBEC3G accounts for minus-strand deamination of the HIV genome. Nat Struct Mol Biol 2004;11:435–442. 397. Yu S, Ou Y, Xiao H, et al. Experimental treatment of SIV-infected Macaques via autograft of CCR5-disrupted hematopoietic stem and progenitor cells. Mol Ther Methods Clin Dev 2020;17:520–531. 398. Yuste E, Reeves JD, Doms RW, et al. Modulation of Env content in virions of simian immunodeficiency virus: correlation with cell surface expression and virion infectivity. J Virol 2004;78:6775–6785. 399. Zhang B, Jin S, Jin J, et al. A tumor necrosis factor receptor family protein serves as a cellular receptor for the macrophage-tropic equine lentivirus. Proc Natl Acad Sci U S A 2005;102:9918–9923. 400. Zhang B, Montelaro RC. Replication of equine infectious anemia virus in engineered mouse NIH 3T3 cells. J Virol 2009;83:2034–2037. 401. Zhang F, Bieniasz PD. HIV-1 Vpr induces cell cycle arrest and enhances viral gene expression by depleting CCDC137. Elife 2020;9:e55806. 402. Zhang F, Wilson SJ, Landford WC, et al. Nef proteins from simian immunodeficiency viruses are tetherin antagonists. Cell Host Microbe 2009;6:54–67. 403. Zheng X, Wang X, Tu F, et al. TRIM25 is required for the antiviral activity of zinc finger antiviral protein. J Virol 2017;91(9):e00088-17. 404. Zhu Y, Chen G, Lv F, et al. Zinc-finger antiviral protein inhibits HIV-1 infection by selectively targeting multiply spliced viral mRNAs for degradation. Proc Natl Acad Sci U S A 2011;108:15834–15839.

765

CHAPTER 20 Foamy Viruses Dirk Lindemann • Ottmar Herchenröder Overview Foamy virus isolation and diagnosis of infection Natural history and trans-species transmissions Evolution of foamy viruses Replication in vitro Replication in the natural host in vivo Apathogenicity of foamy viruses Virion structure Genome structure and organization Virion-associated proteins Gag Pol Env Nonstructural proteins Tas Bet Stages of replication The early phase: establishing the provirus (Fig. 20.14) The late phase: generation of progeny viruses (Fig. 20.15)

OVERVIEW Foamy viruses (FVs) are found in numerous mammalian species. Basically all simian and prosimian species investigated to date harbor exogenous FVs. Otherwise, FVs were found for instance in cattle, horses, and felines. Human beings are no natural hosts for FV, although transmissions from apes or monkeys to man do occur since these viruses exhibit a broad tropism with respect to both, cell and species type. Like all other retroviruses, FVs establish lifelong persistent infections but according to current knowledge do not cause any obvious disease, be it within the species of origin or after zoonotic transmission. Although the FV genomes are typical for complex retroviruses, their polymerases are phylogenetically related to those of other retroviruses, and their reverse transcription mechanism employs a tRNA primer. Consequently, FVs were classified as a separate subfamily in the Retroviridae family, the Spumaretrovirinae, because their replication cycle deviates from that observed in all other retroviruses. Therefore, and with respect to their potential use as vectors systems and to gain valuable insights into zoonotic virus transmissions, this “Cinderella” group of viruses is worth to be thoroughly investigated.283,284

The term “foamy virus” was coined in the 1950s to acknowledge the spontaneous formation of a typical foamy cytopathic effect (CPE) observed in FV infected cell cultures (Fig. 20.1; Video 20.1). This CPE is characterized by multinucleated syncytia and vacuolization in primary monkey kidney cell cultures causing a “foamy” appearance.53,174,214,230 The CPE observed before destruction of the cell cultures was later attributed to the fact that the donor monkeys were latently infected with a transmittable agent, and it became the light microscopy hallmark of in vitro FV propagation. Following the discovery of reverse transcriptase, FVs were identified to be retroviruses.193 The first 30 years of FV research dealt mainly with the identification of infected monkeys, prior to sacrifice, as sources of primary cell cultures used to diagnose human transmissible diseases.41 Molecular cloning of the first FV—at that time believed to be a human isolate—permitted initial functional studies on their replication.216 FV research gained momentum following the discovery that these viruses replicate differently from all other retroviruses.151,298 These studies culminated in the finding that the FV infectious genome consists of DNA rather than RNA.180,225,303 In brief, the FV replication strategy combines those of retroviruses with some characteristics of hepadnaviruses, such as hepatitis B virus (HBV), with other properties unique to FVs.133,212,214 Consequently, retroviral taxonomy was updated into two retroviral subfamilies, the Orthoretrovirinae, which encompass all other retrovirus genera, and the Spumaretrovirinae, which currently contain five genera of spumaviruses (spuma, latin for foam).125,152 This chapter will summarize our knowledge of FV epidemiology and biology. We shall focus on FVs of nonhuman primates (NHPs) and mention the nonprimate viruses only when they become relevant depending on the scientific context. For particular aspects such as FV vectors, the reader is referred to more specialized reviews.148,149,204,206,252

766

767

vector (lane 2, mock), lysed, and precipitated with a foamy virus-positive chimpanzee serum. Immunodominant proteins are indicated by arrowheads (left) and molecular weight markers are shown on the right.

NATURAL HISTORY AND TRANS-SPECIES TRANSMISSIONS Various vertebrate species are naturally infected with FVs. Figure 20.3 gives an overview of the currently known exogenous spumavirus genera and FV species. Extensively studied are simian foamy viruses (SFVs) of Great Apes, Old and New World monkeys, and prosimians.104,105,174,214,235 Probably all monkey species harbor a FV.113,154 Otherwise, FV infections appear to occur worldwide in bovines and other Artiodactyla, equines, and felidae, and one report shows FVs in a bat species.66,125,134,167,174,214,218,271 The prevalence in the natural hosts in the wild is usually over 30% and may reach 100%.104,154,174,214 It is generally assumed that FVs are transmitted among NHPs through saliva via social contacts, including aggressive activities such as biting among young animals.32,154 These contacts and lactation are also suspected to be the main transmission route of the feline foamy virus (FFV), whereas bovine foamy virus (BFV) probably is mainly transmitted via milk from infected cows to their offspring.220,291 It is, therefore, not unlikely that FVs are, in terms of prevalence, the most successful of all retroviruses. Although host restriction factors show some species restriction (see later discussion), FVs have been reported to cross the species barrier between monkeys and apes in captivity or in the wild.104,139,154 In their hosts, FVs cause lifelong persistent infections of benign nature, often in the presence of neutralizing antibodies.104,174,214 Laboratory animals, such as mice and rabbits, have also been infected in the absence of overt disease.28,232,240 However, none of these reports yielded a suitable animal model that mimics replication in the natural host.

Humans are not a natural host of FVs.214 Indeed, the most intensely studied FV isolate was once believed to be of human origin but is meanwhile designated as the prototype foamy virus (PFV).1,161,213 Initial reports on naturally occurring human infections162,166 were not confirmed after large-scale screening of more than 5,000 samples using sophisticated methodological combinations of serology and PCR.3,247 Formerly postulated associations of FVs with human diseases such as autoimmune disorders or neurologic conditions of unknown origin could never be validated.45,97,223,248 Because of the close nucleotide sequence homology to FVs from the Pan troglodytes verus chimpanzee subspecies, initially called SFVcpz and now designated SFVpve,99,154 the first documented human isolate from a Kenyan patient is believed to have resulted from a zoonotic transmission.58 Primate FVs are not circulating in the human population, but humans are susceptible to zoonotic transmissions of NHP FVs.204,244,261 Worldwide, over a hundred human infections with NHP FVs have been thoroughly confirmed. In a survey, the Centers for Disease Control and Prevention identified approximately 2% seropositives and virus DNA positives among several hundred samples from persons occupationally exposed to NHPs, and virus could be isolated in several instances. Some of these infections dated back to severe monkey bites decades before.24,98 FV infections were also identified in African bushmeat hunters.31,71,292 Other persons at risk are those living in close proximity to quasi-wild NHPs (e.g., at Asian temple sites) or individuals possessing NHP as pets.114

FIGURE 20.3 Spumaretrovirus phylogeny. The tree was generated using aligned nucleotide sequences beginning approximately in the middle of the pol gene and extending to approximately the middle of the env gene, corresponding to positions 5089–7927 of SFVpsc. Unrooted tree is depicted as rooted for ease of visualization. Branches are labeled with virus names; corresponding host names are given in parentheses. Branches corresponding to viruses within the same genus are indicated with colored boxes and genus names. Percent bootstrap support (100 replicates) is indicated for each node. Sequences were aligned using MUSCLE 3.8.425 and phylogeny inferred using PhyML 3.2.2 (HKY85 substitution model and the NNI search option) as implemented in Geneious Prime 2020.2.3. Image was created using FigTree 1.4.4 and Adobe Illustrator CC. Virus genome accession numbers are SFVcni (JQ867466); SFVcae (NC_010820); SFVpan (MK241969); SFVmfa (LC094267); SFVmcy (MN585198.1); SFVmmu (MF280817); SFVmfu (AB923518); SFVpve (NC_001364); SFVpsc (KX087159); SFVptr (JQ867463); SFVggo (HM245790); SFVppy (AJ544579); SFVhpi (MF621235); SFVsxa (KP143760); SFVcja (GU356395); SFVaxx (EU010385); SFVssc (GU356394); SFVocr (KM233624); FFVfca (Y08851); EFVeca (AF201902); BFVbta (NC_001831); and CFVraf (JQ814855). (Courtesy of A.S. Khan, Silver Spring, USA and W.E. Johnson, Chestnut Hill, USA.) 768

Human infections are lifelong; however, none induced any disease and remained unrecognized prior to respective investigations. The viral load in buffy-coat cells of infected humans revealed very low FV DNA copy numbers.31 Moreover, in contrast to lentiviruses, in vivo adaptation to what could be called a human FV has not occurred. Even decades after infection, FV nucleotide sequences in humans barely change, and the transmitting donor species can be readily identified as over time, a baboon FV will remain a baboon FV in a human host.24,98,250 There is no indication of human-to-human transmission even between close contacts.204 But since FV can be transmitted by transfusion,27,124 infected persons are advised not to donate blood to avoid virus spreading.96 Special cases were FV transmissions to end-stage diseased recipients of NHP xenotransplants.5 For reasons not fully understood, humans appear to be “dead-end” hosts for primate FVs. Concerning infections in humans with other than NHP FVs, it is worth mentioning that antibody screening of more than 200 veterinarians at risk of acquiring FFV did not reveal a single positive case.29 Based on this result, it appears also unlikely that bovine or equine foamy virus can infect humans.204 Why throughout the evolution of humans FVs have not adapted to this species remains a mystery.

EVOLUTION OF FOAMY VIRUSES Exogenous FVs are ancient retroviruses. While the prominent pathogenic retroviruses like simian immunodeficiency viruses (SIVs) and simian T-lymphotropic viruses (STLVs), or the D-type retroviruses including Mason-Pfizer monkey virus (MPMV) are only inherent in African or Asian monkeys, respectively, the higher evolutionary age of primate FV is reflected in that they are the only known exogenous primate retroviruses also found in the Neotropis, precisely in South and Central America.235 Over the last decade, researchers have uncovered a number of endogenous foamy viral sequences in several species testifying that exogenous FVs were around before the advent of mammals. First, Katzourakis et al.122 reported the detection of endogenous FVs in the South American sloths genome (Choloepus hoffmanni). Han and Worobey described endogenous FVs in a lemur species, the Madagascan aye-aye (Daubentonia madagascariensis), in the Coelacanth genome (Latimeria chalumnae), and the Cape golden mole (Chrysochloris asiatica), suggesting that endogenous FVs existed more than 400 million years ago.82 An evolutionary gap was filled after detecting endogenous FV in the reptile species tuatara (Sphenodon punctatus).281 Currently, and on the basis of additional foamy-like endogenous sequences in fish and amphibians, it is believed that this retroviral lineage and probably retroviruses as a whole originated in ancient marine life.2

Apparently, when an exogenous FV has adapted to its host, it mutates only slightly faster than the host mitochondrial DNA. The substitution rate of SFVs has been estimated to be around 1.7 × 10−8 per site and year.263 This makes FVs the most genetically stable of viruses with an RNA phase in their replication. For instance, the FV mutation rate is approximately 10 times lower than STLV-1, a virus that replicates primarily by a proviral expansion mechanism, that is, through DNA (see Chapter 16). Therefore, FVs are of extraordinary genetic stability and, if not acquired by trans-species transmissions, always point to the host species from which they were derived. Curiously, the fidelity of the reverse transcriptase enzyme (RT) does not reflect this enormous genetic stability. If analyzed in vitro, the PFV RT was found to be of exceptional high processivity but of low fidelity.25,26,219 The mutation rate of PFV RT (~1.7 × 10−4 per site and replication round) is similar to that of human immunodeficiency virus (HIV). Most mutations found were small deletions and insertions.26 If analyzed in cell culture, such deletions and insertions were not found. However, a point mutation error rate of 1.1 × 10−5 per site and round of replication remained.68 Thus, the genetic stability of FVs at the molecular level is not currently understood, and the involvement of yet unidentified specific cellular factor(s) or the very limited replication in specific tissues in persistently infected hosts may play a role in this process.

REPLICATION IN VITRO The host cell range for FVs is quite broad and includes species-independent primary cells or cell lines of fibroblastoid, epithelial, and lymphoblastoid origin, such as various B and T lymphocytes, and cells of erythroid and of myeloid lineages.104,174,214,302 Upon replication in adherent cell cultures, FVs induce massive multinucleated giant cell CPE (Fig. 20.1; Video 20.1), and apoptosis is thought to be the ultimate cause of cell death.178 Vacuolization of cells is often only observed using primary isolates. The paucity of cell lines resistant to FV infections or FV glycoprotein complex (GPC)-mediated membrane fusion has hindered the identification of the cellular receptor(s) required for entry by classical approaches. It is now appreciated that all FVs use the same cellular receptor, including those present on bird, reptile, and fish cells.13,101,258 Two cell lines reported to be refractory to infection, Pac-2 zebrafish embryonic fibroblasts and the G1E-ER4 human erythroid precursors,258 were later found to block virus replication intracellularly. Thus, systematic screenings of complementary DNA (cDNA) libraries for FV receptor-related genes are still pending.

Whereas the characteristic CPE develops in adherent cells, this hallmark of FVs is often absent in cells of lymphoblastoid origin, in which FVs appear to become latent but intermittently capable of being reactivated. Latently infected cells do not undergo syncytium formation and death but proliferate with normal kinetics and produce low amounts of virus.302 Interestingly, chemical treatment of lymphocytes, for example, with phorbol esters may induce the latent virus and cause cell death owing to activated viral replication.177,302 Although this is reminiscent of the lymphotropic herpesviruses, the molecular basis for FV reactivation has not been investigated nor have sites within the cellular or FV DNA genome responsive to the drug-mediated reactivation been mapped. Whether the methylation of FV DNA that has been observed in a cell culture model245 contributes to in vivo latency remains unresolved, because there is no evidence of transcriptional down-regulation of FV vectors by methylation following their introduction in vivo.102,190

REPLICATION IN THE NATURAL HOST IN VIVO Liu et al.154 used methods similar to those employed to demonstrate that HIV-1 was derived from SIV from chimpanzees (SIVcpz; i.e., the authors collected and analyzed fecal samples from wild chimpanzees; see Chapter 19). It was found that SFVs are widely distributed among chimpanzees in the wild with a phylogeographic distribution. In these animals, FVs are transmitted horizontally, because babies younger than 2 years were free from FVs and infection rates increased with age. Moreover, superinfection of chimpanzees by SFVs from lower primates and subsequently recombination events occur. The most interesting finding by these authors has been the detection of viral RNA (vRNA) but not viral DNA (vDNA) in the fecal samples.154 This issue directly relates to the FV replication pathway (see later discussion). However, Liu et al.154 did not investigate whether the RNA-containing virus transmits the infection, and it is not known in which cell type these viruses were produced. It is possible that the DNA content in the fecal samples may have been too low to be detected even with very sensitive methods.

769

The tissue distribution of SFV and sites of in vivo replication have been investigated using another approach. As expected from the broad host cell range of FVs seen in vitro, vDNA was detected at a frequency of one genome copy per 102 to 103 cells in every organ examined.62 Because animals in this study were perfused prior to the analysis of nucleic acids, infected lymphocytes were not detected, although these cells were the probable vehicles of virus dissemination in vivo.62,63 As judged from in situ hybridization experiments, vRNA, indicative of active virus replication, was confined to superficial cells of the oral mucosa.62,185 Thus, it appears that only cells, which are destined to be shed, are productively infected and undergo lytic replication in vivo. Differential expression of yet undisclosed host factors restricting viral replication in other tissues is a likely explanation for this observation.

It was subsequently found that in severely immunosuppressed, dually SFV- and SIV-infected macaques, the predominant site of FV replication changed from the oral mucosa to the small intestine.184 However, diseases attributable to SFV did not occur. This was also observed in cats dually infected with feline immunodeficiency virus (FIV) and FFV.11,306 One case of a human coinfected by HIV-1 and SFV from mandrills has also been reported, without clinical consequences that could be attributed to the SFV infection.262

APATHOGENICITY OF FOAMY VIRUSES For decades in the last century, several pathogenic conditions in humans were attributed to PFV, formerly called human foamy virus (HFV) or in some earlier publications human spumaretrovirus (HSRV). Prompted by a gathering in the mid-1990s of all laboratories involved in FV research and subsequent cross-examinations on sera, two essential perceptions were agreed upon: humans do not naturally harbor FV and neither in the natural hosts nor after zoonotic transmission to man do FV cause any disease.284 After it was proven that PFV, originally isolated from an African patient,1 had originated from a chimpanzee, and thorough analyses of samples from patients with diseases once attributed to FV infections, this consensus was sustained.99,243 This notion seems strange, since to the layperson, a virus infection and a disease seem to belong together. Instead, the pathogenicity of an infectious agent may require an explanation, not the apathogenicity of what has been called a “perfect parasite” whose “interest” is to multiply its genome without doing harm, with the FVs as a prominent example.276 It appears that FVs have coevolved with their hosts over millions of years to do exactly this.2

Certainly, some arguments can be made in favor of this hypothesis. The site of active replication in vivo determines to a large extent the pathogenicity of an infectious agent. FVs appear to mainly replicate in cells of the oral and occasionally the intestinal mucosa, both tissues, that are destined to be desquamated.62,63 However, during the establishment of persistence, it is likely that lymphocytes become infected and shed low amounts of virus that disseminate throughout the body. Once persistence is established, FV genes may no longer be expressed in lymphocytes. A second factor contributing to the benign character of FV infections is that the Tas transactivator appears to be specific for its autologous cognate viral promoters.100,155 The off-target activation of gene expression upon zoonotic transmission may probably be a rare event, although Tas has been demonstrated to activate transcription of some cellular genes in vitro.128,278 Furthermore, the integration of FVs does not induce the activation of cellular oncogenes because they lack strong enhancer elements and their LTRs were recently reported to harbor insulator sequences.61,78 In addition, a strong polyadenylation signal in the LTR seems to prevent a read-through of viral transcripts into cellular genes for activation.95,242

Meanwhile, newer reports challenge to some extent the fully innocent nature of FVs, however, just as a cofactor in conjunction with other conditions. Cameroonian and French researchers thoroughly examined hunters residing in the Cameroonian rain forest of whom numerous contracted zoonotic transmissible agents and among those NHP FV. Gorilla FV for instance seems to alter hematological parameters inversely correlated to the presence of strong neutralizing antibodies.132 A case–control study revealed that the cases had numerous deviations in their immune parameters compared to the controls.71 In such FV-positive individuals, coinfections with other retroviruses do occur and the viruses may influence each other.226 After experimental infection of two rhesus monkey cohorts with the pathogenic SIVmac239 strain, the naturally FV-positive animals developed higher SIV plasma loads, their CD4-positive T cells decay was steeper, and they died earlier than the FV-negative monkeys.40 Appreciating the latter reports and during the continuous inventory of the planet’s virome, FVs should, therefore, not be released from the watch list.

VIRION STRUCTURE By ultrastructural analysis, FVs appear as immature-looking core particles surrounded by a lipid bilayer with embedded prominent Env proteins104,174,214,287 (Fig. 20.4; Video 20.2). Investigations using electron microscopy (EM) or electron tomography (ET) techniques suggest that the FV virions have a diameter of approximately 110 nm and contain a core of about 60 nm diameter.52,104,174,214,287 The core has an isometric, angular, and immature morphology owing to the very limited cleavage of the Gag precursor protein by the viral protease (PR) (see later discussion). The cores of infectious FV virions are made up of the Gag precursor (pr71Gag for PFV) and its large processing product (p68Gag for PFV) at a ratio of 1:1 up to 1:435 (Fig. 20.2). This Gag doublet is inherent in all FVs,79 although the nonprimate FV capsid proteins are considerably smaller than their primate relatives.103,134,221,271 Whether the FV core is made up of a hexameric lattice as observed for mature orthoretroviral capsids is currently unknown. However, the recently determined 3D structure of the central, capsid-like Gag domain, which share the same core fold as the N- (NtDCA) and C-terminal domains (CtDCA) of archetypal orthoretroviral CAs, may suggest so. In addition, the FV core is characterized by being surrounded by an unusual, less ordered shell of density, termed matrix (MA) layer or intermediate shell (Fig. 20.4B and C). It follows the virus membrane at budding, subsequently relocates to the capsid’s edges in released virions, and displays different width characteristics for individual FV genera.287 The intermediate shell is likely formed by the Gag N-terminal domain (NtD) for which a subdomain structure is available,77 which points outward to the cytoplasmic domains (CyDs) of the FV GPC.52,287

770

FIGURE 20.4 Structure of foamy virus (FV) virions. A: Schematic representation of the prototype foamy virus (PFV) particle structure prior to reverse transcription initiation. pr, precursor protein; p, protein; gp, glycoprotein. B: Cryo-electron tomography of a wild-type PFV virion. The left image has the central, polyhedral capsid structure highlighted in red and two oligomeric glycoprotein spike structures in green. Scale bar: 50 nm. (Courtesy of P. Rosenthal and T. Calcraft, London, UK.) C: Electron micrograph of negative stained PFV budding structures at cellular membranes of 293T cells transfected with proviral expression constructs. Scale bar: 100 nm. (Panel A adapted from Lindemann D, Rethwilm A. Foamy virus biology and its application for vector development. Viruses 2011;3(5):561–585.)

771

FIGURE 20.5 Principle replication strategies of reverse transcribing viruses. Orthoretroviruses (left panel) are RNA viruses that reverse transcribe early in replication, replicate through a DNA intermediate, and exhibit obligate virus-mediated integration into the cellular genome. Hepadnaviruses (right panel) are essentially DNA viruses, replicate through an RNA intermediate, and do not integrate. The retroviral subfamily of spumaretroviruses (middle panel) functionally combines both pathways by being DNA viruses that reverse transcribe late in replication like hepadnaviruses and integrate like orthoretroviruses. In addition to RTr in late phases of FV particle morphogenesis, it has been shown that, similar to orthoretroviruses, RTr also occurs during the early phases of FV replication upon target cell entry in vitro.46,304 Given the aforementioned ratios of RNA to DNA in extracellular viruses generated by in vitro propagation, RTr during the early phase of FV replication in vitro may only become relevant at a very low MOI, when the amount of virion DNA may be too low to sustain a productive infection.304 Furthermore, the discrepant results reported about the relevance of the differentially timed RTr events for FV infectivity may reflect the inherent differences in the cell types used for virus production.46,171,180,225,303,304 In essence, the generally accepted view that virions contain either an RNA or DNA genome, but not both, may not apply to FVs.

The physical stability of FVs has not been directly compared with that of orthoretroviruses. However, owing to their immature core and a particular Env topology (see later discussion), these virions are probably quite stable. This is illustrated by the fact that vector particles can be concentrated more than 100-fold (i.e., by ultracentrifugation) without significant loss of infectivity.72,101

GENOME STRUCTURE AND ORGANIZATION All FV genomes share common genome structures.215 The schematic representation of the PFV genome is shown in Figure 20.6. The general organization of the canonical gag, pol, and env genes and accessory open reading frames (ORFs) downstream of env reaching into the 3′ LTR resembles other complex retroviruses. FVs encode two such ORFs, which in older literature were designated bel1 and bel2. The first ORF (tas) encodes the transcriptional transactivator Tas, and orf-2 is partially used to express a fusion protein designated Bet. In PFV, Bet consists of the first 88 amino acids of Tas fused to the bulk of the coding capacity of orf-2. It is noteworthy that no independent protein is expressed from orf-2 alone in any of the known FVs.

772

FIGURE 20.6 Foamy virus provirus organization, genomic and structural gene transcripts, and essential cis-acting viral RNA sequence elements. Top: Schematic illustration of the prototype foamy virus (PFV) proviral DNA genome structure with long terminal repeats (LTRs) and ORFs indicated as boxes. For ORFs encoding Gag, Pol, and Env precursor proteins, the regions encompassing the mature subunits generated by proteolytic processing are indicated in differential colors and labeled accordingly. Underneath, unspliced and spliced transcripts originating at the LTR or internal promoter (IP) including the full-length viral RNA genome (vgRNA) or giving rise to the viral structural (Gag, Pol, Env) and accessory (Tas, Bet) proteins are indicated. The respective coding capacities of individual transcripts are indicated to the right. The bimodal transcriptional stimulation of FV transcription by Tas at both, the IP and the LTR promoter, is indicated by dashed yellow arrows. Bottom: Cis-acting sequence (CAS) elements localized within the full-length vgRNA, which are essential for viral replication, are indicated by black bars underneath. Functionally important or essential RNA sequence motifs are marked in each enlarged CAS element below. Numbers represent nucleotide positions of the PFV vgRNA (HSRV2 isolate). Individual regions within the vgRNA essential for specific functions in viral replication as indicated to the right are marked as differentially colored bars. U3, unique 3′ LTR region; R, repeat LTR region; U5, unique 5′ LTR region; miRNAs, microRNA Pol III expression cassettes; p68, Gag p68 subunit; p3, Gag p3 subunit; PR, Pol protease domain; POLY, Pol polymerase domain; RH, Pol RNase H domain; IN, Pol integrase subunit; LP, Env leader peptide subunit; SU, Env surface subunit; TM, Env transmembrane subunit; ©, cap structure; An, poly-A tail; mSD, major splice donor; PBS, primer binding site; DIS, dimerization sites; PARM, protease activating RNA motif; SRE, splicing regulatory element; cPPT, central poly-purine tract; 3′ PPT, 3′ poly-purine tract; pA, polyadenylation signal; A–D, purine-rich (PuR) sequence motifs A through D; RTr, reverse transcription. (Adapted from Lindemann D, Rethwilm A. Foamy virus biology and its application for vector development. Viruses 2011;3:561–585.)

Unique for all FVs is a second, internal promoter (IP) near the 3′ end of the env gene, which mainly drives expression of the accessory ORFs155,158 (Fig. 20.6). FV proviruses are 12 to 13 kb long and thereby larger than those of most other retroviruses. Their size is partially attributed to the extraordinary long U3 regions of the LTR, which can be explained in part by the overlapping orf-2 (Fig. 20.6). However, in the case of PFV, the accessory orf-2 reaches only 300 bp into the 5′ end of the more than 1.4 kb long U3 region. In addition, very few enhancer elements, such as those for AP-1 and Ets-1,173,239 and the short sequence motifs responsive to the viral transactivator (see later discussion), are present at the 3′ end of U3. The enhancer elements are involved in regulating LTR-derived expression of the structural genes. For many years, several hundred bases in the U3 region remained devoid of known functions until the discovery of one or more RNA Pol III-directed microRNA (miRNA) cassettes129,285 (Fig. 20.6). For BFV and SFVcae, miRNA expression was experimentally confirmed and a first study examining the possible functions of BFV miRNAs suggest a role in modulating cellular antiviral defense mechanisms.33 Bioinformatic analysis suggests that nearly all FV genomes contain at least one miRNA cassette.129,170

There are length differences in the gag genes in different FVs, with those in FVs from cats, bovines, and equines being shorter than those from primates.103,172,271,290 In sharp contrast to their orthoretroviral cousins, the FV Gag proteins are more variable than the Env proteins. Primate lentiviruses for instance have an amino acid conservation of roughly 60% in Gag and 40% in Env compared to 45% in Gag and 65% in Env among primate FVs.214,267 It is likely that this curiosity of FV biology is a consequence of the process of adaptation to and coevolution with their natural hosts. Furthermore, this finding is consistent with the use of the same cellular receptor(s) by all FVs.

Like orthoretroviruses, FV genomes contain elements, designated as cis-active sequences (CAS) that harbor secondary and tertiary structures essential for various aspects of viral replication (Fig. 20.6). The first region, CAS-I, is located in the 5′ untranslated leader of the vgRNA and extends into the gag gene. It includes the R-U5 region of the LTR, the primer binding site (PBS), which is complementary to the cellular transfer RNA(Lys1,2) and conserved among all FVs and elements conferring dimerization of the vgRNA.30,60,172 CAS-I has functions in RTr and splice regulation as well as in vgRNA and Pol encapsidation.92–94,153,293

However, FVs require another cis-acting genomic RNA element, CAS-II, for viral infectivity and FV vector function59,91,93,153,293 (Fig. 20.6). CAS-II is reported to harbor additional RNA packaging elements in its 5′ part, while its 3′ portion contains the major Pol encapsidation signal (PES) and four purine-rich sequence elements (PuR-A to -D).197,198,286 PuR-A and -B appear to be essential elements of the PES element of CAS-II and have been reported to promote FV PR-RT dimerization (protease-activating RNA motif, PARM) and thereby probably Pol precursor encapsidation.86 PuR-C was shown to represent a splicing regulatory element (SRE) preventing further processing of full-length Gag and singlespliced Pol mRNA.181 PuR-D, found in all sequenced FV genomes, has been demonstrated to serve as a second internal or central poly-purine 773

tract (PPT), similarly to what has been reported for HIV, as an initiation site for plus-strand DNA synthesis during RTr, in addition to the 3′ PPT upstream of the 3′ LTR7,197,246,272,286 Finally, a third element, CAS-III, is located far downstream on the vgRNA. It contains the 3′ PPT and the LTR (U3-R) sequences needed for RTr and integration as well as transcription initiation and polyadenylation upon proviral integration148 (Fig. 20.6).

VIRION-ASSOCIATED PROTEINS Gag The FV capsid proteins have several unusual characteristics compared to orthoretroviral Gag proteins. They are neither cleaved into the canonical matrix (MA), capsid (CA), and nucleocapsid (NC) subunits nor are several sequence motifs present, which are conserved in all orthoretroviral Gag proteins (Fig. 20.7). These include the N-terminal myristoylation signal of the MA domain, the major homology region in the CA domain, or the cysteine-histidine (CH) boxes in the NC domain. Instead of Gag subunit processing observed with orthoretroviruses, at least half of the particle-associated FV Gag molecules are truncated approximately 3 to 4 kD C-terminally by pol-encoded PR processing at a singular cleavage site. In the case of PFV, this generates a large p68Gag and a small p3Gag product from the pr71Gag precursor molecule67 (Fig. 20.7A). Whereas the p68Gag cleavage product together with the pr71Gag precursor forms the capsid of free PFV virions, the smaller p3Gag so far could not be detected in extracellular virus particles (Figs. 20.2 and 20.4A). Whether this is due to limitations in the sensitivity of the detection methods employed, or may indicate that Gag precursor processing starts before the PFV capsid is fully assembled and this small processing product thereby evades encapsidation, is unclear. The p3Gag postprocessing cellular localization, fate, function, and possible functions remain to be elucidated.55 FV Gag cleavage is required for infectivity, as FVs expressing only the pr71Gag precursor are not infectious and often produce aberrantly formed capsids.55,130,305 Mutants expressing only the large p68Gag cleavage product are infectious albeit at 10- to 50-fold lower titers.55,108,257,305 Secondary protease cleavage sites, located in the central part of FV Gag, have been identified in vitro, using recombinant proteins and peptides200 (Fig. 20.7A). They are proposed to be utilized for a viral disassembly process involving proteolytic processing of Gag by the FV PR and cellular proteases following entry into target cells.73,140,200 However, the essential requirement of the Gag processing by FV PR at the secondary cleavage sites for viral replication is a matter of debate.108,140 Thus, FV capsid disassembly appears to be a unique process that involves viral and cellular protease-mediated processing.

In the FV Gag precursors, currently five major regions or domains can be structurally and functionally distinguished (Fig. 20.7A): an NtD; a proline-rich (P-rich) region of variable length mainly responsible for the size variation among Gag precursor proteins of different FV genera (52 to 71 kDa); a central capsid (CA) domain; a glycine–arginine–rich (GR-rich) region; and the C-terminal p3Gag domain, with the first four located in the p68Gag subunit. For PFV, of two of those, 3D structures were recently solved, namely the 180 aa NtD and the 178 aa central CA10,77 (Fig. 20.7B and C).

Various peptide and structural motifs have been identified and characterized to variable extent in FV Gag proteins (Fig. 20.7). Four coiled-coil domains (CC1-4) are predicted to be present in PFV Gag,199,269 and functions have been assigned to the first three located in the NtD (Fig. 20.7A and B). The N-terminal CC1 (aa 14–24) has been suggested to interact with the CyD of the Env leader peptide (LP) according to initial biochemical analyses.34,146,207,288 A direct interaction was confirmed later by 3D structural data of Gag NtD-Env LP peptide cocrystals77 (Fig. 20.7B). CC2 (aa 130–146) has been reported to harbor a domain necessary for Gag multimerization,269 and CC3 (aa 161–180) is believed to be required for incoming capsids to interact with the dynein light chain 8 for retrograde movement along the cellular microtubule network to the microtubule organizing center (MTOC).199 The Gag NtD 3D structure revealed an extended, single coiled-coil structure comprised by CC2 and CC3 essential for dimerization of the NtD.77 The function of CC4 (aa 436–453), which is located at the C-terminus of the PFV Gag CA domain (Fig. 20.7C), whose 3D structure was recently determined,10 remains unknown.

774

FIGURE 20.7 Schematic representation of the prototype foamy virus (PFV) Gag protein organization and subdomain 3D structures. A: PFV Gag protein organization and selected functional motifs highlighted by the colored boxes. The organization of the N-terminal domain (NtD) and the central capsid-like (CA) domain are shown in the enlargements in B and C. Numbers indicate amino acid (aa) sequence positions of PFV Gag. The black arrow indicates the viral protease cleavage site of pr71Gag for processing into p68Gag and p3Gag utilized during virus morphogenesis, gray arrows mark secondary cleavage sites of potential processing upon virus entry. Gray boxes represent the proline-rich (P-rich) and glycine–arginine–rich (GR-rich) regions, respectively. CC1 to CC4, coiled-coil domains 1 to 4; CTRS, cytoplasmic targeting and retention signal; NES, nuclear export sequence; PLKBS, polo-like kinase binding site; L, late budding domain motif; A, assembly motif YxxLGL; A′, assembly motif PGQA; CBS, chromatin binding sequence. B: Crystal structure of PFV Gag NtD-Env LP complex (PDB ID: 4JMR). Upper, PFV Gag enlargement showing just the PFV Gag NtD, secondary structure elements indicated above. Lower, cartoon representation of the crystal structure of the PFV Gag NtD homodimer. The functional sequence motifs, CC1 Env-binding region (blue), CTRS (orange), NES (green) and CC2-3 dimer interface coiled-coil (cyan) are color coded as in the schematic. Residue side chains in the GWWGQ core of the CTRS are shown in stick representation. The helical regions of the Env-LP bound at the periphery of the Gag-NtD head domain are colored gold. C: 3D solution NMR structure of the central PFV Gag capsid (CA) domains (PDB ID: 5M1H). Upper, PFV Gag CA enlargement with secondary structure elements indicated above. Lower, cartoon representation of the PFV Gag CA structure. The backbone of the N-terminal capsid domain (CA-NtD) and C-terminal capsid domain (CA-CtD) are shown in pink and brown. Helices in the structure are numbered sequentially from N- to C-terminus in the left- and right-hand views. The CC4, PGQA, and YxxLGL sequence motifs are color coded as in the schematic. Residue side chains in the PGQA and YxxLGL sequence motifs in the α7-α8 connecting loop and C-terminal of α9 are shown in stick representation. (Panel A adapted from Lindemann D, Rethwilm A. Foamy virus biology and its application for vector development. Viruses 2011;3(5):561–585. https://creativecommons.org/licenses/by/3.0/. panels B and C 3D-ribbon structure cartoons of panels B and C courtesy of Ian A. Taylor, London, UK)

C-terminally of CC4 the PFV Gag CA domain also harbors an evolutionary conserved YxxLGL sequence termed assembly motif (A-motif; aa 464–469) (Fig. 20.7A and C) shown to be essential for both, capsid assembly and RTr.168 The PFV Gag CA structure revealed the presence of two all α-helical domains (CA-NtD) and (CA-CtD) that, although having no sequence similarity, both share the same core fold as the N- (NtDCA) and C-terminal domains (CtDCA) of archetypal orthoretroviral CAs (Fig. 20.7C). The tyrosine residue of the A-motif at the C-terminus of an α-helix 9 in the PFV CA-CtD appears to be involved in hydrophobic interactions forming part of the core of the CA-CtD bundle. In contrast, the LGL portion of the A-motif is exposed and forms a continuous hydrophobic surface patch together with a second, A′-motif (aa 431–434, PGQA) in an α-helix 8 (Fig. 20.7C). It is speculated that it is involved in interactions that give rise to hexameric assemblies analogous to those formed in orthoretrovirus capsids.10 This study has also identified a previously unknown hydrophobic interface between PFV CA-NtD α-helices 2 and 4 and CA-CtD α-helices 5 and 6 (Fig. 20.7C). The relevance of this interface for correct capsid assembly and generation of infectious virions has been demonstrated by cryo-electron tomography and infectivity analysis of viral mutants with alterations in key residues of this hydrophobic interface.10

In primate FVs, the GR-rich region is organized in three 11 to 13 aa-long clusters, termed GR boxes I to III, which is not the case in other FVs including the ancient endogenous FVs103,237,271,290 (Fig. 20.7A). Initial studies assigned various functions to individual PFV Gag GR boxes, like vgRNA or Pol protein encapsidation, RTr, capsid assembly, viral infectivity, or nuclear localization of PFV Gag.182,237,270,300 However, in respect to vgRNA packaging, a more recent report suggests that encapsidation of the PFV vgRNA and cellular RNAs are mediated by the cooperative action of arginine residues in the GR-rich domain.80

775

Nuclear localization of the Gag protein is a common feature of most FVs proven by a strong nuclear fluorescence using homologous serum in IFA. Intracellular distribution and/or trafficking of FV Gag proteins is affected by several peptide motifs. The NtD harbors a nuclear export signal (NES) at aa 95–112 (Fig. 20.7A and B). It is proposed to be responsible for the active nuclear export of Gag after its nuclear interaction with vgRNA208 and to antagonize another trafficking signal located in the GR-rich region, the chromatin binding signal (CBS) at aa 536–549.145,182,270 Furthermore, GR-I and GR-III sequences were reported to promote nucleolar targeting of PFV Gag, a function that appears to be naturally antagonized by the CBS located in GR-II.192 By this regulatory circuit, a putative temporal nuclear trafficking of Gag is achieved, which may be involved in selective encapsidation of vgRNA during assembly of the virion.208 However, this proposed function of PFV Gag is challenged by other studies. Either, involvement of Gag in nuclear vgRNA export was not observed,23 or Gag nuclear localization was seen as a passive process achieved only by CBS-mediated chromatin tethering upon nuclear membrane breakdown during host cell mitosis, rather than its active nuclear import into interphase cell nuclei.145,182 These studies rather suggest a primary role of the CBS during virus entry by enabling Gag association with cellular chromosomes through direct CBS–nucleosome interaction and appears to be important for tethering the FV preintegration complex to cellular chromatin, thereby determining the integration site profile of FVs.145

Similar to MPMV, a cytoplasmic targeting and retention signal (CTRS, aa 43–60) is also located at the NtD of PFV Gag34,51,249 (Fig. 20.7A and B). However, unlike MPMV, mutation of a conserved arginine in the CTRS did not lead to a switch from a B/D to a C-type capsid assembly strategy. Instead, mutation of the analogous arginine in PFV Gag completely abolished particle release.51,146 This suggests that the CTRS of MPMV and FV Gag are functionally different.

A late assembly (L) domain specified by the motif PSAP (aa 284–287 of PFV Gag) and located at the C-terminus of the P-rich region (Fig. 20.7A) interacts with the cellular export machinery vacuolar protein sorting (VPS) via TSG101 to mediate release of virus particles from the plasma membrane.195,255 However, PSAP motifs are absent in nonprimate FV Gag proteins, and their functional L-domains remain to be identified.255 Interestingly, ubiquitination of PFV Gag—a common feature of orthoretroviral capsids upon interaction with the VPS machinery—has not been observed as discussed below.256 This suggests that ubiquitin conjugation to transacting cellular factors, not the Gag protein itself, may be critical for ubiquitin-dependent particle release of enveloped viruses.307,308

In addition to the proteolytic processing of the FV Gag precursor, several additional posttranslational modifications of PFV Gag were reported. The latter is phosphorylated at unspecified serine residues with unknown functional consequences.55 Furthermore, particle-associated PFV Gag was found to be phosphorylated at threonine 225 (T225), which is located in a polo-like kinase (PLK) S-T/S-P consensus binding sequence310 (Fig. 20.7A). Upon host cell entry, Gag phosphorylated at T225 interacts with PLK1 and/or PLK2 and relocalizes the PLKs to mitotic condensed chromatin, a process which is dependent on a functional Gag CBS. PFV mutants deficient in PLK interaction displayed reduced infectivity due to a delayed and decreased integration efficiency and an increased preference for heterochromatin integration sites. This demonstrates that PFV Gag–PLK interactions are important for early viral replication steps and may be involved in the mitosis-dependent disassembly and integration process of FVs. Furthermore, PFV Gag was reported to interact with cellular protein arginine methyltransferases (PMRTs) 1 and 5, which results in Gag methylation at several arginine residues.192 Methylation of PFV Gag arginine 540 (R540), which is a key residue of the CBS (Fig. 20.7A), was reported to mask its chromatin tethering function and result in a GR-I and GR-III dependent nucleolar relocalization of Gag in interphase nuclei. Other posttranslational modifications, like N-terminal myristoylation or ubiquitination, reported for orthoretroviral Gag proteins, could not be detected for FV Gag proteins. The absence of ubiquitination is not surprising since an unusual and distinguishing feature of FV Gag proteins is their paucity of lysine residues.171,256,308 PFV Gag contains only a single lysine residue essential for replication in primary cells.171 This notable feature may be the reason for the FVs ability to remain biologically active for a quite prolonged time after infecting resting cells.

In summary, FV capsids share various features with the cores of hepadnaviruses (i.e., glycoprotein dependence for budding, nuclear localization, and the presence of arginine-rich motifs) as well as with orthoretroviruses (i.e., presence of an L-domain and a CTRS). There are also Gag characteristics that are unique to FVs, particularly their unusual cleavage pattern. Furthermore, alterations of conserved FV Gag motifs result in morphologic defects that affect the ability of capsids to support intraparticle RTr and render such mutants noninfectious.

Pol Unlike orthoretroviruses, FV Pol is translated from its own spliced mRNA, using the major splice donor (mSD) in the R region of the LTR and a suboptimal splice acceptor in the gag ORF, instead from the full-length vgRNA (Fig. 20.6). The latter prevents pol mRNA from becoming too abundant.22,56,116,136,156,298 Historically, the discovery of a spliced pol mRNA and large amounts of vgDNA in extracellular virions represent a landmark in retrovirus research.298 PFV Pol is translated separately from Gag as a large, approximately 127-kD polyprotein harboring the four enzymatic domains of PR, polymerase (POLY), RNase H (RH), and IN from N- to C-terminus (Fig. 20.8A). Deviating from other retroviruses, the PFV pr127Pol precursor is processed by the viral PR during or following capsid assembly into only two mature subunits: p85PR-RT and p40IN.67 FV PR does not exist as a separate subunit and instead remains covalently attached to the N-terminus of the RT POLY domain through a peptide linker (Fig. 20.8A). This unique element (aa 93–120) is not present in other RTs and consists of an unstructured region followed by an alpha-helix.188 Studies with deletion variants indicated that it is an integral part of the RT domain, which is important for activity and solubility.241 Both PR-RT and IN localize to the nucleus in infected cells.110 FV Pol precursor processing is required for virus replication.130,224 For the PFV IN subunit, NLS sequence motifs have been characterized.6,109

On the other hand, FV mutants have been reported to be replication competent when Pol was expressed in-frame with the preceding gag ORF or by an orthoretroviral-like frameshift mechanism.137,260 From orthoretroviruses, it is known that expression of a Gag-Pol fusion protein alone is incompatible with viral replication owing to severe particle assembly or release defects.280 Furthermore, these defects often involved the orthoretroviral PR that was either found to be inactive or hyperactive. The finding that FV replication tolerates expression of an in-frame GagPol fusion protein indicates a mode of Pol encapsidation and regulation of PR domain activity that is unique to these viruses and different from that observed for orthoretroviruses (see later discussion). In the BFV system, equal amounts of gag and pol mRNAs have been reported.103 Whether this also leads to similar amounts of intracellular Gag and Pol proteins has not been investigated.

All retroviral PRs including the FV PR belong to the family of aspartic PRs and are active as dimers. Each subunit of the homodimer provides one catalytic aspartate residue situated in the conserved motif Asp-Thr-Ser-Gly in order to create the active site. So the question arises, how FV Pol dimerization is accomplished. In orthoretroviruses, this is facilitated by Gag oligomerization of the Gag-Pol fusion protein, which is not possible in FVs. Biochemical and biophysical evidence point to transient dimer formation of FV Pol, as the enzyme is always purified as a 776

monomer, except under nonphysiologic conditions of high salt.87 A role of RNA in PR activation was proposed by Hartl et al.86 by identifying a pol ORF-located RNA PARM present on vgRNA (Fig. 20.6). By binding to PARM, the full-length p85PR-RT subunit including its C-terminal RH domain dimerizes and PR is activated.86,241 Although the entire process has not yet been elucidated, this mechanism could explain the replication competence of viral mutants expressing an in-frame Gag-Pol protein,137,260 because they can also rely on activation of PR by PARM, which would be the rate-limiting step for PR dimerization. The mechanism of PR activation by binding to RNA is unique among retroviruses and explains the necessity of a PR-RT fusion protein. Thus, premature PR activity before virus assembly can be avoided.86

FV RTs bear the motif YxDD in the active center and are sensitive to nucleoside analog RT inhibitors, such as AZT131,138,219,222 (Fig. 20.8B). Both, the RH domain and the connection subdomain of RT substantially contribute to polymerase integrity and stability as well as polymerase activity and substrate binding241 (Fig. 20.8B–D). RH is an endonuclease covalently coupled to the POLY domain of Pol and hydrolyzes the RNA template strand in RNA/DNA hybrids during RTr. This activity is essential for virus replication.210,268 Like the murine leukemia virus (MLV) enzyme, but unlike HIV RT, FV RH possesses a protruding basic loop and the so-called C-helix25,142 (Fig. 20.8E). This has been validated in the solution structure of the PFV RH domain alone143 and in the complete White-tufted-ear marmoset SFV (SFVcja) PR-RT subunit crystal structure,188 indicating a function of the basic loop in substrate binding. Strikingly, PFV RH is inhibited by HIV-1 RH inhibitors, suggesting a similar inhibitor binding pocket of the two proteins.42

Only very recently, the first 3D structures of complete, mature PR-RT subunits from SFVcja in complex with different substrates were solved by X-ray crystallography and cryo-electron microscopy reconstruction188 (Fig. 20.8C and D). Surprisingly, differential oligomeric states were observed depending on the type of nucleic acid substrate used. Full-length SFVcja PR-RT monomers bound to RNA/DNA hybrid substrates (Fig. 20.8C), whereas dsDNA substrates were found in complex with asymmetric homodimers of either RH-deleted or full-length SFVcja PR-RT (Fig. 20.8D), resembling HIV-1 p66/p51 RT heterodimers. This not only is the first report of a retroviral RT to adopt different oligomeric configurations but may suggest that FV −strand and +strand DNA synthesis may be executed by different enzyme configurations. Perhaps the monomeric or dimeric state of this enzymatic complex may also be attributed to the sequential order of FV polyprotein processing by the viral PR, of which we unfortunately know so little until now.

FIGURE 20.8 Schematic representation of the foamy virus Pol protein organization and PR-RT subunit/subdomain 3D structures. A: Schematic of the prototype foamy virus (PFV) Pol precursor protein. Numbers indicate amino acid positions of PFV Pol. The black arrow marks the cleavage site of pr127Pol for processing into p85PR-RT and p40IN. (Subdomains of PR-RT: PR, protease; L, linker; POLY, polymerase; con, connection; RH, RNase H; IN: integrase subunit.) B: Schematic representation of the PFV Pol p85PR-RT subunit organization. Individual subdomains are indicated by differently colored boxes. PR (light blue); POLY palm (dark red); POLY fingers (dark blue); POLY thumb (green); con (yellow); RH (orange). Active site residues of PR, POLY, and RH are marked. C: Cartoon representation of monomeric White-tufted-ear marmoset simian foamy virus (SFVcja) PR-RT in complex with an RNA/DNA hybrid substrate (pdb:7O0G). D: Cryo-EM reconstruction of dimeric SFVcja PR-RT in complex with a 22 bp/2 nt overhang dsDNA substrate. Lighter shades of colors are used for domains/subdomains of subunit B (pdb:7O24). E: Cartoon representation of the solution structure of the Taiwanese macaque simian foamy virus (SFVmcy) protease (PR) monomer. The flap region (red) and the C-terminal α-helix (blue) are highlighted. The conserved amino acids D24, S25, and G26 forming the active site in the dimer are depicted in red as sticks (pdb:2JYS). F: Ribbon diagram of the solution structure of the PFV RNase H. The C-helix is highlighted in blue; the basic loop in green. The active site residues D599, E646, D669, and D740 are depicted in red as sticks (pdb:2LSN). (Panel A adapted from Lindemann D, Rethwilm A. Foamy virus biology and its application for vector development. Viruses 2011;3(5):561–585; panels C and D courtesy of 777

Marcin Nowotny, Warsaw, Poland; panels E and F courtesy of Birgitta M. Wöhrl, University of Bayreuth, Germany.) FV replication depends on integration mediated by the active IN. A 4-bp duplication of staggered chromosomal nucleotides occurs at the site of integration.57,176 Orthoretroviruses utilize 3′ end processing as the initial step of the integration reaction. This involves the removal of two nucleotides from each terminus of the blunt-ended linear vDNA. During FV integration, only the 3′ terminus (within the U5 region) of the vDNA undergoes processing, whereas the 5′ end (the U3 region of the LTR) remains unprocessed, possibly because it is already suitable for integration.57,118 In 2010, a seminal study was published that described the crystal structure of full-length PFV IN bound to its cognate DNA as a tight complex, termed the intasome83 (Fig. 20.9). This achievement was possible because, unlike orthoretroviral IN proteins, the recombinant PFV IN is uniquely soluble and catalytically active in vitro.275 These properties contrast with the aggregation and poor enzymatic activity observed with other retroviral INs, regardless of the expression system.

FIGURE 20.9 The architecture of the prototype foamy virus (PFV) intasome. A: Schematic representation of the PFV Pol p40IN subunit organization. Individual subdomains are indicated by differentially colored boxes. Amino-terminal domain (NTD, green); NTD extension domain (NED, green); catalytic core domain (CCD, yellow); carboxyterminal domain (CTD, green). Active site residues of the D,D-35-E motif of the CCD are marked. B: The crystal structure (PDB ID 3OY9) is shown as viewed along (bottom panel) or perpendicular (top panel) to its twofold axis. Viral DNA (vDNA) chains are shown as cartoons and colored by chain (magenta: reactive strand; orange: nontransferred strand); vDNA bases and active site IN residues are shown as sticks. Gray spheres are metal cations. Locations of IN domains (NTD, CTD, and CCD) are indicated. C: Segmented electron density map as semitransparent surface with docked PFV intasome and nucleosome structures shown as ribbons. H2B, the N-terminal tail of H2A (H2A-N), the CTD and one of the CCD dimers are indicated. D: Nucleosomal DNA within the tDNA-binding cleft of the intasome. DNA conformations as in available nucleosome structures (left) and as in the crystals of the PFV target capture complex (right) produce local electron density crosscorrelation scores of 0.36 and 0.70, respectively. Protein Data Bank accessions are indicated in brackets. (Panel B courtesy of Peter Cherepanov, Francis Crick Institute, London, UK; panels C and D reprinted by permission from Nature: Maskell DP, Renault L, Serrao E, et al. Structural basis for retroviral integration into nucleosomes. Nature 2015;523(7560):366–369. Copyright © 2015 Springer Nature.)

Classically, retroviral INs are subdivided into three domains (Fig. 20.9A): an N-terminal Zn2+ binding domain (NTD), characterized by pairs of His and Cys residues (HHCC motif), that is expanded by an approximately 40 aa residue NTD extension domain (NED) in FV-, γ-, and ε-retrovirus INs; a catalytic core domain (CCD), harboring the Asp, Asp-35-Glu (DD35E) motif; and a nucleic acid–binding Arg/Lys-rich C-terminal domain (CTD).144 These domains are connected by nonconserved flexible linkers. Early studies reported unspecific DNA-binding activity by the CTD, and it was assumed that retroviral INs would adopt a dimeric or tetrameric structure when engaged with the vDNA ends.144 It was also hypothesized that multimers were highly flexible, and several contrasting structures of the retroviral intasome had been proposed. Crystallization of the PFV intasome has revealed the definitive answer to a long-standing puzzle. The foamyviral integration apparatus contains a tetramer of IN, assembled on a pair of vDNA ends, in which all three IN domains and interdomain linkers are involved in intimate protein– protein and protein–DNA interactions cross-linking the complex in a rigid structure83 (Fig. 20.9B). Further cocrystallization of the PFV intasome with target DNA to mimic of the host cell chromosomal DNA revealed the assembly of the entire retroviral synaptic integration complex prior to and following strand transfer.165 Structures of the PFV intasome–nucleosome complex revealed a multivalent intasome– nucleosome interface involving both gyres of nucleosomal DNA and one H2A-H2B histone heterodimer169 (Fig. 20.9C and D; Video 20.3). Without altering the histone octamer, the cellular target DNA is lifted from the surface of the H2A-H2B heterodimer by a looping-and-sliding mechanism to allow integration at preferred locations.169,289 Moreover, because of the structural and functional similarity of PFV and HIV-1 INs, the mechanism of action of several HIV IN strand transfer inhibitors in clinical use (raltegravir, elvitegravir, and dolutegravir) was elucidated.84,85 Using the PFV intasome as a surrogate for its HIV counterpart, it was shown that these small molecule inhibitors bind to the active site of IN and displace the reactive 3′ hydroxyl group of the vDNA, thereby preventing strand transfer.144 After solving the PFV intasome structure, it was believed that retroviral integrases generally function as a tetrameric complex. However, subsequent elucidation of other retroviral intasome structures revealed that α- and β-retroviruses require eight, and lentiviruses up to sixteen integrase subunits, to assemble the intasome core structure.54

778

FIGURE 20.10 Schematic representation of the prototype foamy virus (PFV) Env protein domain organization and membrane topology. A: Schematic representation of the prototype foamy virus (PFV) Env protein organization. The furin cleavage sites within the gp130Env precursor used to generate the mature gp18LP, gp80SU, and gp48TM subunits are indicated by gray arrows. The individual subunits are shown as boxes in different shades of green. Hydrophobic sequences spanning the membrane in the gp18LP (h) and the gp48TM (membrane-spanning domain, MSD) subunit as well as the fusion peptide (FP) are indicated. The amino acid sequence of the PFV Env gp18LP subunit and the cytoplasmic domain of the gp48TM subunit are shown in the enlargements below. The conserved WxxW and RxxR motif in LP are highlighted in red and the lysine residues potentially ubiquitinated in blue. The KKxx ER retrieval signal at the C-terminus is highlighted. The approximate positions of PFV Env N-glycosylation sites are marked by Y-shaped symbols. B: Schematic membrane topology of the monomeric unprocessed Env precursor protein with ubiquitination (Ub) sites in the leader peptide regulating subviral particle release. The N- and C-termini of the protein are indicated. C: Schematic view of the trimeric PFV glycoprotein complex assembled from mature LP, SU, and TM subunits. Color coding of subunits and hydrophobic h, MSD, and FP peptides in B and C is identical to A. (Panels A to C adapted from Lindemann D, Rethwilm A. Foamy virus biology and its application for vector development. Viruses 2011;3(5):561–585. https://creativecommons.org/licenses/by/3.0/.)

779

FIGURE 20.11 Prototype foamy virus (PFV) glycoprotein structures on cell-free virions. A and B: 0.8-nm-thick tomographic slice perpendicular to the glycoprotein long axis and its corresponding schematic of interlocked hexagonal assemblies of trimers. Numbers are indicated at the center of each hexagon, and triangles represent the positions of each trimer of Env in the hexagonal network. B: Top view of intertwined hexagonal assemblies. C: Side view of a single trimer. D and E: In situ single particle 3D reconstruction of PFV glycoprotein by cryo-EM. Full (D) and cut-away (E) side views of a single PFV Env trimer (sharpened map) after threefold symmetry application (~9 A resolution at FSC = 0.143). The densities corresponding to the extracellular domains and the viral membrane are colored salmon and gray, respectively in (D). The three central helices attributed to gp48 fusion peptide are represented by three green α-helices each being 22 residues long. The transmembrane helices (TMHs) are represented by three inner (colored blue) and three outer (colored orange) α-helices. In (E), the densities surrounding the three central helices and the three inner and outer TMHs are colored green, blue, and orange, respectively, while the remaining of the spike is gray colored. (Panels A to E adapted from Effantin G, Estrozi LF, Aschman N, et al. Cryo-electron microscopy structure of the native prototype foamy virus glycoprotein and virus architecture. PLoS Pathog 2016;12(7):e1005721. https://creativecommons.org/licenses/by/4.0/.) As a consequence, the mature LP subunit is an integral component of the mature tripartite and trimeric Env GPC present on released FV particles52,287 (Figs. 20.4A; 20.10B and C; 20.11). It transverses the viral membrane at the N-terminus (a type II transmembrane protein), as does the TM subunit at the C-terminus of Env (a type I transmembrane protein)150,288 (Figs. 20.10B and C, 20.11D and E). All three Env subunits are heavily glycosylated. Fourteen N-linked glycosylation sites have been mapped, only two of which, N8 and N13 located in PFV SU and PFV TM, respectively, are essential for viral infectivity163 (Fig. 20.10A).

The C-terminal CyD of the TM subunit comprising 16 aa is rather short, and its presence is not required for particle egress202 (Fig. 20.10A). In contrast, the N-terminal CyD of the LP subunit, comprising approximately 68 aa, is considerably longer. Alteration of conserved tryptophan residues herein, at aa positions 10 and 13 of the PFV LP, abolished interaction with the Gag protein.77,150,288 Not only is FV Env required for export of capsids, but Gag expression is also necessary for the transport of Env to the cell surface.203 This observation implied that highly specific interactions influence the intracellular distribution and trafficking of both proteins.34,65,202 Coimmunoprecipitation analyses and surface plasmon resonance to define an N-terminal LP “budding domain” suggested and demonstrated that a direct interaction of LP and Gag with both tryptophan residues is critical and essential.150,207,288 Indeed, this has been confirmed by solving the crystal structure of an N-terminal MA-like domain of PFV Gag comprising aa 1–179 and variants thereof cocrystallized with peptides representing the first 20 aa of the PFV LP N-terminus77 (Fig. 20.7B).

Aside from its interaction with Gag, additional factors appear to regulate Env intracellular trafficking and transport to the cell surface. First, a dilysine motif, known to be responsible for retrieval of glycoproteins to the endoplasmic reticulum (ER), is present near the C-terminus of the TM of most FV Env proteins75 (Fig. 20.10A). Although this signal can sort Env to the ER, it is not required for efficient virus replication of PFV76 and, in comparison to the other factors, has only a weak effect on Env intracellular distribution. Second, posttranslational ubiquitination of four of five lysine residues located within the LP subunit N-terminal CyD also appears to mediate efficient Env removal from the cell surface as observed for PFV as well as for SFVmcy, a monkey FV formerly designated SFVmac254,256 (Fig. 20.10A). In contrast, FFV Env LP appears not to be ubiquitinated.69 Whether this type of posttranslational modification occurs in nonprimate FVs like BFV or EFV and has a crucial function in their intracellular GP trafficking has not been investigated. Third, BFV Env lacking the dilysine ER retrieval signal was recently shown to be palmitoylated at two conserved cysteine residues located close to the lipid membrane in the N-terminal CyD of the LP subunit.36 BFV Env mutants with inactivated palmitoylation sites showed an impaired Env cell surface expression and membrane fusion activity.36

The PFV Env has been shown to support not only viral particle release from cells but also release of SVP from other cellular membranes harboring the viral glycoprotein.251 This is again analogous to a similar process observed in hepadnaviruses, which secrete vast amounts of SVPs, the so-called Australia antigen (see Chapter 18, Volume 2 DNA Viruses). Ubiquitination appears to suppress the intrinsic activity of the primate FV glycoprotein to induce SVP release, and mutants of the lysine-specifying codons in the LP CyD release large amounts of SVP.256 These PFV Env mutants appear to be particularly well suited to pseudotype orthoretroviral capsids, in contrast to wild-type PFV Env, probably owing to its low level of cell surface expression.81

Surprisingly, the gp130Env is not the only FV glycoprotein synthesized. Using conserved splice sites within env ORF of the FV genome (Fig. 20.6), alternatively spliced env transcripts are generated. In case of PFV, this leads to translation of proteins consisting of Env lacking the membrane-spanning domain (MSD) and CyD of TM fused in-frame with the Bet protein (Env-Bet)74,147 or, as in the case of FFV, Env lacking only the MSD of TM fused to the orf-2 encoded peptide sequence (Env-Bel2).21 The Env-Bet fusion protein and its processing products LP, SU, and δTM-Bet are secreted into the supernatant of PFV-infected cells but are not associated with the viral particle.147 This fusion protein is synthesized at about 50% of the level of particle-associated gp130Env, suggesting that it may have a useful function in vivo. In cell culture, however, replication-competent PFV mutants deficient in Env-Bet synthesis do not exhibit a distinctive phenotype, and no revertants with restored Env-Bet expression have been observed.147

In summary, the biosynthesis and membrane topology of the FV Env are highly unusual for retroviral GPs. The Env LP subunit is an integral 780

component of the particle-associated Env complex and harbors in its CyD the major interaction domain with FV capsid essential for viral particle budding. The interaction of Env with Gag and ubiquitination of the CyD of the Env LP subunit seem to be the main determinants of its intracellular transport and are probably dominant over the C-terminal dilysine motif. Furthermore, some properties of orthoretroviral Gag proteins, such as ubiquitination and budding functions, have been delegated to Env in FVs. In addition, the function of the unusual EnvBet/Bel-2 fusion proteins is unknown.

FIGURE 20.12 Schematic outline of prototype foamy virus (PFV) Tas protein. Functional domains are indicated as differentially colored boxes. SR, Tas-Bet shared N-terminal region; MMD-1 to 3, multimerization domains 1 to 3; CD, conserved domain; NLS, nuclear localization signal; AD, activation domain.

NONSTRUCTURAL PROTEINS Tas Tas is the transactivator of spumaviruses and essential for replication.160,217 PFV Tas is a 35-kD nuclear protein that binds to upstream DNA elements in, and augments gene expression from, both the IP and the U3 LTR promoter37,123,157 (Fig. 20.6). Most Tas protein is translated from a spliced mRNA initiated at the IP.12,21,155,183 Among the FVs, Tas is variable in size with 209 aa in FFV and 300 aa in PFV and has a modular organization155,179,209,211 (Fig. 20.12). Its N-terminus contains a region of variable length that is shared with Bet (SR) and harbors several multimerization domains (MMDs). Unique to Tas are a DNA-binding central domain (CD) of approximately 100 aa, a basic NLS, and a C-terminal acidic activation domain (AD) of around 30 aa.

Except for the AD, Tas shows no homology to known cellular proteins, and there is little or no cross-transactivation between different FVs.100,155 The reason for this is the species specificity of the Tas DNA-binding CD, which is highly variable in aa sequence among different FV Tas proteins. This is consistent with the highly divergent DNA targets that mediate Tas function among different FVs. In contrast, the Tas AD shares key amino acids with other viral and cellular transcription activators and is also active in yeast.16 For transcriptional activation, the Tas protein has to multimerize. This process that is apparently facilitated by residues of the MMDs mainly located within the N-terminal SR. This property has been demonstrated experimentally for BFV Tas and can probably be generalized to all FV Tas proteins.266 Little is known about likely cellular factors that might interact with Tas.127 Phosphorylation by DNA-PK is required for full Tas activity, and only acetylated Tas protein has full DNA-binding capacity.20,38 In addition, the yeast ADA2 adaptor molecule is required for Tas AD-mediated activation in yeast.16

Research carried out on cellular factors engaged with BFV Tas has led to the identification of RelB as its interaction partner to activate the nuclear factor κB (NF-κB) pathway.279 However, whether these results also apply to Tas proteins other than BFV Tas is not known. Furthermore, the NF-κB–mediated transcriptional enhancement seen is unlikely to explain the full activity of Tas. It can also activate cellular genes if they happen to harbor Tas responsive DNA elements. This has been investigated for various human genes activated by PFV Tas.155,278

Bet Bet is the least conserved of all FV proteins (Fig. 20.13). It is translated predominantly from multiply spliced mRNAs originating at the IP and to a minor extent from mRNA variants initiating at the LTR promoter12,183 (Fig. 20.6). The splice sites located within the tas ORF utilized for generation of the bet mRNA are highly efficient, resulting in Bet to be always made in vast excess over Tas. In transfected or infected cells, Bet is found in the cytoplasm and the nucleus. The nuclear localization is mediated by a C-terminal NLS (Fig. 20.13). Because SR at the N-terminus contains the MMD of Tas, it is likely that Bet also multimerizes via this domain. 3D structures of Bet are not available. A recent bioinformatic protein structure modeling of PFV Bet consisting of 482 aa predicts an organization in two domains. The protein consists of a smaller NtD with 164 aa, which includes the N-terminal 88 aa shared region, and a larger C-terminal domain (CtD) with 318 aa.111

For a long time, no clear function could be attributed to Bet. For PFV, Bet was shown to be dispensable for in vitro replication in most cell types, with only a minimal decrease in viral titers.12,297 The first generation of FV vectors actually had the orf-2 region encoding for most of Bet replaced by sequences of interest putting them under Tas-dependent transcriptional control of the IP or of an inserted heterologous promoter.238 For FFV, a more drastic reduction in viral titers was observed for Bet-deficient viruses when grown on feline CRFK, but not on human 293T cells.4 This observation eventually led to the identification of Bet being an antagonist of APOBEC3 (A3) proteins, similar to HIV Vif.159,196 Whereas SR of FFV Bet was shown to be dispensable, conserved motifs (CMs) within its C-terminus are essential for A3 antagonization164 (Fig. 20.13).

The splice sites that lead to generation of the Bet protein appear to be so efficient that they are used also in vgRNA. An integrated FV has been described that carries the characteristic deletion leading to the generation of Bet (δTas)233 (Fig. 20.6). As the vgRNA leading to δTas carries all features necessary for successful packaging and RTr, δTas infects new cells where it integrates. δTas has been found in vitro and also in vivo in a rabbit infection model and in the monkey to a considerable extent.21,62,232,233 δTas provirus is replication incompetent because of its tas gene deletion; however, it is not transcriptionally silent. Owing to the basal activity of the IP (see later discussion), there is still some residual bet gene expression. The magnitude of viral transcription depends on the number of integrated copies of δTas and probably also depends on the site of integration where cellular promoter/enhancer elements could augment levels of Bet mRNA.233 Cells expressing Bet become resistant to superinfection by homologous virus, a feature that has so far not been further investigated.17,233 Furthermore, a role in promoting viral persistence has been discussed for Bet in general and δTas in particular.175,233 Functionally, δTas behaves like a defective interfering genome. However, whether the typical oscillating frequency of DI viruses occurs with δTas has not been investigated. Because only either Bet or Tas can be made, and since FV gene expression starts with the translation of Tas (see later discussion), it has been speculated that Bet synthesis represents the molecular switch, which determines viral latency.175

781

FIGURE 20.13 Schematic outline of prototype foamy virus (PFV) Bet protein. Functional domains are indicated as differentially colored boxes. SR, Tas-Bet shared N-terminal region; CM1 to 6, conserved motifs 1 to 6; NLS, bipartite nuclear localization signal.

STAGES OF REPLICATION The Early Phase: Establishing the Provirus (Fig. 20.14) Attachment FVs bind via a receptor-binding domain located in the Env SU subunit48 (Fig. 20.10), to yet unknown, probably ubiquitously expressed and evolutionary conserved cellular receptor molecules. Overexpression of FV Env in target cells results in superinfection resistance toward the parental virus as well as FVs derived from other species. This suggests that all FVs use same entry receptor(s).13 Proteoglycans contribute significantly to FV entry but do not appear to be the major cellular receptor.186,205,258

Entry and Intracellular Trafficking Uptake of most FVs predominantly seems to involve endocytosis and a pH-dependent FV Env-mediated fusion process.50,201,259 Only PFV Env displays a significant fusion activity at neutral pH and enables fusion to take place at the plasma membrane as well. Single particle tracing of FVs entering the target cells revealed a unique intermediate fusion step characterized by tethering of glycoprotein and capsid but separation of up to 400 nm before final separation of both viral components.50 After release into the cytoplasm, intact FV capsids migrate along the microtubular network to the centrosome, where they accumulate.50,234,259 Retrograde movement of FV capsids appears to utilize dynein motor complexes and involves an interaction of its dynein light chain 8 component with the CC3 domain of FV Gag.199

A few other host cell factors interacting with FV components during the early replication phases have been identified. TRIM5a, a well-known restriction factor for various retroviruses, was shown to restrict FVs in a species-specific manner.191,296 The specificity of TRIM5a has been mapped to variable residues of the B30.2 domain, which are important for neutralization of lentiviruses, and to the N-terminal half of FV Gag.296 The activity of TRIM5a against several retroviral capsid proteins, including those of FVs, which do not mature into the canonical orthoretrovial MA, CA, and NC subunits, implies an even wider structural recognition pattern than previously assumed.

Uncoating Intact FV capsids that accumulate at the centrosome in G0-resting cells can remain functionally active for weeks, allowing productive infection to proceed upon their re-entry into the cell cycle.141 It is generally agreed upon that mitosis is required for FV replication and that the latency period from cell entry to integration and gene expression, owing to the facultative vDNA genome, can be very long.15,47,141,194,273 Most productive infections are attributed to FV particles that have their vgRNA reverse transcribed into vgDNA prior to target cell infection.171,180 However, some genome RTr takes place upon FV host cell entry and seems to add to viral infectivity predominantly under conditions of a low MOI.46,304 The trigger that initiates RTr during FV entry has not been identified.

Disassembly of FV capsids accumulated at the centrosome is reported to involve Gag cleavage by yet uncharacterized cellular protease(s), and potentially the viral protease, in a cell cycle–dependent manner.73,108,140,259 Upon mitotic breakdown of the nuclear membrane, the vDNA and fragments of Gag gain access to the chromosome, whereas active nuclear import of Gag into interphase nuclei is not observed.182,259

Capsid disassembly is essential for further steps in FV replication but also makes the virus vulnerable to cellular defense mechanisms. It leads to exposure of a key pathogen-associated molecular pattern, the viral nucleic acids, recognized by the innate immune system and triggering antiviral responses in different immune cell types. Toll-like receptor 7 (TLR7) expressed in plasmacytoid dendritic cells was shown to be the likely factor in endosomal sensing of FV RNA resulting in the induction of type I IFN.227 Replicating virus was not found to be required for this type of IFN induction. Furthermore, FV RTr products already present in FV particles taken up by various myeloid immune cells induced an efficient innate immune response.14 It required FV particles with an active RT, was largely unaffected by RTr inhibition during viral entry, and was dependent on RTr products to be derived from full-length vgRNA. RTr products were sensed in the cytoplasm in a cGAS and STINGdependent manner by the innate immune system in host cells of the myeloid lineage.

Integration Productive viral replication requires insertion of the provirus into the host genome mediated by the viral integrase,57,176 although unintegrated vDNA appears to be transcriptionally active, similarly as reported for orthoretroviruses.47 Like other retroviruses, there are no preferred sites of FV integration. Analyses of PFV integration site patterns have revealed that in sharp contrast to HIV-1 and MLV, PFV disfavores integration into genes.145,169,189,274 Recent studies using Gag mutants of PFV, SFVmcy,145 or FFV282 demonstrated a functional role for the CBS in proviral integration and integration site distribution. PFV and SFVmcy Gag CBS mutant viruses are characterized by a wild-type–like integration efficiency but a massive redistribution of integration sites toward centromeres in different cell lines,145 whereas FFV Gag CBS mutants display a strongly reduced integration efficiency.282 Lesbats and colleagues145 showed that the PFV Gag CBS is essential and sufficient for a direct interaction with nucleosomes. By determining a crystal structure of the PFV Gag CBS bound to mononucleosomes, they found that this viral entity directly interacts with the histone octamer, engaging the H2A-H2B acidic patch in a manner similar to other acid patch–binding proteins such as herpesvirus latency-associated nuclear antigen. How PFV Gag is embedded within the extensive PFV intasome–nucleosome interface,169 which involves a lift off of cellular DNA from the H2A-H2B heterodimer during the integration process, remains to be determined.

Unlike other retroviruses, the contribution of cellular proteins to FV integration and integration site distribution is largely unknown. Only 782

cellular PLK, and PLK-2 in particular, was shown to interact with PFV Gag during virus entry.310 The PLK–Gag interaction is dependent on the phosphorylation of a PLKBS in Gag (Fig. 20.7A) by a yet uncharacterized cellular kinase. Thereby cellular PLKs are relocalized to mitotic chromatin in a PFV Gag CBS-dependent manner. PFV Gag PLKBS mutants display a delayed and reduced integration, which is accompanied with an enhanced preference to integrate into heterochromatin.

The Late Phase: Generation of Progeny Viruses (Fig. 20.15) Transcription Like orthoretroviruses, FVs exploit the cellular transcription machinery to initiate virus propagation. FV transcription is special, as a cascade of events is launched by the action of two viral promoters and one transcriptional transactivator. FV gene expression (Fig. 20.6) begins with the production of the tas and bet genes’ transcripts directed by and initiated at the IP located in the env ORF.158 This is mediated by the weak basal transcriptional activity of the IP, whereas the U3 promoter in the LTR of the provirus has no, or almost no, basal activity.123,157

In FFV, enhancer elements upstream of the IP that are implicated in its basal transcriptional activity include sites for SP-118,290; however, their biological function has not been characterized in any depth. In BFV, AP-1 sites in this location have been partially characterized294 and may control the basal gene expression that it directs.

As a consequence of the IP’s low basal activity, some Tas protein is made, which subsequently binds with high affinity to specific DNA elements —namely, Tas-binding sites (TBS) upstream of the IP—resulting in a positive feedback loop of tas gene expression (Fig. 20.6). Once sufficient amounts of Tas have been synthesized by this mechanism, the transactivator also binds with lower affinity but higher avidity to upstream promoter elements in the 5′ LTR U3 region. At this step, the expression of all structural genes and, to some extent, additional LTR-directed Tas and Bet expression set on and virus production is initiated.18,155

The IP and LTR U3 TBSs are essential for Tas-mediated transactivation and have been determined and characterized in detail, for example, by electromobility shift assays for PFV, SFVmcy, FFV, and BFV.19,88,121,264,309 However, no real TBS consensus sequence has been pinpointed. Furthermore, even within each specific FV species, the IP and U3 TBS show only very weak homology. In general, it can be noted that the finer details of Tas-mediated regulation of FV gene expression, in particular the involvement of cellular proteins, remain to be elucidated. The IFNinduced leucine zipper protein human IFN-induced 35 kDa protein (IFP35) has been previously reported to down-regulate BFV and PFV transcription and replication by interacting with the respective transactivators.265 Furthermore, N-Myc interactor (Nmi) of man and cattle, an IFN-stimulated gene (ISG), was found to bind the respective Tas proteins of PFV and BFV. Binding interferes with viral replication by sequestering the protein in the cytoplasm.106 More recently, PHD finger domain protein-11 (PHF11), another ISG identified in a screen as an antiviral factor of PFV replication, was reported to inhibit the basal expression from the IP, thereby preventing Tas expression.119,120

FIGURE 20.14 Early steps of FV replication. Pathways of FV particle attachment, uptake, intracellular trafficking, virion disassembly, and proviral integration are illustrated. Putative early steps of FV replication that may include RTr in vgRNA bearing capsids are marked in a dashed area as follows: RTr ↑?. Known cellular cofactors involved in early steps of FV replication are mentioned at the respective subcellular locations. Further details can be found in the inserted legend and throughout the main text. The LTR structure of FVs is unique with respect to the location of the mSD, which is located upstream of the polyadenylation signal (pA) (Fig. 20.6). For the expression of full-length genomic transcripts, the vgRNA, FVs suppress premature polyadenylation at the 5′ LTR pA site by U1 small nuclear ribonucleoprotein (U1snRNP) binding to the 5′ LTR mSD.242 At their 3′ LTR, FV transcripts fold in a different secondary structure that presumably blocks access of U1snRNP and thereby activates polyadenylation at the 3′ end of the RNAs.

Nuclear RNA Export FV nuclear RNA export also appears to be unique. Similar to other complex retroviruses, the foamyviral regulatory protein Tas acts at the transcriptional level. A posttranscriptional regulator, such as Rev of HIV or Rex of T-lymphotropic retroviruses, has never been identified in 783

FVs. The peculiarities of FV gene regulation—one transcriptional activator and two active promoters—allow a biphasic mode of FV gene expression analogous to other complex retroviruses.43 However, this does not circumvent a central problem of all retroviruses: regulation of nuclear export of intron-bearing mRNAs containing functional splice sites.44 As detailed elsewhere in this book (see Chapters 15, 16, and 17), complex retroviruses solve this problem by interacting with the karyopherin CRM1 by using a viral regulatory protein, which binds to an RNA secondary structure embedded in viral mRNAs. In contrast, some simple retroviruses utilize the NXF1/NXT1-mediated cellular mRNA export pathway by means of a constitutive transport element (CTE) located within their genomic mRNAs.44 PFV vgRNA and Gag-ecoding mRNA appears to make use of yet another, so far undisclosed pathway.23 Nuclear export of FV RNA is, on one hand, CRM1 dependent and, on the other, relies on the presence of additional cellular proteins. The host cell HuR protein binds to the unspliced PFV vgRNA, and two cellular adapter molecules, ANP32A and ANP32B, mediate the interaction between the RNA-bound complex and CRM1.18,23 The FV RNA elements in question have not yet been characterized. The consequences of this viral nuclear RNA export mechanism on the synthesis of early response cellular proteins, whose mRNAs normally use this export pathway, are unknown. It is tempting to speculate that FVs, by making use of a nuclear mRNA export pathway involving cellular proteins also required for the synthesis of early response proteins, may outcompete the latter.

FIGURE 20.15 Late steps of FV replication. Pathways of FV RNA and protein trafficking resulting in capsid assembly and particle release are illustrated. Known and putative trafficking pathways (marked by the dashed area with the question mark in the center) of viral RNAs from the nucleus into the cytoplasm to the capsid assembly site at the microtubule organizing center (MTOC) are shown. Putative RNA–protein interactions and assembly intermediates are depicted. Four different variants of vgRNA dimerization throughout the transport pathway to the assembly site are illustrated. Known cellular cofactors involved in late steps of FV replication are mentioned at the respective subcellular locations. Dashed arrows indicate the potential pathway leading to reintegration of vgDNA originating from preassembled capsids, which underwent reverse transcription prior to particle release. Further details can be found in the inserted legend and throughout the main text. Translation The Gag and Pol proteins, as well as the accessory gene products Tas and Bet, are translated on free ribosomes in the cytoplasm. It should be remembered that pol translation of FVs as a separate protein from a spliced mRNA is unique among retroviruses (see above). The Env protein is targeted to the secretory pathway by ribosomal translation in the rough ER and displays a highly unusual biosynthesis and membrane topology150,288 (see above).

Translation of pr71Gag, encoded in the vgRNA containing the authentic gag 5′ UTR, appears to use a ribosomal shunting mechanism236 (Fig. 20.6). This is a mechanism that is used by plant pararetroviruses to translate their Gag homolog and involves the selective jumping of ribosomes to the translation initiation site.231 In addition, FV Gag translation is regulated at different levels. At least three genetic elements were reported to influence expression of Gag, in context of its natural environment of the full-length vgRNA (Fig. 20.6). The first one is the mSD found in the gag 5′ UTR sequences, which regulates 5′ LTR pA site suppression242 (see above). Its presence appears to be crucial for the translation of the Gag protein from vgRNA or from expression constructs harboring the authentic 5′ UTR sequences and FV gag ORF.91–93,153,229,242 The second element, although poorly characterized, appears to be located within the gag ORF itself and inhibits expression of authentic gag ORFs in the absence of its 5′ UTR.92,93,153,229 Its inhibitory effect can be neutralized by replacing the authentic FV gag 5′ UTR with a heterologous SD site or a complete intron sequence but not by adding such sequences downstream of an authentic PFV gag ORF with 5′ UTR sequences lacking an SD site. The third element is found in the 3′ UTR sequence of Gag-encoding mRNAs. It is one of the four PuR sequences, element C (PuR-C), which was initially reported to be essential for Gag expression, without knowing the underlying mechanism.197 Recently, PuR-C was shown to represent an SRE overlapping the branch point of a strong env-specific 3′ ss (SA3).181 Most likely, serine-rich protein binding the PuR-C results in suboptimal recognition of the major env 3′ ss BP by SF1/mBBP, thereby permitting retention of the env intron and formation of unspliced gag and singly spliced pol transcripts.

Capsid Assembly FV capsid assembly follows a retrovirus type B/D morphotype strategy. Gag is the driving force of FV in this process, since neither Pol nor vgRNA are essential for intracellular capsid assembly. C-terminal FV Gag truncation mutants as well as full-length PFV Gag with 23 arginine 784

residues of the C-terminal GR-rich domain changed to alanine are unable to assemble normal shaped capsids.34,80 This suggests that interaction of Gag with nucleic acids in general, but not necessarily with vgRNA, through its C-terminal GR-rich domain, is a prerequisite for the assembly of FV capsids with normal morphology or virus-like structures. FV morphogenesis includes the microtubule-dependent transport of Gag proteins to the centrosome involving the CTRS within Gag51,299 (Fig. 20.7). The Gag CC2 domain mediates FV capsid preassembly at the centrosome.269

An important question concerning assembly deals with the mechanism of Pol protein encapsidation. Pol packaging via protein–protein interaction of the fused Gag precursor, as it occurs with orthoretroviruses, cannot apply to FVs since Pol is separately expressed from its own spliced mRNA56,116,136,156,298 (Fig. 20.6). FV Pol precursor, but not its mature cleavage products PR-RT and IN, are packaged into assembling capsids after provirus-derived expression.108,198,224 Two competing views on this mechanism exist for the initial step of Pol incorporation. One favors Gag-Pol protein–protein interactions when proviral mutants were analyzed. Another concept arose from subgenomic vector analyses, where Gag and Pol seem to interact in that the viral vgRNA serves as a bridging molecule.94,108,135,198 Currently, it cannot be excluded that both mechanisms are involved in packaging the Pol precursor. However, details of the temporal and spatial regulation of FV vgRNA and Pol encapsidation as well as the contribution of cellular factors have not been well characterized. Only a cellular DEAD-box RNA helicase, DDX6, was described to promote vgRNA encapsidation by a yet unclear mechanism independent of a direct interaction with FV Gag.301 Furthermore, it is yet unclear whether all three viral components, vgRNA, Gag, and Pol, are transported separately to the centrosome and capsid assembly initiates and proceeds at this key organelle in the FV replication cycle, or if assembly intermediates encompassing two or all components are formed before or during transport to the centrosome. The order of protein–nucleic acid interactions is also still unknown. Whether Gag or Pol first interact with vgRNA, or whether both bind simultaneously, perhaps enhanced by additional viral and/or cellular protein–protein interactions, remains to be elucidated.

Although the exact time point of RTr with respect to capsid assembly and maturation is unknown, preassembled intracellular FV capsids, generated upon propagation in vitro, contain significant amounts of already reverse transcribed, infectious vgDNA.171,180,225,303 In this respect, FVs are unique among retroviruses but show a similarity to HBV replication (see Chapter 18, Fields Virology Volume 2, DNA Viruses). Gag, but not Pol precursor protein cleavage, is a prerequisite for intraparticle RTr that occurs late in the FV replication cycle.55,108,224 Therefore, Gag precursor cleavage appears to be the initiating event for intraparticle RTr in assembling FV capsids.108

Orthoretroviruses can occasionally behave like retrotransposons and reshuttle their genome to the nucleus without an extracellular phase. The frequency of such intracellular retrotransposition (IRT) has been estimated to be 1 per 106 proviruses and is thought to reflect the frequency at which the RNA genome is prematurely reverse transcribed.89,225 For PFV, this frequency is much higher with approximately 5% and thereby concordant with late RTr of the vgRNA.89 It was subsequently found that IRT strongly depended on the particular FV isolate and the cell type used for analysis.90,225 Thus, IRT is not a general phenomenon of FVs. However, the late phase of RTr of the vgRNA is required for this process.

Some cellular defense mechanisms tackle viruses at late replication steps. One of few factors known to interfere with FV replication are members of a family of evolutionarily conserved cytidine deaminases, the apolipoprotein B mRNA editing enzyme, catalytic polypeptide-like 3 (APOBEC3; A3) proteins.111 As observed for other retroviruses, the A3s are encapsidated into the nascent virus particle due to specific Gag-A3 interactions. The antiviral activity is mediated by cytidine deamination of the viral reverse transcript during RTr that results in G-to-A hypermutations of the viral genome. FVs encode the accessory Bet protein (Fig. 20.13) to preserve genome stability and counteract A3s.159,228 Unlike the HIV-1 Vif protein, which prevents A3 particle incorporation by routing it to the proteasomal degradation pathway, Bet prevents A3 encapsidation by binding, inhibiting its dimerization, and quantitatively trapping the deaminase.39,112,196 The Bet-specific orf-2–encoded domain of FFV Bet, but not the shared N-terminal Tas domain, was found to be essential for feline A3 binding and inactivation.164 The PFV and FFV Bet function was found to be broadly active against various primate or feline A3s.196 However, some species specificity was also observed, as PFV Bet was found to be inactive against all or some mouse, feline, and rhesus monkey A3 proteins, as well as against human A3DE and A3H.196 Because RTr of FV RNA takes place to a significant degree in virus-producing cells, A3 restriction of FVs occurs at a different point in the replication cycle than that reported for orthoretroviruses.159

Membrane Envelopment and Release One of the most distinctive features of FVs is the failure of their capsids to spontaneously bud from cellular membranes and generate virus-like particles. This is owing to the absence of a membrane-targeting signal in Gag9,65,202 (Fig. 20.7A). For cellular egress, FVs require the cognate envelope protein, with which their own capsids specifically interact. Coexpression of FV Gag and FV Env is required to detect capsids secreted into the cell culture supernatant. Biochemical and biophysical investigations with PFV and FFV70,207,288 suggested that a direct interaction between Gag and Env occurs (Fig. 20.7B). This aspect was later confirmed by determination of 3D structures derived from of Gag and Env subdomain cocrystals.77 This interaction is mediated by the N-terminal Gag CC1 of capsids preassembled at the centrosome, and the connection with the N-terminus of the envelope LP subunit CyD34,150,288 presumably takes place at the trans-Golgi network.299

Budding of FVs occurs at intracellular membranes and the plasma membrane with differences in the relative contribution of the sites depending on the FV species studied.134,150,299 Primate FVs, like other retroviruses, have been shown to exploit the cellular VPS machinery, through a TSG101-Gag L-domain interaction, for this process195,255 (Fig. 20.7A). It appears likely that also nonprimate FVs make use of the same mechanism, though they lack the classical PSAP L domain motif conserved in primate FV Gag proteins. Only recently, in BFV Gag, nonclassical PLPI and YGPL motifs with L-domain function were identified and shown to promote BFV budding in a TSG101- and ALIX-dependent manner.279a Whether release of FFV that depends on the host cell VPS machinery has yet to be demonstrated.

CD317/tetherin is an integral membrane protein with an N-terminal MSD and a C-terminal glycosyl-phosphatidylinositol anchor. It interacts directly with the actin cytoskeleton and blocks the release of various enveloped viruses, including orthoretroviruses, from infected cells. CD317 is also active against FVs.117,295 Unlike in HIV-1 and HIV-2, a FV protein antagonizing tetherin has not yet been identified. The activity of tetherin against PFV shows some mechanistic differences in comparison to HIV-1, because dimerization-deficient tetherin inhibits PFV replication with the same efficiency as the wild-type factor.295

Perspectives 785

Above we learned, that foamy viruses are “the other complex retroviruses” since they are apparently apathogenic and follow a different replication strategy as their cousins. The latter issue justified their classification as an own subfamily within the family of retroviruses. For the future, there remain many questions to be addressed, both in the field and in the laboratory. Certainly, more exogenous FVs await their discovery, possibly in species beyond those known so far in simians, prosimians, and probably, other classes of mammalia. Researchers in the field and in primate facilities should investigate possible impacts of coinfections with FV and other viruses. It will be interesting to elucidate further in how the immune system maintains the constant replication for instance in the oral mucosa without harming the host. An animal model with well-characterized tools for immunologic studies and smaller than primates or cats would be very helpful for these studies. Will we ever understand why mouse and man are apparently spared from foamy viruses? This question leads to whether more endogenous FV remnant sequence stretches will be identified in the genome of vertebrates. Could such new discoveries find a mutualistic contribution of ancient FVs to modern life? On the molecular side, the yet unknown receptor(s) still await discovery. Certainly, the intracellular fate of both, the incoming FVs and the assembly and release of virus offspring, may hold further surprises in comparison to other more extensively studied retroviruses. FVs are characterized by accumulation of intact capsids at the centrosome of nondividing cells that can remain biologically active for weeks. What triggers uncoating of these latent capsids upon host cell entry into mitosis? In orthoretroviruses, genome packaging is quite well understood. How do FVs selectively package their vgRNA upon capsid assembly? Another peculiar aspect of the FV replication cycle is the formation of a truncated provirus lacking the bulk part of the transactivator gene only allowing low expression levels of Bet. Does the shortened provirus play any role in FV persistence? Aside from basic research on their replication, FVs have been exploited as useful tools for gene transfer purposes. It will be interesting to see a first gene therapy clinical trial being initiated using FV-derived vectors or components thereof for treatment of patients. Ultimately, foamy viruses, like Cinderella, may surprise scientists once more at second glance. Further research will show.

ACKNOWLEDGMENTS We like to express our thanks to all the scientists who permitted us to use their figures or parts thereof for this edition. Also, we apologize to those in the yet limited foamy virus researcher’s community whose contributions did not find their way into this chapter. We shall reconvene in person, once the other virus That-Must-Not-Be-Named, which restricted our freedom, including ours while assembling this chapter, will be under tight control!

786

References 1. Achong BG, Mansell PW, Epstein MA, et al. An unusual virus in cultures from a human nasopharyngeal carcinoma. J Natl Cancer Inst 1971;46:299–307. 2. Aiewsakun P, Katzourakis A. Marine origin of retroviruses in the early Palaeozoic Era. Nat Commun 2017;8:13954. 3. Ali M, Taylor GP, Pitman RJ, et al. No evidence of antibody to human foamy virus in widespread human populations. AIDS Res Hum Retroviruses 1996;12:1473–1483. 4. Alke A, Schwantes A, Kido K, et al. The bet gene of feline foamy virus is required for virus replication. Virology 2001;287:310–320. 5. Allan JS, Broussard SR, Michaels MG, et al. Amplification of simian retroviral sequences from human recipients of baboon liver transplants. AIDS Res Hum Retroviruses 1998;14:821–824. 6. An DG, Hyun U, Shin CG. Characterization of nuclear localization signals of the prototype foamy virus integrase. J Gen Virol 2008;89:1680–1684. 7. Arhel N, Munier S, Souque P, et al. Nuclear import defect of human immunodeficiency virus type 1 DNA flap mutants is not dependent on the viral strain or target cell type. J Virol 2006;80:10262–10269. 8. Bähr A, Singer A, Hain A, et al. Interferon but not MxB inhibits foamy retroviruses. Virology 2016;488:51–60. 9. Baldwin DN, Linial ML. The roles of Pol and Env in the assembly pathway of human foamy virus. J Virol 1998;72:3658–3665. 10. Ball NJ, Nicastro G, Dutta M, et al. Structure of a spumaretrovirus Gag central domain reveals an ancient retroviral capsid. PLoS Pathog 2016;12:e1005981. 11. Bandecchi P, Matteucci D, Baldinotti F, et al. Prevalence of feline immunodeficiency virus and other retroviral infections in sick cats in Italy. Vet Immunol Immunopathol 1992;31:337–345. 12. Baunach G, Maurer B, Hahn H, et al. Functional analysis of human foamy virus accessory reading frames. J Virol 1993;67:5411–5418. 13. Berg A, Pietschmann T, Rethwilm A, et al. Determinants of foamy virus envelope glycoprotein mediated resistance to superinfection. Virology 2003;314:243–252. 14. Bergez M, Weber J, Riess M, et al. Insights into innate sensing of prototype foamy viruses in myeloid cells. Viruses 2019;11:1095. 15. Bieniasz PD, Weiss RA, McClure MO. Cell cycle dependence of foamy retrovirus infection. J Virol 1995;69:7295–7299. 16. Blair WS, Bogerd H, Cullen BR. Genetic analysis indicates that the human foamy virus Bel-1 protein contains a transcription activation domain of the acidic class. J Virol 1994;68:3803–3808. 17. Bock M, Heinkelein M, Lindemann D, et al. Cells expressing the human foamy virus (HFV) accessory Bet protein are resistant to productive HFV superinfection. Virology 1998;250:194–204. 18. Bodem J. Regulation of foamy viral transcription and RNA export. Adv Virus Res 2011;81:1–31. 19. Bodem J, Kang Y, Flügel RM. Comparative functional characterization of the feline foamy virus transactivator reveals its species specificity. Virology 2004;318:32–36. 20. Bodem J, Kräusslich HG, Rethwilm A. Acetylation of the foamy virus transactivator Tas by PCAF augments promoter-binding affinity and virus transcription. J Gen Virol 2007;88:259–263. 21. Bodem J, Löchelt M, Delius H, et al. Detection of subgenomic cDNAs and mapping of feline foamy virus mRNAs reveals complex patterns of transcription. Virology 1998;244:417–426. 22. Bodem J, Löchelt M, Winkler I, et al. Characterization of the spliced pol transcript of feline foamy virus: the splice acceptor site of the pol transcript is located in gag of foamy viruses. J Virol 1996;70:9024–9027. 23. Bodem J, Schied T, Gabriel R, et al. Foamy virus nuclear RNA export is distinct from that of other retroviruses. J Virol 2011;85:2333–2341. 24. Boneva RS, Switzer WM, Spira TJ, et al. Clinical and virological characterization of persistent human infection with simian foamy viruses. AIDS Res Hum Retroviruses 2007;23:1330–1337. 25. Boyer PL, Stenbak CR, Clark PK, et al. Characterization of the polymerase and RNase H activities of human foamy virus reverse transcriptase. J Virol 2004;78:6112–6121. 26. Boyer PL, Stenbak CR, Hoberman D, et al. In vitro fidelity of the prototype primate foamy virus (PFV) RT compared to HIV-1 RT. Virology 2007;367:253–264. 27. Brooks JI, Merks HW, Fournier J, et al. Characterization of blood-borne transmission of simian foamy virus. Transfusion 2007;47:162–170. 28. Brown P, Moreau-Dubois MC, Gajdusek DC. Persistent asymptomatic infection of the laboratory mouse by simian foamy virus type 6: a new model of retrovirus latency. Arch Virol 1982;71:229–234. 29. Butera ST, Brown J, Callahan ME, et al. Survey of veterinary conference attendees for evidence of zoonotic infection by feline retroviruses. J Am Vet Med Assoc 2000;217:1475–1479. 30. Cain D, Erlwein O, Grigg A, et al. Palindromic sequence plays a critical role in human foamy virus dimerization. J Virol 2001;75:3731–3739. 31. Calattini S, Betsem EBA, Froment A, et al. Simian foamy virus transmission from apes to humans, rural Cameroon. Emerg Infect Dis 2007;13:1314–1320. 32. Calattini S, Wanert F, Thierry B, et al. Modes of transmission and genetic diversity of foamy viruses in a Macaca tonkeana colony. Retrovirology 2006;3:23. 33. Cao W, Stricker E, Hotz-Wagenblatt A, et al. Functional analyses of bovine foamy virus-encoded miRNAs reveal the importance of a defined miRNA for virus replication and host-virus interaction. Viruses 2020;12:1250. 34. Cartellieri M, Herchenröder O, Rudolph W, et al. N-terminal Gag domain required for foamy virus particle assembly and export. J Virol 2005;79:12464–12476. 35. Cartellieri M, Rudolph W, Herchenröder O, et al. Determination of the relative amounts of Gag and Pol proteins in foamy virus particles. Retrovirology 2005;2:44. 36. Chai K, Wang Z, Xu Y, et al. Palmitoylation of the bovine foamy virus envelope glycoprotein is required for viral replication. Viruses 2020;13:31. 37. Chang J, Lee KJ, Jang KL, et al. Human foamy virus Bel1 transactivator contains a bipartite nuclear localization determinant which is sensitive to protein context and triple multimerization domains. J Virol 1995;69:801–808. 38. Chang R, Tan J, Xu F, et al. Lysine acetylation sites in bovine foamy virus transactivator BTas are important for its DNA binding activity. Virology 2011;418:21–26. 39. Chareza S, Slavkovic Lukic D, Liu Y, et al. Molecular and functional interactions of cat APOBEC3 and feline foamy and immunodeficiency virus proteins: different ways to counteract host-encoded restriction. Virology 2012;424:138–146. 40. Choudhary A, Galvin TA, Williams DK, et al. Influence of naturally occurring simian foamy viruses (SFVs) on SIV disease progression in the rhesus macaque (Macaca mulatta) model. Viruses 2013;5:1414–1430. 41. Clarke JK, Attridge JT, Gay FW. The morphogenesis of simian foamy agents. J Gen Virol 1969;4:183–188. 787

42. Corona A, Schneider A, Schweimer K, et al. Inhibition of foamy virus reverse transcriptase by human immunodeficiency virus type 1 RNase H inhibitors. Antimicrob Agents Chemother 2014;58:4086–4093. 43. Cullen BR. Retroviruses as model systems for the study of nuclear RNA export pathways. Virology 1998;249:203–210. 44. Cullen BR. Nuclear RNA export. J Cell Sci 2003;116:587–597. 45. Debons-Guillemin MC, Valla J, Gazeau J, et al. No evidence of spumaretrovirus infection markers in 19 cases of De Quervain’s thyroiditis [letter]. AIDS Res Hum Retroviruses 1992;8:1547. 46. Delelis O, Saïb A, Sonigo P. Biphasic DNA synthesis in spumaviruses. J Virol 2003;77:8141–8146. 47. Deyle DR, Li Y, Olson EM, et al. Nonintegrating foamy virus vectors. J Virol 2010;84:9341–9349. 48. Duda A, Lüftenegger D, Pietschmann T, et al. Characterization of the prototype foamy virus envelope glycoprotein receptor-binding domain. J Virol 2006;80:8158–8167. 49. Duda A, Stange A, Lüftenegger D, et al. Prototype foamy virus envelope glycoprotein leader peptide processing is mediated by a furin-like cellular protease, but cleavage is not essential for viral infectivity. J Virol 2004;78:13865–13870. 50. Dupont A, Glück IM, Ponti D, et al. Identification of an intermediate step in foamy virus fusion. Viruses 2020;12:1472. 51. Eastman SW, Linial ML. Identification of a conserved residue of foamy virus Gag required for intracellular capsid assembly. J Virol 2001;75:6857–6864. 52. Effantin G, Estrozi LF, Aschman N, et al. Cryo-electron microscopy structure of the native prototype foamy virus glycoprotein and virus architecture. PLoS Pathog 2016;12:e1005721. 53. Enders JF, Peebles TC. Propagation in tissue cultures of cytopathogenic agents from patients with measles. Proc Soc Exp Biol Med 1954;86:277–286. 54. Engelman AN, Cherepanov P. Retroviral intasomes arising. Curr Opin Struct Biol 2017;47:23–29. 55. Enssle J, Fischer N, Moebes A, et al. Carboxy-terminal cleavage of the human foamy virus Gag precursor molecule is an essential step in the viral life cycle. J Virol 1997;71:7312–7317. 56. Enssle J, Jordan I, Mauer B, et al. Foamy virus reverse transcriptase is expressed independently from the Gag protein. Proc Natl Acad Sci USA 1996;93:4137–4141. 57. Enssle J, Moebes A, Heinkelein M, et al. An active foamy virus integrase is required for virus replication. J Gen Virol 1999;80(Pt 6):1445–1452. 58. Epstein MA. Simian retroviral infections in human beings. Lancet 2004;364:138–139; author reply 139–140. 59. Erlwein O, Bieniasz PD, McClure MO. Sequences in pol are required for transfer of human foamy virus-based vectors. J Virol 1998;72:5510–5516. 60. Erlwein O, Cain D, Fischer N, et al. Identification of sites that act together to direct dimerization of human foamy virus RNA in vitro. Virology 1997;229:251–258. 61. Everson EM, Olzsko ME, Leap DJ, et al. A comparison of foamy and lentiviral vector genotoxicity in SCID-repopulating cells shows foamy vectors are less prone to clonal dominance. Mol Ther Methods Clin Dev 2016;3:16048. 62. Falcone V, Leupold J, Clotten J, et al. Sites of simian foamy virus persistence in naturally infected African green monkeys: latent provirus is ubiquitous, whereas viral replication is restricted to the oral mucosa. Virology 1999;257:7–14. 63. Falcone V, Schweizer M, Neumann-Haefelin D. Replication of primate foamy viruses in natural and experimental hosts. Curr Top Microbiol Immunol 2003;277:161–180. 64. Falcone V, Schweizer M, Toniolo A, et al. Gamma interferon is a major suppressive factor produced by activated human peripheral blood lymphocytes that is able to inhibit foamy virus-induced cytopathic effects. J Virol 1999;73:1724–1728. 65. Fischer N, Heinkelein M, Lindemann D, et al. Foamy virus particle formation. J Virol 1998;72:1610–1615. 66. Flanagan M. Isolation of a spumavirus from a sheep. Aust Vet J 1992;69:112–113. 67. Flügel RM, Pfrepper KI. Proteolytic processing of foamy virus Gag and Pol proteins. Curr Top Microbiol Immunol 2003;277:63–88. 68. Gärtner K, Wiktorowicz T, Park J, et al. Accuracy estimation of foamy virus genome copying. Retrovirology 2009;6:32. 69. Geiselhart V, Bastone P, Kempf T, et al. Furin-mediated cleavage of the feline foamy virus Env leader protein. J Virol 2004;78:13573–13581. 70. Geiselhart V, Schwantes A, Bastone P, et al. Features of the Env leader protein and the N-terminal Gag domain of feline foamy virus important for virus morphogenesis. Virology 2003;310:235–244. 71. Gessain A, Montange T, Betsem E, et al. Case–control study of the immune status of humans infected with zoonotic gorilla simian foamy viruses. J Infect Dis 2020;221:1724–1733. 72. Gharwan H, Hirata RK, Wang P, et al. Transduction of human embryonic stem cells by foamy virus vectors. Mol Ther 2007;15:1827–1833. 73. Giron ML, Colas S, Wybier J, et al. Expression and maturation of human foamy virus Gag precursor polypeptides. J Virol 1997;71:1635–1639. 74. Giron ML, Rozain F, Debons-Guillemin MC, et al. Human foamy virus polypeptides: identification of env and bel gene products. J Virol 1993;67:3596–3600. 75. Goepfert PA, Shaw KL, Ritter GD Jr, et al. A sorting motif localizes the foamy virus glycoprotein to the endoplasmic reticulum. J Virol 1997;71:778–784. 76. Goepfert PA, Shaw K, Wang G, et al. An endoplasmic reticulum retrieval signal partitions human foamy virus maturation to intracytoplasmic membranes. J Virol 1999;73:7210–7217. 77. Goldstone DC, Flower TG, Ball NJ, et al. A unique spumavirus Gag N-terminal domain with functional properties of orthoretroviral matrix and capsid. PLoS Pathog 2013;9:e1003376. 78. Goodman MA, Arumugam P, Pillis DM, et al. Foamy virus vector carries a strong insulator in its long terminal repeat which reduces its genotoxic potential. J Virol 2018;92:e01639-17. 79. Hahn H, Baunach G, Bräutigam S, et al. Reactivity of primate sera to foamy virus Gag and Bet proteins. J Gen Virol 1994;75(Pt 10):2635–2644. 80. Hamann MV, Müllers E, Reh J, et al. The cooperative function of arginine residues in the Prototype Foamy Virus Gag C-terminus mediates viral and cellular RNA encapsidation. Retrovirology 2014;11:87. 81. Hamann MV, Stanke N, Müllers E, et al. Efficient transient genetic manipulation in vitro and in vivo by prototype foamy virus-mediated nonviral RNA transfer. Mol Ther 2014;22:1460–1471. 82. Han GZ, Worobey M. Endogenous viral sequences from the Cape golden mole (Chrysochloris asiatica) reveal the presence of foamy viruses in all major placental mammal clades. PLoS One 2014;9:e97931. 83. Hare S, Gupta SS, Valkov E, et al. Retroviral intasome assembly and inhibition of DNA strand transfer. Nature 2010;464:232–236. 84. Hare S, Smith SJ, Metifiot M, et al. Structural and functional analyses of the second-generation integrase strand transfer inhibitor dolutegravir (S/GSK1349572). Mol Pharmacol 2011;80:565–572. 85. Hare S, Vos AM, Clayton RF, et al. Molecular mechanisms of retroviral integrase inhibition and the evolution of viral resistance. Proc Natl Acad Sci USA 2010;107:20057–20062. 788

86. Hartl MJ, Bodem J, Jochheim F, et al. Regulation of foamy virus protease activity by viral RNA: a novel and unique mechanism among retroviruses. J Virol 2011;85:4462–4469. 87. Hartl MJ, Schweimer K, Reger MH, et al. Formation of transient dimers by a retroviral protease. Biochem J 2010;427:197–203. 88. He F, Blair WS, Fukushima J, et al. The human foamy virus Bel-1 transcription factor is a sequence-specific DNA binding protein. J Virol 1996;70:3902–3908. 89. Heinkelein M, Pietschmann T, Jarmy G, et al. Efficient intracellular retrotransposition of an exogenous primate retrovirus genome. EMBO J 2000;19:3436–3445. 90. Heinkelein M, Rammling M, Juretzek T, et al. Retrotransposition and cell-to-cell transfer of foamy viruses. J Virol 2003;77:11855–11858. 91. Heinkelein M, Schmidt M, Fischer N, et al. Characterization of a cis-acting sequence in the Pol region required to transfer human foamy virus vectors. J Virol 1998;72:6307–6314. 92. Heinkelein M, Thurow J, Dressler M, et al. Complex effects of deletions in the 5′ untranslated region of primate foamy virus on viral gene expression and RNA packaging. J Virol 2000;74:3141–3148. 93. Heinkelein M, Dressler M, Jarmy G, et al. Improved primate foamy virus vectors and packaging constructs. J Virol 2002;76:3774–3783. 94. Heinkelein M, Leurs C, Rammling M, et al. Pregenomic RNA is required for efficient incorporation of pol polyprotein into foamy virus capsids. J Virol 2002;76:10069–10073. 95. Hendrie PC, Huo Y, Stolitenko RB, et al. A rapid and quantitative assay for measuring neighboring gene activation by vector proviruses. Mol Ther 2008;16:534–540. 96. Heneine W, Kuehnert MJ. Preserving blood safety against emerging retroviruses. Transfusion 2006;46:1276–1278. 97. Heneine W, Musey VC, Sinha SD, et al. Absence of evidence for human spumaretrovirus sequences in patients with Graves’ disease [letter]. J Acquir Immune Defic Syndr Hum Retrovirol 1995;9:99–101. 98. Heneine W, Switzer WM, Sandstrom P, et al. Identification of a human population infected with simian foamy viruses [see comments]. Nat Med 1998;4:403–407. 99. Herchenröder O, Renne R, Loncar D, et al. Isolation, cloning, and sequencing of simian foamy viruses from chimpanzees (SFVcpz): high homology to human foamy virus (HFV). Virology 1994;201:187–199. 100. Herchenröder O, Turek R, Neumann-Haefelin D, et al. Infectious proviral clones of chimpanzee foamy virus (SFVcpz) generated by long PCR reveal close functional relatedness to human foamy virus. Virology 1995;214:685–689. 101. Hill CL, Bieniasz PD, McClure MO. Properties of human foamy virus relevant to its development as a vector for gene therapy. J Gen Virol 1999;80:2003–2009. 102. Hirata RK, Miller AD, Andrews RG, et al. Transduction of hematopoietic cells by foamy virus vectors. Blood 1996;88:3654–3661. 103. Holzschu DL, Delaney MA, Renshaw RW, et al. The nucleotide sequence and spliced pol mRNA levels of the nonprimate spumavirus bovine foamy virus. J Virol 1998;72:2177–2182. 104. Hooks JJ, Gibbs CJ Jr. The foamy viruses. Bacteriol Rev 1975;39:169–185. 105. Hooks JJ, Gibbs CJ Jr, Cutchins EC, et al. Characterization and distribution of two new foamy viruses isolated from chimpanzees. Arch Gesamte Virusforsch 1972;38:38–55. 106. Hu X, Yang W, Liu R, et al. N-Myc interactor inhibits prototype foamy virus by sequestering viral Tas protein in the cytoplasm. J Virol 2014;88:7036–7044. 107. Hussain AI, Shanmugam V, Bhullar VB, et al. Screening for simian foamy virus infection by using a combined antigen Western blot assay: evidence for a wide distribution among Old World primates and identification of four new divergent viruses. Virology 2003;309:248–257. 108. Hütter S, Müllers E, Stanke N, et al. Prototype foamy virus protease activity is essential for intraparticle reverse transcription initiation but not absolutely required for uncoating upon host cell entry. J Virol 2013;87:3163–3176. 109. Hyun U, Lee DH, Shin CG. Minimal size of prototype foamy virus integrase for nuclear localization. Acta Virol 2011;55:169–174. 110. Imrich H, Heinkelein M, Herchenröder O, et al. Primate foamy virus Pol proteins are imported into the nucleus. J Gen Virol 2000;81:2941–2947. 111. Jaguva Vasudevan AA, Becker D, Luedde T, et al. Foamy viruses, bet, and APOBEC3 restriction. Viruses 2021;13:504. 112. Jaguva Vasudevan AA, Perkovic M, Bulliard Y, et al. Prototype foamy virus bet impairs the dimerization and cytosolic solubility of human APOBEC3G. J Virol 2013;87:9030–9040. 113. Johnston PB. Taxonomic features of seven serotypes of simian and ape foamy viruses. Infect Immun 1971;3:793–799. 114. Jones-Engel L, May CC, Engel GA, et al. Diverse contexts of zoonotic transmission of simian foamy viruses in Asia. Emerg Infect Dis 2008;14:1200–1208. 115. Jones-Engel L, Steinkraus KA, Murray SM, et al. Sensitive assays for simian foamy viruses reveal a high prevalence of infection in commensal, free-ranging Asian monkeys. J Virol 2007;81:7330–7337. 116. Jordan I, Enssle J, Güttler E, et al. Expression of human foamy virus reverse transcriptase involves a spliced pol mRNA. Virology 1996;224:314–319. 117. Jouvenet N, Neil SJ, Zhadina M, et al. Broad-spectrum inhibition of retroviral and filoviral particle release by tetherin. J Virol 2009;83:1837–1844. 118. Juretzek T, Holm T, Gärtner K, et al. Foamy virus integration. J Virol 2004;78:2472–2477. 119. Kane M, Mele V, Liberatore RA, et al. Inhibition of spumavirus gene expression by PHF11. PLoS Pathog 2020;16:e1008644. 120. Kane M, Zang TM, Rihn SJ, et al. Identification of interferon-stimulated genes with antiretroviral activity. Cell Host Microbe 2016;20:392–405. 121. Kang Y, Blair WS, Cullen BR. Identification and functional characterization of a high-affinity Bel-1 DNA binding site located in the human foamy virus internal promoter. J Virol 1998;72:504–511. 122. Katzourakis A, Gifford RJ, Tristem M, et al. Macroevolution of complex retroviruses. Science 2009;325:1512. 123. Keller A, Partin KM, Löchelt M, et al. Characterization of the transcriptional trans activator of human foamy retrovirus. J Virol 1991;65:2589–2594. 124. Khan AS, Kumar D. Simian foamy virus infection by whole-blood transfer in rhesus macaques: potential for transfusion transmission in humans. Transfusion 2006;46:1352–1359. 125. Khan AS, Bodem J, Buseyne F, et al. Spumaretroviruses: updated taxonomy and nomenclature. Virology 2018;516:158–164. 126. Khan IH, Mendoza S, Yee J, et al. Simultaneous detection of antibodies to six nonhuman-primate viruses by multiplex microbead immunoassay. Clin Vaccine Immunol 2006;13:45–52. 127. Kido K, Bannert H, Gronostajski RM, et al. Bel1-mediated transactivation of the spumaretroviral internal promoter is repressed by nuclear factor I. J Biol Chem 2003;278:11836–11842. 128. Kido K, Doerks A, Löchelt M, et al. Identification and functional characterization of an intragenic DNA binding site for the spumaretroviral trans-activator in the human p57Kip2 gene. J Biol Chem 2002;277:12032–12039. 129. Kincaid RP, Chen Y, Cox JE, et al. Noncanonical microRNA (miRNA) biogenesis gives rise to retroviral mimics of lymphoproliferative and 789

immunosuppressive host miRNAs. mBio 2014;5:e00074. 130. Konvalinka J, Löchelt M, Zentgraf H, et al. Active foamy virus proteinase is essential for virus infectivity but not for formation of a Pol polyprotein. J Virol 1995;69:7264–7268. 131. Kretzschmar B, Nowrouzi A, Hartl MJ, et al. AZT-resistant foamy virus. Virology 2008;370:151–157. 132. Lambert C, Couteaudier M, Gouzil J, et al. Potent neutralizing antibodies in humans infected with zoonotic simian foamy viruses target conserved epitopes located in the dimorphic domain of the surface envelope protein. PLoS Pathog 2018;14:e1007293. 133. Lecellier CH, Saïb A. Foamy viruses: between retroviruses and pararetroviruses. Virology 2000;271:1–8. 134. Lecellier CH, Neves M, Giron ML, et al. Further characterization of equine foamy virus reveals unusual features among the foamy viruses. J Virol 2002;76:7220–7227. 135. Lee EG, Linial ML. The C terminus of foamy retrovirus Gag contains determinants for encapsidation of Pol protein into virions. J Virol 2008;82:10803–10810. 136. Lee EG, Kuppers D, Horn M, et al. A premature termination codon mutation at the C terminus of foamy virus Gag downregulates the levels of spliced pol mRNA. J Virol 2008;82:1656–1664. 137. Lee EG, Sinicrope A, Jackson DL, et al. Foamy virus Pol protein expressed as a Gag-Pol fusion retains enzymatic activities, allowing for infectious virus production. J Virol 2012;86:5992–6001. 138. Lee CC, Ye F, Tarantal AF. Comparison of growth and differentiation of fetal and adult rhesus monkey mesenchymal stem cells. Stem Cells Dev 2006;15:209–220. 139. Leendertz FH, Zirkel F, Couacy-Hymann E, et al. Interspecies transmission of simian foamy virus in a natural predator–prey system. J Virol 2008;82:7741–7744. 140. Lehmann-Che J, Giron ML, Delelis O, et al. Protease-dependent uncoating of a complex retrovirus. J Virol 2005;79:9244–9253. 141. Lehmann-Che J, Renault N, Giron ML, et al. Centrosomal latency of incoming foamy viruses in resting cells. PLoS Pathog 2007;3:e74. 142. Leo B, Hartl MJ, Schweimer K, et al. Insights into the structure and activity of prototype foamy virus RNase H. Retrovirology 2012;9:14. 143. Leo B, Schweimer K, Rösch P, et al. The solution structure of the prototype foamy virus RNase H domain indicates an important role of the basic loop in substrate binding. Retrovirology 2012;9:73. 144. Lesbats P, Engelman AN, Cherepanov P. Retroviral DNA integration. Chem Rev 2016;116:12730–12757. 145. Lesbats P, Serrao E, Maskell DP, et al. Structural basis for spumavirus GAG tethering to chromatin. Proc Natl Acad Sci USA 2017;114:5509–5514. 146. Life RB, Lee EG, Eastman SW, et al. Mutations in the amino terminus of foamy virus Gag disrupt morphology and infectivity but do not target assembly. J Virol 2008;82:6109–6119. 147. Lindemann D, Rethwilm A. Characterization of a human foamy virus 170-kilodalton Env-Bet fusion protein generated by alternative splicing. J Virol 1998;72:4088–4094. 148. Lindemann D, Rethwilm A. Foamy virus biology and its application for vector development. Viruses 2011;3:561–585. 149. Lindemann D, Hütter S, Wei G, et al. The unique, the known, and the unknown of spumaretrovirus assembly. Viruses 2021;13:105. 150. Lindemann D, Pietschmann T, Picard-Maureau M, et al. A particle-associated glycoprotein signal peptide essential for virus maturation and infectivity. J Virol 2001;75:5762–5771. 151. Linial ML. Foamy viruses are unconventional retroviruses. J Virol 1999;73:1747–1755. 152. Linial M, Fan H, Hahn B, et al. Retroviridae. In: Fauquet C, Mayo M, Maniloff J, et al., eds. Virus Taxonomy. 2nd ed. Elsevier Inc.; 2005:421–440. 153. Liu W, Backes P, Löchelt M. Importance of the major splice donor and redefinition of cis-acting sequences of gutless feline foamy virus vectors. Virology 2009;394:208–217. 154. Liu W, Worobey M, Li Y, et al. Molecular ecology and natural history of simian foamy virus infection in wild-living chimpanzees. PLoS Pathog 2008;4:e1000097. 155. Löchelt M. Foamy virus transactivation and gene expression. Curr Top Microbiol Immunol 2003;277:27–61. 156. Löchelt M, Flügel RM. The human foamy virus pol gene is expressed as a Pro-Pol polyprotein and not as a Gag-Pol fusion protein. J Virol 1996;70:1033–1040. 157. Löchelt M, Flügel RM, Aboud M. The human foamy virus internal promoter directs the expression of the functional Bel 1 transactivator and Bet protein early after infection. J Virol 1994;68:638–645. 158. Löchelt M, Muranyi W, Flügel RM. Human foamy virus genome possesses an internal, Bel-1-dependent and functional promoter. Proc Natl Acad Sci USA 1993;90:7317–7321. 159. Löchelt M, Romen F, Bastone P, et al. The antiretroviral activity of APOBEC3 is inhibited by the foamy virus accessory Bet protein. Proc Natl Acad Sci USA 2005;102:7982–7987. 160. Löchelt M, Zentgraf H, Flügel RM. Construction of an infectious DNA clone of the full-length human spumaretrovirus genome and mutagenesis of the bel 1 gene. Virology 1991;184:43–54. 161. Loh PC, Achong BC, Epstein MA. Further biological properties of the human syncytial virus. Intervirology 1977;8:204–217. 162. Loh PC, Matsuura F, Mizumoto C. Seroepidemiology of human syncytial virus: antibody prevalence in the Pacific. Intervirology 1980;13:87–90. 163. Lüftenegger D, Picard-Maureau M, Stanke N, et al. Analysis and function of prototype foamy virus envelope N glycosylation. J Virol 2005;79:7664–7672. 164. Lukic DS, Hotz-Wagenblatt A, Lei J, et al. Identification of the feline foamy virus Bet domain essential for APOBEC3 counteraction. Retrovirology 2013;10:76. 165. Maertens GN, Hare S, Cherepanov P. The mechanism of retroviral integration from X-ray structures of its key intermediates. Nature 2010;468:326–329. 166. Mahnke C, Kashaiya P, Rössler J, et al. Human spumavirus antibodies in sera from African patients. Arch Virol 1992;123:243–253. 167. Malmquist WA, Van der Maaten MJ, Boothe AD. Isolation, immunodiffusion, immunofluorescence, and electron microscopy of a syncytial virus of lymphosarcomatous and apparently normal cattle. Cancer Res 1969;29:188–200. 168. Mannigel I, Stange A, Zentgraf H, et al. Correct capsid assembly mediated by a conserved YXXLGL motif in prototype foamy virus Gag is essential for infectivity and reverse transcription of the viral genome. J Virol 2007;81:3317–3326. 169. Maskell DP, Renault L, Serrao E, et al. Structural basis for retroviral integration into nucleosomes. Nature 2015;523:366–369. 170. Materniak-Kornas M, Tan J, Heit-Mondrzyk A, et al. Bovine foamy virus: shared and unique molecular features in vitro and in vivo. Viruses 2019;11:1084. 171. Matthes D, Wiktorowicz T, Zahn J, et al. Basic residues in the foamy virus Gag protein. J Virol 2011;85:3986–3995. 172. Maurer B, Bannert H, Darai G, et al. Analysis of the primary structure of the long terminal repeat and the gag and pol genes of the human spumaretrovirus. J Virol 1988;62:1590–1597. 173. Maurer B, Serfling E, ter Meulen V, et al. Transcription factor AP-1 modulates the activity of the human foamy virus long terminal repeat. J Virol 1991;65:6353–6357. 174. Meiering CD, Linial ML. Historical perspective of foamy virus epidemiology and infection. Clin Microbiol Rev 2001;14:165–176. 790

175. Meiering CD, Linial ML. Reactivation of a complex retrovirus is controlled by a molecular switch and is inhibited by a viral protein. Proc Natl Acad Sci USA 2002;99:15130–15135. 176. Meiering CD, Comstock KE, Linial ML. Multiple integrations of human foamy virus in persistently infected human erythroleukemia cells. J Virol 2000;74:1718–1726. 177. Meiering CD, Rubio C, May C, et al. Cell-type-specific regulation of the two foamy virus promoters. J Virol 2001;75:6547–6557. 178. Mergia A, Blackwell J, Papadi G, et al. Simian foamy virus type 1 (SFV-1) induces apoptosis. Virus Res 1997;50:129–137. 179. Mergia A, Renshaw-Gegg LW, Stout MW, et al. Functional domains of the simian foamy virus type 1 transcriptional transactivator (Taf). J Virol 1993;67:4598–4604. 180. Moebes A, Enssle J, Bieniasz PD, et al. Human foamy virus reverse transcription that occurs late in the viral replication cycle. J Virol 1997;71:7305–7311. 181. Moschall R, Denk S, Erkelenz S, et al. A purine-rich element in foamy virus pol regulates env splicing and gag/pol expression. Retrovirology 2017;14:10. 182. Müllers E, Stirnnagel K, Kaulfuss S, et al. Prototype foamy virus gag nuclear localization: a novel pathway among retroviruses. J Virol 2011;85:9276–9285. 183. Muranyi W, Flügel RM. Analysis of splicing patterns of human spumaretrovirus by polymerase chain reaction reveals complex RNA structures. J Virol 1991;65:727–735. 184. Murray SM, Picker LJ, Axthelm MK, et al. Expanded tissue targets for foamy virus replication with simian immunodeficiency virusinduced immunosuppression. J Virol 2006;80:663–670. 185. Murray SM, Picker LJ, Axthelm MK, et al. Replication in a superficial epithelial cell niche explains the lack of pathogenicity of primate foamy virus infections. J Virol 2008;82:5981–5985. 186. Nasimuzzaman M, Persons DA. Cell Membrane-associated heparan sulfate is a receptor for prototype foamy virus in human, monkey, and rodent cells. Mol Ther 2012;20:1158–1166. 187. Neumann-Haefelin D, Rethwilm A, Bauer G, et al. Characterization of a foamy virus isolated from Cercopithecus aethiops lymphoblastoid cells. Med Microbiol Immunol 1983;172:75–86. 188. Nowacka M, Nowak E, Czarnocki-Cieciura M, et al. Structures of substrate complexes of foamy viral protease-reverse transcriptase. J Virol 2021;95:e0084821. 189. Nowrouzi A, Dittrich M, Klanke C, et al. Genome-wide mapping of foamy virus vector integrations into a human cell line. J Gen Virol 2006;87:1339–1347. 190. Ohmine K, Li Y, Bauer TR Jr, et al. Tracking of specific integrant clones in dogs treated with foamy virus vectors. Hum Gene Ther 2011;22:217–224. 191. Pacheco B, Finzi A, McGee-Estrada K, et al. Species-specific inhibition of foamy viruses from South American monkeys by New World Monkey TRIM5{alpha} proteins. J Virol 2010;84:4095–4099. 192. Paris J, Tobaly-Tapiero J, Giron ML, et al. The invariant arginine within the chromatin-binding motif regulates both nucleolar localization and chromatin binding of Foamy virus Gag. Retrovirology 2018;15:48. 193. Parks WD, Todaro GJ, Scolnick EM, et al. RNA dependent DNA polymerase in primate syncytium-forming (foamy) viruses. Nature 1971;229:258–260. 194. Patton GS, Erlwein O, McClure MO. Cell-cycle dependence of foamy virus vectors. J Gen Virol 2004;85:2925–2930. 195. Patton GS, Morris SA, Chung W, et al. Identification of domains in gag important for prototypic foamy virus egress. J Virol 2005;79:6392–6399. 196. Perkovic M, Schmidt S, Marino D, et al. Species-specific inhibition of APOBEC3C by the prototype foamy virus protein bet. J Biol Chem 2009;284:5819–5826. 197. Peters K, Barg N, Gärtner K, et al. Complex effects of foamy virus central purine-rich regions on viral replication. Virology 2008;373:51–60. 198. Peters K, Wiktorowicz T, Heinkelein M, et al. RNA and protein requirements for incorporation of the Pol protein into foamy virus particles. J Virol 2005;79:7005–7013. 199. Petit C, Giron ML, Tobaly-Tapiero J, et al. Targeting of incoming retroviral Gag to the centrosome involves a direct interaction with the dynein light chain 8. J Cell Sci 2003;116:3433–3442. 200. Pfrepper KI, Löchelt M, Rackwitz HR, et al. Molecular characterization of proteolytic processing of the Gag proteins of human spumavirus. J Virol 1999;73:7907–7911. 201. Picard-Maureau M, Jarmy G, Berg A, et al. Foamy virus envelope glycoprotein-mediated entry involves a pH-dependent fusion process. J Virol 2003;77:4722–4730. 202. Pietschmann T, Heinkelein M, Heldmann M, et al. Foamy virus capsids require the cognate envelope protein for particle export. J Virol 1999;73:2613–2621. 203. Pietschmann T, Zentgraf H, Rethwilm A, et al. An evolutionarily conserved positively charged amino acid in the putative membranespanning domain of the foamy virus envelope protein controls fusion activity. J Virol 2000;74:4474–4482. 204. Pinto-Santini DM, Stenbak CR, Linial ML. Foamy virus zoonotic infections. Retrovirology 2017;14:55. 205. Plochmann K, Horn A, Gschmack E, et al. Heparan sulfate is an attachment factor for foamy virus entry. J Virol 2012;86:10028–10035. 206. Rajawat YS, Humbert O, Kiem HP. In-vivo gene therapy with foamy virus vectors. Viruses 2019;11:1091. 207. Reh J, Stange A, Götz A, et al. An N-terminal domain helical motif of Prototype Foamy Virus Gag with dual functions essential for particle egress and viral infectivity. Retrovirology 2013;10:45. 208. Renault N, Tobaly-Tapiero J, Paris J, et al. A nuclear export signal within the structural Gag protein is required for prototype foamy virus replication. Retrovirology 2011;8:6. 209. Renne R, Friedl E, Schweizer M, et al. Genomic organization and expression of simian foamy virus type 3 (SFV-3). Virology 1992;186:597–608. 210. Repaske R, Hartley JW, Kavlick MF, et al. Inhibition of RNase H activity and viral replication by single mutations in the 3′ region of Moloney murine leukemia virus reverse transcriptase. J Virol 1989;63:1460–1464. 211. Rethwilm A. Regulation of foamy virus gene expression. Curr Top Microbiol Immunol 1995;193:1–24. 212. Rethwilm A. Unexpected replication pathways of foamy viruses. J Acquir Immune Defic Syndr Hum Retrovirol 1996;13(Suppl 1):S248–253. 213. Rethwilm A. The replication strategy of foamy viruses. Curr Top Microbiol Immunol 2003;277:1–26. 214. Rethwilm A. Foamy viruses. In: Mahy BWJ, ter Meulen V, eds. Topley & Wilson’s Microbiology Virology. Edward Arnold; 2005:1304–1321. 215. Rethwilm A, Baunach G, Netzer KO, et al. Infectious DNA of the human spumaretrovirus. Nucleic Acids Res 1990;18:733–738. 216. Rethwilm A, Darai G, Rösen A, et al. Molecular cloning of the genome of human spumaretrovirus. Gene 1987;59:19–28. 217. Rethwilm A, Erlwein O, Baunach G, et al. The transcriptional transactivator of human foamy virus maps to the bel 1 genomic region. Proc Natl Acad Sci USA 1991;88:941–945. 791

218. Riggs JL, Oshirls, Taylor DO, et al. Syncytium-forming agent isolated from domestic cats. Nature 1969;222:1190–1191. 219. Rinke CS, Boyer PL, Sullivan MD, et al. Mutation of the catalytic domain of the foamy virus reverse transcriptase leads to loss of processivity and infectivity. J Virol 2002;76:7560–7570. 220. Romen F, Backes P, Materniak M, et al. Serological detection systems for identification of cows shedding bovine foamy virus via milk. Virology 2007;364:123–131. 221. Romen F, Pawlita M, Sehr P, et al. Antibodies against Gag are diagnostic markers for feline foamy virus infections while Env and Bet reactivity is undetectable in a substantial fraction of infected cats. Virology 2006;345:502–508. 222. Rosenblum LL, Patton G, Grigg AR, et al. Differential susceptibility of retroviruses to nucleoside analogues. Antivir Chem Chemother 2001;12:91–97. 223. Rösener M, Hahn H, Kranz M, et al. Absence of serological evidence for foamy virus infection in patients with amyotrophic lateral sclerosis. J Med Virol 1996;48:222–226. 224. Roy J, Linial ML. Role of the foamy virus Pol cleavage site in viral replication. J Virol 2007;81:4956–4962. 225. Roy J, Rudolph W, Juretzek T, et al. Feline foamy virus genome and replication strategy. J Virol 2003;77:11324–11331. 226. Rua R, Betsem E, Montange T, et al. In vivo cellular tropism of gorilla simian foamy virus in blood of infected humans. J Virol 2014;88:13429–13435. 227. Rua R, Lepelley A, Gessain A, et al. Innate sensing of foamy viruses by human hematopoietic cells. J Virol 2012;86:909–918. 228. Russell RA, Wiegand HL, Moore MD, et al. Foamy virus Bet proteins function as novel inhibitors of the APOBEC3 family of innate antiretroviral defense factors. J Virol 2005;79:8724–8731. 229. Russell RA, Zeng Y, Erlwein O, et al. The R region found in the human foamy virus long terminal repeat is critical for both Gag and Pol protein expression. J Virol 2001;75:6817–6824. 230. Rustigian R, Johnston P, Reihart H. Infection of monkey kidney tissue cultures with virus-like agents. Proc Soc Exp Biol Med 1955;88:8–16. 231. Ryabova LA, Hohn T. Ribosome shunting in the cauliflower mosaic virus 35S RNA leader is a special case of reinitiation of translation functioning in plant and animal systems. Genes Dev 2000;14:817–829. 232. Saïb A, Neves M, Giron ML, et al. Long-term persistent infection of domestic rabbits by the human foamy virus. Virology 1997;228:263–268. 233. Saïb A, Peries J, de The H. A defective human foamy provirus generated by pregenome splicing. EMBO J 1993;12:4439–4444. 234. Saïb A, Puvion-Dutilleul F, Schmid M, et al. Nuclear targeting of incoming human foamy virus Gag proteins involves a centriolar step. J Virol 1997;71:1155–1161. 235. Santos AF, Cavalcante LTF, Muniz CP, et al. Simian foamy viruses in Central and South America: a new world of discovery. Viruses 2019;11:967. 236. Schepetilnikov M, Schott G, Katsarou K, et al. Molecular dissection of the prototype foamy virus (PFV) RNA 5′-UTR identifies essential elements of a ribosomal shunt. Nucleic Acids Res 2009;37:5838–5847. 237. Schliephake AW, Rethwilm A. Nuclear localization of foamy virus Gag precursor protein. J Virol 1994;68:4946–4954. 238. Schmidt M, Rethwilm A. Replicating foamy virus-based vectors directing high level expression of foreign genes. Virology 1995;210:167–178. 239. Schmidt M, Herchenröder O, Heeney J, et al. Long terminal repeat U3 length polymorphism of human foamy virus. Virology 1997;230:167–178. 240. Schmidt M, Niewiesk S, Heeney J, et al. Mouse model to study the replication of primate foamy viruses. J Gen Virol 1997;78(Pt 8):1929–1933. 241. Schneider A, Peter D, Schmitt J, et al. Structural requirements for enzymatic activities of foamy virus protease-reverse transcriptase. Proteins 2014;82:375–385. 242. Schrom EM, Moschall R, Hartl MJ, et al. U1snRNP-mediated suppression of polyadenylation in conjunction with the RNA structure controls poly (A) site selection in foamy viruses. Retrovirology 2013;10:55. 243. Schweizer M, Neumann-Haefelin D. Phylogenetic analysis of primate foamy viruses by comparison of pol sequences. Virology 1995;207:577–582. 244. Schweizer M, Falcone V, Gänge J, et al. Simian foamy virus isolated from an accidentally infected human individual. J Virol 1997;71:4821–4824. 245. Schweizer M, Fleps U, Jäckle A, et al. Simian foamy virus type 3 (SFV-3) in latently infected Vero cells: reactivation by demethylation of proviral DNA. Virology 1993;192:663–666. 246. Schweizer M, Renne R, Neumann-Haefelin D. Structural analysis of proviral DNA in simian foamy virus (LK-3)-infected cells. Arch Virol 1989;109:103–114. 247. Schweizer M, Turek R, Hahn H, et al. Markers of foamy virus infections in monkeys, apes, and accidentally infected humans: appropriate testing fails to confirm suspected foamy virus prevalence in humans. AIDS Res Hum Retroviruses 1995;11:161–170. 248. Schweizer M, Turek R, Reinhardt M, et al. Absence of foamy virus DNA in Graves’ disease. AIDS Res Hum Retroviruses 1994;10:601–605. 249. Sfakianos JN, LaCasse RA, Hunter E. The M-PMV cytoplasmic targeting-retention signal directs nascent Gag polypeptides to a pericentriolar region of the cell. Traffic 2003;4:660–670. 250. Shankar A, Shanmugam V, Switzer WM. Complete genome sequence of a baboon simian foamy virus isolated from an infected human. Microbiol Resour Announc 2020;9:e00522-20. 251. Shaw KL, Lindemann D, Mulligan MJ, et al. Foamy virus envelope glycoprotein is sufficient for particle budding and release. J Virol 2003;77:2338–2348. 252. Simantirakis E, Tsironis I, Vassilopoulos G. FV vectors as alternative gene vehicles for gene transfer in HSCs. Viruses 2020;12:332. 253. Smiley Evans T, Gilardi KV, Barry PA, et al. Detection of viruses using discarded plants from wild mountain gorillas and golden monkeys. Am J Primatol 2016;78:1222–1234. 254. Stange A, Lüftenegger D, Reh J, et al. Subviral particle release determinants of prototype foamy virus. J Virol 2008;82:9858–9869. 255. Stange A, Mannigel I, Peters K, et al. Characterization of prototype foamy virus gag late assembly domain motifs and their role in particle egress and infectivity. J Virol 2005;79:5466–5476. 256. Stanke N, Stange A, Lüftenegger D, et al. Ubiquitination of the prototype foamy virus envelope glycoprotein leader peptide regulates subviral particle release. J Virol 2005;79:15074–15083. 257. Stenbak CR, Linial ML. Role of the C terminus of foamy virus Gag in RNA packaging and Pol expression. J Virol 2004;78:9423–9430. 258. Stirnnagel K, Lüftenegger D, Stange A, et al. Analysis of prototype foamy virus particle-host cell interaction with autofluorescent retroviral particles. Retrovirology 2010;7:45. 259. Stirnnagel K, Schupp D, Dupont A, et al. Differential pH-dependent cellular uptake pathways among foamy viruses elucidated using dualcolored fluorescent particles. Retrovirology 2012;9:71. 260. Swiersy A, Wiek C, Reh J, et al. Orthoretroviral-like prototype foamy virus Gag-Pol expression is compatible with viral replication. 792

Retrovirology 2011;8:66. 261. Switzer WM, Bhullar V, Shanmugam V, et al. Frequent simian foamy virus infection in persons occupationally exposed to nonhuman primates. J Virol 2004;78:2780–2789. 262. Switzer WM, Garcia AD, Yang C, et al. Coinfection with HIV-1 and simian foamy virus in West Central Africans. J Infect Dis 2008;197:1389–1393. 263. Switzer WM, Salemi M, Shanmugam V, et al. Ancient co-speciation of simian foamy viruses and primates. Nature 2005;434:376–380. 264. Tan J, Hao P, Jia R, et al. Identification and functional characterization of BTas transactivator as a DNA-binding protein. Virology 2010;405:408–413. 265. Tan J, Qiao W, Wang J, et al. IFP35 is involved in the antiviral function of interferon by association with the viral tas transactivator of bovine foamy virus. J Virol 2008;82:4275–4283. 266. Tan J, Qiao W, Xu F, et al. Dimerization of BTas is required for the transactivational activity of bovine foamy virus. Virology 2008;376:236–241. 267. Thümer L, Rethwilm A, Holmes EC, et al. The complete nucleotide sequence of a New World simian foamy virus. Virology 2007;369:191–197. 268. Tisdale M, Schulze T, Larder BA, et al. Mutations within the RNase H domain of human immunodeficiency virus type 1 reverse transcriptase abolish virus infectivity. J Gen Virol 1991;72(Pt 1):59–66. 269. Tobaly-Tapiero J, Bittoun P, Giron ML, et al. Human foamy virus capsid formation requires an interaction domain in the N terminus of Gag. J Virol 2001;75:4367–4375. 270. Tobaly-Tapiero J, Bittoun P, Lehmann-Che J, et al. Chromatin tethering of incoming foamy virus by the structural Gag protein. Traffic 2008;9:1717–1727. 271. Tobaly-Tapiero J, Bittoun P, Neves M, et al. Isolation and characterization of an equine foamy virus. J Virol 2000;74:4064–4073. 272. Tobaly-Tapiero J, Kupiec JJ, Santillana-Hayat M, et al. Further characterization of the gapped DNA intermediates of human spumavirus: evidence for a dual initiation of plus-strand DNA synthesis. J Gen Virol 1991;72(Pt 3):605–608. 273. Trobridge G, Russell DW. Cell cycle requirements for transduction by foamy virus vectors compared to those of oncovirus and lentivirus vectors. J Virol 2004;78:2327–2335. 274. Trobridge GD, Miller DG, Jacobs MA, et al. Foamy virus vector integration sites in normal human cells. Proc Natl Acad Sci USA 2006;103:1498–1503. 275. Valkov E, Gupta SS, Hare S, et al. Functional and structural characterization of the integrase from the prototype foamy virus. Nucleic Acids Res 2009;37:243–255. 276. Vassilopoulos G, Rethwilm A. The usefulness of a perfect parasite. Gene Ther 2008;15:1299–1301. 277. Voss M, Fukumori A, Kuhn PH, et al. Foamy virus envelope protein is a substrate for signal peptide peptidase-like 3 (SPPL3). J Biol Chem 2012;287:43401–43409. 278. Wagner A, Doerks A, Aboud M, et al. Induction of cellular genes is mediated by the Bel1 transactivator in foamy virus-infected human cells. J Virol 2000;74:4441–4447. 279. Wang J, Tan J, Guo H, et al. Bovine foamy virus transactivator BTas interacts with cellular RelB to enhance viral transcription. J Virol 2010;84:11888–11897. 279a.Wang Z, Li R, Liu C, et al. Characterization of bovine foamy virus gag late assembly domain motifs and their role in recruiting ESCRT for budding. Viruses 2022;13:522. 280. Weaver TA, Talbot KJ, Panganiban AT. Spleen necrosis virus gag polyprotein is necessary for particle assembly and release but not for proteolytic processing. J Virol 1990;64:2642–2652. 281. Wei X, Chen Y, Duan G, et al. A reptilian endogenous foamy virus sheds light on the early evolution of retroviruses. Virus Evol 2019;5:vez001. 282. Wei G, Kehl T, Bao Q, et al. The chromatin binding domain, including the QPQRYG motif, of feline foamy virus Gag is required for viral DNA integration and nuclear accumulation of Gag and the viral genome. Virology 2018;524:56–68. 283. Weiss RA. Foamy retroviruses. A virus in search of a disease [news]. Nature 1988;333:497–498. 284. Weiss RA. Reverse transcription. Foamy viruses bubble on [news]. Nature 1996;380:201. 285. Whisnant AW, Kehl T, Bao Q, et al. Identification of novel, highly expressed retroviral microRNAs in cells infected by bovine foamy virus. J Virol 2014;88:4679–4686. 286. Wiktorowicz T, Peters K, Armbruster N, et al. Generation of an improved foamy virus vector by dissection of cis-acting sequences. J Gen Virol 2009;90:481–487. 287. Wilk T, de Haas F, Wagner A, et al. The intact retroviral Env glycoprotein of human foamy virus is a trimer. J Virol 2000;74:2885–2887. 288. Wilk T, Geiselhart V, Frech M, et al. Specific interaction of a novel foamy virus Env leader protein with the N-terminal Gag domain. J Virol 2001;75:7995–8007. 289. Wilson MD, Renault L, Maskell DP, et al. Retroviral integration into nucleosomes through DNA looping and sliding along the histone octamer. Nat Commun 2019;10:4189. 290. Winkler I, Bodem J, Haas L, et al. Characterization of the genome of feline foamy virus and its proteins shows distinct features different from those of primate spumaviruses. J Virol 1997;71:6727–6741. 291. Winkler IG, Löchelt M, Flower RL. Epidemiology of feline foamy virus and feline immunodeficiency virus infections in domestic and feral cats: a seroepidemiological study. J Clin Microbiol 1999;37:2848–2851. 292. Wolfe ND, Switzer WM, Carr JK, et al. Naturally acquired simian retrovirus infections in central African hunters. Lancet 2004;363:932–937. 293. Wu M, Chari S, Yanchis T, et al. cis-Acting sequences required for simian foamy virus type 1 vectors. J Virol 1998;72:3451–3454. 294. Wu Y, Tan J, Su Y, et al. Transcription factor AP1 modulates the internal promoter activity of bovine foamy virus. Virus Res 2010;147:139–144. 295. Xu F, Tan J, Liu R, et al. Tetherin inhibits prototypic foamy virus release. Virol J 2011;8:198. 296. Yap MW, Lindemann D, Stanke N, et al. Restriction of foamy viruses by primate Trim5alpha. J Virol 2008;82:5429–5439. 297. Yu SF, Linial ML. Analysis of the role of the bel and bet open reading frames of human foamy virus by using a new quantitative assay. J Virol 1993;67:6618–6624. 298. Yu SF, Baldwin DN, Gwynn SR, et al. Human foamy virus replication: a pathway distinct from that of retroviruses and hepadnaviruses. Science 1996;271:1579–1582. 299. Yu SF, Eastman SW, Linial ML. Foamy virus capsid assembly occurs at a pericentriolar region through a cytoplasmic targeting/retention signal in Gag. Traffic 2006;7:966–977 300. Yu SF, Edelmann K, Strong RK, et al. The carboxyl terminus of the human foamy virus Gag protein contains separable nucleic acid binding and nuclear transport domains. J Virol 1996;70:8255–8262. 793

301. Yu SF, Lujan P, Jackson DL, et al. The DEAD-box RNA helicase DDX6 is required for efficient encapsidation of a retroviral genome. PLoS Pathog 2011;7:e1002303. 302. Yu SF, Stone J, Linial ML. Productive persistent infection of hematopoietic cells by human foamy virus. J Virol 1996;70:1250–1254. 303. Yu SF, Sullivan MD, Linial ML. Evidence that the human foamy virus genome is DNA. J Virol 1999;73:1565–1572. 304. Zamborlini A, Renault N, Saïb A, et al. Early reverse transcription is essential for productive foamy virus infection. PLoS One 2010;5:e11023. 305. Zemba M, Wilk T, Rutten T, et al. The carboxy-terminal p3Gag domain of the human foamy virus Gag precursor is required for efficient virus infectivity. Virology 1998;247:7–13. 306. Zenger E, Brown WC, Song W, et al. Evaluation of cofactor effect of feline syncytium-forming virus on feline immunodeficiency virus infection. Am J Vet Res 1993;54:713–718. 307. Zhadina M, Bieniasz PD. Functional interchangeability of late domains, late domain cofactors and ubiquitin in viral budding. PLoS Pathog 2010;6:e1001153. 308. Zhadina M, McClure MO, Johnson MC, et al. Ubiquitin-dependent virus particle budding without viral protein ubiquitination. Proc Natl Acad Sci USA 2007;104:20031–20036. 309. Zou JX, Luciw PA. The transcriptional transactivator of simian foamy virus 1 binds to a DNA target element in the viral internal promoter. Proc Natl Acad Sci USA 1996;93:326–330. 310. Zurnic I, Hütter S, Rzeha U, et al. Interactions of prototype foamy virus capsids with host cell polo-like kinases are important for efficient viral DNA integration. PLoS Pathog 2016;12:e1005860.

794

CHAPTER 21 Severe Acute Respiratory Syndrome Coronavirus 2 (SARS-CoV-2) Stephen A. Goldstein • Brenda G. Hogue • Julian L. Leibowitz • Susan R. Weiss History of human coronaviruses Evolution of SARS-CoV-2–related viruses Phylogenetic structure of SARS-related coronaviruses (SARSr-CoV) Origin and evolution of SARS-CoV Evolutionary history of SARS-CoV-2 Evolution of ACE2 usage by SARS-related coronaviruses (SARSr-CoVs) Recombination drives diversification of SARS-CoV-2 lineage viruses SARS-CoV-2 replication and structural proteins (Fig. 21.5) Spike (S) protein Membrane (M) protein Envelope (E) protein Nucleocapsid (N) protein SARS-CoV-2 assembly and release Nonstructural proteins (Table 21.1) Nsp1 Nsp2 Nsp3 Nsp4 and Nsp6 Nsp5 The replicase complex: Nsp7-Nsp16 SARS-CoV-2 accessory proteins ORF3a ORF3b ORF6 ORF7a ORF7b ORF8 ORF9b Recombinant viruses SARS-CoV-2 variants—transmission and immune evasion SARS-CoV-2: secondary transmission to wild and domestic animals

HISTORY OF HUMAN CORONAVIRUSES While the first highly lethal human coronavirus (HCoV), severe acute respiratory syndrome coronavirus (SARS-CoV), was identified in late 2002, human respiratory coronaviruses were isolated more than 50 years before that. HCoV-229E and related strains were isolated in human embryonic intestine cell cultures. HCoV-OC43 and related viruses were isolated human tracheal from organ cultures and later adapted to grow in suckling mice. Early literature suggested that HCoVes may be associated with enteric disease104,172 and also multiple sclerosis,104,260 but neither has been confirmed. 795

From that time forward, there was extensive research performed on the basic mechanisms of coronavirus replication, including cell entry, cell to cell fusion, mRNA synthesis, protein expression, and host–virus interactions including innate and adaptive immune responses. Most of these studies were performed on the model murine coronavirus mouse hepatitis virus (MHV), the avian infectious bronchitis virus (IBV), and bovine coronavirus (BCV). All of these systems were more amenable to molecular biology studies than the human viruses since these were more difficult to work with in cell culture and there were no useful animal models for the human viruses. In late 2002, severe acute respiratory syndrome (SARS)-CoV emerged in southern China, causing an epidemic, shocking coronavirologists to the reality that members of this virus family could be etiologic agents of highly lethal respiratory disease in humans and furthermore making coronaviruses known to the general public (Fig. 21.1A). SARS-CoV disappeared after 6 or 7 months, but during that time caused 8,069 infections and 774 deaths. In the years after the SARS epidemic, it was discovered that bats were the hosts for many SARS-like coronaviruses197,210 and that SARS-CoV was likely transmitted to humans through an intermediate host, the palm civet.

Following the SARS epidemic, two other HCoVes were identified, NL63299 and HKU1,198 causing respiratory symptoms (Fig. 21.1A). Approximately 10 years later in 2012, the deadlier Middle East respiratory syndrome (MERS)-CoV emerged in Saudi Arabia. MERS-CoV was found also to have its origin in bats, and to have a reservoir in the intermediate animal, the camel, and continues to spread from camel to human (Fig. 21.1A). While less transmissible among humans, MERS-CoV has a higher mortality rate and continues to cause infections in humans (https://Orf.who.int/emergencies/disease-outbreak-news/item/2021-DON317). In late 2019, a SARS-CoV–like virus emerged in Wuhan China, far from the site of the first SARS-CoV infections (Fig. 21.1A). SARS-CoV-2, the cause of coronavirus disease 2019 (COVID-19), causes severe respiratory disease with a lower fatality rate than SARS-CoV, even though it spreads more quickly, sometimes from asymptomatic individuals around the world. Since its emergence, SARS-CoV-2 has caused the largest and most serious global pandemic since the 1918 influenza. It is still not completely understood why this virus, which is similar to SARS-CoV, was able to spread more readily to cause the global pandemic.

Coronaviruses are divided into four genera designated as alpha, beta, gamma, and delta. All genera have similar genome organizations (structural proteins and nonstructural proteins including replicase proteins) but differ in their nonessential accessory proteins that are often involved in evading or antagonizing host cell responses. The less pathogenic, HCoVes are grouped in the Alphacoronavirus genus (229E, NL63, and related viruses) or the Betacoronavirus genus (Fig. 21.1B). The latter genus is further divided into subgroups and lineages based on the phylogenetic relatedness. These include the nonhighly pathogenic embeco or lineage a (OC43, HKU-1) and the highly pathogenic sarbeco or lineage b (SARS-CoV, SARS-CoV-2 and related bat CoVs) and merbeco or lineage c (MERS-CoV and related bat CoVs) (Fig. 21.1B).

EVOLUTION OF SARS-COV-2–RELATED VIRUSES The discovery of SARS-CoV-2 and ensuing pandemic439 has focused attention on the evolution and origins of this viral lineage. SARS-CoV-2 and the 2002–2003 epidemic SARS-CoV fall within the SARSr(elated)-CoV species in the Sarbecovirus subgenus of Betacoronavirus62 (Fig. 21.1B). This group of SARSr-CoVs was unknown prior to the SARS-CoV epidemic but is now known to comprise at least several hundred viruses143,196 present not only throughout China but also in southeast Asia,383 Japan,259 South Korea,201 Europe,81 and sub-Saharan Africa.291,396 SARSrCoVs are evidently not narrowly restricted from a geographic standpoint, though the degree of public health concern emanating from different viral reservoirs remains unclear. Both 21st century spillovers of SARSr-CoVs have occurred in mainland China.

Phylogenetic Structure of SARS-Related Coronaviruses (SARSr-CoV) In the period since the emergence of SARS-CoV (detailed below), dozens of related full-length genomes and additional partial genomes have been described, largely through metagenomic analyses. SARSr-CoV sequences have been identified primarily in East Asia spanning Japan, South Korea, China, Thailand, and Cambodia. Based on geographic distribution of the reservoir species, primarily Rhinolophus bats, such viruses surely circulate elsewhere in southeast Asia. In addition, distantly related SARSr-CoVs have been identified in Europe and Africa. The wide distribution suggests an ancient origin of this group of viruses. A 2021 study by Wells et al.396 conducted a phylogenetic analysis of the RNA-dependent RNA polymerase (RdRp)-encoding region of ORF1ab (described in detail below) and identified five distinct lineages within SARSr-CoV (Fig. 21.2). Four of these are distributed throughout East Asia while the fifth comprises the African and European SARSr-CoVs. Notably, three of the Asian lineages fall onto sub-branches within a single branch, including SARS-CoV and related viruses. The SARS-CoV-2– like viruses are on a distant branch and exhibit greater than 10% genetic distance in RdRp from SARS-CoV–like viruses; they are comparably distant as the European and African viruses and the relative phylogenetic positions of the Africa/Europe and SARS-CoV-2–like viruses remain somewhat uncertain. This suggests that the split between the SARS-CoV and SARS-CoV-2 branches was quite ancient despite their common receptor [angiotensin converting enzyme −2 (ACE2)] usage, possibly approaching 1,000 years ago.396 SARS-CoV and SARS-CoV-2 are much more closely related to different non–ACE2-utilizing viruses than they are to each other, suggesting a complex evolutionary history of ACE2binding spike proteins. Interestingly NL63, an alphacoronavirus also uses ACE2 as its receptor251 indicating receptor utilization is not always useful for determining relatedness of viruses.

FIGURE 21.1 A: Timeline of detection of human coronaviruses. Common cold human coronaviruses HCoV-OC43 and HCoV-229E have 796

been known since the 1960s; HCoV-NL63 and HCoV-HKU-1 were identified in 2004–2005. The first highly pathogenic human SARS-CoV coronavirus was identified in 2002, MERS-CoV in 2012, and SARS-CoV-2 in 2019. B: Human coronavirus genome structure. Human coronaviruses are found in the alphavirus genus and in three lineages of betacoronaviruses, embeco (lineage a), sarbeco (lineage b), and merbeco (lineage c). The genomes have 5′ capped ends and leader (L) sequences. The intergenic short light brown bars are transcription regulatory sequence, which mark the initiation site of transcription of each subgenomic mRNA. All coronaviruses encode 16 nonstructural proteins (nsps) in 5′ open reading frames 1a and 1b. Genes encoding structural proteins are arrayed in the 3′ portion of the genome in the order spike (S), small membrane (E), membrane (M), and nucleocapsid (N). Lineage specific accessory genes are found interspersed among the structural genes (dark brown bars). ORF8 of SARS-CoV evolved into ORF8a and ORFb but remains as an intact ORF8 in SARS-CoV-2 (as shown in the figure). (Created by Alejandra Fausto using Biorender.com.)

FIGURE 21.2 RdRp phylogenetic tree. Maximum-likelihood phylogenetic tree of the RdRp-encoding region of ORF1ab. This tree demonstrated the segregation of SARSrelated(r)-CoV into 5 RdRp lineages with SARS-CoV and SARS-CoV-2 each defining two widely divergent lineages. Viruses shown able to use human ACE-2 are bolded. To construct the tree, representative SARSr-CoV sequences were obtained from GenBank or GISAID and aligned to the RdRp-encoding region of SARS-CoV-2 using MAFFT. A tree was then inferred using FastTree.

ORIGIN AND EVOLUTION OF SARS-COV Though the definitive origin of SARS-CoV-2 remains unknown, the historical example of SARS-CoV provides valuable context and guidance given the likely similar ecological niche occupied by the two viruses. The SARS-CoV epidemic began as a cluster of unusual pneumonia cases in spring 2003 in Hong Kong200,369 and retrospectively was determined to have begun the preceding fall in Guangdong in mainland China.434 Via both electron microscopy and genomic analyses, a coronavirus was identified as the etiological agent of these outbreaks.82,285 Ultimately, greater than 8,000 cases and nearly 800 deaths due to SARS-CoV were recorded globally (https://Orf.who.int/publications/m/item/summaryof-probable-sars-cases-with-onset-of-illness-from-1-november-2002-to-31-july-2003).

Field investigators quickly determined that small mammals in live animal markets but not on rural farms were infected, suggesting they were an intermediate rather than reservoir host.123 By 2005, viruses related to SARS-CoV had been identified in Rhinolophus (horseshoe) bats in mainland China,210 establishing them as the reservoir host for these viruses. Although numerous viruses were quickly identified with greater than 90% nucleotide identity to SARS-CoV, only in 2013 was a bat virus, BtCoV/WIV1, isolated that is able to infect human cells via the SARSCoV receptor ACE2.102,209 Over the next several years, additional such viruses were isolated and shown able to replicate in primary human airway epithelial (HAE) cells.244,246,418,427,437 Most recently, dozens of novel members of the species SARSr-CoV have been identified in 797

Chinese horseshoe bats196 though many are characterized based only on partial sequences of the ORF1ab region encoding the RdRp. Although SARSr-CoVs have been found throughout China, the closest related viruses to SARS-CoV have been identified in the southern Chinese province of Yunnan.

The discovery and genome analysis of diverse SARSr-CoVs in bats have revealed a critical role for recombination in the evolution of SARS-CoV. Despite the isolation of BtCoV/WIV1 in 2013, several genomic mysteries remained regarding the evolutionary history of SARS-CoV. Although BtCoV/WIV1 infects human cells via ACE2, the receptor binding domain (RBD) of its spike protein is distinct from SARS-CoV,102,418 and the ORF3b and ORF8 accessory proteins (putative functions described below) of SARS-CoV are highly divergent from the orthologs encoded by its otherwise closest relatives, indicative of recombination. The ORF8 of SARS-CoV famously experienced a 29 nucleotide deletion, splitting it into two open-reading frames, early in the human epidemic.261 A comprehensive 2017 report from a bat sampling study conducted by the Wuhan Institute of Virology reported bat SARSr-CoVs with nearly identical ORF8 genes to SARS-CoV, whereas other viruses such as BtCoV/WIV1 were most closely related to SARS-CoV in other regions of the genome.143 The same study identified the first close relative of SARS-CoV ORF3b in a bat SARSr-CoV in a different genome from the SARS-CoV–like ORF8, indicating multiple recombination events produced the immediate progenitor virus to SARS-CoV, which has nevertheless not been identified in a natural reservoir.

Evolutionary History of SARS-CoV-2 Despite intense sampling of bat populations for SARSr-CoVs between 2003 and 2019, the SARS-CoV-2 lineage remained largely undiscovered prior to early 2020. The genome of SARS-CoV-2 was first published on Virological.org on January 10, 2020 and characterized in greater detail the following month.440 This study also provided the first insight into the evolutionary origins of the pandemic virus, describing a virus, BtCoV/RaTG13, with approximately 96% nucleotide identity to SARS-CoV-2, a comparable similarity to SARS-CoV and its closest known relative. A partial RdRp sequence of BtCoV/RaTG13 was previously reported in 2016 as a possibly new lineage within SARSr-CoV.103 The sample containing this viral sequence was collected from a Rhinolophus affinis bat in an abandoned mineshaft in Yunnan province in southern China, the same region of China from which the closest relatives of SARS-CoV have been identified, though farther south. These viruses exhibit only 79% to 80% nucleotide identity to SARS-CoV across the entire genome and qualify as a novel lineage of SARSr-CoV.

Since the initial discovery of SARS-CoV-2, several other viral genome sequences in this lineage have been identified in previously collected samples (Fig. 21.2). Shortly after the full genomic characterization of SARS-CoV-2 in February 2020, related viral sequences were identified in pangolins seized in antitrafficking operations in the southern Chinese provinces of Guangxi and Guangdong. In particular, a sequence from a pangolin seized in Guangdong province yielded a complete genome sequence exhibiting high (>90%) identity to SARS-CoV-2 and an even higher identity to SARS-CoV-2 in the spike RBD than BtCoV/RaTG13, with all six critical residues for ACE2 binding conserved.192,384,408 Despite the apparent association of SARS-CoV-2–like viruses with pangolins, whether they are a reservoir host or incidentally infected during wildlife trafficking remains unclear. A 2009–2019 study in Malaysia sampled greater than 300 pangolins entering the wildlife trade and found no evidence of coronavirus infection.202 In contrast, pangolins with evidence of coronavirus infection have been seized “downstream” in trafficking networks in Thailand and southern China and in some cases shown evidence of severe lung disease,408 suggesting that at least under stress conditions pangolins are susceptible to disease caused by these viruses.

Discoveries of additional related viruses reported later in 2020 and early in 2021 have further populated the SARS-CoV-2–like branch of the Sarbecovirus phylogenetic tree (Fig. 21.2). Whereas most viruses on the SARS-CoV branch have been identified in Rhinolophus sinicus, the SARS-CoV-2 branch exhibits an apparently more diverse host range. BtCoV/RmYN02 was sequenced from a R. malayanus sample taken in southern Yunnan and exhibits even higher nucleotide identity to SARS-CoV-2 in ORF1ab than does BtCoV/RaTG13,440 with recombination apparently accounting for lower genome-wide identity (Fig. 21.3). Based on the relatedness of SARS-CoV-2 with closely related viruses in ORF1ab, it is estimated to have had a common ancestor with BtCoV/RaTG13 approximately 52 years ago and with BtCoV/RmYN02 approximately 37 years ago,389 though the confidence intervals for these estimates overlap, producing considerable uncertainty as to the exact evolutionary relationships. In March 2021, a new virus sequence was reported from Yunnan province with high identity to SARS-CoV-2. BtCoV/RpYN06 exhibits 94.5% genome-scale nucleotide identity to SARS-CoV-2 and 97.2% identity in the entirety of ORF1ab, making it equivalently close to SARS-CoV-2 as BtCoV/RmYN02 (Fig. 21.3).441 Although all of these viruses are too distantly related to SARS-CoV-2 to have been the progenitor of the pandemic virus, the discovery of these viruses within a limited geographic area raises the possibility that the SARS-CoV-2 progenitor naturally circulates in southern Yunnan province. However, sampling bias and the recently expanded known range of SARS-CoV-2–related virus lends considerable uncertainty to this hypothesis.

FIGURE 21.3 Average nucleotide identity analysis of SARS-CoV-2–related virus genomes using SARS-CoV-2 as the reference genome. Sliding window analysis showing nucleotide identity (ANI) of related viruses to SARS-CoV-2 generated in IDPlot. Large decreases in ANI are often indicative of genomic recombination with divergent SARSr-CoVs. Putative recombination hotspots at approximately 22 kb and 28 kb correspond to the spike and ORF8 genes, respectively. Numbers on the y axis correspond to nucleotides of the coronavirus genome.

798

FIGURE 21.4 Receptor binding motif (RBM) alignment. Multiple sequence alignment of SARS-CoV-2–like virus (plus SARS-CoV) receptor binding motifs within the receptor binding domain. Six critical residues for interaction with ACE-2 are indicated by red arrows. RmYN02, which is not predicted to use ACE-2, contains deletions, one of which removes two of the critical residues. All 6 residues are conserved in BANAL-20-52 and PangolinCoV/GD19 and 5 in RShSTT200. The lack of conservation with SARS-CoV demonstrates the tolerance of ACE-2 binding to mutations in this region. While most interest on the origins of SARS-CoV-2 has focused on Yunnan province, the early 2021 report of a closely related virus in Thai Rhinopholus bats383 expands the known geographic range of this viral lineage. BtCoV/RacCS203 exhibits similar identity to SARS-CoV-2 as BtCoV/RmYN02 and is the most closely related virus to the latter. Another southeast Asian SARS-CoV-2–like virus was identified retrospectively from Rhinolophus shameli samples collected in 2010 in Cambodia. This virus exhibits 92.6% genome-wide identity to SARSCoV-2 and contains a SARS-CoV-2–like RBD, including conservation of five out of six critical ACE2 contact residues (Fig. 21.4).148 Continued efforts to identify the origin of SARS-CoV-2 are likely to include wildlife sampling in southern China and in southeast Asia, which has seen extremely limited virus surveillance previously. At the time of this writing, the progenitor of SARS-CoV-2 has not been discovered, but the best estimates are that the virus emerged from a genetic pool of viruses circulating widely across southern China and southeast Asia. To this point, recently a group of viruses were isolated in Laos with very high identity to SARS-CoV-2; BANAL-52 has 96.8 % genome wide identity and conservation of all six critical ACE2 binding residues (Figs. 21.3 and 21.4).361

Evolution of ACE2 Usage by SARS-Related Coronaviruses (SARSr-CoVs) The emergence in humans of SARS-CoV and SARS-CoV-2 owes principally to their ability to utilize the human cell surface protease ACE2 as a receptor for cell attachment and entry. ACE2 binding is mediated by the RBD (Fig. 21.4), located in the C-terminal portion of the spike protein S1 subunit. ACE2 usage is far from ubiquitous among SARSr-CoVs. A 2020 study by Letko et al. that demonstrated ACE2 utilization by the SARS-CoV-2 spike207 segregated SARSr-CoV spike proteins into three groups based on ACE2 usage. Non–ACE2-utilizing viruses, in comparison, contain one or two deletions that encompass some of the ACE2 contact residues207,305 and the receptor used by these viruses for entry is currently unknown.

The phylogenetic distance between SARS-CoV-2 and SARS-CoV is in contrast to the similarity of their RBDs and use of a common receptor. Wells et al. found that although SARS-CoV was discovered first and this lineage is best represented in currently sampled diverse viruses, ACE2 binding is ancestral in the SARS-CoV-2 lineage and introgressed via recombination into the SARS-CoV lineage where it is widespread but not ubiquitous. Similarly, in the SARS-CoV-2 lineage, the ancestral RBD has been lost by multiple viruses via recombination.23

Notably with respect to the potential for human transmission, the binding affinity of numerous bat SARSr-CoV spikes is higher for human than R. sinicus ACE2,125 reflecting rapid evolution of the gene due to selective pressure imposed by SARSr-CoVs. Further evidence supports a generalist evolutionary trajectory of ACE2 binding SARSr-CoV spikes, calling into question the need for adaptation of the RBD to facilitate cross-species transmission. BtCoV/WIV1 readily mediates infectivity in a range of mammalian species.437 Additionally, a 2021 study by MacLean et al. found evidence for powerful diversifying selection in the SARS-CoV-2 lineage RBDs, suggesting that rather than converging on an ACE2 binding solution highly specific to particular hosts, evolutionary pressures diversify their sequences resulting in generalist viruses.232

Indeed, analysis of the ACE2-interaction region of the RBD of these viruses (Fig. 21.4) suggests high tolerability to amino acid diversity of ACE2 binding. Six residues in the SARS-CoV spike, Y442, L472, N479, D480, T487, and Y491, were previously identified as critical for binding to ACE211,387 corresponding to amino acids L455, F486, Q493, S494, N501, and Y505 in SARS-CoV-2; only the final tyrosine is conserved. Despite this sequence variation, affinity for human ACE2 is sufficiently high for both spike proteins to effectively mediate infection. In the SARSCoV-2 lineage, only Y505 is conserved in BtCoV/RaTG13, whereas in PangolinCoV/GD19, all six residues are conserved (Fig. 21.4), confirming that the SARS-CoV-2 RBD is highly conserved with that of viruses circulating in wildlife. It has recently been discovered that viruses with RBD containing all six critical human ACE2 binding residues are also found in bats, as illustrated by the recently identified BtCoV BANAL-50 isolates (Fig. 21.4).361 Other related viruses, such as BtCoV/RmYN02, contain deletions that eliminate some of these residues and presumably must use an alternative receptor. Their potential for human infection is unknown.

Recombination Drives Diversification of SARS-CoV-2 Lineage Viruses Genomic recombination is a common feature in coronavirus evolution and clear signatures are evident among SARSr-CoVs. Given the apparent recombinant nature of SARS-CoV,143,199 immediate interest arose in whether SARS-CoV-2 similarly arose via recombination in a reservoir or intermediate host. Recombination is most frequent in the 3′ end of the genome121 and particularly in the spike gene,107,116 which may produce differences in tropism, receptor usage, and host range. This is easily observed on a large scale in the incongruence between RdRp and spike phylogenies396 as well as in large changes in average nucleotide identity within the RdRp-defined SARS-CoV-2 lineage of SARSr-CoV (Fig. 21.2).

Although a flurry of preliminary studies suggested a recent recombinant origin of SARS-CoV-2, studies by Boni et al.23 and Nielsen et al.389 found no evidence supporting this hypothesis. The consistently high genome-wide nucleotide identity with RaTG13 (Fig. 21.3) suggests little 799

change in genome composition since the last common ancestor of these viruses but does not preclude recombination in deeper evolutionary history. However, evidence of recombination is striking in other viruses on the SARS-CoV-2 branch of SARSr-CoV, including in RpYN06 and RmYN02, which are more closely related to SARS-CoV-2 than RaTG13 in large regions of their genome (Fig. 21.3). The considerable genomic distance between these viruses and SARS-CoV-2 in the spike gene (and ORF8 in the case of RmYN02) is clear evidence that SARS-CoV-2–like viruses circulate alongside and coinfect host animals with more distantly related SARSr-CoVs. Therefore, viruses outside the SARS-CoV-2 branch may contribute genetic material to viruses within it, expanding the genetic diversity with the potential to contribute to future pandemics.

SARS-COV-2 REPLICATION AND STRUCTURAL PROTEINS (FIG. 21.5) SARS-CoV-2, like other coronaviruses, has a positive-sense, single-stranded RNA genome consisting of almost 30,000 nucleotides.109,112,227 The genome is capped and also has a leader sequence at the 5′ end of the RNA (Fig. 21.1B). The original SARS-CoV-2 isolates exhibit 79.6% identity to the SARS-CoV genome at the nucleotide level.404,439 Approximately two-thirds of the 5′ end of the genome contains open reading frames (ORFs) 1a and 1b that encode primarily proteins involved in replication. The 3′ one-third of the genome encodes the structural proteins and eight accessory genes (Fig. 21.1B).404,439 SARS-CoV-2 virions have four structural proteins, the spike (S), envelope (E), membrane (M) anchored in the viral envelope, and the nucleocapsid (N), which encapsidates the single-stranded RNA genome (Fig. 21.6). Spike trimers prominently extend from the envelope and the helical nucleocapsid consisting of the viral genome and N protein form a dense core when viewed by EM (Fig. 21.6B and C). Coronavirus entry into cells is receptor mediated through interactions of the S protein with host cell ACE2. Virus replication and assembly takes place in the cytoplasm (Fig. 21.5).

FIGURE 21.5 Model of coronavirus life cycle. 1. Virus enters the cell by both direct fusion of the viral membrane with the cellular plasma membrane or is endocytosed and enters the cell by fusion between the viral and endosomal membranes, both processes mediated by spike that has been activated by proteolytic cleavage. 2. Viral genome RNA is uncoated. 3. Genome RNA is translated from ORFs1a and 1b into polyproteins pp1a and pp1ab, which are processed into replicase proteins. 4. The replicase complex is assembled on double membrane vesicles (DMV) at the endoplasmic reticulum (ER). 5. Genome RNA is replicated through a negative strand intermediate and subgenomic negative strand RNAs are transcribed from genome and then copied into subgenomic mRNAs. 6. Subgenomic (sg) mRNAs are translated into viral structural and accessory proteins. 7. Viral membrane proteins spike (S), membrane (M), and envelope (E) are inserted into ER membranes. 8. Viral nucleocapsid protein (N) complexes with genome RNA to form nucleocapsids. 9. Nucleocapsids bud into the ER–Golgi intermediate compartment (ERGIC) in regions containing viral membrane proteins to form progeny virions. 10. Spike protein on virions is cleaved by furin in the Golgi. 11. Virus is transported to the cell surface in exocytic vesicles and/or through the lysosomal pathway. 12. Progeny virus is released into the extracellular space. (Created by Alejandra Fausto using Biorender.com.)

Spike (S) Protein Spike (S) proteins extend from the viral envelope as trimers to give coronaviruses their characteristic crown-like appearance. The proteins play important roles in infectivity, host range, and pathogenesis.208,287 SARS-CoV-2 S protein is the host receptor binding protein that, like SARSCoV and HCoV-NL63 S proteins, mediates entry into host cells by binding to hACE2.137,207,386 The S protein exhibits significant structural and sequence homology with SARS-CoV S. Initial SARS-CoV-2 S isolates shared approximately 76% amino acid identity with SARS-CoV S.227,404 Like other coronaviruses, SARS-CoV-2 S proteins are the major target for neutralizing antibodies, therapeutic monoclonal antibodies, and vaccine development.

SARS-CoV-2 S exhibits the same overall structural features as other coronavirus S proteins. The proteins are type I transmembrane proteins that form homotrimers. S protein monomers consist of two subunits, the N-terminal S1 extracellular amino end head region and the S2 carboxy end stalk, transmembrane domain (TMD) and short viral endo-domain tail (Fig. 21.7). The S1 subunit consists of the N-terminal domain (NTD) and the C-terminal domain (CTD), which contains the RBD.137,207,386,402,439 The S2 subunit contains two heptad-repeat (HR) domains and a hydrophobic fusion peptide that mediates membrane fusion.207,208,367,386,402 The proteins are class I fusion glycoproteins.208 SARS-CoV-2 S protein is highly glycosylated at 66 N-linked sites on each trimeric S, which shields the RBD in its prefusion conformation.394,435

Coronavirus S proteins are cleaved sequentially by host cellular proteases at the S1/S2 junction and S2′ sites (Fig. 21.7) to promote viral virus– host membrane fusion and entry,134 although cleavage at the S1/S2 junction may not be required for activation of all coronavirus S proteins, for example in alphacoronaviruses.398 The S2′ cleavage site is located in the S2 subunit between the amino end helix region and the fusion peptide (Fig. 21.7). SARS-CoV-2 S differs from SARS-CoV S in that it has a multibasic furin cleavage site (R682RAR↓S686) at the S1/S2 junction.136,386,402 MERS-CoV S is also cleaved by furin in producer cells.136,325 Less pathogenic hCoV-HKU1 and hCoV-OC43, also have furin cleavage sites at the S1/S2 junction.63,252 Furin is a Golgi-associated protease that cleaves the S1/S2 junction during virus assembly or 800

transport through the secretory pathway.252 The SARS-CoV-2 S1/S2 multibasic cleavage site with the additional arginine residues is presented in an exposed loop.386,402

FIGURE 21.6 SARS-CoV-2 virions contain four structural proteins, spike (S), envelope (E), membrane (M), and nucleocapsid (N). A: Schematic of the virion shows the S trimers, M and E proteins anchored in the viral lipid envelope. The positivesense, single-strand RNA genome (ssRNA) encapsidated by the N protein is packaged inside the particle. B: SARS-CoV-2 transmission electron micrograph of virus particles that is color-enhanced (NIAID). C: EM tomographic slice of a SARS-CoV-2 virion showing granular density of the ssRNA encapsidated by the N protein and S trimers extending from the viral envelope. (Panel A adapted by permission from Nature: V'kovski P, Kratzel A, Steiner S, et al. Coronavirus biology and replication: implications for SARS-CoV-2. Nat Rev Microbiol 2021;19(3):155–170. Copyright © 2020 Springer Nature, Ref.379. Panel C adapted by permission from Nature: Ke Z, Oton J, Qu K, et al. Structures and distributions of SARS-CoV-2 spike proteins on intact virions. Nature 2020;588(7838):498–502. Copyright © 2020 Springer Nature.)

Cleavage at the S2' site primes the protein by revealing the fusion peptide in the S2 domain that is necessary for fusion of viral and cellular membranes during entry. SARS-CoV S2′ site is cleaved by either cell surface protease TMPRSS2, a member of the type II transmembrane serine protease 2 family of enzymes, or endosomal cysteine protease cathepsin B or L.106,239,336 SARS-CoV-2 S is primed by TMPRSS2 cleavage at S2′ and the TMPRSS2 inhibitor camostat mesylate blocks infection in Calu-3 and primary human lung cells.137 Cleavage at the S2′ site is necessary for fusion and entry into human lung cells.136 SARS-CoV-2 S, like MERS-CoV, requires cleavage by furin at the S1/S2 site for downstream activation by membrane-associated transmembrane serine protease 2 (TMPRSS2) cleavage at S2′ in lung cells.22,136

FIGURE 21.7 Summary of the functions and domain organization of full-length (1,273 amino acids) SARS-CoV-2 spike (S) protein. Amino acid sequence alignments of the S1/S2 and S2′ cleavage sites of human alphacoronavirus (HCoV-229E, HCoVNL63), betacoronavirus embeco (a, HCoV-OC43, HCoV-HKU1), sarbeco (b, SARS-CoV-2, SARS-CoV) and merbeco (c, MERS-CoV) lineages, and bat RaTG13 and pangolin MP789 are shown. Basic residues are highlighted in red. Multibasic S1/S2 cleavage sites are highlighted with gray boxes. SS, signal sequence; NTD, N-terminal domain; RBD, receptor binding domain; CTD; carboxy-terminal domains 1 and 2; S1/S2, furin cleavage site; S2′, protease cleavage site; FP, fusion peptide; HR, heptad repeats 1 and 2; CH, central helix; CD, connector domain; TMD, transmembrane domain; CT, cytoplasmic tail (intravirion domain). Aside from the furin cleavage site at the S1/S2 junction that is not present in SARS-CoV S, SARS-CoV-2 S protein exhibits other distinctive characteristics compared with that other sarbecoviruses.136,325 The S proteins have NTD loops that are divergent from other sarbecoviruses.386 The S1 RBD is structurally divergent and has been reported to bind hACE2 receptor with higher affinity than SARS-CoV S RBD.386,402 The RBD consists of a core structure and a receptor-binding motif (RBM) located in an extended loop, flanked by alpha-helices193,326

Coronavirus S proteins are very dynamic structures during entry and infection. Prefusion S exists in a metastable state, which transitions between conformations with all three RBDs in the down conformation where the RBM is not in a position to bind to its cognate ACE2 receptor, to conformations in which one or more of the trimers adopt a conformation with the RBD in up position with their RBM positioned to engage with ACE2.402 S undergoes changes during entry and infection as the viral envelope membrane fuses with host cell membranes.208 In the postfusion state, the trimers have all three RBDs in the up conformation.34,90,181,386 S1 binding to the receptor results in conformational changes that trigger S2 extension to reveal the hydrophobic fusion peptide that promotes virus and cell membrane fusion during entry.208,238 The protein assumes a postfusion stabilization of helical bundles as the protein refolds.34,386

Most coronavirus S proteins, including SARS-CoV-2, transport to some extent to the cell surface in infected cells. This promotes cell-to-cell fusion that results in multinucleated syncytia that provides a mechanism for virus spread.370,442 This is seen in infected Calu-3 or A549 cells derived from the respiratory tract.136,262

Coronaviruses can enter cells by two different routes (Fig. 21.5).157,338 Following binding of virus to a cellular receptor, virion can be endocytosed or alternatively, the viral envelope fuses directly with the host cell surface membrane. Previous studies with other coronavirus that use similar entry mechanism provided significant insight that helped inform how SARS-CoV-2 enters cells. In the direct plasma membrane 801

route, the SARS-CoV-2 S protein, as well as MERS-CoV S, has already been cleaved at the S1/S2 site by furin during synthesis in infected host producer cells.136,325 Following attachment to the receptor, S cleavage at the S2′ site by membrane-associated TMPRSS2 initiates fusion between the viral and host cell membranes. Cleavage allows the fusion peptide to insert into the host cell membrane to promote fusion between the viral and cell membrane. Fusion between the membranes results in release of the viral nucleocapsid (viral genome coated by nucleocapsid (N) protein) into the cytoplasm to initiate replication.

In the case of entry by the endosomal route, virus attaches to its receptor and is then endocytosed. The low pH environment of the endosome activates cleavage by the protease cathepsin L, resulting in fusion of viral and endosomal membranes and release of the nucleocapsid into the cytoplasm to initiate infection.21 The pathway(s) used depends on the proteases available, which can vary among cell types and the sequence of the S protein.252,338 Some cell types may favor one pathway or the other. In lung-derived cells, SARS-CoV-2 entry is dependent on both furin for S1/S2 junction cleavage and TMRPSS2 for the second S2′ cleavage.22 It is possible that in other cell types different proteases and one or the other pathways is primarily utilized or alternatively both pathways can be used by the same virus in a particular cell type.

Spike cleavage is important for entry but can also play a role in virus spread. Following synthesis at endoplasmic reticulum membranes, coronavirus S proteins traffics through the exocytic pathway and for those with a furin cleavage site, such as SARS-CoV-2, cleavage occurs in the Golgi. In the endoplasmic reticulum Golgi intermediate compartment (ERGIC)/Golgi region, some S molecules are retained through interaction with the membrane (M) protein where virions are assembled (see additional details in the assembly section). The S protein that is not retained is transported to the plasma membrane where it can mediate fusion with receptor bearing neighboring cells, thus allowing cell to cell spread of virus shielded from the extracellular immune responses. This promotes cell-to-cell fusion that results in multinucleated syncytia that provides a mechanism for virus spread.370,442 This is seen in infected Calu-3 or A549 cells derived from the respiratory tract.136,213,262 However, it is not known to what extent, if at all, this promotes viral spread in vivo. While no obvious cell to cell fusion was observed in induced pluripotent stem cell (iPSC)-derived AT2 cells, robust cell fusion was observed in iPSC-derived cardiomyocytes.213 The dynamics of this spread are determined at least in part by the levels of ACE2, TMPRSS2, and furin in each cell line.

Significant progress toward understanding the structural biology of SARS-CoV-2 S was made quickly after the virus emerged. Multiple cryo-EM structures have been determined using X-ray crystallography or cryo-electron microscopy (cryo-EM) for SARS-CoV-2 S protein in prefusion and postfusion conformations. SARS-CoV-2 S bound to ACE2 has been determined at the atomic level.193,243,326,386,410 The ectodomain of SARSCoV-2 S was determined by cryo-EM at 3.5-angstrom resolution of a prefusion trimer.402 The crystal structure of SARS-CoV-2 RBD complexed with ACE2 shows that the binding ridge is more compact than the SARS-CoV binding ridge and that residue differences stabilize two binding hotspots compared to SARS-CoV RBD, which results in higher receptor binding affinity than previously measured for SARS-CoV RBD.326 S trimers exists in both down and up conformations on virions when examined by cryo-EM tomography.174,218,371,419 Trimers were observed to tilt relative to the viral membrane due to flexibility of the membrane-proximal stalk region that functions like a hinge.174

Coronaviruses evolution is slowed as a result of 3′-5′ exonuclease proofreading activity of nonstructural protein 14 (nsp14). Nonetheless, mutations in SARS-CoV-2 S have emerged during the COVID-19 pandemic, referred to as “variants of concern.” Changes in the spike protein are particularly important since these can affect both receptor and antibody binding, resulting in changes in infectivity, transmission, antibody binding, and vaccine escape. The strongest spike neutralizing epitopes are located in the RBD. SARS-CoV-2 RBD has three nonoverlapping antigenic sites.365 A major change during evolution of S since the emergence of COVID-19 is a D614G (Asp614 to Gly) change in S1. Multiple studies suggest that the change increases infectivity and transmission.54,187,429,432

Membrane (M) Protein Coronavirus membrane (M) proteins play a major role in assembly of virus particles. The proteins are highly conserved in their overall architecture. SARS-CoV-2 M consists of 222 amino acids with a theoretical molecular weight of approximately 25 kDa, which aligns with the overall characteristics of other coronavirus M proteins. The protein is 91% identical to SARS-CoV M protein at the amino acid level. M proteins are type III membrane proteins, consisting of three TMDs, a short N-terminal ectodomain and a long, approximately 100 amino acid C-terminal intravirion tail.140,310 Coronavirus M proteins are glycosylated on the NTD with either N-linked or O-linked glycans.140,264,286 SARS-CoV-2 M has a predicted N-glycosylation site in the N-terminal ectodomain. Prior to incorporation into assembled virions, the C-terminal tail is located in the cytoplasm and where it plays a role in trafficking to and localization in the Golgi.69,214,223,264,288 The tail associates with the inner leaflet of the plasma membrane where it interacts with the viral nucleocapsid during budding.68 M proteins can homo-oligomerize and interact with other virion components, including S, N, and E.68,69,242 For all coronaviruses, virus-like particles (VLPs) can be efficiently assembled with protein M, S, and E.372,376 Recent studies indicate that SARS-CoV-2 M protein is also required for VLP formation and plays a role in retention of the S protein for incorporation into the particles.26,295

Envelope (E) Protein Coronavirus E proteins are small, 8.4 to 12 kDa (74 to 109 amino acids), multifunctional membrane proteins that play roles in virus assembly, egress, inflammation, and pathogenesis.140,318 The proteins localize at the site of assembly in the ERGIC.270,376 SARS-CoV-2 E consists of 75 amino acids that share 96% identity with SARS-CoV E protein but exhibit very low identity with other coronavirus E proteins. Nonetheless, SARS-CoV-2 E exhibits the same overall structure and key conserved amino acids characteristic of the proteins across the virus family. All E proteins consist of an NTD, a long alpha helical TMD and a C-terminal hydrophilic domain, including conservation of cysteine and proline residues in the C-terminus.140,311 Structural data and in vitro measurement have shown that E proteins multimerize to form a pentamer in the membrane bilayer and form ion channels that are known as viroporins.18,283,289 SARS-CoV-2 E, like SARS-CoV E, also assembles into pentameric ion channels in membranes.237,318 Structural models based on NMR studies in either micelles or liposomes have been produced for SARS-CoV and SARS-CoV-2 E proteins.237,353 Comparison of the models suggest that SARS-CoV-2 ion channels exhibit a more simple architecture, but it was suggested that based on the high sequence homology between the two E proteins that the structural difference may be due to experimental differences in the NMR studies.18 This indicates that understanding of the proteins will benefit from additional structural and functional studies that directly compare the E proteins from multiple coronaviruses. E proteins are posttranslationally modified by palmitoylation at conserved cysteine residues located on the C-terminal side of the third TMD, which affects its interactions with the membrane bilayer.25,225 The proteins are expressed at a high level in infected cells, but only a few molecules are assembled into virus particles.140,311

802

Of the four coronavirus structural proteins, the E viroporin protein and its functions are the least understood. Studies have shown that E proteins play diverse roles, including modulating virion assembly/membrane budding, trafficking, and release by exocytosis, as well as virion stability and pathogenesis.140,212,311,318,319 It is well established that E proteins colocalize and interact with M proteins during virus assembly.286,318 Mechanistically how E proteins contribute to virus assembly is still not fully understood since the requirement for E during the process varies among coronaviruses.72,278 The roles of E may be associated with the viroporin activity and/or contributions to membrane curvature or scission during assembly and budding.271,311,318

SARS-CoV-2 E protein and SARS-CoV E both contain PDZ-domain binding motifs (PBMs) in the C-terminus. The E PBMs interact with PALS1, a PDZ containing protein that is important for epithelial polarity in mammalian cells.44,162,362 The C-terminal PBMs have four residues (DLLV) that are conserved between SARS-CoV and SARS-CoV-2. C-terminal peptides containing the conserved residues show that the SARS-CoV-2 peptide exhibits enhanced binding to the PALS1 PDZ domain.368 A cryo-EM structure of PALS1 with SARS-CoV-2 E CTD containing the PBM shows that conserved residues DLLV bind a pocket formed by PALS1 PDZ and SH3 domains.44 A crystal structure of SARS-CoV and SARSCoV-2 E protein PBMs with PALS1 PDZ has also shown binding to a pocket in the latter domain.159 These structural studies suggest that the E protein C-terminal PMD interactions with PALS1 may play a role in pathogenesis by interference with lung epithelial cell junction integrity.73,362 E protein is expressed at high levels in SARS-CoV-2 infected lung epithelial cells.401 Previously, it was suggested that SARS-CoV E localizes at the site of virus assembly where recruitment of PALS1 may disrupt lung cell polarity and vascular structure.362 SARS and SARSCoV-2 E proteins also interact with other cellular junction proteins, including PDZ-containing syntenin adhesion junction protein, and tight junction ZO-1 protein.35,42,162 Thus, E protein interactions with these proteins and its viroporin activity likely contribute to significant inflammation, tight junction breakdown, vascular leakage, and immune-mediated damage that results in acute respiratory distress syndrome (ARDS). Indeed, a recent study suggests that SARS-CoV-2 E contributes to pathological damage that resembles ARDS.406

Nucleocapsid (N) Protein Coronavirus nucleocapsid (N) proteins are multifunctional phosphoproteins that encapsidate the viral RNA to form a ribonucleoprotein complex (vRNP) that mediates packaging into virus particles.188 N protein is recruited to replication–transcription complexes where it participates in RNA synthesis and is thought to promote RNA template switching to facilitate addition of the 5′ leader sequence during subgenomic mRNA synthesis.60,243,377,444 Full-length SARS-CoV-2 N protein consists of 419 amino acids that has a theoretical molecular weight of 45.6 kDa. SARS-CoV-2 N protein shares 90.52% identify at the amino acid level with SARS-CoV N. The protein exhibits the same overall modular architecture as other CoV N proteins with conserved, folded N-terminal (NTD) and C-terminal (CTD) domains. The NTD interacts with the viral RNA genome packaging signal and the CTD associates to form dimers that are thought to facilitate vRNP assembly.145,282,352,420 The ordered NTD and CTD are flanked by intrinsically disordered regions (IRDs) and separated by a conserved central IDR that contains a serine/arginine-rich (SR) region that is phosphorylated in infected cells. A high-resolution structure of SARS-CoV-2 N dimerization domain shows a compact structure similar to other betacoronaviruses, whereas extension to include the CTD results in homotetramer formation.420 SARS-CoV N crystal structures previously showed a helical assembly of dimer domains that was proposed to form the basis for helical nucleocapsid filaments in virions.49 SARS-CoV-2 N dimer–dimer packing modes, revealed in the crystal structures, shows that the N protein would not assemble into helical filaments. This suggests that CTD dimers may not form the basis for interaction with the viral RNA and assembly of N protein helical filaments or globular vRNPs that were recently also described using cryoEM tomography.182,420

SARS-CoV-2 Assembly and Release Coronaviruses assemble at the endoplasmic reticulum Golgi intermediate compartment (ERGIC) and Golgi where virions bud into to the lumen (Fig. 21.5).184,349,366 In situ cryo-EM tomography studies have shown that SARS-CoV-2 virions bud into high vesicle density areas in close proximity to ER and Golgi-like membranes.182 As with other coronaviruses, SARS-CoV-2 structural proteins (S, M, N, E) interact to drive budding at ERGIC membranes. The M protein plays a significant role in virus assembly through its interactions with the other structural proteins.140 Coexpression of the E and M proteins is sufficient in many cases for the production of noninfectious VLPs, which forms the basic structure for the viral envelope.30,311 SARS-CoV-2 and SARS-CoV E and M proteins are required for formation of VLPs, but coexpression with the N protein enhances particle production.26,30,295 SARS-CoV-2 E and M proteins are required for retention and maturation of S proteins at the site of assembly, as has been shown for other coronaviruses.26,30,295 SARS-CoV-2 S trimers localized in ERGIC members do not alone drive membrane curvature but appear to laterally reorganize in the envelope during virion assembly.182 Viral ribonucleoprotein complexes consisting of the genomic RNA and N protein accumulate at the curved membranes where budding takes place, possibly recruited by bending of the membranes to help drive the budding process.182

SARS-CoV-2 efficiently infects differentiated HAE cells.442 Peak SARS-CoV-2 release from the apical surface of HAE cells occurs 48 to 72 hours postinfection, whereas HCoV-NL63 and SARS-CoV exhibit slower kinetics with peak virus production occurring 72 to 96 hours pi.340

Less is known about how coronaviruses are released, or egress, from cells. After coronavirus particles are assembled, these traffic through the Golgi where glycosylation and other posttranslational processing occur.241,274 Previous studies have supported that virus particles are released through the exocytic secretory pathway.30,231,366 New information has emerged with SARS-CoV-2 that describe the use a lysosomal, Arl8bdependent pathway for egress and release.105 In this study, SARS-CoV-2 and MHV particles were observed to bud into ERGIC membranes, transport to the Golgi, as previously seen, but to diverge by trafficking to lysosomes for egress and release from cells. Additionally, lysosomal deacidification was observed that resulted in disruption of antigen presentation pathways.105 SARS-CoV-2 ORF3a protein has been implicated to promote lysosomal exocytosis, but the mechanism is not understood.53 Further studies are warranted to fully understand coronavirus egress and release from infected cells.

NONSTRUCTURAL PROTEINS (TABLE 21.1) SARS-CoV-2 and other betacoronaviruses encode 16 nonstructural proteins, nsp1-nsp16 within ORF1a and ORF1b (Fig. 21.1B). They are synthesized similarly to other coronaviruses; as two long overlapping polyproteins that are cotranslationally processed to individual nsps by either a papain-like protease domain contained within nsp3 (nsp1, nsp2, and nsp3) or by the 3C-like protease domain contained in nsp5 803

(nsp4-16).

Nsp1 SARS-CoV-2 Nsp1 is the first protein encoded in the ORF1a/ORF1ab polyproteins. It is proteolytically cleaved from the ORF1a precursor polyprotein by the papain-like protease domain in nsp3, as is the nsp1 of other coronaviruses. This results in a protein that is 180 amino acids long and is 84.4% sequence identical with SARS-CoV nsp1. This high degree of sequence conservation between SARS-CoV and SARS-CoV-2 nsp1 proteins (Table 21.1) results in a high degree of functional and structural similarity. The nsp1 proteins of both viruses interfere with host gene expression (reviewed in Ref.263) and consequently antagonize the development of an interferon response in infected cells.158,168,206,263,265,395,405

The molecular basis for the inhibition of host protein synthesis by nsp1 has been extensively investigated for SARS-CoV. Studies utilizing ectopic overexpression of nsp1 and purified bacterially expressed SARS-CoV nsp1 demonstrated that nsp1 promotes degradation of host mRNAs, binds to the 40S ribosomal subunit, inhibits formation of the 80S ribosome, and inhibits translation of capped RNAs during the initiation process.167,168,224 Binding of nsp1 to the 40S ribosome preinitiation complex results in endonucleolytic cleavage of 5′ capped host mRNAs near their 5′ end during translation elongation.146,167,168,356 Nsp1 does not contain a recognized nuclease motif and a direct ribonuclease activity has not been shown with purified protein, suggesting that nsp1 recruits a host nuclease.146,411 SARS-CoV nsp1 does not induce cleavage of SARS-CoV viral mRNAs, and protection from endonucleolytic cleavage was conferred on reporter mRNAs by adding the SARS-CoV 5′ leader sequence to their 5′ ends.146,356 However, when purified recombinant nsp1 protein was added to in vitro translation reactions programmed with a viral mRNA or a host mRNA carrying the 5′leader sequence and thus resistant to nsp1-mediated cleavage, protein synthesis was still inhibited, suggesting that nsp1 directly inhibited protein synthesis independently of its cleavage at the 5′ termini of host mRNAs.147 However, cotransfection of a nsp1 expression plasmid and a SARS-CoV-5′UTR-luciferase construct failed to significantly inhibit luciferase expression.356 Additional studies demonstrated that the first stem-loop in the 5′UTR bound to nsp1 and was sufficient to protect mRNAs from nucleolytic degradation.356 NMR analysis of the SARS-CoV nsp1 protein indicated that the N-terminal 12 amino acids are unstructured, that amino acids 13 to 128 form a well-folded globular domain, followed by a second C-terminal unstructured domain.8 The globular domain is made up of a 6-stranded beta-barrel with a long alpha helical segment, residues 36 to 49, across one barrel opening, and a second short alpha helical segment, residues 62 to 64 positioned next to the barrel. Additionally, there are two locally disordered regions at residues 77 to 86 and 121 to 128. Reverse genetic studies by Narayanan265 implicated the CTD in the inhibition of translation. Viruses carrying K164A/H165A mutations in this CTD inhibited host protein synthesis and degraded host mRNAs much less efficiently than wild type virus and allowed greater expression of interferon-β, ISG15 and ISG56, implying lesser inhibition of innate immune responses to infection.265 A deletion mutant within this region of the CTD was attenuated in a mouse model of SARS.163 In addition to inhibiting host protein synthesis, Wathelet et al.395 showed that overexpression of SARS-CoV nsp1 interferes with induction of interferon-β synthesis by Sendai virus infection by inhibiting activation of the transcription factors IRF-3, IRF-7. NF-КB and c-Jun that are required for induction of interferon-β mRNA synthesis.395 In the same work, these authors also showed that overexpression of nsp1 inhibited downstream signaling by interferons by decreasing phosphorylation of STAT1 after exposure of cells to interferon. Additionally, overexpression of nsp1 inhibited progression of cells through the cell cycle.395 Introduction of mutations in two basic surface exposed residues, R124 and K125, greatly attenuated these effects of nsp1.395 SARS-CoV carrying these nsp1 mutations are more sensitive to the antiviral effect of interferon than wild-type virus and replicate less well than wild-type in interferon competent cells. SARS-CoV nsp1 has also been shown to bind to Nup93 and thus disrupt the nucleocytoplasmic transport of host cell proteins.108

TABLE 21.1 Nonstructural proteins (NSPs) of SARS-CoV-2 Protein

Size (AA)

Sequence Identity With SARS-CoV Functional Roles

Nsp1

180

84.4%

Immune evasion, inhibitor of translation

Nsp2

638

68.3%

Unknown

Nsp3

1945

76.0%

Papain-like protease releases nsp1nsp4 from pp1a, immune evasion, DMV formation

Nsp4

500

80%

DMV formation

Nsp5

306

96.1%

Main protease (Mpro) releases nsp5-nsp16 from pp1a

Nsp6

290

88.2%

DMV formation

Nsp7

83

98.8%

Component of core replication/transcription complex

804

Nsp8

198

97.5%

Component of core replication/transcription complex

Nsp9

113

97.3%

Acceptor for nsp12 NiRAN nucleotide transferase activity; component of extended replication/transcription complex

Nsp10

139

99.3%

Cofactor for nsp14 exonuclease activity; cofactor for nsp16 2-2′-Omethyl transferase activity

Nsp11

13

84.6%

Unknown if functional, also contained in nsp12 N-terminal NiRAN domain due to programmed ribosomal frameshifting

Nsp12

932

96.4%

RNA-dependent RNA polymerase; nucleotidyl transferase

Nsp13

601

99.8%

Helicase, nucleotide triphosphatase

Nsp14

527

99.1%

3′ exonuclease with RNA proofreading activity; 5′Cap synthesis, N-methyl transferase activity

Nsp15

346

88.7%

Endonuclease, immune evasion

Nsp16

298

93.3%

5′ cap synthesis, 2′-O-methyl transferase activity

SARS-CoV-2 and other betacoronaviruses contain 16 nonstructural proteins, nsp1-nsp16. They are synthesized similarly in other coronaviruses as two long overlapping polyproteins, which are cotranslationally processed to individual nsps by either a papain-like protease domain contained within nsp3 (nsp1, nsp2, and nsp3) or by the 3C-like protease domain contained in nsp5 (nsp4-16). DMV, double membrane vesicles. Similarly, to SARS-CoV nsp1, SARS-CoV-2 nsp1 inhibits the translation of host mRNAs in both cell-free and cell-based assays.17,194,320,363,364,423 Studies of SARS-CoV-2 nsp1 interactions with the ribosome and the 5′UTR of the virus have advanced our understanding of the mechanism by which nsp1 inhibits protein synthesis while allowing translation of viral mRNAs.17,194,320,333,363,364,423 Cryoelectron microscopy (cryo-EM) revealed that nsp1 bound to 40S ribosomal subunits with two alpha helices of the previously structurally uncharacterized CTD positioned in the mRNA entry channel, obstructing it.320,363,423 The conserved residues K164 and H165 essential for nsp1 inhibition of protein synthesis and induction of an interferon response are located in a loop flanked by two alpha helices.17,363 These amino acids interact with 18S rRNA helix h18 and the two nsp1 helices interact with ribosomal proteins uS5 of the ribosomal body and with uS3 of the ribosomal head.17,320,363,423 These interactions do not block binding of the mRNA to the ribosome, but rather prevent the physiologic conformation of the 48S preinitiation complex by restricting rotation of the 40S ribosomal head and the loading of the mRNA into the entry channel.423 SL1, the first stem loop of the 5′UTR, when present at the 5′end of an mRNA, interacts with the globular NTD of nsp1, allowing entry of mRNA into the entry channel and subsequent translation.17,194,333,364 The structure of the NTD (amino acids 10 to 127) has been determined by X-ray crystallography57,321 and is similar overall to the structure of the SARS-CoV nsp1 NTD, with some alterations in surface exposed loops. A previously unreported activity of nsp1 is its inhibition of mRNA export from the nucleus. Zhang et al.433 have shown that nsp1 interacts with the nuclear mRNA export machinery by binding to the nuclear export receptor NFX1, and that infection with SARS-CoV2 inhibits the translocation of polyadenylated RNAs from the nucleus to the cytoplasm, a similar effect to that observed with NFX depletion. This effect could be replicated by nsp1 overexpression and the inhibition of mRNA nuclear export could be overcome by overexpressing NFX1 in SARSCoV-2–infected cells. Furthermore, nsp1 overexpression was shown to displace NFX1 from its binding partners at the nucleopore complex, providing a functional basis for these observations.

Nsp2 A relatively limited number of studies have been done on coronavirus nsp2, including SARS and SARS-CoV-2 nsp2, making it one of the poorly understood coronavirus proteins. The 638 amino acid nsp2 protein is proteolytically cleaved from the ORF1a precursor polyprotein by the papain-like protease domain in nsp3.297 Nsp2 proteins of SARS-CoV and SARS-CoV-2 are not as well conserved as many of the other nonstructural proteins, with only 68.3% sequence identity. The protein is not essential for SARS-CoV, and presumably SARS-CoV-2, or for MHV replication in cell cultures as viable mutants of SARS-CoV and MHV in which the entire nsp2 coding sequence has been deleted have been recovered using reverse genetics.117 These mutants were attenuated for viral replication in permissive cell cultures reaching titers 1 to 1.5 logs 805

lower than the corresponding isogenic wild-type virus. There have been few functional studies of this protein, but Prentice et al.297 showed that SARS-CoV nsp2 colocalizes with nsp3 and nsp8 in double membrane vesicular structures (DMVs) associated with viral RNA synthesis. Using the MHV system, Hagemeijer et al.128 demonstrated that fluorescently tagged nsp2 expressed by transient transfection was diffusely located in the cytoplasmic in uninfected cells but was recruited to DMVs, the site of coronavirus RNA synthesis, in MHV-infected cells. The association of nsp2 with DMVs was confirmed by immune-cryo-EM. Experiments expressing overlapping truncations of nsp2 fused with a fluorescent reporter revealed that the C-terminal fragment spanning amino acids 247 to 585 was sufficient for this localization to DMVs. Photobleaching experiments demonstrated that once recruited to DMVs nsp2 was not freely exchangeable after photobleaching. A yeast two-hybrid screen followed by confirmatory coprecipitation assays of overexpressed proteins in mammalian cells to identify binding partners among SARS-CoV proteins identified nsp4 and nsp6, two proteins associated with DMVs, nsp8, nsp11, and nsp16, associated with viral RNA synthesis and capping, and the SARS-CoV ORF3a protein, a virion associated accessory protein,154,330 as binding partners for nsp2.380

A study to identify potential SARS-CoV nsp2 host binding partners by an overexpression pull-down mass spectroscopy strategy identified 11 proteins, and the binding of two of these protein PHB1 and PHB2 were verified by western blotting assays.61 These two proteins have been associated with a variety of cellular processes: cell cycle progression, mitochondrial biogenesis, cell differentiation, and apoptosis.61 A comparative study using a similar affinity purification mass spectroscopy strategy to probe host protein interactions with nsp2 identified 4 proteins, ERLIN1, ERLIN2, RNF170, and TMEM, that interacted with both SARS-CoV and SARS-CoV-2 nsp2 proteins and 16 proteins that bound uniquely to each one.66 This study also confirmed the binding of SARS-CoV nsp2 to PHB2. Immunofluorescence colocalization indicated that considerable nsp2 localized to ER membranes in uninfected cell.66 Protein binding network and gene enrichment analyses suggest a role for nsp2 in interacting with the mitochondrial ER interface.66 As yet, there have been no studies to confirm these interactions with host proteins in infected cells nor to assess the functional importance of any of the nsp2–host protein interactions in virus infected cells.

Two studies of genetic variation in isolates of SARS-CoV-2 have identified an interesting pattern of sequence variation involving nsp2.296,426 Zeng et al.426 identified two epistatic linkages of a nsp2 T85I mutation to both a Q57H mutation in the ORF3a protein and to a noncoding sequence change in nsp14. Pohl et al.296 examined 14 low passage isolates for differences in their ability to replicate in Vero cells or bronchial epithelial cells. The three isolates that replicated well in the bronchial epithelial cells but relatively poorly in Vero cells all contained the linked nsp2 T85I and ORF3a Q57H mutations. The frequency of these changes reflected the frequency with which this linkage is found in SARS-CoV-2 and they were not further selected in culture by passage of the 14 viruses studied. The mechanism(s) underlying this linkage are unknown.

Nsp3 Nsp3 is the largest and most structurally complex of the ORF1a encoded nonstructural proteins (Fig. 21.8). It is proteolytically cleaved from the primary OF1a translation product, by the papain-like protease domain contained within nsp3 itself.443 It is a multidomain protein of 1945 amino acids in SARS-CoV-2, a 1922 amino acid protein in SARS-CoV, and contains 15 recognized domains.206 The difference in size between the SARS-CoV and SARS-CoV-2 nsp3 proteins is largely due to a 19 amino acid deletion in the hypervariable domain in the N-terminal region plus two smaller, two and three amino acid deletions in this same region. The functional significance, if any, of these deletions is unknown. The domain organization and functions of SARS-CoV nsp3 are summarized in Figure 21.8. The various domains are discussed below moving from the N-terminus to the C-terminus of nsp3.

Structural and functional characterizations for many these domains have been performed for SARS-CoV and for the more distantly related betacoronavirus, MHV, and will be briefly summarized. The nsp3 most N-terminal domain contains a ubiquitin-like fold and is designated Ubl1 since it is the first of two ubiquitin-like domains in coronavirus nsp3 proteins. The structure of the SARS-CoV Ubl1 domain has been determined by NMR spectroscopy.322 Two functions have been assigned to this domain: a ssRNA binding activity323 and, for the MHV nsp3 Ubl1 domain, a nucleocapsid (N) protein binding activity, which plays an important role in the initial stages of viral replication and is postulated to tether the viral genome to newly translated replicase– transcriptase complex.60,150 The hypervariable domain is an intrinsically disordered, glutamic acid rich region of unknown function which is poorly conserved among coronaviruses.322 It is dispensable for MHV in that recombinant viruses containing deletions in this region could be generated by reverse genetics and had no discernable phenotype.150

FIGURE 21.8 Summary of the functions and domain organization of SARS-CoV Nsp3. Nsp3 is bound to double-membrane vesicles recruited from the endoplasmic reticulum membrane. The protein passes through this membrane twice, via two transmembrane regions, TM1 and TM2. AH1 is possibly an amphipathic helix attached to the ER membrane, next to TM2. Except for the 3Ecto domain, all other Nsp3 domains are located in the cytosol. All domains with known three-dimensional structures are indicated in light green (Xray structures) or orange (NMR structures), whereas parts with unknown structure are in red. The best-characterized functions of each domain of Nsp3 are shown. Asterisks indicate the Asn1431 and Asn1434 glycosylation sites in the 3Ecto domain. Ubl, ubiquitin-like fold; HVR, hypervariable region; Mac, macrodomain; DPUP, domain preceding Ubl; NAB, nucleic acid binding domain; betaSM, betacoronavirus-specific marker. (Adapted from Lei J, Kusov Y, Hilgenfeld R. Nsp3 of coronaviruses: Structures and functions of a large multi-domain protein. Antiviral Res 2018;149:58–74. Copyright © 2017 Elsevier. With permission.) Following the hypervariable region is a conserved macro domain, initially named the X domain. It was bioinformatically predicted to have an 806

ADP-ribose-1″-phosphate (ADPR) phosphatase activity,345 and this activity was subsequently demonstrated for the SARS macro domain by Saikatendu et al. and a structure determined by X-ray crystallography.314 The enzyme contains a deep cleft in its surface containing the substrate binding pocket. Asparagine 41 is essential for macro domain phosphatase activity88; however, recombinant SARS-CoV containing the same asparagine 41 alanine mutation is viable and replicates to the same titer as wild-type virus in Vero cells, albeit slightly more slowly.189 In addition to its rather weak ADPR phosphatase activity, coronavirus macro domains have been reported to have hydrolase activity removing mono- and poly-ADP-ribose conjugated to proteins (deMARylation and dePARylation, respectively), posttranslational modifications that regulate innate immune responses.92 Recently, the SARS-CoV, SARS-CoV-2 and MERS-CoV macrodomains were all determined to have a deMARylation activity rather than a dePARylation activity.7 Recombinant mouse adapted SARS-CoV carrying mutations inactivating ADPR enzymatic activity were attenuated in mice and induced a greater interferon response than the parental mouse adapted virus.92

Following the macro domain is a region of nsp3 that was originally designated as the SARS unique domain or SUD.345 This region contains 3 subdomains, SUD-N, SUD-M, and SUD-C. SUD-N and SUD-M are macro domains, designated Mac2 and Mac3, and have been structurally characterized, are devoid of enzymatic activity, but bind RNAs containing G-quadraplexes.355 This binding activity is essential for viral RNA replication and transcription.190 Mac2Mac3 also binds to and stabilizes the RCHY1 protein, an E3-ubiquitin ligase targeting p53, an inhibitor of SARS-CoV replication.235 The SUD-C subdomain, also known as DPUP, is not required for replication of a SARS-replicon, but its deletion leads to greatly decreased replication.190 The functional role of the Ubl2 domain in virus replication and pathogenesis is unclear as it does not appear to affect the catalytic activity of the adjacent papain-like protease domain, and there are conflicting results in the literature regarding its role in antagonizing innate immune responses.59,98

Adjacent to the Ubl2 domain is the papain-like protease (PLpro) domain. The single SARS-CoV PLpro domain proteolytically cleaves the ORF1a encoded precursor polypeptide (pp1a) at LXGG sites found at the boundaries between nsp1, nsp2, nsp3, and nsp4, an activity required for viral replication. The structure of the SARS-CoV PLpro protease domain has been determined by X-ray crystallography and contains four Cys residues that coordinate zinc and the Cys-His-Asp catalytic triad of papain-like proteases.19,303 In addition to its role in generating nsps1-4 from its polyprotein precursor, PLpro also cleaves ubiquitin and ISG15 (an antiviral protein induced by interferon and containing two ubiquitinlike domains) at their C-terminal LXGG motifs and this cleavage event removes them from proteins to which they have been conjugated.19,216 Both ubiquitination and ISG15ylation play important roles in innate immune signaling pathways, thus deubiquitination (DUB) and deISGylation by overexpressed SARS-CoV PLpro unsurprisingly inhibited innate immune signaling.302 The PLpro domain contains two binding pockets for ubiquitin and the structurally related ISG15 that are separate from the 4 substrate binding pockets for their coronavirus substrates.302,303 Devaraj et al. have reported that interference with IRF3 by overexpressed SARS-CoV PLpro does not require it to be enzymatically active.78 In contrast, Deng et al.75 created recombinant MHV strains with mutations, which decreased binding of ubiquitin and ISG15 to the MHV PLpro2 domain, greatly decreasing DUB and deISGylation activities, yet still allowing MHV polyprotein processing by PLpro2 and thus recovery of infectious virus. These viruses activated an interferon response more quickly than the parental wild-type virus in macrophages and significantly decreased virus titer in the livers of infected mice. The SARS PLpro has been a target for the development of antivirals blocking its proteolytic activity and several compounds with IC50s in the low micromolar range with good selectivity in biochemical assays have been developed (reviewed in Refs.15,205).

The next two domains, the nucleic acid binding (NAB) domain and betacoronavirus specific domain (βSM), have not been extensively studied. The NAB domain binds to ssRNA and its structure has been determined by NMR.268,323 Even less is known about the betacoronavirus specific domain. Nsp3 contains two TMDs277 separated by a 3Ecto domain, which is situated on the luminal side of the ER membrane.277 When Nsp3 is coexpressed with nsp4, they induce curvature of ER-derived membranes,130 and when coexpressed with nsp4 and nsp6, they induce the formation of double membrane vesicles (DMVs) similar to the coronavirus replication organelle.13,101 The nsp3 3Ecto domain interacts with the luminal loops of nsp4 driving membrane curvature necessary for DMV formation.129,130 The second nsp3 TMD is followed by a hydrophobic amphipathic helical domain (AH1) that is located on the cytosolic side of the ER membrane. The C-terminal region contains two domains designated as the Y1 and Y-CoV domains. Their function is unknown.

The SARS-CoV-2 nsp3 protein has largely been studied in the context of inhibitors of its two enzymatic activities, the macro domain’s deMARylation activity and the PLpro domain’s proteinase activity. The structure of the SARS-CoV-2 PLpro domain has been determined by X-ray crystallography and is quite similar to the previously determined structure of the SARS-CoV PLpro domain.101 Both enzymes cleave a model peptide substrate mimicking the cleavage site between nsp2 and nsp3 with similar efficiencies.101 However, the two enzymes differ in their preferences toward ubiquitinated and ISGylated substrates with the SARS-CoV enzyme preferring ubiquitinated substrates and the SARSCoV-2 enzyme preferring ISGylated substrates.101,334 The basis for this difference has been determined by cocrystallization of these enzymes with ubiquitin or ISG15 and solving the structures of PLpro complexed with these molecules.334 Additionally, it has been reported that purified SARS-CoV-2 PLpro cleaves IRF3 at an LGGG motif in vitro,256 correlating with the observation that decreased levels of IRF3 are observed in SARS-CoV-2 infected cells in tissue culture. Several SARS-CoV protease inhibitors, including the noncovalent inhibitor GRL-0617,15,304 are also effective against SARS-CoV-2183,334 inhibiting both viral replication and the PLpro deubiquitinating and deISGylase activities. Additionally, GRL0167 treatment of SARS-CoV-2 infected cells increased the level of ISG15ylated IRF3 and increased the levels of interferon stimulated proteins over that detected in infected but untreated cells, suggesting that the inhibitor was effective in reversing the SARS-CoV-2 inhibition of the antiviral interferon response.334

The second enzymatically active domain in SARS-CoV-2 nsp3, the Mac1 ADP-ribosylhydrolase, has also been studied as a potential drug target. It has been suggested that the deMARylation of STAT1 may contribute to the cytokine storm associated with severe COVID-1958 making the Mac1 domain a possible therapeutic target. Its three-dimensional structure has been determined by four different groups7,31,248,300 and its interaction with various substrates characterized. As mentioned above, the SARS-CoV-2 macrodomain has deMARylating activity but not dePARylating activity, with the SARS-CoV-2 enzyme being somewhat more active than the SARS-CoV Mac1 domain. Potential binding pockets for both the adenosyl and ribose moieties of substrates were identified7,31,248,300 and potential inhibitors identified by virtual screening and optimization of inhibitors of a structurally related human poly(ADP-ribose) glycohydrolase.31 Screening against various nucleotides and nucleotide analogues demonstrated considerable flexibility of the adenosyl binding pocket and a surprising interaction with active metabolites of remdesivir suggesting another possible approach to developing inhibitors.269

807

Nsp4 and Nsp6 The nsp4 and nsp6 proteins are 500 and 290 amino acids long, respectively. At its N-terminus, nsp4 is cleaved from nsp3 by PLpro, and at its C-terminus, it is cleaved from nsp5 by Mpro, the protease contained within nsp5. Nsp6 is released from the coronavirus pp1a polyprotein by Mpro. Nsp4 contains four TMDs with both the N- and CTDs exposed to the cytoplasm, an approximately 100 amino acid CTD, and a relatively large luminal loop between the first and second TMDs.276 The luminal loop of nsp3 and the first large luminal loop of nsp4 are necessary for DMV formation.130,347 Nsp3, nsp4, and nsp6 all localize to the endoplasmic reticulum when expressed separately; coexpression of nsp3, nsp4, and nsp6 result in relocalization of these proteins to perinuclear foci.12,129,130,276 Electron tomography studies of cells coexpressing nsp3 and nsp4 in the absence of nsp6 demonstrated that these two proteins are sufficient to form structures, which resemble the DMV replication organelle found in infected cells.279 This is supported by the earlier colocalization of these proteins in perinuclear foci containing nsp8,297 a component of the RNA replication machinery (see below) in infected cells.

There have been a limited number of studies of the SARS-CoV-2 nsp4 and nsp6 proteins. An overexpression screen for proteins that antagonize the interferon response has shown that nsp6 suppresses type I interferon secretion in response to poly(I:C).405 Further examination of the effect of nsp6 overexpression on interferon induction and responses to interferon showed that this inhibited phosphorylation of TBK1 and its downstream target, IRF3, two events necessary for the induction of interferon-β, and down-regulated interferon receptor signaling by inhibiting phosphorylation of STAT1 and STAT2. As yet, the mechanism by which nsp6 produces these effects is unknown; nor have these effects of nsp6 on the interferon response been verified in infected cells. A screen for host binding partners with individually overexpressed SARS-CoV-2 proteins identified the ER receptor protein sigma-1 as an interaction partner with nsp6 and several mitochondrial proteins that are part of the TIM complex were identified as binding partners for nsp4.113 Small molecule ligands (inhibitors) of sigma-1, including a number of compounds in clinical use such as haloperidol and hydroxychloroquine, inhibited viral replication in Vero cells, as did inhibitors of the sigma-2 receptor.113 Genome sequencing of SARS-CoV-2 isolates have shown that a nsp6 L37F mutation is epistatically linked to a mutation in the accessory ORF3a gene.426 This mutation has recurred in different countries on multiple occasions and an association with milder disease has been suggested.373,388

Nsp5 Nsp5 is a highly conserved 306 amino acid protein in SARS-CoV and SARS-CoV-2, with 96.1% amino acid identity between the two proteins. This protein contains the main protease (Mpro), also referred to as the 3C-like protease (3CLpro), due to its similar substrate specificity and some structural similarities to the picornavirus 3C protease. Mpro is responsible for the majority of the processing of the pp1a and pp1b polyproteins, releasing nsp4-nsp16 from these polyproteins.9,345 Its enzymatic activity is essential for coronavirus replication. The structures of the SARS-CoV and SARS-CoV-2 Mpro have been solved by X-ray crystallography9,414,430 and reveal a protein with a chymotrypsin-like betabarrel domain (domain I, amino acids 10 to 99) and an adjacent picornavirus 3C-like beta-barrel domain (domain II, residues 100 to 182), which together make up a chymotrypsin-like fold but contain a catalytic dyad of His41 and Cys 145 in the active site, rather than the serine– histidine–aspartate catalytic triad of chymotrypsin (Fig. 21.9). A substrate binding cleft separates domains I and II, and there are three substrate binding pockets that provide substrate specificity. A third domain consists of 5 alpha helices and mediates dimerization, which is required for enzymatic activity.331 Mpro has strong sequence specificity requiring a glutamine in the P1 position (just N-terminal of the cleavage site), a strong preference for leucine immediately upstream (P2 position) of the glutamine, although other bulky hydrophobic amino acids can be recognized, no real amino acid preference in the P3 position, a preference for small neutral amino acids (valine, serine, threonine, proline) in the P4 position, and a strong preference for serine or alanine just C-terminal to the cleavage site (P1′ position). Substrate binding pockets (termed S1-S4, and S1′) have been identified for each of these positions. In addition to its essential role in viral replication, SARS-CoV and SARS-CoV-2 Mpro have been suggested to have a role in pathogenesis. Proteins that are components of the innate immune response expressed in vitro and incubated with purified recombinant SARS-CoV-2 Mpro identified NLRP12, a negative regulator of the release of inflammatory cytokines, and TAB1, another regulator of the inflammatory response, NRP, as potential targets for cleavage by Mpro.256 Levels of both proteins were decreased in SARS-CoV-2–infected ACE2-expressing 293 cells, consistent with these proteins being targeted by Mpro, although specific Mpro cleavage products were not reported.256

FIGURE 21.9 The structure of the SARS-CoV-2 Mpro, the protease contained within nsp5, and Mpro inhibitors. A: The three-dimensional structure of a dimer of SARS-CoV-2 Mpro. The protomers of the dimeric molecule are shown in light blue (protomer A) and orange (protomer B). Domains are labeled by Roman numerals and those in protomer B are marked with asterisks. The N- and C-terminals are labeled and the positions of the catalytic His41 and Cys145 are indicated by blue and yellow spheres, respectively. (From Zhang L, Lin D, Sun X, et al. Crystal structure of SARS-CoV-2 main protease provides a basis for design of improved α-ketoamide inhibitors. Science 2020;368(6489):409–412. Reprinted with permission from AAAS.) B: The structures of the most advanced Mpro inhibitors. (Reprinted from Vandyck K, Deval J. Considerations for the discovery and development of 3-chymotrypsin-like cysteine 808

protease inhibitors targeting SARS-CoV-2 infection. Curr Opin Virol 2021;49:36–40. Copyright © 2021 Elsevier. With permission.) Because of its essential role in viral replication and the structural information that is available, Mpro has been a therapeutic target for SARS, MERS, and SARS-CoV-2 (reviewed in Ref.219). Coronavirus Mpro is well conserved and some broadly active compounds inhibiting Mpro from multiple coronaviruses have been developed. Inhibitors can generally be classified as being peptidomimetic, which bind in the S1, S2, S3, and S1′ substrate binding pockets or allosteric inhibitors binding elsewhere and altering the substrate binding interface sites.219 Peptidomimentic compounds may contain ketones, ketoamide, aldehyde, or Michael receptors as reactive moieties, which covalently link to the active site cysteine or be noncovalent reversible inhibitors. Prior to the COVID-19 pandemic, a number of inhibitors of Mpro were developed and inhibit viral replication in cell cultures; several had broad anticoronavirus activity as well.9,24,175,176,415 The compound GC376, a bisufite adduct of a dipepityl-aldehyde Mpro inhibitor (GC373), has been shown to be an effective inhibitor of replication for multiple coronaviruses175,176 and has been used to successfully treat an otherwise fatal infection of cats with the feline coronavirus, infectious peritonitis virus.177 A derivative of GC376 has also been used to protect mice expressing the human MERS-CoV receptor from otherwise lethal infection with MERS-CoV and inhibit the replication of SARS-CoV-2 in cultured primary human airway cells301 and in Vero cells.382 PF-07321332374,436 is an oral Mpro inhibitor containing a nitrile warhead (Fig. 21.9B), which when combined with ritonavir, was shown to reduce hospitalization in SARS-CoV-2 infected persons in the interim analysis of the EPIC-HR phase 2/3 clinical trial (NCT04960202).280 A third peptidomimetic Mpro inhibitor developed against SARS-CoV, N3,415 contains a Michael receptor warhead, and is also active against SARS-CoV-2.164 An X-ray crystallographic screen of a library of drugs and drug-like compounds identified two allosteric inhibitors, which bound at or near the Mpro dimer interface resulting in a narrowing of the substrate binding cleft and altering the substrate binding site.124 Computational and biochemical screens of libraries of approved or investigational drugs and natural products identified a number of additional inhibitors of SARS-CoV-2 Mpro, which also inhibited SARS-CoV-2 replication.164 Among these are cinanserin, a serotonin antagonist, disulfiram, an inhibitor of aldehyde dehydrogenase used to treat alcoholism, and ebselem, a selenorganic compound, and carmofur, a 5-fluorouracil derivative used as a chemotherapeutic agent. Cinanserin48 had previously been identified as an inhibitor of the SARS enzyme and was an effective antiviral in tissue culture. Inhibitors of other viral proteases such as boceprevir, a ketoaminde inhibitor of hepatitis C virus ns3 protease, is also effective against SARS-CoV-2 Mpro.100 Lopinavir, a drug targeting the HIV protease, also inhibited SARS-CoV Mpro, SARS-CoV replication in vitro, and had modest effectiveness in an open label clinical trial during the 2002–2003 SARS outbreak.56,403 However, a clinical trial during the COVID-19 outbreak indicated that this drug was not effective in hospitalized patients.39 Clinical trials of ebselem (NCT04484025 and NCT04483973) and disulfiram (NCT04485130) have been registered at ClinicalTrials.gov.

The Replicase Complex: Nsp7-Nsp16 Overview Coronavirus RNA replication and transcription is a complex process that requires enzymatic machinery able to carry out the discontinuous coronavirus transcription process and replicate the very large coronavirus genome. The nine proteins that carry out these processes, nsp7-nsp10 and nsp12-16, carry a number of enzymatic and nonenzymatic functions (summarized in Table 21.1) and they have been colocalized to the DMVs that are the site of coronavirus RNA synthesis in the cell.27,77,115,127 During the 2002–2003 SARS outbreak, many of the enzymatic activities were predicted by a comparative bioinformatic analysis prior to being demonstrated biochemically.345 Nsp12 contains the typical seven sequence motifs (A-G) of an RNA-dependent RNA polymerase (RdRp)110,409 and has been reported to have primer-dependent358 and primer-independent RdRp activity.6 Bioinformatic analyses of nsp12 suggested that it lacked priming loops needed for positioning NTPs for initiating RNA synthesis and sequence comparison grouped it with primer-dependent RdRps such as the picornavirus 3Dpol based on the presence or absence of a motif implicated in primer recognition.409 Additional nsps with identified enzymatic activities in coronavirus RNA synthesis and modification are nsp13, which has 5′-3′ helicase and nucleotide triphosphatase activities155; nsp14, which has a N7methyltransferase activity that methylates the N7 position of guanosine in the 5′ cap of viral positive-sense RNAs producing a Cap-0 structure50 and a 3′ 5′ exoribonuclease activity which carries out a proof-reading function during coronavirus RNA synthesis84,254; and nsp16, which has a 2′-O-methyl transferase activity which methylates the 2′-O position of the ribose of the first nucleotide of the 5′ end of coronavirus mRNAs to form a Cap-1 structure.70 Nsp15 has an endoribonuclease activity, which when expressed cleaves dsRNA with a strong preference for U residues.156 Mutants with catalytically inactive nsp15 promote increased accumulation of dsRNA and consequent activation of innate immune antiviral pathways including IFN production and signaling, OAS/RNase L and PKR pathways; such mutants are reported to have modest170 to severe179 defects in viral replication.170 While the exact mechanisms underlying how nsp15 functions in the evasion of the host innate immune response are unknown, it is proposed to act by either degrading poly (U) on minus strand RNA126 or by degrading positive sense genome RNA.10 Nsp8 has been reported to have an RdRp activity that synthesizes, in a primer-independent manner, short oligoribonucleotides and thus was speculated to function as primase for nsp12 RNA synthesis.151 Subsequent work with a complex of nsp7, nsp8, and nsp12 has called this conclusion into question (see below).351 Nsps 7, 9, and 10 are devoid of enzymatic activities but form a number of different complexes with nsp8, nsp12, nsp13, nsp14, and nsp16 and have important regulatory roles in coronavirus RNA synthesis and modification.344

Core Replicase/Transcriptase Complex Proteins (Nsp12, Nsp8, and Nsp7) Nsp12 The SARS-CoV and SARS-CoV-2 nsp12 proteins are 932 amino acids in length and are proteolytically cleaved from their pp1ab precursor by the nsp5 Mpro.345 The two proteins are 96.4% identical in amino acid sequence. Nsp12 contains two domains, an N-terminal domain of approximately 300 amino acids, the NiRAN (nidovirus RdRp-associated nucleotidyltransferase) domain found in all nidovirus RdRp proteins, a roughly 500 amino acid C-terminal RdRp domain, with these two domains being joined by a linker region.204 The coronavirus RdRp domain has a similar architecture to other viral RdRps and can be described as resembling a cupped right hand with a thumb region, fingers and a palm, and contains a conserved SDD motif as part of the polymerase active site rather than the GDD motif found in most other RdRps, as well as the other seven (A-G) motifs associated with RdRp activity.110,180,409 Purified recombinant SARS-CoV nsp12 has a very modest level of in vitro polymerase activity with low processivity when presented with a template RNA that has been annealed to an RNA primer.358 A complex of nsp12 with nsp8 and nsp7 greatly increased polymerase activity and processivity.351 This tetrameric complex consists of one molecule of nsp12 809

bound to an nsp7-nsp8 heterodimer and a single molecule of nsp8. The complex exhibits both primer-independent de novo initiation of RNA synthesis and primer-dependent RdRp activity, and both these activities depend on the presence of an intact catalytic site in the RdRp domain of nsp12.351 The SARS-CoV nsp12 was refractory to crystallization but its structure was determined in 2019 by cryo-EM of the complex of nsp12 bound to nsp7 and nsp8 (Fig. 21.10).180 In this structure, the N-terminal NiRAN domain and the linker (interface) domain contact the RdRp domain at the outer surface of the fingers and the base of the palm subdomains. RNA entry and exit channels were identified in the RdRp domain as was the NTP binding channel. In addition, two previously unsuspected metal-binding sites were identified, one in the linker domain (H295-C301-C306-C310) and the second in the RdRp fingers subdomain (C487-H642-C645-C646). Although zinc was found ligated at these metal binding sites in the cryo-EM generated structure, subsequent work233 showed that these metal binding sites were occupied by Fe-S clusters in infected cells, that binding of the Fe-S clusters to RdRp is essential for polymerase activity, and that treatment of infected cells with a compound that oxidizes and disassembles FeS clusters strongly inhibited SARS-CoV-2 replication in infected cells. The nsp7-nsp8 heterodimer is bound above the RdRp thumb subdomain and the second molecule of nsp8 contacts nsp12 at the outside surface near the top of the finger subdomain of the RdRp domain and at the linker domain, which is adjacent to the to the RdRp fingers subdomain. The complex of nsp12 with nsp7 and nsp8 is the core structure of the coronavirus replication machinery. It provides a platform for larger complexes containing additional nsps involved in RNA synthesis and modification and has been used in assays of potential RdRp inhibitors (see below).

FIGURE 21.10 The structure of the SARS-CoV nsp12-nsp7-(nsp8)2 complex. SARS-CoV nsp12 contains a large N-terminal extension composed of the NiRAN domain (dark red) and an interface domain (purple) adjacent to the polymerase domain (orange). nsp12 binds to a heterodimer of nsp7 (blue) and nsp8 (green) as well as to a second subunit of nsp8. (Adapted from Kirchdoerfer RN, Ward AB. Structure of the SARS-CoV nsp12 polymerase bound to nsp7 and nsp8 co-factors. Nat Commun 2019;10(1):2342. https://creativecommons.org/licenses/by/4.0/.) The nsp12 NiRAN domain is unique to nidoviruses and was first characterized by Lehmann et al.204 This domain is conserved in all nidoviruses and a bioinformatic analysis identified three conserved motifs designated AN, BN, and CN. The NiRAN domain has a nucleotidyl transferase activity with a preference for UTP over other nucleotide triphosphates.204,342 When the SARS nsp12 was incubated with a UTP or GTP substrate, this activity transfers a UMP or GMP to the epsilon amino group of a conserved lysine residue at amino acid 73 in SARS-CoV nsp12 by formation of a phosphoramidate bond. In this assay, there was a strong preference for UTP over GTP and other nucleotide triphosphates were not utilized. The transferase activity was shown to be essential for nidovirus replication by a series of reverse genetic experiments with SARSCoV and for the related arterivirus, equine arteritis virus, in which infectious viruses carrying mutations in the conserved residues in motifs AN, BN, and CN required for the transferase activity could not be recovered. Three possible functions for this transferase activity were put forward: it could function as an RNA ligase, it could provide the as yet unidentified guanylyltransferase activity needed for 5′cap formation, or by transferring a UMP to a protein target it could have a protein priming function for minus-strand RNA synthesis. A subsequent set of experiments using HCoV-229E and SARS-CoV-2 nsp12 demonstrated that the NiRAN domain nucleotidyltransferase activity was much more active when it was provided with nsp9 as a substrate, that the N-terminal asparagine of nsp9 was the target of the nucleotide transfer reaction, and that this was vital for virus replication.342 Recent studies with the SARS-CoV-2 nsp12 demonstrated that in the presence of nsp13, a protein that binds to nsp12 (see discussion of replication/transcription complexes below) and contains both helicase and NTPase activity, the NiRAN domain was capable of transferring GMP to ppA-RNA. Thus the nsp12 NiRAN domain appears to be able to function as a guanylyl transferase, carrying out one of the early steps in the synthesis of a 5′ cap.412 This activity was diminished by the addition of nsp9 to the complex.

The RdRp and nucleotidyl transferase activities exhibited by nsp12 are attractive targets for antivirals. Prior to the COVID-19 outbreak, remdesivir (GS-5734), a prodrug of a nucleoside analogue of adenine (GS-441524) and a broad-spectrum inhibitor of viral RdRps, was shown to be an effective inhibitor of coronavirus replication in cell culture and protective in a mouse model of SARS.328 Its activity against coronaviruses is notable as most other nucleoside analogues are relatively ineffective because of the coronavirus nsp14 exonuclease proofreading activity (see section on nsp14 below).3 With the advent of the COVID-19 pandemic, remdesivir or its biochemically active triphosphate form was rapidly shown to be an effective inhibitor of the SARS-CoV-2 RdRp in biochemical assays,114 of SARS-CoV-2 replication in cell culture,298,391 and showed efficacy in mouse298 and macaque400 models of COVID-19. Incorporation of remdesivir into an elongating RNA by the SARS-CoV-2 RdRp results in delayed chain termination of RNA synthesis at the +3 position, three nucleotides after the site of remdesivir incorporation,114 similar to the mechanism observed in SARS and MERS coronaviruses.111 The cryo-EM structure of the RdRp domain in complex with an elongating RNA in the presence of remdesivir demonstrated a steric clash with nsp12 serine-861 that prevented the incorporation of a fourth nucleotide after the incorporated remdesivir monophosphate.185,421 Mutation of serine 861 to a less bulky glycine eliminates chain termination.357 A second mechanism of action has been proposed in which remdesivir that has been incorporated into a successfully elongated RNA acts as a poor template for subsequent rounds of RNA synthesis due to mispositioning of the incorporated remdesivir monophosphate in the RdRp active site.357 Passage of SARS-CoV and MHV in cell culture in the presence of remdesivir resulted in the accumulation of mutant viruses that had acquired resistance to the drug. These mutations were F480L and V557L in the SARS-CoV RdRp.3 Both mutations are in the fingers subdomain of the RdRp, and V557L is in RdRp motif F, which forms a channel for incoming NTPs, suggesting that the mutation may affect RdRp fidelity.

The COVID-19 pandemic spurred a series of computational and high throughput screens to identify potential inhibitors of SARS-CoV-2 replication. Multiple nucleoside analogues currently used as antivirals for other RNA viruses were tested as their triphosphate derivatives as inhibitors of primer elongation by the SARS-CoV-2 core polymerase complex.55 The triphosphate derivatives of sofosbuvir, tenofovir, alovudine, 810

and abacavir are incorporated in the elongating RNA by the RdRp and terminate further elongation. Sofosbuvir has been shown to have activity in inhibiting SARS-CoV-2 replication in cell culture, as was daclatasvir, a second nucleoside inhibitor of HCV replication.312 A meta-analysis of a number of small clinical trials of sofosbuvir in combination daclatasvir in COVID-19 patients suggested that this combination improved survival and shortened the time to clinical recovery,339 although the modest number of patients enrolled in the trials and differences in trial design precluded definitive conclusions on efficacy. Three additional nucleoside analogs, favipiravir, β-D-N4-hydroxycytidine (EIDD-1931), and a prodrug of EIDD-1931, EIDD-2801, inhibit coronavirus replication by a different mechanism, inducing an increase in the rate of transition mutations in replicating genomes resulting in error catastrophe.4,327,329 EIDD-2801 (molnupiravir) was developed to address the rapid catabolism of EIDD-1931 in the intestine of nonhuman primates.281 EIDD-2081 improved pulmonary function, decreased viral titer in the lungs, and decreased weight loss in mouse models of SARS, MERS, and in mouse and hamster models of COVID-19.308,329,385 EIDD-2801 can be given orally, was effective in preclinical studies, and reduced hospitalization in SARS-CoV-2–infected persons.281

Nsp7 and Nsp8 These two relatively small (83 to 198 amino acids) proteins are part of the coronavirus RNA replication/transcription machinery and are localized to the DMV RNA replication compartment in coronavirus infected cells.27 They are released from the pp1a and pp1ab precursor polypeptides by the nsp5 Mpro domain.74,345 Nsp7 and nsp8 are highly conserved with 98.8% and 97.5% sequence identity between the respective SARS-CoV and SARS-CoV-2 proteins. Structures of the SARS-CoV nsp7 have been determined by NMR290 and by X-ray crystallography,428 with the latter structure being determined in an in vitro assembled hexadecameric ring-like complex consisting of 8 molecules of nsp7 complexed with 8 molecules of nsp8. Nsp7 contains four alpha helices in both structures, although the helices are oriented somewhat differently in space in the two structures. The nsp8 molecules in the hexameric complex take on two conformations, one which generally has been described as a golf club with the N-terminal shaft (residues 6 to 104) consisting of three helices and the C-terminal head consisting of a beta-barrel and three helical segments folded into a tight hydrophobic domain. In the second nsp8 conformation, described as a bent golf club, the longest helix in the shaft domain contains a bend and thus is divided into two shorter helical segments.428 In the ring-like complex, the inner surface of this ring is positively charged and provides a surface for binding dsRNA with the long helices of nsp8 interacting with the RNA. As mentioned earlier, purified recombinant SARS nsp8 was shown to have a template-dependent RNA polymerase activity producing short, 6 nucleotides long, RNA products and has been suggested to act as primase for the nsp12 RNA-dependent RNA polymerase (RdRp), nsp12.151 Formation of the complex of nsp8 with nsp7 stimulates this polymerase activity and also allowed nsp8 to perform primer extension RNA synthesis as well as the de novo RNA synthesis described earlier although it was still about 20-fold less active than the nsp12 RdRp.359 Mutations of basic residues in the helical shaft of nsp8 abolished RNA binding and polymerase activity and mutagenesis of conserved D/ExD/E motifs identified residues in the nsp8 N-terminal helical domain required for polymerase activity of the complex.359,428 It should be kept in mind that the physiological relevance of the hexadecameric ring-like structure in viral replication is unclear since complexes with nsp12 have nsp7:nsp8:nsp12 ratios of 1:2:1 and polymerase activity is dependent on the nsp12 catalytic center.351

Nsp9 Nsp9 is a 113 amino acid protein that is a component of extended transcription/replication complexes.412 The protein is released from the pp1a and pp1ab precursor polypeptides by the nsp5 Mpro domain.74,345 In MHV-infected cells, nsp9 was localized to the DMV along with other components of the replicative machinery.27 The sequences of the SARS-CoV and SARS-CoV-2 proteins are 97.3% identical and bacterially overexpressed SARS-CoV protein was crystallized and its three-dimensional structure determined.88,354 Nsp9 was dimerized in the crystal structure with each molecule of nsp9 having the overall conformation of a six-stranded conical-shaped β-barrel with projecting loops flanked by a C-terminal α-helix bound to the β-barrel through hydrophobic interactions. The outward facing side of the helix is the dimerization interface between two molecules of nsp9. Dynamic light scattering measurements suggested that nsp9 was dimeric in solution while gel permeation chromatography suggested that nsp9 was monomeric in solution. Equilibrium sedimentation analysis suggested that the dimer and the monomeric forms of nsp9 were in equilibrium with each other.354 Further experiments employing site-directed mutagenesis demonstrated that a conserved GXXXG protein interaction motif spanning residues 100 to 104 was critical for dimerization and crucial for successful virus replication.250 Dimerization positions a patch of basic amino acids in each monomer adjacent to each other providing an area of high electrostatic potential to serve as binding region for RNA. Surface plasma resonance and electrophoretic mobility shift studies demonstrated that nsp9 was a ssRNA binding protein.88,354

With the advent of the COVID-19 pandemic, the SARS-CoV-2 nsp9 was overexpressed, purified, crystallized, and its structure determined by X-ray crystallography fairly early in the course of the pandemic.217 Gel permeation chromatography demonstrated that the purified protein was a dimer in solution and that its affinity for short polyU or polyT oligonucleotides was very low. The structure of the SARS-CoV-2 nsp9 was very similar to that of the SARS molecule as could be expected from their high degree of sequence identity. As noted above, nsp9 is a target for the nsp12 NiRAN domain catalyzed nucleotide transferase activity resulting in NMPylation of the nsp9 N-terminal asparagine residue, which is vital for virus replication.342 Cryo-EM of an extended replicase/transcriptase complex consisting of nsp7-nsp82-nsp9-nsp12-nsp132 showed that nsp9 was positioned with its N-terminus near the catalytic center of the nsp12 NiRAN domain.412

Nsp13 Nsp13 is a 601 amino acid protein that is a component of extended transcription/replication complexes.412 The SARS-CoV-2 nsp13 is 99.8% identical with the SARS-CoV nsp13, only differing from SARS-CoV nsp13 at residue 570 where a valine is substituted for an isoleucine. It is released from the pp1ab precursor polypeptide by the nsp5 Mpro domain74,345 and localizes to the DMV, along with other components of the replicative machinery.77,155 Nsp13 contains C-terminal helicase and N-terminal zinc finger (ZF) domains identified by bioinformatics analyses of the IBV and SARS-CoV genomes.110,345 The helicase domain contains 11 motifs associated with the SF1 family of helicases, with motifs I and II, also known as Walker A and B boxes, being required for NTP binding and hydrolysis.324 It most closely resembles helicases of the Upf1 family.203 Biochemical analyses of purified recombinant nsp13 from several coronaviruses, including SARS-CoV, were shown to have both NTPase and helicase activities.135,155,324 Studies with the HCoV-229E324 and the SARS-CoV nsp13155 established that the helicase was capable of unwinding both DNA and RNA duplexes with protruding 5′ ends in a 5′ to 3′ direction. The presence of a substrate for the helicase greatly stimulated its NTPase activity and a K288A mutation in motif I that interfered with NTP binding abolished both helicase NTPase and helicase activity, providing evidence that the NTPase activity provided the energy for NSP13 unwinding RNA duplexes. Further investigation of the 811

NTPase activity demonstrated that it was able to hydrolyze the 5′ triphosphate of a single-stranded RNA molecule to produce an RNA with a 5′ diphosphate. This 5′ NTPase activity was hypothesized to carry out the first step in synthesizing the 5′cap found on coronavirus mRNAs and the genome. A K288A mutation abolished this triphosphatase activity as well the NTPase and helicase activities. A kinetic analysis of nsp13 helicase activity demonstrated that unwinding a nucleic acid duplex took place in discrete steps of 9.3 nucleotides at a rate of 30 steps per second, and the addition of nsp12 to the reaction doubled the step size thus increasing the rate of unwinding two-fold, suggesting an association between the two proteins.1 Structures were determined for the MERS-CoV and SARS-CoV nsp13 molecules in 2017 and 2019, respectively.133,160 Both molecules were dimers in the crystallographic asymmetric unit, although the dimers were packed differently in the two structures. The structures of the two enzymes were very similar with both structures containing five domains: an N-terminal Cys-His rich domain binding 3 zinc atoms (designated ZBD or CH domains) and corresponding to the ZF domain identified bioinformatically,110,345 an α-helical stalk domain, a β-barrel domain (designated the 1B domain) followed by two RecA-like domains designated RecA1 (residues 241 to 443) and RecA2 (residues 444 to 596). These domains are arranged to form a pyramidal shaped molecule with the two RecA-like domains containing the first seven core SF1 family helicase motifs plus the 1B domain forming the base of the pyramid, the α-helical stalk domain lying atop the RecA1 and 1B domains with the zinc binding domain sitting above the α-helical stalk. The seven conserved SF1 family helicase motifs are located in a cleft between the two RecA-like domains, and this cleft contains the nucleotide binding pocket. A comparison of the nsp13 structure with the related arterivirus helicase and the Upf1 helicase suggested that the 3′ end of a bound ssRNA is located in a 6 to 12 angstrom wide tunnel formed by the 1B, stalk and RecA1 domains, with the 5′end of the RNA sitting atop the RecA2 domain. A binding site for nsp12 on the adjacent surfaces of the nsp13 ZBD and 1B domains was identified in 2019.133 Mutagenesis studies with the HCoV-229E nsp13 and a related arterivirus nsp10 indicated that an intact and functional ZBD was essential for ATPase and helicase activity and thus for viral replication.324 Unsurprisingly, a compound that inhibited the unwinding activity of the SARS-CoV helicase also inhibited the replication of a SARS-CoV RNA replicon.2

Studies of the SARS-CoV-2 nsp13 have either focused on identifying inhibitors of the NTPase and helicase activities424 or have been cryo-EM studies of nsp13 in complex with the replication/transcription complex (nsp7-nsp82-nsp12-nsp132) or in a extended replication complex additionally containing nsp9.52,236,411,412 Although there have been multiple computational and enzymatic screens for inhibitors of the NTPase and helicase activities, there have been relatively few which investigated viral replication in infected cells and SARS-CoV-2 infection of animals. Prior to the COVID-19 pandemic, bismuth compounds had been identified as inhibitors of the SARS-CoV nsp13 helicase and viral replication in infected cells.416 The compound ranitidine bismuth citrate is a potent inhibitor of purified nsp13 SARS-CoV-2 helicase and NTPase activities and inhibited virus replication in both infected cells and in SARS-CoV-2–infected Syrian hamsters while also decreasing the severity of pneumonitis in these animals.424 Treatment of purified nsp13 with bismuth compounds displaces zinc ions from the nsp13 zinc binding domain suggesting that this alters the interaction of this domain with the enzymatically active helicase. Cryo-EM studies52,417 of a replication/transcription complex (nsp7-nsp82-nsp12-nsp132) with a partially double-stranded RNA template-primer demonstrated that the two nsp13 molecules interacted through their 1B domains. The 1B domain of one molecule of nsp13, designated nsp13-1, bound to the nsp12 interface domain (links the RdrRp and NiRAN domains) while its ZBD domain bound to nsp8-1. The ZBD of the second molecule of nsp13, designated nsp13-2, bound to the nsp8-2 helical domain and the RdRp thumb domain, while its 1B domain made additional contacts with nsp8-2. The unpaired 5′ extension of the template RNA protrudes from the nsp12 catalytic site into the nsp13-2 RNA binding channel.52,236,411 It should be noted that as nsp12 RdRp progresses along the template RNA in a 3′ to 5′ direction and the nsp13 helicase unwinds RNA in the 5′ to 3′ direction, these two molecules’ movements along the template RNA oppose each other. If the helicase prevails, the RdRp will move backward on the template,52,236 a process called backtracking. Molecular dynamic simulations suggested that one or more misincorporated nucleotides at the elongating 3′end of the newly synthesized RNA will spontaneously fray away from the template and enter the nucleotide binding tunnel of the RdRp, a result supported by RNA-protein cross-linking experiments.236 Structural studies support a model where the RdRp motif F provides a strand-separating structure directing the frayed end of the elongating RNA to the nucleotide binding tunnel during backtracking. Backtracking has been suggested to play a role in the template switching event that is central to leader-body joining, an essential step in coronavirus transcription.236 It has also been suggested that this provides a mechanism to extrude an elongating RNA with a misincorporated nucleotide or nucleotide analogue through the nucleotide entry tunnel, providing access to the nsp14 exonuclease that provides a proof-reading function for coronavirus RNA synthesis (see below for a discussion of nsp14).

Coronavirus Capping Machinery Proteins (Nsp10, Nsp14, and Nsp16) The capping reactions There are five coronavirus proteins involved in the addition of a 5′ cap to positive sense virus-specific RNAs that are synthesized by the replication/transcription complex: nsp13, nsp12, nsp14, nsp16, and nsp10. The NTPase activity of nsp13 removes the gamma-phosphate from the 5′-triphosphate of nascent RNA transcripts and the nsp12 NiRAN domain appears to provide a guanylyltransferase activity to produce a GpppN at the 5′end of the newly synthesized RNAs (see sections on nsp12 and nsp13 above). The next two steps in producing a 5′ cap are the addition of a methyl group at the N7 position of the guanine by nsp14’s N7-methyl transferase activity to produce a Cap-0 structure, followed by a second methyl transferase reaction mediated by nsp16, adding methyl groups to the 2′-O position of the first RNA nucleotide (produces a Cap-1 structure) or to the first and second RNA nucleotides to produce a Cap-2 structure. The presence of a 5′ cap structure is essential for mRNA recognition by translation initiation factor eIF4E and thus translation of viral RNA and for replication,93,95 and it also plays a role in avoiding recognition by the innate immune system.445 Both nsp14 and nsp16 form heterodimers with nsp10, and these interactions and their functional effects are described in more detail below.

Nsp10 The SARS-CoV and SARS-CoV-2 nsp10 proteins are 139 amino acids in length and are proteolytically cleaved from their pp1ab precursor by the nsp5 Mpro.345 The two proteins are 99.3% identical in amino acid sequence, with the two differences being a nonconservative change from a proline (SARS-CoV) to an alanine (SARS-CoV-2) at position 23 and a conservative arginine (SARS-CoV) to lysine change (SARS-CoV-2) at amino acid 113. Nsp10 has an important role in viral replication and RNA synthesis, as demonstrated by a biochemical analysis of a MHV temperature sensitive mutant, tsLA6, that was mapped to nsp10 by sequencing the mutant and spontaneous revertants315 and by reverse genetic experiments with MHV and SARS-CoV.29,79 A yeast two-hybrid screen of the 16 SARS-CoV ORF1ab encoded proteins demonstrated that nsp10 could self-associate and strongly interacted with both nsp14 and nsp16.152 X-ray crystallography of purified SARS-CoV nsp10 provided a single domain structure composed of two N-terminal antiparallel alpha-helices stacked against a beta-sheet core and three additional alpha-helical regions followed by a coiled C-terminus.165 The protein contains two conserved zinc fingers and in MHV these were shown to play 812

an essential role in RNA synthesis.79 A cluster of basic amino acids on its surface is thought to mediate its modest single- and double-stranded NAB activity.165 Structures of the SARS-CoV nsp10-nsp1671 and nsp10-nsp14230 heterodimers have been determined and interaction surfaces mapped for both by their X-ray crystallographic structures. The interaction of nsp10 with nsp14 is exclusively with the nsp14 exonuclease domain and significantly activates nsp14 exonuclease activity.230 The nsp10 interaction surfaces for these two proteins partially overlap, thus excluding a single nsp10 molecule simultaneously interacting with both nsp14 and nsp16. The functional role of these interactions will be discussed in more detail under nsp14 and nsp16 below.

The SARS-CoV-2 nsp10 structure has been resolved by X-ray crystallography and was virtually identical to the structures determined for SARSCoV nsp10.306 The nsp10 structure in complexes with nsp14,215,220 with nsp16,307,378 and with an extended replication/transcription complex have been determined by either cryo-EM or X-ray crystallography studies and will be discussed below.

Nsp14 The SARS-CoV and SARS-CoV-2 nsp14 proteins are 527 amino acids in length and are proteolytically cleaved from their pp1ab precursor by the nsp5 Mpro.345 Nsp14 contains two domains each with a different enzymatic activity, an N-terminal exonuclease activity, designated as ExoN,254,345 and an N7-methyl transferase activity.50 The ExoN domain was predicted to be a 3-to-5′ exonuclease belonging to the DEDD super family of DNA and RNA exonucleases, so named because of the conserved amino acids in three motifs (motifs I, II, and III)345 essential for nsp14’s nuclease activity.254 A fifth residue, a histidine, is conserved 4 amino acids upstream of the last conserved aspartate in motif III, making ExoN a member of the DEDDh subfamily of the DEDD superfamily. The four acidic residues are part of the catalytic site and form two metal binding sites needed for catalytic activity.344 The ExoN domain also contains a zinc finger-like domain inserted between motifs I and II of the DEDDh helicases.345 Based on the role of some members of the DEDD family of exonucleases in DNA proofreading and the conservation of this domain in coronaviruses, toroviruses, and roniviruses, viruses with very large genomes in the Nidovirus family, and its absence in the smaller arteriviruses, it was speculated that this nsp14 domain might have a role in RNA proofreading or template switching during subgenomic RNA synthesis.345

Biochemical characterization of overexpressed and purified SARS-CoV ExoN demonstrated that the enzyme digested both single- and doublestranded RNA substrates in the predicted 3′-to- 5′ direction, but not DNA.254 The pattern of cleavage products obtained suggested that the target of this enzyme might be partially single-stranded RNAs. As noted under nsp10, binding of nsp10 to nsp14 greatly stimulated nsp14 ExoN nuclease activity.230 This complex is able to efficiently carry out the excision of mismatches at the 3′ end of a dsRNA, an activity consistent with a role in RNA proofreading. Additionally, the complex was shown to be able to excise remdesivir-MP from the 3′end of a synthetic dsRNA substrate, suggesting a role in resistance of coronaviruses to nucleoside analogues.94 Coexpression of nsp10 and nsp14 allowed the purification of nsp10-nsp14 complexes and determination of their structure by X-ray crystallography.230 Nsp14 contained two domains, an N-terminal ExoN domain encompassing amino acids 1 to 287 and a C-terminal N7-methyltransferase domain of amino acids 288 to 527. The structure defined the nsp10-nsp14 interaction surface showing that nsp10 interacts exclusively with the ExoN domain. The structure revealed that the ExoN catalytic residues are D (aspartatic acid) 90, E (glutamic acid) 92, E191 rather than the conserved D at position 243 in SARS-CoV nsp14 that was thought to be part of the ExoN active site, H (histidine) 268, and D273. The structure also revealed two zinc fingers with the first zinc finger positioned between motifs I and II with the second zinc finger overlapping domain III. Both are required for nuclease activity. The interaction of nsp14 with nsp10 produced a change in nsp14 conformation that stabilized the ExoN active site with a concomitant increase in ExoN enzymatic activity.94 In addition to interacting with nsp10, nsp14 interacts with nsp12 and the core RdRp complex consisting of nsp7nsp82-nsp12, suggesting that a super-molecular replication/transcriptase complex might exist to carry out both RNA synthesis, RNA proofreading, and the enzymatic reactions necessary for cap formation on mRNAs.94,351 In this complex, nsp12 interacts with both the ExoN and N7methyltransferase domains of nsp14.

Reverse genetic experiments with the alphacoronavirus HCoV-229E demonstrated that replacing any of the four conserved DDED residues in the exonuclease catalytic site was lethal for virus replication due to greatly reduced viral RNA synthesis, suggesting that the exonuclease activity was a potential therapeutic target.254 Mutations in the MHV and SARS-CoV nsp14 ExoN active site were not lethal but resulted in viruses with impaired viral RNA synthesis and modest growth deficits84,85 while significantly increasing mutation rates, supporting the idea that the ExoN domain had a functional role in RNA proofreading. Interestingly, similar mutations in the MERS-CoV nsp14 ExoN domain severely impaired enzymatic activity and markedly reduced viral RNA synthesis and were lethal to MERS-CoV, suggesting a possible role in replication beyond proofreading.273 This was confirmed in a study of recombination in three betacoronaviruses, MHV, MERS-CoV, and SARS-CoV-2, which demonstrated that recombination events were extensive during RNA synthesis, and that inactivating mutations in the ExoN domain decreased the frequency and altered the spectrum of recombination products.121 Importantly, the ExoN nuclease activity plays a key role in the relative resistance of coronaviruses to some nucleoside analogues. Recombinant MHV and SARS-CoV carrying catalytically inactivating mutations in the ExoN domain had a 300-fold decrease in growth in response to 5-fluorouracil (5-FU) compared to the more modest effect of the drug on wildtype viruses.343 The increased response to 5-FU is accompanied by a 24-fold increase in the mutation rate over that observed during infections with wild-type viruses treated with an equivalent dose of 5-FU. The majority of the increased number of mutations were the A-to-G and U-to-C transitions that are expected from incorporation of 5-FU into replicating genomes followed by mispairing of the incorporated 5-FUMP during subsequent rounds of RNA synthesis. Additionally, recombinant MHV strains with a catalytically inactive ExoN were more than 5-fold more sensitive to remdesivir than wild-type virus.3 These observations support the idea that at least one function of the nsp14 ExoN domain is RNA proofreading, and it may mediate relative resistance of coronaviruses to nucleoside analogues.3,343 In addition to its proofreading function, mutations rendering ExoN catalytically inactive increase MHV’s sensitivity to interferon; the nsp14 ExoN activity is necessary for MHV to replicate in macrophages and overcome the cell’s innate immune response,41 suggesting a role for this domain in immune evasion by coronaviruses. This is consistent with the observation that SARS-CoV carrying a catalytically inactive ExoN domain is attenuated in an aged mouse model of SARS.118

With the onset of COVID-19 pandemic, the role of the ExoN domain in SARS-CoV-2 replication and potential resistance to nucleoside analogues was investigated. Inactivating mutations in the catalytic site of ExoN were lethal, similarly to this effect in MERS-CoV.273 This contrasts with the effect of similar mutations in the SARS-CoV enzyme where infectious virus could be recovered in reverse genetic experiments84,85 and is surprising considering the much closer similarity of the SARS-CoV-2 ExoN domain to the SARS-CoV enzyme (99.7% identity) than to the MERS-CoV sequence (63% identity). Similar to findings for other coronaviruses, binding of SARS-CoV-2 nsp10 to nsp14 activates its ExoN activity.229 The substrate specificity of the SARS-CoV-2 ExoN domain is similar to that of the SARS-CoV nsp14 in that it removes mismatched 813

nucleotides at the 3′ end of a dsRNA, with the exception that it is also able to hydrolyze RNAs in the context of a 3′ mismatched RNA:DNA hybrid.229 The structural basis of mismatch recognition has been investigated by cryo-EM studies of synthetic templates bound to SARS-CoV-2 nsp10-nsp14 heterodimers (Fig. 21.11).220 Binding of a partially double-stranded RNA with a 5′ single-stranded extension and a mismatched 3′-terminal base pair to the nsp10-nsp14 heterodimer resulted in local conformational changes of the ExoN domain, slightly narrowing the RNA binding site and shifting the position of the catalytic residues stabilizing an active conformation of the enzyme. The binding of ExoN to the substrate RNA encompasses the last 2 residues of the fully base-paired nucleotides, flipping the template strand unpaired nucleotide out of the RNA helix, leaving the template mis-incorporated nucleotide as a 3′ single strand extension in the active site. This same study demonstrated that ExoN was capable of removing remdesivir-MP that had been incorporated into substrate RNA, consistent with earlier work showing that sensitivity to remdesivir is increased in coronaviruses carrying mutations in the ExoN catalytic site.3 Another cryo-EM study of a complex with the stoichiometry of nsp7-nsp82-nsp9-nsp12-nsp132-nsp10-nsp14 and a partially base-paired template RNA with a 5′ unpaired extension revealed that the nsp10-nsp14 complex bound to the extended replication complex (nsp7-nsp82-nsp9-nsp12-nsp132) in a shallow valley present between nsp9 and the nsp12 NiRAN domain with both of those domains contacting the nsp14 ExoN domain and nsp10 being in contact with the NiRAN domain (Fig. 21.11A and B).413 In the images captured by cryo-EM, a greater proportion of the molecules of this complex were side by side antiparallel dimers rather than monomers (Fig. 21.11C). In this dimer, the catalytic site of the ExoN domain is in relative proximity to the RdRp NTP entry tunnel suggesting that backtracking of the RdRp could allow extrusion of the 3′end of a mis-paired backtracked primer strand RNA where it would be accessible to the ExoN bound to the opposite nsp12 molecule. The cryo-EM data does not exclude the possibility that RNA proofreading is carried out by a nsp10-nsp14 heterodimer acting on a monomeric replication/transcription complex composed of nsp7-nsp82-nsp9-nsp12-nsp132 or nsp82-nsp9-nsp12-nsp132-nsp10-nsp14. The data demonstrating that ExoN activity was necessary for SARS-CoV-2 viability and that other coronaviruses that were able to tolerate mutational inactivation of ExoN were more sensitive to antivirals such as remdesivir has led to computational and biochemical screens for inhibitors of ExoN activity. One screen has identified two compounds, patulin and aurintricarboxylic acid, that inhibit SARS-CoV-2 nsp10-nsp14 ExoN activity in a concentration-dependent manner and inhibited SARS-CoV-2 replication at low micromolar concentration37 and synergized with remdesivir in inhibiting virus replication.

FIGURE 21.11 The cryoelectron microscopy structures of monomeric and dimeric SARS-CoV-2 extended replication/transcription complexes containing nsp10-nsp14 heterodimer. A: A schematic diagram of the domain structure of each component of the complex. Nsp7, deep purple; nsp8-1, gray; nsp8-2, green cyan; nsp9, purple blue; nsp10, slate; nsp12 NiRAN, yellow; nsp12 Interface, orange; nsp12 fingers, blue; nsp12 palm, red; nsp12 thumb, forest green; nsp13 ZBD, light green; nsp13 S, salmon; nsp13 1B, violet; nsp13 1A, sand; nsp13 2A, hot pink; nsp14 ExoN, pale green; nsp14 N7-MTase, brown. B: Three perpendicular views of the cryoelectron microscopy densities of the monomeric complex. C: Three perpendicular views of the cryoelectron microscopy densities of the dimeric complex. The dashed lines roughly indicate the boundary between the two complexes making up the dimer. (Adapted from Yan L, Ge J, Zheng L, et al. Cryo-EM Structure of an extended SARS-CoV-2 replication and transcription complex reveals an intermediate state in cap synthesis. Cell 2021;184(1):184–193.e10. Copyright © 2020 Elsevier. With permission.) Chen et al.50 first identified SARS-CoV nsp14 as having a guanine N7-methyltransferase (N7-MTase) activity by a functional genetic screen in yeast seeking enzymes needed for 5′cap formation. N7-MTase catalyzes the reaction that is the third step in cap formation. Biochemical analyses with purified nsp14 confirmed that the protein transferred a methyl group from S-adenosyl methionine (SAM) to the guanine N7 position of a GpppRNA substrate to synthesize a Cap-0 structure. They further mapped the activity to the CTD of nsp14 by mutagenesis. Alignment with other N7-MTases and an analysis of predicted secondary structure allowed the identification of a conserved DxG motif that is part of the SAM binding site in other methyl transferases. Mutations in this site abolished nsp14 N7-MTase activity in the yeast trans-complementation assay without affecting its ExoN activity. Similarly, mutations in the nsp14 ExoN catalytic site did not inactivate N7-MTase activity. Further experiments using a SARS-CoV replicon demonstrated that RNA replication and transcription were significantly impaired in replicons carrying inactivating mutations in the DxG motif, demonstrating the importance of the N7-MTase activity in viral replication. Biochemical characterization of purified bacterially expressed nsp14 confirmed the previous results of the yeast complementation assay and demonstrated the requirement for SAM as a the methyl donor.28 Binding of nsp10 to nsp14 failed to augment N7-MTase activity.28 The crystal structure of the nsp14-nsp10 heterodimer230 showed that the N7-MTase domain contained an atypical MTase fold. It contains a beta-sheet containing five betastrands rather than the usual seven strands, with the first four strands oriented parallel to one another with the last strand in an antiparallel orientation. Additionally, there is a small three-stranded antiparallel beta-sheet that is inserted perpendicularly to the central beta-sheet between the last two strands of the central sheet, which together with an N-terminal alpha-helix forms a pocket that tightly accommodates GpppA with the guanine N7 proximal to the methyl group of SAM in a position to effect the methyl transfer reaction. Mutation of residues making up the GpppA binding pocket confirmed their role in positioning the guanine in the appropriate position and orientation to accept the methyl group. There is also a zinc finger located distil to the N7-MTase active site. 814

The structural studies of the SARS-CoV-2 extended replication complex (nsp7-nsp82-nsp9-nsp12-nsp132) have shed some additional

understanding of how the 5′ ends of the coronavirus mRNAs are generated.413 In this complex, the catalytic site of the nsp12 NiRAN domain responsible for the guanylyl transfer reaction to the dephosphorylated 5′ end of the of elongating RNA is opposite the catalytic site of the nsp14 N7-MTase. The N7-MTase zinc finger of unknown function is positioned on a potential transfer path of the GpppRNA to the N7-MTase active site and is hypothesized to play a role in this transfer. The outbreak of the COVID-19 pandemic triggered a search for inhibitors of the SARSCoV-2 N7-MTase. Two biochemical screens of chemical libraries using purified nsp14 identified a number of inhibitors of the N7-MTase.20,173 The most potent inhibitors without toxicity inhibited viral replication at low micromolar concentrations. A study on the role of SARS-CoV-2 nsps1-16 as inhibitors of protein synthesis and the interferon response identified ectopically overexpressed nsp14 as potential inhibitor of these processes.142 Mutational inactivation of either the SARS-CoV-2 nsp14 ExoN endonuclease or the N7-MTase activities abrogated nsp14’s ability to inhibit translation, and the inhibition of translation was augmented by coexpression of nsp10, suggesting that nsp14 enzymatic activities are important in this effect. It has not yet been shown that nsp14 mediates an inhibitory effect on translation during virus infection, nor is the mechanism understood.

Nsp16 Nsp16 is a 298 amino acid protein and is released from the pp1ab precursor polypeptide by the nsp5 Mpro domain.74,345 The sequences of the SARS-CoV and SARS-CoV-2 proteins are 93.3% identical. Nsp16 was predicted to have a 2′O-methyltrransferase (2′O-MTase) activity based on comparative sequence analysis.345,381 It contains the four conserved catalytic amino acids, K-D-K-E, and a conserved SAM binding site that is characteristic of this family of enzymes. Biochemical assays of bacterially expressed and purified feline coronavirus (FeCoV) nsp16 demonstrated that it was able to bind to a Cap-0 (7meGpppRNA) containing short RNA substrate and transfer a methyl group from SAM to the 2′-O position of the first transcribed nucleotide, the last step in the synthesis of a fully functional 5′ cap (Cap-1). The enzyme had a requirement for the guanine N7-methyl group to be present for nsp16 to bind the RNA substrate, positioning it downstream of the N7-methyltransferase reaction catalyzed by nsp14 in 5′cap synthesis.70 Mutational analysis confirmed that the K-D-K-E residues were essential for enzyme activity. Although the FeCoV nsp16 is able to carry out this reaction in the absence of other viral proteins, the 2′O-MTase activity of SARS-CoV nsp16 is absolutely dependent on nsp10 binding.28,228 Binding of nsp10 to nsp16 enabled SARS-CoV nsp16 to bind both its substrates, 7meGpppRNA and SAM.51 The structure of the SARS-CoV nsp16 complexed with nsp10 was determined by X-ray crystallography and contains a canonical seven-stranded beta-sheet methyltransferase fold.51,71 Mutations of the SARS-CoV nsp10-nsp16 binding interface that abolish their interaction also abolish nsp16 2′O-methyltransferase activity.51,71,228 S-adenosylhomocysteine (SAH), a product of the methyltransferase reaction after the methyl group has been transferred from SAM to the RNA, was visualized and identified in the SAM binding pocket. The residues making up the SAM binding pocket are conserved among coronaviruses nsp16s. Binding of SAM in its nsp16 binding pocket increased the formation of enzymatically active nsp10-nsp16 complexes through an allosteric effect.14 Conversely, dissociation of SAH, a product of the methyl transferase reaction, from the SAM binding pocket promoted the dissociation of the nsp10-nsp16 complex. Mutation of the K-D-K-E residues to alanine abolished SARS-CoV nsp16 methyltransferase activity; similarly, mutation of the amino acids lining the SAM binding pocket severely compromised activity.71 A hydrophobic putative RNA binding groove leading to the catalytic site was identified using the vaccinia virus VP39 methyltransferase bound to RNA as a template.

The function of 2′-O methylation on mammalian and viral mRNAs was uncertain for many years as it is not required for mRNA translation.65 A role for this RNA modification in distinguishing host from nonhost mRNAs was first described in West Nile virus, a flavivirus, vaccinia virus, a poxvirus, and the coronavirus MHV, where mutational inactivation of these viruses’ 2′O-MTase activities made these viruses more sensitive to interferon and the interferon-stimulated IFIT proteins.65 This implicated the coronavirus nsp16 in immune evasion. Mutational inactivation of nsp16 in MHV or HCoV-229E resulted in increased interferon production in infected macrophages accompanied by decreased viral replication compared to recombinant viruses with a wild-type nsp16 gene.445 The induction of interferon upon infection with a 2′O-MTase deficient recombinant mutant MHV was dependent upon the macrophage having a functioning MDA5 protein. The role of nsp16 in immune evasion in vivo was demonstrated by the failure of a recombinant 2′-OMT mutant of MHV-A59 to productively infect the liver and spleen of mice challenged by intraperitoneal inoculation.445 These observations were extended to SARS-CoV using the MA15 mouse adapted SARS-CoV model, where mutational inactivation of the SARS-CoV nsp16 2′OMTase activity resulted in an attenuated infection both in vitro and in vivo and the attenuated viruses were more sensitive to interferon.245 As was the case for MHV, the attenuation of nsp16 mutants was dependent on intact MDA5 and IFIT proteins. The ability of attenuated 2′OMTase negative nsp16 mutants to serve as an attenuated live virus vaccine platform was also examined. Immunization of mice with mouse adapted SARS-CoV with an enzymatically inactive nsp16 elicited protective immunity to challenge with wild-type MA15. Similar results were obtained with MERS-CoV carrying a 2′OMTase mutation in nsp16247 suggesting that this approach to the development of live-attenuated vaccines might be applicable to a wide range of coronaviruses. Because the enzyme activity of nsp16 is dependent upon its allosteric activation by binding to nsp10, short peptides or peptidomimetic compounds targeting the interaction interface of these two proteins is a potential approach to an antiviral therapy. This approach was tested experimentally by the development of a short peptide targeting the nsp10-nsp16 interaction and demonstrated that it inhibited 2′-OMTase activity of a number of coronaviruses in vitro and inhibited MHV replication in cell cultures and attenuated infection and pathogenesis in vivo.390

With the onset of the COVID-19 pandemic, four groups determined the structure of SARS-CoV-2 nsp10-nsp16 complexes by X-ray crystallography.253,307,378,399 The structures were similar to that determined for the SARS-CoV nsp16. Structures in the presence and absence of a cap analogue, 7meGpppA, SAM, SAH, or the SAH analogue sinefungin, were also determined and allowed identification of the Cap-0 binding and SAM binding pockets.253,307,378,399 Nsp16 undergoes a conformational change upon binding SAM and its RNA substrate bringing the nsp16 catalytic residues and its substrates into closer proximity.378 The presence of divalent cations stabilizes interactions between nsp16 and its RNA substrate enhancing the alignment of the RNA in the active site.253 A comparison of the SARS-CoV-2 nsp16 structure to mammalian methyltransferases revealed a four amino acid insertion, which is unique to coronaviruses and promotes nsp16’s catalytic activity by altering the conformation of the RNA backbone in its binding groove.253 This four residue insertion was suggested as a promising target for the design of coronavirus-specific inhibitors of the nsp16 2′OMTase activity. In addition to the well-described role of nsp16’s 2′OMTase activity in immune evasion described earlier, it has been reported that overexpressed nsp16 binds to U1/U2 snRNAs and thus inhibits splicing.16 The observation that nsp16 can be identified in the nucleus of SARS-CoV-2–infected cells by immunofluorescent staining supports the idea that nsp16 may suppress the interferon response by interfering with the proper splicing of interferon and interferon responsive transcripts. Identification of the structural basis for binding to U1/U2 snRNAs followed up by reverse genetic experiments is needed to provide confirmation of the effect of nsp16 on splicing during SARS-CoV-2 infection.

815

SARS-COV-2 ACCESSORY PROTEINS Coronavirus accessory proteins differ among the lineages of human viruses. The proteins encoded by these genes often function to antagonize the host cell innate immune responses. Like other RNA viruses, coronaviruses produce double-stranded (ds)RNA early during the infection cycle as an intermediate in genome replication and messenger mRNA transcription.346 Host cell pattern recognition receptors (PRRs) sense viral dsRNA as pathogenic nonself and respond by activating several antiviral pathways critical for early defense against viral invasion. DsRNA sensing by cytosolic PRRs leads to activation of three key pathways, interferon (IFN) production and signaling, oligoadenylate-ribonuclease L (OAS-RNase L), and protein kinase R (PKR).149 Detection of dsRNA by MDA5 during coronavirus infection309 leads to signaling through mitochondrial antiviral signaling (MAVS) protein leading to activation and nuclear translocation of transcription factor IRF3 and production of type I (α/β) and type III (λ) interferon (IFN). Upon binding to its specific cell surface receptor, IFN promotes phosphorylation by Janus kinase (JAK) and translocation to the nucleus of signal transducers and activators of transcription (STAT)1 and STAT2, which induce expression of IFN stimulated genes (ISGs) with antiviral activities.226,294 In parallel, dsRNA is also sensed by oligoadenylate synthetases (OASs), primarily OAS3, which synthesize 2′,5′-linked oligoadenylates (2-5A),211,397 which induce dimerization and activation of RNase L, leading to degradation of viral and host single-stranded sRNA and protein synthesis inhibition.80 Finally, also in parallel, dsRNA sensing by PKR induces PKR autophosphorylation, permitting PKR to phosphorylate the translation initiation factor eIF2α leading to protein synthesis inhibition.313 While RNase L and PKR antiviral activities are not dependent on IFN production,397 OASs and PKR are encoded by ISGs; therefore, these pathways can be activated and/or further upregulated by IFNs. Similarly, RNase L and PKR activation can promote cellular stress, inflammation, and/or apoptotic death.16,43,45,169,234,438 Thus, these antiviral pathways are also destructive to the cell, thus further reducing host cell viability.

TABLE 21.2 Accessory proteins of SARS-CoV-2 compared to SARS-CoV

Coronaviruses utilize a large portion of their genomes to encode proteins that antagonize host defenses by many diverse mechanisms. These include some of the ORF1a/1b encoded proteins conserved among all coronaviruses (discussed above). In addition, each lineage of CoV encodes a unique set of accessory genes encoding proteins that promote evasion and/or antagonism of dsRNA-induced pathways described above (Fig. 21.1B; Table 21.2). These proteins are named by the ORFs encoding them. CoV proteins are usually translated from the 5′ ORF of each individual mRNA. However, there are less often two CoV proteins translated from separate ORFs encoded by the same RNA and in those cases the downstream ORFs are designated “b,” for example, ORF3a,b and ORFs7a,7b. The ORFs for accessory proteins in the SARS-CoV-2 genome are named with the same numbers as their homologues in SARS-CoV.166,249 These accessory proteins will be described below and summarized in Table 21.2.

ORF3a The SARS-CoV-2 ORF3a protein is similar to that of SARS-CoV, containing three transmembrane domains (TMDs),153 potassium ion channel activity, and a role in virion assembly and budding. The SARS-CoV ORF3a encoded protein accumulates and localizes to vesicles containing markers for late endosomes and is necessary for SARS-CoV–induced Golgi fragmentation.97 Consistent with this, a recombinant mutant SARSCoV-2 with a knockout of ORF3a was attenuated for weight loss and mortality of K18 mice expressing the human ACE2 receptor.337

ORF3b The very short (22 amino acid) ORF3b encoded protein of SARS-CoV-2, when overexpressed, was found to antagonize transcription from an IFNβ promoter construct and to prevent IRF3 localization to the nucleus, while the corresponding SARS-CoV 153 amino acid proteins does not have this activity.186 IFN antagonism was associated with the cytoplasmic localization noted for SARS-CoV-2 ORF3b but not that of SARS-CoV. Paradoxically, longer versions of ORF3b were isolated from two COVID-19 patients with severe disease and were associated with stronger antagonism of transcription from an IFNβ promoter in an in vitro assay. Interestingly, short forms of ORF3b are found in bat viruses related to both SARS-CoV and SARS-CoV-2. Confirmation of the role of this protein in the virus life cycle is needed.

ORF6 The ORF6 encoded protein was initially described for SARS-CoV and shown to prevent translocation of STAT1 to the nucleus, by sequestering 816

nuclear import factors on the membranes of the rough endoplasmic reticulum and Golgi.99 Subsequently, the SARS-CoV ORF6 encoded protein was shown to more generally antagonize karyopherin-dependent nuclear import of additional transcription factors needed for an effective host response to viral infections.341 Interestingly, a similar mechanism is utilized by MERS-CoV in that the NS4b protein was shown to antagonize the translocation of NF-κB, a transcription factor important for induction of inflammatory cytokines by competing for binding to karyopherins.38 The SARS-CoV-2 ORF6 encoded protein localizes to the nuclear pore complex (NPC) where it binds directly to the Nup98-Rae1 complex via its carboxyterminal end, to target the nuclear import pathway and in doing so reduces docking of karyopherin/importin and the attached transcription factors.255 The SARS-CoV protein also binds to this complex; however, SARS-CoV-2 more strongly disrupts nucleocytoplasmic transport than its SARS-CoV homolog. Not surprisingly a recombinant SARS-CoV-2 ORF6 knockout virus was attenuated inducing less weight loss and mortality during infection of K18 mice expressing the human ACE2 receptor.337

ORF7a The SARS-CoV ORF7a encodes a protein of 122 amino acids containing a TMD and cytoplasmic tail, which is localized to the Golgi complex and in lesser amounts to the endoplasmic reticulum (ER) and ER–Golgi intermediate compartment.317 Its luminal portion is similar in topology to members of the immunoglobulin superfamily.266 The ORF7a encoded protein has been reported to be incorporated into SARS-CoV particles by interacting with viral structural proteins E and M144 and in addition to contribute to apoptosis.317,375

The SARS-CoV-2 ORF7a encoded protein, when overexpressed, antagonizes type 1 IFN signaling as evidenced by inhibition of expression from an interferon-sensitive response element containing promoter (ISRE) driving expression of luciferase. This occurs by preventing phosphorylation of STAT2, a transcription factor required for IFN signaling and this depends on ubiquitination the ORF7a protein.40 Interestingly, mutations in the carboxy terminal portion of ORF7a protein were identified in clinical isolates of SARS-CoV-2 and shown to have a replication defect as well reduce the antagonism of IFN.267

Since the ORF7a encoded proteins of SARS-CoV and SARS-Cov2 are 85.2% identical and 95.9% similar, it is very likely that they have similar functions. However, studies of each protein, usually by overexpression, as described above, have revealed different activities. Thus, further studies utilizing infectious virus are needed to determine the actual function of the ORF7a encoded protein during the life cycle.

ORF7b The 44 amino acid SARS-CoV ORF7b encoded protein is an integral membrane protein that is localized to the Golgi and has been shown, like the ORF7a protein, to participate in inducing apoptosis.292,316 The SARS-CoV-2 ORF7b encoded protein, like the ORF7a protein, is very similar to that of SARS-CoV and likely to have similar functions.

ORF8 ORF8 encodes a protein of 121 amino acids with a signal sequence. As such it is found ER membrane associated and also secreted into the extracellular space and in serum in humans. Antibodies to the ORF8 encoded protein were detected early in patients before the detection of antinucleocapsid antibodies and proposed to be an early marker for infection and perhaps have diagnostic value.392 The ORF8 of SARS-CoV evolved into two smaller ORFs, 8a and 8b, following a 29 nucleotide deletion early in the SARS epidemic,275 suggesting the protein encoded in ORF8 was not required for viral replication in humans. SARS-CoV-2 has thus far retained ORF8, which is conserved among related bat coronaviruses. However, interestingly, isolates with deletions or mutations have been reported in different parts of the world. As early as January 2020, a variant with a 382 nucleotide deletion encompassing part of ORF7b and ORF8, including its transcriptional regulatory sequence (TRS), was reported in Singapore, accounting for 23.6% (45/191) of the sequenced viruses.350 These viruses replicated to a similar extent in cell culture and produced similar viral loads in people. However, infection with this variant was associated with milder disease422 and this variant died out due to control of viral spread in Singapore. Variants with deletions in parts of ORFs7b and 8 from 62 to 345 nucleotides were also identified in Taiwan, Australia, Bangladesh, and Spain,350 and the genome of the alpha variant (B.1.1.7) also contains a stop codon in ORF8 (C27972T; Q27stop) producing a truncated protein.

While it is clear that SARS-CoV-2 ORF8 encoded protein is not required for replication in cell culture or humans, there are several studies addressing its function. The ORF8 encoded protein when overexpressed bound to MHC class I and directed its degradation resulting in less antigen presentation on the cell surface, a type of immune evasion.431 ORF8 overexpression, but not the overexpression of other SARS-CoV-2 accessory proteins, has also been reported to induce ER stress responses within transfected cells; however, the consequences of such activation have not yet been characterized using viral mutants with deletion of ORF8 n.83 This finding is derived from data in cell culture and the relevance to infection of humans needs to be validated. However, a recombinant virus in which ORF8 was deleted replicated in cell culture with similar kinetics to wild-type virus and demonstrated similar pathogenesis in K18 human ACE2 expressing mice indicating ORF8 expression is not required for pathogenesis in this mouse model.337

ORF9b The ORF9b encoded protein of SARS-CoV-2 is quite similar in sequence and structure to that of SARS-CoV.332 The ORF9b encoded protein is localized to the membrane of mitochondria where it associates with TOM70, an outer membrane mitochondrial protein that serves as receptor of the mitochondrial antiviral-signaling (MAVS) protein. When overexpressed, the ORF9b encoded protein suppresses type I interferon responses.161 As with many other accessory gene products, this has yet to be confirmed in the context of infection.

Recombinant Viruses As described above, often initial investigations with a new pathogen identify host antagonists by overexpression of each individual protein and assessment of effects on IFN production or signaling. While this is a good first step, the most definitive way to prove this function is to produce a recombinant virus and identify the steps in virus–host interactions that are altered. Several groups have developed reverse genetics systems needed to construct mutant viruses. Using a Bacterial Artificial Chromosome (BAC) cloning system, a group of mutant recombinant viruses were

817

constructed in which ORF3a, ORF6, ORF7a, ORF7b, or ORF8 were deleted.337 Most of these viruses demonstrated smaller plaque morphology compared to parental virus, suggesting they may have defects in viral spread, and mutants with deletions of ORF7a and ORF8 showed lower levels of replication in several cell lines. When K18 mice bearing human ACE2 receptor were infected with each of these mutant viruses, those deleted for ORF3a, ORF7a, ORF7b, or ORF6 were attenuated in morbidity and mortality with the most striking attenuation exhibited by viruses lacking expression of the ORF3a or ORF6 encoded proteins. These data suggest that these proteins most likely contribute to SARS-CoV-2 virulence.

SARS-COV-2 VARIANTS—TRANSMISSION AND IMMUNE EVASION From the earliest time of detection in December 2019, SARS-CoV-2 has efficiently been transmitted between people. Without genome sequences and/or isolates from the earliest period of human transmission, it is impossible to know definitively whether adaptation in an intermediate host before transmission to humans modified the transmissibility. One line of evidence suggests a single spike mutation, threonine to alanine at amino acid in position 372 (T372A) occurred during this period and may confer a significant fitness benefit during infection of human cells.171 In the subsequent first several months of the pandemic phase of SARS-CoV-2, emergence of an additional mutation occurred, with viruses bearing G614D (glycine at position 614 rather than aspartic acid) rising to dominance worldwide.187 Functional studies in cell culture and animals have confirmed that this amino acid change alters spike protein functionality425,432 and enhances upper respiratory tract replication in animal models, conferring increased transmissibility.293

In the several months following the emergence of D614G, SARS-CoV-2 exhibited a remarkable degree of genetic stability, with variation due solely to genetic drift and an apparent absence of further adaptation to humans. In late 2020, coincident with a global acceleration in the number of cases, several variants of suspected altered biological properties emerged—some have been formally classified as variants of concern (VOC). The first of these to be identified was initially detected by robust genomic sequencing in the United Kingdom and classified as lineage B.1.1.7 (alpha), defined by 14 amino acid changes and 3 deletions, including an N501Y substitution in the spike RBD.67 Early studies identified a 30% to 70% transmission advantage for the alpha variant,67 which has been replicated in settings other than the United Kingdom including during spread of this lineage in the United States.393 Two other significant VOCs containing N501Y have emerged: B.1.351 (beta) first identified in South Africa360 and P.1 (gamma)64,91 identified in Brazil. Both of these lineages, like the alpha variant, contain an unusually large number of mutations overall, suggesting they may have evolved during prolonged infection of an immunocompromised individual, and both share a concerning suite of RBD mutations.

Beta and gamma variants contain substitutions at spike positions 417 (K417N/T), 484 (E484K), and 501 (N501Y) in addition to unique, lineage-defining mutations. Of these, E484K is of particular concern due to its potential to mediate immune escape, increasing the possibility of reinfection and potentially the frequency of breakthrough infections of vaccinated individuals. Substantial research earlier in the pandemic included deep mutational scanning of the spike RBD to identify changes that alter affinity for ACE2348 and reduce serum neutralization.119 In these assays, changes at position 484, specifically to K, P, or Q, produced the most substantial reductions in neutralization potency by human convalescent serum, though the effect was highly variable between individuals. In contrast, N501Y increases affinity for ACE2, possibly contributing to its higher transmissibility but does not appear to mediate any notable antibody escape.

Laboratory studies to determine how these variants might impact vaccine efficacy suggest that the alpha variant is fully susceptible to sera from vaccines,86 whereas beta and gamma variants are neutralized less potently by serum from vaccinated individuals.87,138 Most vaccines are expected to retain substantial efficacy against these established variants, although whether efficacy is retained at the maximal possible level remains unclear. Data from Israel reported in April 2021 suggest the beta variant may produce an increased likelihood of breakthrough infection in vaccinated individuals, but there are no data to support increased severity or transmission of such infections.191 Finally, as of spring 2021, the potential for booster vaccines matched to VOCs has already been contemplated and appears promising. Serum from individuals infected with the beta variant potently neutralizes gamma and wild-type spike proteins, suggesting a vaccine encoding the beta spike may produce broad efficacy257 against a variety of SARS-CoV-2 variants.

In addition to these three VOCs identified in fall and winter 2020, a fourth variant of concern was identified in early 2021 in California, identified as lineage B.1.427/429 (epsilon) containing RBD mutation L452R. This lineage appears to exhibit approximately 20% enhance transmissibility compared to variants previously circulating in the United States76 but does not appear to pose a significant threat to vaccine efficacy120 and in June 2021 was downgraded to a variant of interest as its frequency declined in competition of alpha and delta variants. As SARS-CoV-2 continues to circulate at high levels globally, additional variants continue to emerge. Amino acid changes at spike positions 484 and 681, most commonly E484K and P681H/R, respectively, repeatedly emerge in various lineages. This signal of convergent evolution is consistent with experimental support for E484K conferring a fitness advantage in the face of individual and population immunity, so as global vaccine coverage increases, viruses containing K484 may rise to dominance globally. Continued surveillance activities will be required to ensure vaccine formulations are optimized for dominant circulating viruses, and infrastructure put in place to deliver updated vaccines as warranted.

In late spring 2021, an additional variant of concern, B.1.617.2 (delta), was identified associated with the catastrophic SARS-CoV-2 epidemic in India. The delta variant contains RBD amino acid changes L452R and T478K, as well as a P681R substitution preceding the S1/S2 furin cleavage site. Following introduction to the United Kingdom, this variant rapidly displaced the alpha variant as the dominant lineage.36,46 Compared to other VOC, delta is estimated to have the largest increase in reproductive number over nonvariant of concern lineages, explaining its displacement of the alpha variant where the two lineages have circulated in the same populations.36 However, as with other variants, the role of specific amino acid changes in spike and other genes in enhancing transmission remains unclear. The P681R change in the delta variant enhances spike protein cleavage at S1/S2284 and the furin cleavage site plays an important role in transmission and pathogenesis in ferrets. However, the significance of this with respect to transmission in humans is unknown. Finally, the delta variant exhibits enhanced neutralization resistance to convalescent and vaccine sera, but of a lower magnitude than the beta variant221,222 and estimates of the impact on vaccine effectiveness are ongoing at the time of writing but expected to be modest in fully vaccinated individuals.

SARS-COV-2: SECONDARY TRANSMISSION TO WILD AND DOMESTIC ANIMALS 818

In addition to its high infectivity in humans, SARS-CoV-2 exhibits a broad documented and predicted host range. Experimental infections of nonhuman primates,258 hamsters,308 ferrets,178 and raccoon dogs96 result in robust viral replication and, in some cases, disease. Although most of these represent experimental animal models, the susceptibility of raccoon dogs bears on their potential role as intermediate hosts, and they were widely solid in Wuhan markets in the fall of 2019.407

In silico structural modeling combined with functional binding studies have further expanded our understanding of the potential SARS-CoV-2 host range. Generally, in silico modeling has been informative but not perfectly correlated with functional binding assays. One comprehensive study, for example, predicted high binding affinity for human ACE-2 and very low affinity to Chinese pangolin (M. pentadactyla) ACE-2, a species closely related to the Malayan pangolin (Manis javanica), which is susceptible to viruses closely related to SARS-CoV-2.192,384,407 The ACE-2 proteins of the two species share 99% amino acid identity. A more recent study, however, found that SARS-CoV-2 spike binds to Malayan pangolin ACE-2 with equivalent high identity as to human ACE-2, suggesting in silico studies may not be perfectly predictive of in vivo susceptibility.272

In numerous instances, humans have transmitted ACE-2 to domestic animals or captive wildlife, highlighting the potential for SARS-CoV-2 to establish new reservoirs outside its presumed natural range of southeast Asia and/or southern China. During the current period of continued epidemic transmission in humans, flare-ups from secondary reservoirs is not a major concern, but in a possible low endemicity future new outbreaks could be sparked from animal reservoirs worldwide, should they become established. Multiple diverse species factor in these potential concerns. While domestic rodents appear not susceptible to SARS-CoV-2, deer mice, a widely distributed wild rodent, are highly susceptible89,122 and are a reservoir for other viruses that infect humans. Thus, the potential infection of wild deer mice represents a potential “spill-back” scenario for the permanent circulation of SARS-CoV-2 in wildlife, with the potential to seed future human outbreaks.47 Additionally, widespread SARS-CoV-2 infection of white-tailed deer has been documented in North America.47,131 Transmission from deer back to humans has not yet been identified, but the potential clearly exists given close contact between humans and these animals. Although the additional public health risk posed by such transmission may be limited while circulation remains high in the human population, as circulation ebbs spillback events may pose a greater risk of sparking local outbreaks.

Among domestic animals, felines and mink represent the most frequent animals infected via contact with infected humans. The most notorious instance of feline SARS-CoV-2 infection has been the infection of captive lions and tigers at the Bronx Zoo in 2020.240 Other studies have found that infected humans routinely transmit SARS-CoV-2 to domestic cats.33,132,141 Mink have proved particularly susceptible, and mink farms a fertile ground for large outbreaks and the emergence of potentially significant amino acid substitutions in the spike RBD. Mink outbreaks have occurred on farms in both North America33,335 and Europe.5,32,195 On multiple occasions, mink-derived viruses have spilled back into the human population resulting in community spread of these variants. This is of particular concern because mink-associated mutations such as spike Y453F are associated with reduced susceptibility to neutralization by some monoclonal antibodies,139 raising the possibility that minkderived outbreaks may produce infections resistant to some therapeutics and increasing the need for the mink farming industry to implement rigorous biosecurity practices. Mink may also represent a potential spillback reservoir as wild and feral minks have repeatedly found to be infected with SARS-CoV-2, though whether such a reservoir would pose significant risk to humans remains unclear.

The broad potential host range of SARS-CoV-2 highlights its generalist properties as described by MacLean et al.232 produced during its evolution in bats. In addition to secondary infections among domestic animals, captive and feral animals such as cats and mink may prove able to sustain SARS-CoV-2 transmission as new reservoir hosts. Given that these animals, in addition to susceptible wild rodents, come into frequent contact with other animals as well as humans the potential exists to spark human outbreaks with variants marked in some cases by altered neutralization susceptibility. This warrants continued surveillance, improved biosecurity in farming industries, and the necessity to suppress human transmission to minimize the risk of spillback.

1.

819

References 1. Adedeji AO, et al. Mechanism of nucleic acid unwinding by SARS-CoV helicase. PLoS One 2012;7:e36521. 2. Adedeji AO, et al. Severe acute respiratory syndrome coronavirus replication inhibitor that interferes with the nucleic acid unwinding of the viral helicase. Antimicrob Agents Chemother 2012;56:4718–4728. 3. Agostini ML, et al. Coronavirus susceptibility to the antiviral remdesivir (GS-5734) is mediated by the viral polymerase and the proofreading exoribonuclease. MBio 2018;9. 4. Agostini ML, et al. Small-molecule antiviral β-d-N (4)-hydroxycytidine inhibits a proofreading-intact coronavirus with a high genetic barrier to resistance. J Virol 2019;93. 5. Aguilo-Gisbert J, et al. First description of SARS-CoV-2 infection in two Feral American Mink (Neovison vison) caught in the wild. Animals (Basel) 2021;11. 6. Ahn DG, Choi JK, Taylor DR, et al. Biochemical characterization of a recombinant SARS coronavirus nsp12 RNA-dependent RNA polymerase capable of copying viral RNA templates. Arch Virol 2012;157:2095–2104. 7. Alhammad YMO, et al. The SARS-CoV-2 conserved macrodomain is a Mono-ADP-ribosylhydrolase. J Virol 2021;95. 8. Almeida MS, Johnson MA, Herrmann T, et al. Novel beta-barrel fold in the nuclear magnetic resonance structure of the replicase nonstructural protein 1 from the severe acute respiratory syndrome coronavirus. J Virol 2007;81:3151–3161. 9. Anand K, Ziebuhr J, Wadhwani P, et al. Coronavirus main proteinase (3CLpro) structure: basis for design of anti-SARS drugs. Science 2003;300:1763–1767. 10. Ancar R, et al. Physiologic RNA targets and refined sequence specificity of coronavirus EndoU. RNA 2020;26:1976–1999. 11. Andersen KG, Rambaut A, Lipkin WI, et al. The proximal origin of SARS-CoV-2. Nat Med 2020;26:450–452. 12. Angeletti S, et al. COVID-2019: the role of the nsp2 and nsp3 in its pathogenesis. J Med Virol 2020;92:584–588. 13. Angelini MM, Akhlaghpour M, Neuman BW, et al. Severe acute respiratory syndrome coronavirus nonstructural proteins 3, 4, and 6 induce double-membrane vesicles. MBio 2013;4. 14. Aouadi W, et al. Binding of the Methyl Donor S-Adenosyl-l-Methionine to Middle East respiratory syndrome coronavirus 2′-OMethyltransferase nsp16 promotes recruitment of the allosteric activator nsp10. J Virol 2017;91. 15. Báez-Santos YM, St John SE, Mesecar AD. The SARS-coronavirus papain-like protease: structure, function and inhibition by designed antiviral compounds. Antiviral Res 2015;115:21–38. 16. Banerjee S, Chakrabarti A, Jha BK, et al. Cell-type-specific effects of RNase L on viral induction of beta interferon. MBio 2014;5:e00856. 17. Banerjee AK, et al. SARS-CoV-2 disrupts splicing, translation, and protein trafficking to suppress host defenses. Cell 2020;183:1325–1339.e1321. 18. Barrantes FJ. Structural biology of coronavirus ion channels. Acta Crystallogr D Struct Biol 2021;77:391–402. 19. Barretto N, et al. The papain-like protease of severe acute respiratory syndrome coronavirus has deubiquitinating activity. J Virol 2005;79:15189–15198. 20. Basu S, et al. Identifying SARS-CoV-2 antiviral compounds by screening for small molecule inhibitors of Nsp14 RNA cap methyltransferase. Biochem J 2021;478:2481–2497. 21. Bayati A, Kumar R, Francis V, et al. SARS-CoV-2 infects cells after viral entry via clathrin-mediated endocytosis. J Biol Chem 2021;296:100306. 22. Bestle D, et al. TMPRSS2 and furin are both essential for proteolytic activation of SARS-CoV-2 in human airway cells. Life Sci Alliance 2020;3. 23. Boni MF, et al. Evolutionary origins of the SARS-CoV-2 sarbecovirus lineage responsible for the COVID-19 pandemic. Nat Microbiol 2020;5:1408–1417. doi: 10.1038/s41564-020-0771-4. 24. Boras B, et al. Preclinical characterization of an intravenous coronavirus 3CL protease inhibitor for the potential treatment of COVID19. Nat Commun 2021;12:6055. 25. Boscarino JA, Logan HL, Lacny JJ, et al. Envelope protein palmitoylations are crucial for murine coronavirus assembly. J Virol 2008;82:2989–2999. 26. Boson B, et al. The SARS-CoV-2 envelope and membrane proteins modulate maturation and retention of the spike protein, allowing assembly of virus-like particles. J Biol Chem 2021;296:100111. 27. Bost AG, Carnahan RH, Lu XT, et al. Four proteins processed from the replicase gene polyprotein of mouse hepatitis virus colocalize in the cell periphery and adjacent to sites of virion assembly. J Virol 2000;74:3379–3387. 28. Bouvet M, et al. In vitro reconstitution of SARS-coronavirus mRNA cap methylation. PLoS Pathog 2010;6:e1000863. 29. Bouvet M, et al. Coronavirus Nsp10, a critical co-factor for activation of multiple replicative enzymes. J Biol Chem 2014;289:25783–25796. 30. Bracquemond D, Muriaux D. Betacoronavirus assembly: clues and perspectives for elucidating SARS-CoV-2 particle formation and egress. mBio 2021;12:e0237121. 31. Brosey CA, et al. Targeting SARS-CoV-2 Nsp3 macrodomain structure with insights from human poly(ADP-ribose) glycohydrolase (PARG) structures with inhibitors. Prog Biophys Mol Biol 2021;163:171–186. doi: 10.1016/j.pbiomolbio.2021.02.002. 32. Burkholz S, et al. Paired SARS-CoV-2 spike protein mutations observed during ongoing SARS-CoV-2 viral transfer from humans to minks and back to humans. Infect Genet Evol 2021;93:104897. 33. Cai Y, et al. CD26/DPP4 cell-surface expression in bat cells correlates with bat cell susceptibility to Middle East respiratory syndrome coronavirus (MERS-CoV) infection and evolution of persistent infection. PLoS One 2014;9:e112060. 34. Cai Y, et al. Distinct conformational states of SARS-CoV-2 spike protein. Science 2020;369:1586–1592. 35. Caillet-Saguy C, et al. Host PDZ-containing proteins targeted by SARS-CoV-2. FEBS J 2021;288:5148–5162. 36. Campbell F, et al. Increased transmissibility and global spread of SARS-CoV-2 variants of concern as at June 2021. Euro Surveill 2021;26. 37. Canal B, et al. Identifying SARS-CoV-2 antiviral compounds by screening for small molecule inhibitors of nsp14/nsp10 exoribonuclease. Biochem J 2021;478:2445–2464. 38. Canton J, et al. MERS-CoV 4b protein interferes with the NF-kappaB-dependent innate immune response during infection. PLoS Pathog 2018;14:e1006838. 39. Cao B, et al. A trial of lopinavir-ritonavir in adults hospitalized with severe Covid-19. N Engl J Med 2020;382:1787–1799. 40. Cao Z, et al. Ubiquitination of SARS-CoV-2 ORF7a promotes antagonism of interferon response. Cell Mol Immunol 2021;18:746–748. 41. Case JB, et al. Murine hepatitis virus nsp14 exoribonuclease activity is required for resistance to innate immunity. J Virol 2018;92. 42. Castaño-Rodriguez C, et al. Role of severe acute respiratory syndrome coronavirus viroporins E, 3a, and 8a in replication and pathogenesis. MBio 2018;9. 43. Castelli JC, et al. A study of the interferon antiviral mechanism: apoptosis activation by the 2-5A system. J Exp Med 1997;186:967–972. 44. Chai J, et al. Structural basis for SARS-CoV-2 envelope protein recognition of human cell junction protein PALS1. Nat Commun 820

2021;12:3433. 45. Chakrabarti A, et al. RNase L activates the NLRP3 inflammasome during viral infections. Cell Host Microbe 2015;17:466–477. 46. Challen R, et al. Early epidemiological signatures of novel SARS-CoV-2 variants: establishment of B.1.617.2 in England. medRxiv 2021. doi: https://doi.org/10.1101/2021.06.05.21258365. 47. Chandler JC, et al. SARS-CoV-2 exposure in wild white-tailed deer (Odocoileus virginianus). Proc Natl Acad Sci U S A 2021;118. 48. Chen L, et al. Cinanserin is an inhibitor of the 3C-like proteinase of severe acute respiratory syndrome coronavirus and strongly reduces virus replication in vitro. J Virol 2005;79:7095–7103. 49. Chen CY, et al. Structure of the SARS coronavirus nucleocapsid protein RNA-binding dimerization domain suggests a mechanism for helical packaging of viral RNA. J Mol Biol 2007;368:1075–1086. 50. Chen Y, et al. Functional screen reveals SARS coronavirus nonstructural protein nsp14 as a novel cap N7 methyltransferase. Proc Natl Acad Sci U S A 2009;106:3484–3489. 51. Chen Y, et al. Biochemical and structural insights into the mechanisms of SARS coronavirus RNA ribose 2′-O-methylation by nsp16/nsp10 protein complex. PLoS Pathog 2011;7:e1002294. 52. Chen J, et al. Structural basis for helicase-polymerase coupling in the SARS-CoV-2 replication-transcription complex. Cell 2020;182:1560–1573.e1513. 53. Chen D, et al. ORF3a of SARS-CoV-2 promotes lysosomal exocytosis-mediated viral egress. Dev Cell 2021;56:3250–3263.e5. doi: 10.1016/j.devcel.2021.10.006. 54. Cheng YW, et al. D614G substitution of SARS-CoV-2 spike protein increases syncytium formation and virus titer via enhanced furinmediated spike cleavage. MBio 2021;12:e0058721. 55. Chien M, et al. Nucleotide analogues as inhibitors of SARS-CoV-2 polymerase, a key drug target for COVID-19. J Proteome Res 2020;19:4690–4697. 56. Chu CM, et al. Role of lopinavir/ritonavir in the treatment of SARS: initial virological and clinical findings. Thorax 2004;59:252–256. 57. Clark LK, Green TJ, Petit CM. Structure of nonstructural protein 1 from SARS-CoV-2. J Virol 2021;95. 58. Claverie JM. A putative role of de-Mono-ADP-Ribosylation of STAT1 by the SARS-CoV-2 Nsp3 protein in the cytokine storm syndrome of COVID-19. Viruses 2020;12. 59. Clementz MA, et al. Deubiquitinating and interferon antagonism activities of coronavirus papain-like proteases. J Virol 2010;84:4619–4629. 60. Cong Y, et al. Nucleocapsid protein recruitment to replication-transcription complexes plays a crucial role in coronaviral life cycle. J Virol 2020;94. 61. Cornillez-Ty CT, Liao L, Yates JR III, et al. Severe acute respiratory syndrome coronavirus nonstructural protein 2 interacts with a host protein complex involved in mitochondrial biogenesis and intracellular signaling. J Virol 2009;83:10314–10318. 62. Coronaviridae Study Group of the International Committee on Taxonomy of Viruses. The species Severe acute respiratory syndromerelated coronavirus: classifying 2019-nCoV and naming it SARS-CoV-2. Nat Microbiol 2020;5:536–544. 63. Coutard B, et al. The spike glycoprotein of the new coronavirus 2019-nCoV contains a furin-like cleavage site absent in CoV of the same clade. Antiviral Res 2020;176:104742. 64. Coutinho RM, et al. Model-based estimation of transmissibility and reinfection of SARS-CoV-2 P.1 variant. Commun Med 2021;1(48). 65. Daffis S, et al. 2′-O methylation of the viral mRNA cap evades host restriction by IFIT family members. Nature 2010;468:452–456. 66. Davies JP, Almasy KM, McDonald EF, et al. Comparative multiplexed interactomics of SARS-CoV-2 and homologous coronavirus nonstructural proteins identifies unique and shared host-cell dependencies. ACS Infect Dis 2020;6:3174–3189. 67. Davies NG, et al. Estimated transmissibility and impact of SARS-CoV-2 lineage B.1.1.7 in England. Science 2021;372:eabg3055. 68. de Haan CA, Kuo L, Masters PS, et al. Coronavirus particle assembly: primary structure requirements of the membrane protein. J Virol 1998;72:6838–6850. 69. de Haan CA, Rottier PJ. Molecular interactions in the assembly of coronaviruses. Adv Virus Res 2005;64:165–230. 70. Decroly E, et al. Coronavirus nonstructural protein 16 is a cap-0 binding enzyme possessing (nucleoside-2′O)-methyltransferase activity. J Virol 2008;82:8071–8084. 71. Decroly E, et al. Crystal structure and functional analysis of the SARS-coronavirus RNA cap 2′-O-methyltransferase nsp10/nsp16 complex. PLoS Pathog 2011;7:e1002059. 72. DeDiego ML, et al. A severe acute respiratory syndrome coronavirus that lacks the E gene is attenuated in vitro and in vivo. J Virol 2007;81:1701–1713. 73. DeDiego ML, et al. Coronavirus virulence genes with main focus on SARS-CoV envelope gene. Virus Res 2014;194:124–137. 74. Deming DJ, Graham RL, Denison MR, et al. Processing of open reading frame 1a replicase proteins nsp7 to nsp10 in murine hepatitis virus strain A59 replication. J Virol 2007;81:10280–10291. 75. Deng X, et al. Structure-guided mutagenesis alters deubiquitinating activity and attenuates pathogenesis of a murine coronavirus. J Virol 2020;94:e01734. 76. Deng X, et al. Transmission, infectivity, and antibody neutralization of an emerging SARS-CoV-2 variant in California carrying a L452R spike protein mutation. medRxiv 2021;2021.03.07.21252647. doi: 10.1101/2021.03.07.21252647. 77. Denison MR, et al. The putative helicase of the coronavirus mouse hepatitis virus is processed from the replicase gene polyprotein and localizes in complexes that are active in viral RNA synthesis. J Virol 1999;73:6862–6871. 78. Devaraj SG, et al. Regulation of IRF-3-dependent innate immunity by the papain-like protease domain of the severe acute respiratory syndrome coronavirus. J Biol Chem 2007;282:32208–32221. 79. Donaldson EF, Sims AC, Graham RL, et al. Murine hepatitis virus replicase protein nsp10 is a critical regulator of viral RNA synthesis. J Virol 2007;81:6356–6368. 80. Dong B, Silverman RH. 2-5A-dependent RNase molecules dimerize during activation by 2-5A. J Biol Chem 1995;270:4133–4137. 81. Drexler JF, et al. Genomic characterization of severe acute respiratory syndrome-related coronavirus in european bats and classification of coronaviruses based on partial RNA-dependent RNA polymerase gene sequences. J Virol 2010;84:11336–11349. 82. Drosten C, et al. Identification of a novel coronavirus in patients with severe acute respiratory syndrome. N Engl J Med 2003;348:1967–1976. 83. Echavarria-Consuegra L, et al. Manipulation of the unfolded protein response: a pharmacological strategy against coronavirus infection. PLoS Pathog 2021;17:e1009644. 84. Eckerle LD, Lu X, Sperry SM, et al. High fidelity of murine hepatitis virus replication is decreased in nsp14 exoribonuclease mutants. J Virol 2007;81:12135–12144. 85. Eckerle LD, et al. Infidelity of SARS-CoV Nsp14-exonuclease mutant virus replication is revealed by complete genome sequencing. PLoS Pathog 2010;6:e1000896. 86. Edara VV, et al. Infection and mRNA-1273 vaccine antibodies neutralize SARS-CoV-2 UK variant. medRxiv 2021;2021.02.02.21250799. 87. Edara VV, et al. Infection- and vaccine-induced antibody binding and neutralization of the B.1.351 SARS-CoV-2 variant. Cell Host Microbe 2021;29:516–521.e513. 821

88. Egloff MP, et al. Structural and functional basis for ADP-ribose and poly(ADP-ribose) binding by viral macro domains. J Virol 2006;80:8493–8502. 89. Fagre A, et al. SARS-CoV-2 infection, neuropathogenesis and transmission among deer mice: implications for spillback to New World rodents. PLoS Pathog 2021;17:e1009585. 90. Fan X, Cao D, Kong L, et al. Cryo-EM analysis of the post-fusion structure of the SARS-CoV spike glycoprotein. Nat Commun 2020;11:3618. 91. Faria NR, et al. Genomics and epidemiology of the P.1 SARS-CoV-2 lineage in Manaus, Brazil. Science 2021;372:815–821. doi: 10.1126/science.abh2644, eabh2644. 92. Fehr AR, et al. The conserved coronavirus macrodomain promotes virulence and suppresses the innate immune response during severe acute respiratory syndrome coronavirus infection. MBio 2016;7. 93. Ferron F, Decroly E, Selisko B, et al. The viral RNA capping machinery as a target for antiviral drugs. Antiviral Res 2012;96:21–31. 94. Ferron F, et al. Structural and molecular basis of mismatch correction and ribavirin excision from coronavirus RNA. Proc Natl Acad Sci U S A 2018;115:E162–E171. 95. Filipowicz W, et al. A protein binding the methylated 5′-terminal sequence, m7GpppN, of eukaryotic messenger RNA. Proc Natl Acad Sci U S A 1976;73:1559–1563. 96. Freuling CM, et al. Susceptibility of raccoon dogs for experimental SARS-CoV-2 infection. Emerg Infect Dis 2020;26:2982–2985. 97. Freundt EC, et al. The open reading frame 3a protein of severe acute respiratory syndrome-associated coronavirus promotes membrane rearrangement and cell death. J Virol 2010;84:1097–1109. 98. Frieman M, Ratia K, Johnston RE, et al. Severe acute respiratory syndrome coronavirus papain-like protease ubiquitin-like domain and catalytic domain regulate antagonism of IRF3 and NF-kappaB signaling. J Virol 2009;83:6689–6705. 99. Frieman M, et al. Severe acute respiratory syndrome coronavirus ORF6 antagonizes STAT1 function by sequestering nuclear import factors on the rough endoplasmic reticulum/Golgi membrane. J Virol 2007;81:9812–9824. 100. Fu L, et al. Both boceprevir and GC376 efficaciously inhibit SARS-CoV-2 by targeting its main protease. Nat Commun 2020;11:4417. 101. Gao X, et al. Crystal structure of SARS-CoV-2 papain-like protease. Acta Pharm Sin B 2021;11:237–245. 102. Ge XY, et al. Isolation and characterization of a bat SARS-like coronavirus that uses the ACE2 receptor. Nature 2013;503:535–538. 103. Ge XY, et al. Coexistence of multiple coronaviruses in several bat colonies in an abandoned mineshaft. Virol Sin 2016;31:31–40. 104. Gerna G, et al. Coronaviruses and gastroenteritis: evidence of antigenic relatedness between human enteric coronavirus strains and human coronavirus OC43. Microbiologica 1984;7:315–322. 105. Ghosh S, et al. β-Coronaviruses use lysosomes for egress instead of the biosynthetic secretory pathway. Cell 2020;183:1520–1535.e1514. 106. Glowacka I, et al. Evidence that TMPRSS2 activates the severe acute respiratory syndrome coronavirus spike protein for membrane fusion and reduces viral control by the humoral immune response. J Virol 2011;85:4122–4134. 107. Goldstein SA, Brown J, Pedersen BS, et al. Extensive recombination-driven coronavirus diversification expands the pool of potential pandemic pathogens. bioRxiv 2021;10.1101/2021.02.03.429646. 108. Gomez GN, Abrar F, Dodhia MP, et al. SARS coronavirus protein nsp1 disrupts localization of Nup93 from the nuclear pore complex. Biochem Cell Biol 2019;97:758–766. 109. Gorbalenya AB, Baric S, de Groot R, et al. The species severe acute respiratory syndrome-related coronavirus: classifying 2019-nCoV and naming it SARS-CoV-2. Nat Microbiol 2020;5:536–544. 110. Gorbalenya AE, Koonin EV, Donchenko AP, et al. Coronavirus genome: prediction of putative functional domains in the non-structural polyprotein by comparative amino acid sequence analysis. Nucleic Acids Res 1989;17:4847–4861. 111. Gordon CJ, Tchesnokov EP, Feng JY, et al. The antiviral compound remdesivir potently inhibits RNA-dependent RNA polymerase from Middle East respiratory syndrome coronavirus. J Biol Chem 2020;295:4773–4779. 112. Gordon DE, et al. Comparative host-coronavirus protein interaction networks reveal pan-viral disease mechanisms. Science 2020;370. 113. Gordon DE, et al. A SARS-CoV-2 protein interaction map reveals targets for drug repurposing. Nature 2020;583:459–468. 114. Gordon CJ, et al. Remdesivir is a direct-acting antiviral that inhibits RNA-dependent RNA polymerase from severe acute respiratory syndrome coronavirus 2 with high potency. J Biol Chem 2020;295:6785–6797. 115. Gosert R, Kanjanahaluethai A, Egger D, et al. RNA replication of mouse hepatitis virus takes place at double-membrane vesicles. J Virol 2002;76:3697–3708. 116. Graham RL, Baric RS. Recombination, reservoirs, and the modular spike: mechanisms of coronavirus cross-species transmission. J Virol 2010;84:3134–3146. 117. Graham RL, Sims AC, Brockway SM, et al. The nsp2 replicase proteins of murine hepatitis virus and severe acute respiratory syndrome coronavirus are dispensable for viral replication. J Virol 2005;79:13399–13411. 118. Graham RL, et al. A live, impaired-fidelity coronavirus vaccine protects in an aged, immunocompromised mouse model of lethal disease. Nat Med 2012;18:1820–1826. 119. Greaney AJ, et al. The SARS-CoV-2 mRNA-1273 vaccine elicits more RBD-focused neutralization, but with broader antibody binding within the RBD. bioRxiv 2021;2021.04.14.439844. 120. Greaney AJ, et al. Comprehensive mapping of mutations in the SARS-CoV-2 receptor-binding domain that affect recognition by polyclonal human plasma antibodies. Cell Host Microbe 2021;29:463–476.e466. 121. Gribble J, et al. The coronavirus proofreading exoribonuclease mediates extensive viral recombination. PLoS Pathog 2021;17:e1009226. 122. Griffin BD, et al. SARS-CoV-2 infection and transmission in the North American deer mouse. Nat Commun 2021;12:3612. 123. Guan Y, et al. Isolation and characterization of viruses related to the SARS coronavirus from animals in Southern China. Science 2003;302:276–278. 124. Günther S, et al. X-ray screening identifies active site and allosteric inhibitors of SARS-CoV-2 main protease. Science 2021;372:642–646. 125. Guo H, et al. Evolutionary arms race between virus and host drives genetic diversity in bat severe acute respiratory syndrome-related coronavirus spike genes. J Virol 2020;94. 126. Hackbart M, Deng X, Baker SC. Coronavirus endoribonuclease targets viral polyuridine sequences to evade activating host sensors. Proc Natl Acad Sci U S A 2020;117:8094–8103. 127. Hagemeijer MC, Vonk AM, Monastyrska I, et al. Visualizing coronavirus RNA synthesis in time by using click chemistry. J Virol 2012;86:5808–5816. 128. Hagemeijer MC, et al. Dynamics of coronavirus replication-transcription complexes. J Virol 2010;84:2134–2149. 129. Hagemeijer MC, et al. Mobility and interactions of coronavirus nonstructural protein 4. J Virol 2011;85:4572–4577. 130. Hagemeijer MC, et al. Membrane rearrangements mediated by coronavirus nonstructural proteins 3 and 4. Virology 2014;458–459:125–135. 131. Hale VL, et al. SARS-CoV-2 infection in free-ranging white-tailed deer (Odocoileus virginianus). bioRxiv 2021. Nature volume 602, pages 481–486 (2022). 132. Hamer SA, et al. SARS-CoV-2 infections and viral isolations among serially tested cats and dogs in households with infected owners in Texas, USA. Viruses 2021;13. 822

133. Hao W, et al. Crystal structure of Middle East respiratory syndrome coronavirus helicase. PLoS Pathog 2017;13:e1006474. 134. Heald-Sargent T, Gallagher T. Ready, set, fuse! The coronavirus spike protein and acquisition of fusion competence. Viruses 2012;4:557–580. 135. Heusipp G, Harms U, Siddell SG, et al. Identification of an ATPase activity associated with a 71-kilodalton polypeptide encoded in gene 1 of the human coronavirus 229E. J Virol 1997;71:5631–5634. 136. Hoffmann M, Kleine-Weber H, Pöhlmann S. A multibasic cleavage site in the spike protein of SARS-CoV-2 is essential for infection of human lung cells. Mol Cell 2020;78:779–784.e5. doi: 10.1016/j.molcel.2020.04.022. 137. Hoffmann M, et al. SARS-CoV-2 cell entry depends on ACE2 and TMPRSS2 and is blocked by a clinically proven protease inhibitor. Cell 2020;181:271–280.e278. 138. Hoffmann M, et al. SARS-CoV-2 variants B.1.351 and P.1 escape from neutralizing antibodies. Cell 2021;184:2384–2393.e23. doi: 10.1016/j.cell.2021.03.036. 139. Hoffmann M, et al. SARS-CoV-2 mutations acquired in mink reduce antibody-mediated neutralization. Cell Rep 2021;35:109017. 140. Hogue BG, Machamer CM. Coronavirus structural proteins and assembly. In: Perlman SG, ed. The Nidoviruses. Washington, DC: American Society for Microbiology Press; 2008:179–200. 141. Hosie MJ, et al. Anthropogenic infection of cats during the 2020 COVID-19 pandemic. Viruses 2021;13. 142. Hsu JC, Laurent-Rolle M, Pawlak JB, et al. Translational shutdown and evasion of the innate immune response by SARS-CoV-2 NSP14 protein. Proc Natl Acad Sci U S A 2021;118. 143. Hu B, et al. Discovery of a rich gene pool of bat SARS-related coronaviruses provides new insights into the origin of SARS coronavirus. PLoS Pathog 2017;13:e1006698. 144. Huang C, Ito N, Tseng CT, et al. Severe acute respiratory syndrome coronavirus 7a accessory protein is a viral structural protein. J Virol 2006;80:7287–7294. 145. Huang Q, et al. Structure of the N-terminal RNA-binding domain of the SARS CoV nucleocapsid protein. Biochemistry 2004;43:6059–6063. 146. Huang C, et al. SARS coronavirus nsp1 protein induces template-dependent endonucleolytic cleavage of mRNAs: viral mRNAs are resistant to nsp1-induced RNA cleavage. PLoS Pathog 2011;7:e1002433. 147. Huang C, et al. Alphacoronavirus transmissible gastroenteritis virus nsp1 protein suppresses protein translation in mammalian cells and in cell-free HeLa cell extracts but not in rabbit reticulocyte lysate. J Virol 2011;85:638–643. 148. Hul V, et al. A novel SARS-CoV-2 related coronavirus in bats from Cambodia. Nat Commun 2021;12(6563). 149. Hur S. Double-stranded RNA sensors and modulators in innate immunity. Annu Rev Immunol 2019;37:349–375. 150. Hurst KR, Koetzner CA, Masters PS. Characterization of a critical interaction between the coronavirus nucleocapsid protein and nonstructural protein 3 of the viral replicase-transcriptase complex. J Virol 2013;87:9159–9172. 151. Imbert I, et al. A second, non-canonical RNA-dependent RNA polymerase in SARS coronavirus. Embo J 2006;25:4933–4942. 152. Imbert I, et al. The SARS-Coronavirus PLnc domain of nsp3 as a replication/transcription scaffolding protein. Virus Res 2008;133:136–148. 153. Issa E, Merhi G, Panossian B, et al. SARS-CoV-2 and ORF3a: nonsynonymous mutations, functional domains, and viral pathogenesis. mSystems 2020;5. 154. Ito N, et al. Severe acute respiratory syndrome coronavirus 3a protein is a viral structural protein. J Virol 2005;79:3182–3186. 155. Ivanov KA, et al. Multiple enzymatic activities associated with severe acute respiratory syndrome coronavirus helicase. J Virol 2004;78:5619–5632. 156. Ivanov KA, et al. Major genetic marker of nidoviruses encodes a replicative endoribonuclease. Proc Natl Acad Sci U S A 2004;101:12694–12699. 157. Jackson CB, Farzan M, Chen B, et al. Mechanisms of SARS-CoV-2 entry into cells. Nat Rev Mol Cell Biol 2022;23:3–20. doi: 10.1038/s41580-021-00418-x, 1–18. 158. Jauregui AR, Savalia D, Lowry VK, et al. Identification of residues of SARS-CoV nsp1 that differentially affect inhibition of gene expression and antiviral signaling. PLoS One 2013;8:e62416. 159. Javorsky A, Humbert PO, Kvansakul M. Structural basis of coronavirus E protein interactions with human PALS1 PDZ domain. Commun Biol 2021;4:724. 160. Jia Z, et al. Delicate structural coordination of the severe acute respiratory syndrome coronavirus Nsp13 upon ATP hydrolysis. Nucleic Acids Res 2019;47:6538–6550. 161. Jiang HW, et al. SARS-CoV-2 Orf9b suppresses type I interferon responses by targeting TOM70. Cell Mol Immunol 2020;17:998–1000. 162. Jimenez-Guardeno JM, et al. The PDZ-binding motif of severe acute respiratory syndrome coronavirus envelope protein is a determinant of viral pathogenesis. PLoS Pathog 2014;10:e1004320. 163. Jimenez-Guardeno JM, et al. Identification of the mechanisms causing reversion to virulence in an attenuated SARS-CoV for the design of a genetically stable vaccine. PLoS Pathog 2015;11:e1005215. 164. Jin Z, et al. Structure of Mpro from SARS-CoV-2 and discovery of its inhibitors. Nature 2020;582:289–293. 165. Joseph JS, et al. Crystal structure of nonstructural protein 10 from the severe acute respiratory syndrome coronavirus reveals a novel fold with two zinc-binding motifs. J Virol 2006;80:7894–7901. 166. Jungreis I, et al. Conflicting and ambiguous names of overlapping ORFs in the SARS-CoV-2 genome: a homology-based resolution. Virology 2021;558:145–151. 167. Kamitani W, Huang C, Narayanan K, et al. A two-pronged strategy to suppress host protein synthesis by SARS coronavirus Nsp1 protein. Nat Struct Mol Biol 2009;16:1134–1140. 168. Kamitani W, et al. Severe acute respiratory syndrome coronavirus nsp1 protein suppresses host gene expression by promoting host mRNA degradation. Proc Natl Acad Sci U S A 2006;103:12885–12890. 169. Kang R, Tang D. PKR-dependent inflammatory signals. Sci Signal 2012;5:pe47. 170. Kang H, et al. Biochemical and genetic analyses of murine hepatitis virus Nsp15 endoribonuclease. J Virol 2007;81:13587–13597. 171. Kang L, et al. A selective sweep in the Spike gene has driven SARS-CoV-2 human adaptation. Cell 2021;184:4392–4400.e4. 172. Kapikian AZ. The coronaviruses. Dev Biol Stand 1975;28:42–64. 173. Kasprzyk R, et al. Identification and evaluation of potential SARS-CoV-2 antiviral agents targeting mRNA cap guanine N7Methyltransferase. Antiviral Res 2021;193:105142. 174. Ke Z, et al. Structures and distributions of SARS-CoV-2 spike proteins on intact virions. Nature 2020;588:498–502. 175. Kim Y, et al. Broad-spectrum antivirals against 3C or 3C-like proteases of picornaviruses, noroviruses, and coronaviruses. J Virol 2012;86:11754–11762. 176. Kim Y, et al. Broad-spectrum inhibitors against 3C-like proteases of feline coronaviruses and feline caliciviruses. J Virol 2015;89:4942–4950. 177. Kim Y, et al. Reversal of the progression of fatal coronavirus infection in cats by a broad-spectrum coronavirus protease inhibitor. PLoS Pathog 2016;12:e1005531. 823

178. Kim YI, et al. Infection and rapid transmission of SARS-CoV-2 in ferrets. Cell Host Microbe 2020;27:704–709. 179. Kindler E, et al. Early endonuclease-mediated evasion of RNA sensing ensures efficient coronavirus replication. PLoS Pathog 2017;13:e1006195. 180. Kirchdoerfer RN, Ward AB. Structure of the SARS-CoV nsp12 polymerase bound to nsp7 and nsp8 co-factors. Nat Commun 2019;10:2342. 181. Kirchdoerfer RN, et al. Pre-fusion structure of a human coronavirus spike protein. Nature 2016;531:118–121. 182. Klein S, et al. SARS-CoV-2 structure and replication characterized by in situ cryo-electron tomography. Nat Commun 2020;11:5885. 183. Klemm T, et al. Mechanism and inhibition of the papain-like protease, PLpro, of SARS-CoV-2. EMBO J 2020;39:e106275. 184. Klumperman J, et al. Coronavirus M proteins accumulate in the Golgi complex beyond the site of virion budding. J Virol 1994;68:6523–6534. 185. Kokic G, et al. Mechanism of SARS-CoV-2 polymerase stalling by remdesivir. Nat Commun 2021;12:279. 186. Konno Y, et al. SARS-CoV-2 ORF3b is a potent interferon antagonist whose activity is increased by a naturally occurring elongation variant. Cell Rep 2020;32:108185. 187. Korber B, et al. Tracking changes in SARS-CoV-2 spike: evidence that D614G increases infectivity of the COVID-19 virus. Cell 2020;182:812–827.e819. 188. Krupovic M, Koonin EV. Multiple origins of viral capsid proteins from cellular ancestors. Proc Natl Acad Sci U S A 2017;114:E2401– E2410. 189. Kuri T, et al. The ADP-ribose-1”-monophosphatase domains of severe acute respiratory syndrome coronavirus and human coronavirus 229E mediate resistance to antiviral interferon responses. J Gen Virol 2011;92:1899–1905. 190. Kusov Y, Tan J, Alvarez E, et al. A G-quadruplex-binding macrodomain within the “SARS-unique domain” is essential for the activity of the SARS-coronavirus replication-transcription complex. Virology 2015;484:313–322. 191. Kustin T, et al. Evidence for increased breakthrough rates of SARS-CoV-2 variants of concern in BNT162b2 mRNA vaccinated individuals. Nat Med 2021;27:1379–1384. 192. Lam TTY, et al. Identifying SARS-CoV-2-related coronaviruses in Malayan pangolins. Nature 2020;583:282–285. 193. Lan J, et al. Structure of the SARS-CoV-2 spike receptor-binding domain bound to the ACE2 receptor. Nature 2020;581:215–220. 194. Lapointe CP, et al. Dynamic competition between SARS-CoV-2 NSP1 and mRNA on the human ribosome inhibits translation initiation. Proc Natl Acad Sci U S A 2021;118. 195. Larsen HD, et al. Preliminary report of an outbreak of SARS-CoV-2 in mink and mink farmers associated with community spread, Denmark, June to November 2020. Euro Surveill 2021;26. 196. Latinne A, et al. Origin and cross-species transmission of bat coronaviruses in China. Nat Commun 2020;11:4235. 197. Lau SK, et al. Severe acute respiratory syndrome coronavirus-like virus in Chinese horseshoe bats. Proc Natl Acad Sci U S A 2005;102:14040–14045. 198. Lau SK, et al. Coronavirus HKU1 and other coronavirus infections in Hong Kong. J Clin Microbiol 2006;44:2063–2071. 199. Lau SKP, et al. Severe acute respiratory syndrome (SARS) coronavirus ORF8 protein is acquired from SARS-related coronavirus from greater horseshoe bats through recombination. J Virol 2015;89:10532–10547. 200. Lee N, et al. A major outbreak of severe acute respiratory syndrome in Hong Kong. N Engl J Med 2003;348:1986–1994. 201. Lee S, et al. Genetic characteristics of coronaviruses from Korean bats in 2016. Microb Ecol 2018;75:174–182. 202. Lee J, et al. No evidence of coronaviruses or other potentially zoonotic viruses in sunda pangolins (Manis javanica) entering the wildlife trade via Malaysia. Ecohealth 2020;17:406–418. 203. Lehmann KC, Snijder EJ, Posthuma CC, et al. What we know but do not understand about nidovirus helicases. Virus Res 2015;202:12–32. 204. Lehmann KC, et al. Discovery of an essential nucleotidylating activity associated with a newly delineated conserved domain in the RNA polymerase-containing protein of all nidoviruses. Nucleic Acids Res 2015;43:8416–8434. 205. Lei J, Kusov Y, Hilgenfeld R. Nsp3 of coronaviruses: structures and functions of a large multi-domain protein. Antiviral Res 2018;149:58–74. 206. Lei X, et al. Activation and evasion of type I interferon responses by SARS-CoV-2. Nat Commun 2020;11:3810. 207. Letko M, Marzi A, Munster V. Functional assessment of cell entry and receptor usage for SARS-CoV-2 and other lineage B betacoronaviruses. Nat Microbiol 2020;5:562–569. 208. Li F. Structure, function, and evolution of coronavirus spike proteins. Annu Rev Virol 2016;3:237–261. 209. Li W, et al. Angiotensin-converting enzyme 2 is a functional receptor for the SARS coronavirus. Nature 2003;426:450–454. 210. Li W, et al. Bats are natural reservoirs of SARS-like coronaviruses. Science 2005;310:676–679. 211. Li Y, et al. Activation of RNase L is dependent on OAS3 expression during infection with diverse human viruses. Proc Natl Acad Sci U S A 2016;113:2241–2246. 212. Li S, et al. Regulation of the ER stress response by the ion channel activity of the infectious bronchitis coronavirus envelope protein modulates virion release, apoptosis, viral fitness, and pathogenesis. Front Microbiol 2019;10:3022. 213. Li Y, et al. SARS-CoV-2 induces double-stranded RNA-mediated innate immune responses in respiratory epithelial-derived cells and cardiomyocytes. Proc Natl Acad Sci U S A 2021;118. 214. Lim KP, Liu DX. The missing link in coronavirus assembly. Retention of the avian coronavirus infectious bronchitis virus envelope protein in the pre-Golgi compartments and physical interaction between the envelope and membrane proteins. J Biol Chem 2001;276:17515–17523. 215. Lin S, et al. Crystal structure of SARS-CoV-2 nsp10 bound to nsp14-ExoN domain reveals an exoribonuclease with both structural and functional integrity. Nucleic Acids Res 2021;49:5382–5392. 216. Lindner HA, et al. The papain-like protease from the severe acute respiratory syndrome coronavirus is a deubiquitinating enzyme. J Virol 2005;79:15199–15208. 217. Littler DR, Gully BS, Colson RN, et al. Crystal structure of the SARS-CoV-2 non-structural protein 9, Nsp9. iScience 2020;23:101258. 218. Liu C, et al. The architecture of inactivated SARS-CoV-2 with postfusion spikes revealed by Cryo-EM and Cryo-ET. Structure 2020;28:1218–1224.e1214. 219. Liu Y, et al. The development of Coronavirus 3C-Like protease (3CL(pro)) inhibitors from 2010 to 2020. Eur J Med Chem 2020;206:112711. 220. Liu C, et al. Structural basis of mismatch recognition by a SARS-CoV-2 proofreading enzyme. Science 2021;373:1142–1146. 221. Liu C, et al. Reduced neutralization of SARS-CoV-2 B.1.617 by vaccine and convalescent serum. Cell 2021;184:4220–4236.e13. doi: 10.1016/j.cell.2021.06.020. 222. Liu J, et al. BNT162b2-elicited neutralization of B.1.617 and other SARS-CoV-2 variants. Nature 2021;596:273–275. doi: 10.1038/s41586-021-03693-y. 223. Locker JK, et al. The cytoplasmic tail of mouse hepatitis virus M protein is essential but not sufficient for its retention in the Golgi complex. J Biol Chem 1994;269:28263–28269. 824

224. Lokugamage KG, Narayanan K, Huang C, et al. Severe acute respiratory syndrome coronavirus protein nsp1 is a novel eukaryotic translation inhibitor that represses multiple steps of translation initiation. J Virol 2012;86:13598–13608. 225. Lopez LA, Riffle AJ, Pike SL, et al. Importance of conserved cysteine residues in the coronavirus envelope protein. J Virol 2008;82:3000–3010. 226. Lopusna K, et al. Interferons lambda, new cytokines with antiviral activity. Acta Virol 2013;57:171–179. 227. Lu R, et al. Genomic characterisation and epidemiology of 2019 novel coronavirus: implications for virus origins and receptor binding. Lancet 2020;395:565–574. doi: 10.1016/s0140-6736(20)30251-8. 228. Lugari A, et al. Molecular mapping of the RNA Cap 2′-O-methyltransferase activation interface between severe acute respiratory syndrome coronavirus nsp10 and nsp16. J Biol Chem 2010;285:33230–33241. 229. Ma Z, Pourfarjam Y, Kim IK. Reconstitution and functional characterization of SARS-CoV-2 proofreading complex. Protein Expr Purif 2021;185:105894. 230. Ma Y, et al. Structural basis and functional analysis of the SARS coronavirus nsp14-nsp10 complex. Proc Natl Acad Sci U S A 2015;112:9436–9441. 231. Machamer CE. Accommodation of large cargo within Golgi cisternae. Histochem Cell Biol 2013;140:261–269. 232. MacLean OA, et al. Natural selection in the evolution of SARS-CoV-2 in bats created a generalist virus and highly capable human pathogen. PLoS Biol 2021;19:e3001115. 233. Maio N, et al. Fe-S cofactors in the SARS-CoV-2 RNA-dependent RNA polymerase are potential antiviral targets. Science 2021;373:236–241. 234. Malathi K, Dong B, Gale M Jr, et al. Small self-RNA generated by RNase L amplifies antiviral innate immunity. Nature 2007;448:816–819. 235. Ma-Lauer Y, et al. p53 down-regulates SARS coronavirus replication and is targeted by the SARS-unique domain and PLpro via E3 ubiquitin ligase RCHY1. Proc Natl Acad Sci U S A 2016;113:E5192–E5201. 236. Malone B, et al. Structural basis for backtracking by the SARS-CoV-2 replication-transcription complex. Proc Natl Acad Sci U S A 2021;118. 237. Mandala VS, et al. Structure and drug binding of the SARS-CoV-2 envelope protein transmembrane domain in lipid bilayers. Nat Struct Mol Biol 2020;27:1202–1208. 238. Matsuyama S, Taguchi F. Two-step conformational changes in a coronavirus envelope glycoprotein mediated by receptor binding and proteolysis. J Virol 2009;83:11133–11141. 239. Matsuyama S, et al. Efficient activation of the severe acute respiratory syndrome coronavirus spike protein by the transmembrane protease TMPRSS2. J Virol 2010;84:12658–12664. 240. McAloose D, et al. From people to panthera: natural SARS-CoV-2 infection in tigers and lions at the bronx zoo. MBio 2020;11. 241. McBride CE, Li J, Machamer CE. The cytoplasmic tail of the severe acute respiratory syndrome coronavirus spike protein contains a novel endoplasmic reticulum retrieval signal that binds COPI and promotes interaction with membrane protein. J Virol 2007;81:2418–2428. 242. McBride CE, Machamer CE. A single tyrosine in the severe acute respiratory syndrome coronavirus membrane protein cytoplasmic tail is important for efficient interaction with spike protein. J Virol 2010;84:1891–1901. 243. McBride R, van Zyl M, Fielding BC. The coronavirus nucleocapsid is a multifunctional protein. Viruses 2014;6:2991–3018. 244. Menachery VD. MERS vaccine candidate offers promise, but questions remain. EBioMedicine 2015;2:1292–1293. 245. Menachery VD, et al. Attenuation and restoration of severe acute respiratory syndrome coronavirus mutant lacking 2′-omethyltransferase activity. J Virol 2014;88:4251–4264. 246. Menachery VD, et al. Corrigendum: a SARS-like cluster of circulating bat coronaviruses shows potential for human emergence. Nat Med 2016;22:446. 247. Menachery VD, et al. Middle East respiratory syndrome coronavirus nonstructural protein 16 is necessary for interferon resistance and viral pathogenesis. mSphere 2017;2. 248. Michalska K, et al. Crystal structures of SARS-CoV-2 ADP-ribose phosphatase: from the apo form to ligand complexes. IUCrJ 2020;7:814–824. 249. Michel CJ, Mayer C, Poch O, et al. Characterization of accessory genes in coronavirus genomes. Virol J 2020;17:131. 250. Miknis ZJ, et al. Severe acute respiratory syndrome coronavirus nsp9 dimerization is essential for efficient viral growth. J Virol 2009;83:3007–3018. 251. Milewska A, et al. Entry of human coronavirus NL63 into the Cell. J Virol 2018;92. 252. Millet JK, Whittaker GR. Host cell proteases: critical determinants of coronavirus tropism and pathogenesis. Virus Res 2015;202:120–134. 253. Minasov G, et al. Mn(2+) coordinates Cap-0-RNA to align substrates for efficient 2′-O-methyl transfer by SARS-CoV-2 nsp16. Sci Signal 2021;14. 254. Minskaia E, et al. Discovery of an RNA virus 3′->5′ exoribonuclease that is critically involved in coronavirus RNA synthesis. Proc Natl Acad Sci U S A 2006;103:5108–5113. 255. Miorin L, et al. SARS-CoV-2 Orf6 hijacks Nup98 to block STAT nuclear import and antagonize interferon signaling. Proc Natl Acad Sci 2020;117:28344–28354. doi: 10.1073/pnas.2016650117, 202016650. 256. Moustaqil M, et al. SARS-CoV-2 proteases PLpro and 3CLpro cleave IRF3 and critical modulators of inflammatory pathways (NLRP12 and TAB1): implications for disease presentation across species. Emerg Microbes Infect 2021;10:178–195. 257. Moyo-Gwete T, et al. Cross-reactive neutralizing antibody responses elicited by SARS-CoV-2 501Y.V2 (B.1.351). N Engl J Med 2021;384:2161–2163. doi: 10.1056/nejmc2104192. 258. Munster VJ, et al. Respiratory disease in rhesus macaques inoculated with SARS-CoV-2. Nature 2020;585:268–272. 259. Murakami S, et al. Detection and characterization of bat sarbecovirus phylogenetically related to SARS-CoV-2, Japan. Emerg Infect Dis 2020;26:3025–3029. 260. Murray RS, MacMillan B, Cabirac G, et al. Detection of coronavirus RNA in CNS tissue of multiple sclerosis and control patients. Adv Exp Med Biol 1990;276:505–510. 261. Muth D, et al. Attenuation of replication by a 29 nucleotide deletion in SARS-coronavirus acquired during the early stages of human-tohuman transmission. Sci Rep 2018;8:15177. 262. Mykytyn AZ, et al. SARS-CoV-2 entry into human airway organoids is serine protease-mediated and facilitated by the multibasic cleavage site. Elife 2021;10. 263. Nakagawa K, Makino S. Mechanisms of coronavirus Nsp1-mediated control of host and viral gene expression. Cells 2021;10:300. 264. Nal B, et al. Differential maturation and subcellular localization of severe acute respiratory syndrome coronavirus surface proteins S, M and E. J Gen Virol 2005;86:1423–1434. 265. Narayanan K, et al. Severe acute respiratory syndrome coronavirus nsp1 suppresses host gene expression, including that of type I interferon, in infected cells. J Virol 2008;82:4471–4479. 266. Nelson CA, Pekosz A, Lee CA, et al. Structure and intracellular targeting of the SARS-coronavirus Orf7a accessory protein. Structure 825

2005;13:75–85. 267. Nemudryi A, et al. SARS-CoV-2 genomic surveillance identifies naturally occurring truncation of ORF7a that limits immune suppression. Cell Rep 2021;35:109197. 268. Neuman BW, et al. Proteomics analysis unravels the functional repertoire of coronavirus nonstructural protein 3. J Virol 2008;82:5279–5294. 269. Ni X, et al. Structural insights into plasticity and discovery of remdesivir metabolite GS-441524 binding in SARS-CoV-2 macrodomain. ACS Med Chem Lett 2021;12:603–609. 270. Nieto-Torres JL, et al. Subcellular location and topology of severe acute respiratory syndrome coronavirus envelope protein. Virology 2011;415:69–82. 271. Nieva JL, Madan V, Carrasco L. Viroporins: structure and biological functions. Nat Rev Microbiol 2012;10:563–574. 272. Niu S, et al. Molecular basis of cross-species ACE2 interactions with SARS-CoV-2-like viruses of pangolin origin. EMBO J 2021;40:e107786. doi: 10.15252/embj.2021107786, e107786. 273. Ogando NS, et al. The enzymatic activity of the nsp14 exoribonuclease is critical for replication of MERS-CoV and SARS-CoV-2. J Virol 2020;94. 274. Oostra M, de Haan CA, de Groot RJ, et al. Glycosylation of the severe acute respiratory syndrome coronavirus triple-spanning membrane proteins 3a and M. J Virol 2006;80:2326–2336. 275. Oostra M, de Haan CA, Rottier PJ. The 29-nucleotide deletion present in human but not in animal severe acute respiratory syndrome coronaviruses disrupts the functional expression of open reading frame 8. J Virol 2007;81:13876–13888. 276. Oostra M, et al. Localization and membrane topology of coronavirus nonstructural protein 4: involvement of the early secretory pathway in replication. J Virol 2007;81:12323–12336. 277. Oostra M, et al. Topology and membrane anchoring of the coronavirus replication complex: not all hydrophobic domains of nsp3 and nsp6 are membrane spanning. J Virol 2008;82:12392–12405. 278. Ortego J, Escors D, Laude H, et al. Generation of a replication-competent, propagation-deficient virus vector based on the transmissible gastroenteritis coronavirus genome. J Virol 2002;76:11518–11529. 279. Oudshoorn D, et al. Expression and cleavage of middle east respiratory syndrome coronavirus nsp3-4 polyprotein induce the formation of double-membrane vesicles that mimic those associated with coronaviral RNA replication. MBio 2017;8. 280. Owen DR, et al. An oral SARS-CoV-2 Mpro inhibitor clinical candidate for the treatment of COVID-19. Science 2021;374:1586–1593. doi: 10.1126/science.abl4784. 281. Painter GR, Natchus MG, Cohen O, et al. Developing a direct acting, orally available antiviral agent in a pandemic: the evolution of molnupiravir as a potential treatment for COVID-19. Curr Opin Virol 2021;50:17–22. 282. Parker MM, Masters PS. Sequence comparison of the N genes of five strains of the coronavirus mouse hepatitis virus suggests a three domain structure for the nucleocapsid protein. Virology 1990;179:463–468. 283. Parthasarathy K, et al. Structural flexibility of the pentameric SARS coronavirus envelope protein ion channel. Biophys J 2008;95:L39– L41. 284. Peacock TP, et al. The furin cleavage site in the SARS-CoV-2 spike protein is required for transmission in ferrets. Nat Microbiol 2021;6:899–909. doi: 10.1038/s41564-021-00908-w. 285. Peiris JS, et al. Coronavirus as a possible cause of severe acute respiratory syndrome. Lancet 2003;361:1319–1325. 286. Perlman SM. Coronaviridae: the viruses and their replication. In: Howley PMK, Whelan DM, eds. Fields Virology. Vol. 1, Chap. 10. Philadelphia, PA: Wolters Kluwer; 2020:410–421. 287. Perlman S, Netland J. Coronaviruses post-SARS: update on replication and pathogenesis. Nat Rev Microbiol 2009;7:439–450. 288. Perrier A, et al. The C-terminal domain of the MERS coronavirus M protein contains a trans-Golgi network localization signal. J Biol Chem 2019;294:14406–14421. 289. Pervushin K, et al. Structure and inhibition of the SARS coronavirus envelope protein ion channel. PLoS Pathog 2009;5:e1000511. 290. Peti W, et al. Structural genomics of the severe acute respiratory syndrome coronavirus: nuclear magnetic resonance structure of the protein nsP7. J Virol 2005;79:12905–12913. 291. Pfefferle S, et al. Distant relatives of severe acute respiratory syndrome coronavirus and close relatives of human coronavirus 229E in bats, Ghana. Emerg Infect Dis 2009;15:1377–1384. 292. Pfefferle S, et al. Reverse genetic characterization of the natural genomic deletion in SARS-Coronavirus strain Frankfurt-1 open reading frame 7b reveals an attenuating function of the 7b protein in-vitro and in-vivo. Virol J 2009;6:131. 293. Plante JA, et al. Spike mutation D614G alters SARS-CoV-2 fitness. Nature 2021;592:116–121. 294. Platanias LC. Mechanisms of type-I- and type-II-interferon-mediated signalling. Nat Rev Immunol 2005;5:375–386. 295. Plescia CB, et al. SARS-CoV-2 viral budding and entry can be modeled using BSL-2 level virus-like particles. J Biol Chem 2021;296:100103. 296. Pohl MO, et al. SARS-CoV-2 variants reveal features critical for replication in primary human cells. PLoS Biol 2021;19:e3001006. 297. Prentice E, McAuliffe J, Lu X, et al. Identification and characterization of severe acute respiratory syndrome coronavirus replicase proteins. J Virol 2004;78:9977–9986. 298. Pruijssers AJ, et al. Remdesivir inhibits SARS-CoV-2 in human lung cells and chimeric SARS-CoV expressing the SARS-CoV-2 RNA polymerase in mice. Cell Rep 2020;32:107940. 299. Pyrc K, Jebbink MF, Berkhout B, et al. Genome structure and transcriptional regulation of human coronavirus NL63. Virol J 2004;1:7. 300. Rack JGM, et al. Viral macrodomains: a structural and evolutionary assessment of the pharmacological potential. Open Biol 2020;10:200237. 301. Rathnayake AD, et al. 3C-like protease inhibitors block coronavirus replication in vitro and improve survival in MERS-CoV-infected mice. Sci Transl Med 2020;12. 302. Ratia K, Kilianski A, Baez-Santos YM, et al. Structural basis for the ubiquitin-linkage specificity and deISGylating activity of SARS-CoV papain-like protease. PLoS Pathog 2014;10:e1004113. 303. Ratia K, et al. Severe acute respiratory syndrome coronavirus papain-like protease: structure of a viral deubiquitinating enzyme. Proc Natl Acad Sci U S A 2006;103:5717–5722. 304. Ratia K, et al. A noncovalent class of papain-like protease/deubiquitinase inhibitors blocks SARS virus replication. Proc Natl Acad Sci U S A 2008;105:16119–16124. 305. Ren W, et al. Difference in receptor usage between severe acute respiratory syndrome (SARS) coronavirus and SARS-like coronavirus of bat origin. J Virol 2008;82:1899–1907. 306. Rogstam A, et al. Crystal structure of non-structural protein 10 from severe acute respiratory syndrome coronavirus-2. Int J Mol Sci 2020;21. 307. Rosas-Lemus M, et al. High-resolution structures of the SARS-CoV-2 2′-O-methyltransferase reveal strategies for structure-based inhibitor design. Sci Signal 2020;13. 308. Rosenke K, et al. Defining the Syrian hamster as a highly susceptible preclinical model for SARS-CoV-2 infection. Emerg Microbes Infect 826

2020;9:2673–2684. 309. Roth-Cross JK, Bender SJ, Weiss SR. Murine coronavirus mouse hepatitis virus is recognized by MDA5 and induces type I interferon in brain macrophages/microglia. J Virol 2008;82:9829–9838. 310. Rottier PJ. The coronavirus membrane glycoprotein. In: Siddell SG, ed. The Coronaviridae. New York: Plenum Press; 1995:115–139. 311. Ruch TR, Machamer CE. The coronavirus e protein: assembly and beyond. Viruses 2012;4:363–382. 312. Sacramento CQ, et al. In vitro antiviral activity of the anti-HCV drugs daclatasvir and sofosbuvir against SARS-CoV-2, the aetiological agent of COVID-19. J Antimicrob Chemother 2021;76:1874–1885. 313. Sadler AJ, Williams BR. Interferon-inducible antiviral effectors. Nat Rev Immunol 2008;8:559–568. 314. Saikatendu KS, et al. Structural basis of severe acute respiratory syndrome coronavirus ADP-ribose-1”-phosphate dephosphorylation by a conserved domain of nsP3. Structure 2005;13:1665–1675. 315. Sawicki SG, et al. Functional and genetic analysis of coronavirus replicase-transcriptase proteins. PLoS Pathog 2005;1:e39. 316. Schaecher SR, Diamond MS, Pekosz A. The transmembrane domain of the severe acute respiratory syndrome coronavirus ORF7b protein is necessary and sufficient for its retention in the Golgi complex. J Virol 2008;82:9477–9491. 317. Schaecher SR, Touchette E, Schriewer J, et al. Severe acute respiratory syndrome coronavirus gene 7 products contribute to virus-induced apoptosis. J Virol 2007;81:11054–11068. 318. Schoeman D, Fielding BC. Coronavirus envelope protein: current knowledge. Virol J 2019;16:69. 319. Schoeman D, Fielding BC. Is there a link between the pathogenic human coronavirus envelope protein and immunopathology? A review of the literature. Front Microbiol 2020;11:2086. 320. Schubert K, et al. SARS-CoV-2 Nsp1 binds the ribosomal mRNA channel to inhibit translation. Nat Struct Mol Biol 2020;27:959–966. doi: 10.1038/s41594-020-0511-8. 321. Semper C, Watanabe N, Savchenko A. Structural characterization of nonstructural protein 1 from SARS-CoV-2. iScience 2021;24:101903. 322. Serrano P, et al. Nuclear magnetic resonance structure of the N-terminal domain of nonstructural protein 3 from the severe acute respiratory syndrome coronavirus. J Virol 2007;81:12049–12060. 323. Serrano P, et al. Nuclear magnetic resonance structure of the nucleic acid-binding domain of severe acute respiratory syndrome coronavirus nonstructural protein 3. J Virol 2009;83:12998–13008. 324. Seybert A, Hegyi A, Siddell SG, et al. The human coronavirus 229E superfamily 1 helicase has RNA and DNA duplex-unwinding activities with 5′-to-3′ polarity. RNA 2000;6:1056–1068. 325. Shang J, et al. Cell entry mechanisms of SARS-CoV-2. Proc Natl Acad Sci U S A 2020;117:11727–11734. 326. Shang J, et al. Structural basis of receptor recognition by SARS-CoV-2. Nature 2020;581:221–224. 327. Shannon A, et al. Rapid incorporation of Favipiravir by the fast and permissive viral RNA polymerase complex results in SARS-CoV-2 lethal mutagenesis. Nat Commun 2020;11:4682. 328. Sheahan TP, et al. Broad-spectrum antiviral GS-5734 inhibits both epidemic and zoonotic coronaviruses. Sci Transl Med 2017;9. 329. Sheahan TP, et al. An orally bioavailable broad-spectrum antiviral inhibits SARS-CoV-2 in human airway epithelial cell cultures and multiple coronaviruses in mice. Sci Transl Med 2020;12:eabb5883. 330. Shen S, et al. The severe acute respiratory syndrome coronavirus 3a is a novel structural protein. Biochem Biophys Res Commun 2005;330:286–292. 331. Shi J, Sivaraman J, Song J. Mechanism for controlling the dimer-monomer switch and coupling dimerization to catalysis of the severe acute respiratory syndrome coronavirus 3C-like protease. J Virol 2008;82:4620–4629. 332. Shi CS, et al. SARS-coronavirus open reading frame-9b suppresses innate immunity by targeting mitochondria and the MAVS/TRAF3 /TRAF6 signalosome. J Immunol 2014;193:3080–3089. 333. Shi M, et al. SARS-CoV-2 Nsp1 suppresses host but not viral translation through a bipartite mechanism. bioRxiv 2020;2020.09.18.302901. doi: 10.1101/2020.09.18.302901. 334. Shin D, et al. Papain-like protease regulates SARS-CoV-2 viral spread and innate immunity. Nature 2020;587:657–662. 335. Shriner SA, et al. SARS-CoV-2 exposure in escaped mink, Utah, USA. Emerg Infect Dis 2021;27:988–990. 336. Shulla A, et al. A transmembrane serine protease is linked to the severe acute respiratory syndrome coronavirus receptor and activates virus entry. J Virol 2011;85:873–882. 337. Silvas J, et al. Contribution of SARS-CoV-2 accessory proteins to viral pathogenicity in K18 hACE2 transgenic mice. bioRxiv 2021. doi: 10.1101/2021.03.09.434696, 2021.2003.2009.434696. 338. Simmons G, Zmora P, Gierer S, et al. Proteolytic activation of the SARS-coronavirus spike protein: cutting enzymes at the cutting edge of antiviral research. Antiviral Res 2013;100:605–614. 339. Simmons B, et al. Sofosbuvir/daclatasvir regimens for the treatment of COVID-19: an individual patient data meta-analysis. J Antimicrob Chemother 2021;76:286–291. 340. Sims AC, Burkett SE, Yount B, et al. SARS-CoV replication and pathogenesis in an in vitro model of the human conducting airway epithelium. Virus Res 2008;133:33–44. 341. Sims AC, et al. Release of severe acute respiratory syndrome coronavirus nuclear import block enhances host transcription in human lung cells. J Virol 2013;87:3885–3902. 342. Slanina H, et al. Coronavirus replication-transcription complex: vital and selective NMPylation of a conserved site in nsp9 by the NiRANRdRp subunit. Proc Natl Acad Sci U S A 2021;118. 343. Smith EC, Blanc H, Surdel MC, et al. Coronaviruses lacking exoribonuclease activity are susceptible to lethal mutagenesis: evidence for proofreading and potential therapeutics. PLoS Pathog 2013;9:e1003565. 344. Snijder EJ, Decroly E, Ziebuhr J. The nonstructural proteins directing coronavirus RNA synthesis and processing. Adv Virus Res 2016;96:59–126. 345. Snijder EJ, et al. Unique and conserved features of genome and proteome of SARS-coronavirus, an early split-off from the coronavirus group 2 lineage. J Mol Biol 2003;331:991–1004. 346. Sola I, Almazan F, Zuniga S, et al. Continuous and discontinuous RNA synthesis in coronaviruses. Annu Rev Virol 2015;2:265–288. 347. Sparks JS, Lu X, Denison MR. Genetic analysis of murine hepatitis virus nsp4 in virus replication. J Virol 2007;81:12554–12563. 348. Starr TN, et al. Deep mutational scanning of SARS-CoV-2 receptor binding domain reveals constraints on folding and ACE2 binding. Cell 2020;182:1295–1310. 349. Stertz S, et al. The intracellular sites of early replication and budding of SARS-coronavirus. Virology 2007;361:304–315. 350. Su YAO, et al. Discovery and genomic characterization of a 382-nucleotide deletion in ORF7b and ORF8 during the early evolution of SARS-CoV-2. MBio 2020;114:e01610. doi: 10.1128/mBio.01610-20. 351. Subissi L, et al. One severe acute respiratory syndrome coronavirus protein complex integrates processive RNA polymerase and exonuclease activities. Proc Natl Acad Sci U S A 2014;111:E3900–E3909. 352. Surjit M, Liu B, Kumar P, et al. The nucleocapsid protein of the SARS coronavirus is capable of self-association through a C-terminal 209 amino acid interaction domain. Biochem Biophys Res Commun 2004;317:1030–1036. 353. Surya W, Li Y, Torres J. Structural model of the SARS coronavirus E channel in LMPG micelles. Biochim Biophys Acta Biomembr 827

2018;1860:1309–1317. 354. Sutton G, et al. The nsp9 replicase protein of SARS-coronavirus, structure and functional insights. Structure 2004;12:341–353. 355. Tan J, et al. The SARS-unique domain (SUD) of SARS coronavirus contains two macrodomains that bind G-quadruplexes. PLoS Pathog 2009;5:e1000428. 356. Tanaka T, Kamitani W, DeDiego ML, et al. Severe acute respiratory syndrome coronavirus nsp1 facilitates efficient propagation in cells through a specific translational shutoff of host mRNA. J Virol 2012;86:11128–11137. 357. Tchesnokov EP, et al. Template-dependent inhibition of coronavirus RNA-dependent RNA polymerase by remdesivir reveals a second mechanism of action. J Biol Chem 2020;295:16156–16165. 358. te Velthuis AJ, Arnold JJ, Cameron CE, et al. The RNA polymerase activity of SARS-coronavirus nsp12 is primer dependent. Nucleic Acids Res 2010;38:203–214. 359. te Velthuis AJ, van den Worm SH, Snijder EJ. The SARS-coronavirus nsp7+nsp8 complex is a unique multimeric RNA polymerase capable of both de novo initiation and primer extension. Nucleic Acids Res 2012;40:1737–1747. 360. Tegally H, et al. Emergence and rapid spread of a new severe acute respiratory syndrome-related coronavirus 2 (SARS-CoV-2) lineage with multiple spike mutations in South Africa. medRxiv 2020. https://doi.org/10.1101/2020.12.21.20248640. 361. Temmam S, et al. Bat coronaviruses related to SARS-CoV-2 and infectious for human cells. Nature 2021. doi: 10.21203/rs.3.rs871965/v1. https://doi.org/10.1038/s41586-022-04532-4. 362. Teoh KT, et al. The SARS coronavirus E protein interacts with PALS1 and alters tight junction formation and epithelial morphogenesis. Mol Biol Cell 2010;21:3838–3852. 363. Thoms M, et al. Structural basis for translational shutdown and immune evasion by the Nsp1 protein of SARS-CoV-2. Science 2020;369:1249–1255. 364. Tidu A, et al. The viral protein NSP1 acts as a ribosome gatekeeper for shutting down host translation and fostering SARS-CoV-2 translation. RNA 2020;27:253–264. 365. Tong P, et al. Memory B cell repertoire for recognition of evolving SARS-CoV-2 spike. Cell 2021;184:4969–4980.e4915. 366. Tooze J, Tooze SA, Fuller SD. Sorting of progeny coronavirus from condensed secretory proteins at the exit from the trans-Golgi network of AtT20 cells. J Cell Biol 1987;105:1215–1226. 367. Tortorici MA, Veesler D. Structural insights into coronavirus entry. Adv Virus Res 2019;105:93–116. 368. Toto A, et al. Comparing the binding properties of peptides mimicking the envelope protein of SARS-CoV and SARS-CoV-2 to the PDZ domain of the tight junction-associated PALS1 protein. Protein Sci 2020;29:2038–2042. 369. Tsang KW, et al. A cluster of cases of severe acute respiratory syndrome in Hong Kong. N Engl J Med 2003;348:1977–1985. 370. Tseng CT, et al. Apical entry and release of severe acute respiratory syndrome-associated coronavirus in polarized Calu-3 lung epithelial cells. J Virol 2005;79:9470–9479. 371. Turoňová B, et al. In situ structural analysis of SARS-CoV-2 spike reveals flexibility mediated by three hinges. Science 2020;370:203–208. 372. Ujike M, Taguchi F. Incorporation of spike and membrane glycoproteins into coronavirus virions. Viruses 2015;7:1700–1725. 373. van Dorp L, et al. Emergence of genomic diversity and recurrent mutations in SARS-CoV-2. Infect Genet Evol 2020;83:104351. 374. Vandyck K, Deval J. Considerations for the discovery and development of 3-chymotrypsin-like cysteine protease inhibitors targeting SARS-CoV-2 infection. Curr Opin Virol 2021;49:36–40. 375. Vasilenko N, Moshynskyy I, Zakhartchouk A. SARS coronavirus protein 7a interacts with human Ap4A-hydrolase. Virol J 2010;7:31. 376. Venkatagopalan P, Daskalova SM, Lopez LA, et al. Coronavirus envelope (E) protein remains at the site of assembly. Virology 2015;478:75–85. 377. Verheije MH, et al. The coronavirus nucleocapsid protein is dynamically associated with the replication-transcription complexes. J Virol 2010;84:11575–11579. 378. Viswanathan T, et al. Structural basis of RNA cap modification by SARS-CoV-2. Nat Commun 2020;11:3718. 379. V’Kovski P, Kratzel A, Steiner S, et al. Coronavirus biology and replication: implications for SARS-CoV-2. Nat Rev Microbiol 2021;19:155–170. 380. von Brunn A, et al. Analysis of intraviral protein-protein interactions of the SARS coronavirus ORFeome. PLoS One 2007;2:e459. 381. von Grotthuss M, Wyrwicz LS, Rychlewski L. mRNA cap-1 methyltransferase in the SARS genome. Cell 2003;113:701–702. 382. Vuong W, et al. Feline coronavirus drug inhibits the main protease of SARS-CoV-2 and blocks virus replication. Nat Commun 2020;11:4282. 383. Wacharapluesadee S, et al. Evidence for SARS-CoV-2 related coronaviruses circulating in bats and pangolins in Southeast Asia. Nat Commun 2021;12:972. 384. Wahba L, et al. An extensive meta-metagenomic search identifies SARS-CoV-2-homologous sequences in pangolin lung viromes. mSphere 2020;5. 385. Wahl A, et al. SARS-CoV-2 infection is effectively treated and prevented by EIDD-2801. Nature 2021;591:451–457. 386. Walls AC, et al. Structure, function, and antigenicity of the SARS-CoV-2 spike glycoprotein. Cell 2020;181:281–292.e286. 387. Wan Y, Shang J, Graham R, et al. Receptor recognition by the novel coronavirus from Wuhan: an analysis based on decade-long structural studies of SARS Coronavirus. J Virol 2020;94. 388. Wang R, Chen J, Hozumi Y, et al. Decoding asymptomatic COVID-19 infection and transmission. J Phys Chem Lett 2020;11:10007–10015. 389. Wang H, Pipes L, Nielsen R. Synonymous mutations and the molecular evolution of SARS-CoV-2 origins. Virus Evol 2021;7. 390. Wang Y, et al. Coronavirus nsp10/nsp16 methyltransferase can be targeted by nsp10-derived peptide in vitro and in vivo to reduce replication and pathogenesis. J Virol 2015;89:8416–8427. 391. Wang M, et al. Remdesivir and chloroquine effectively inhibit the recently emerged novel coronavirus (2019-nCoV) in vitro. Cell Res 2020;30:269–271. 392. Wang X, et al. Accurate diagnosis of COVID-19 by a novel immunogenic secreted SARS-CoV-2 orf8 Protein. MBio 2020;11. 393. Washington NL, et al. Emergence and rapid transmission of SARS-CoV-2 B.1.1.7 in the United States. Cell 2021;184:2587–2594.e7. doi: 10.1016/j.cell.2021.03.052. 394. Watanabe Y, Allen JD, Wrapp D, et al. Site-specific glycan analysis of the SARS-CoV-2 spike. Science 2020;369:330–333. 395. Wathelet MG, Orr M, Frieman MB, et al. Severe acute respiratory syndrome coronavirus evades antiviral signaling: role of nsp1 and rational design of an attenuated strain. J Virol 2007;81:11620–11633. 396. Wells HL, et al. The evolutionary history of ACE2 usage within the coronavirus subgenus Sarbecovirus. Virus Evol 2021;7:veab007. 397. Whelan JN, Li Y, Silverman RH, et al. Zika virus production is resistant to RNase L antiviral activity. J Virol 2019;93. 398. Whittaker GR, Daniel S, Millet JK. Coronavirus entry: how we arrived at SARS-CoV-2. Curr Opin Virol 2021;47:113–120. 399. Wilamowski M, et al. 2′-O methylation of RNA cap in SARS-CoV-2 captured by serial crystallography. Proc Natl Acad Sci U S A 2021;118. 400. Williamson BN, et al. Clinical benefit of remdesivir in rhesus macaques infected with SARS-CoV-2. Nature 2020;585:273–276. 401. Wong NA, Saier MH Jr. The SARS-Coronavirus infection cycle: a survey of viral membrane proteins, their functional interactions and pathogenesis. Int J Mol Sci 2021;22. 828

402. Wrapp D, et al. Cryo-EM structure of the 2019-nCoV spike in the prefusion conformation. Science 2020;367:1260–1263. 403. Wu CY, et al. Small molecules targeting severe acute respiratory syndrome human coronavirus. Proc Natl Acad Sci U S A 2004;101:10012–10017. 404. Wu F, et al. A new coronavirus associated with human respiratory disease in China. Nature 2020;579:265–269. 405. Xia H, et al. Evasion of type-I interferon by SARS-CoV-2. Cell Rep 2020;10:8234. 406. Xia B, et al. SARS-CoV-2 envelope protein causes acute respiratory distress syndrome (ARDS)-like pathological damages and constitutes an antiviral target. Cell Res 2021;31:847–860. 407. Xiao X, Newman C, Buesching CD, et al. Animal sales from Wuhan wet markets immediately prior to the COVID-19 pandemic. Sci Rep 2021;11:11898. 408. Xiao K, et al. Isolation of SARS-CoV-2-related coronavirus from Malayan pangolins. Nature 2020;583:286–289. 409. Xu X, et al. Molecular model of SARS coronavirus polymerase: implications for biochemical functions and drug design. Nucleic Acids Res 2003;31:7117–7130. 410. Yan R, et al. Structural basis for the recognition of SARS-CoV-2 by full-length human ACE2. Science 2020;367:1444–1448. 411. Yan L, et al. Architecture of a SARS-CoV-2 mini replication and transcription complex. Nat Commun 2020;11:5874. 412. Yan L, et al. Cryo-EM structure of an extended SARS-CoV-2 replication and transcription complex reveals an intermediate state in cap synthesis. Cell 2021;184:184–193.e110. 413. Yan L, et al. Coupling of N7-methyltransferase and 3′-5′ exoribonuclease with SARS-CoV-2 polymerase reveals mechanisms for capping and proofreading. Cell 2021;184:3474–3485.e3411. 414. Yang H, et al. The crystal structures of severe acute respiratory syndrome virus main protease and its complex with an inhibitor. Proc Natl Acad Sci U S A 2003;100:13190–13195. 415. Yang H, et al. Design of wide-spectrum inhibitors targeting coronavirus main proteases. PLoS Biol 2005;3:e324. 416. Yang N, et al. Bismuth complexes inhibit the SARS coronavirus. Angew Chem Int Ed Engl 2007;46:6464–6468. 417. Yang Y, et al. The structural and accessory proteins M, ORF 4a, ORF 4b, and ORF 5 of Middle East respiratory syndrome coronavirus (MERS-CoV) are potent interferon antagonists. Protein Cell 2013;4:951–961. 418. Yang XL, et al. Isolation and characterization of a novel bat coronavirus closely related to the direct progenitor of severe acute respiratory syndrome coronavirus. J Virol 2016;90:3253–3256. 419. Yao H, et al. Molecular architecture of the SARS-CoV-2 virus. Cell 2020;183:730–738.e713. 420. Ye Q, West AMV, Silletti S, et al. Architecture and self-assembly of the SARS-CoV-2 nucleocapsid protein. Protein Sci 2020;29:1890–1901. 421. Yin W, et al. Structural basis for inhibition of the RNA-dependent RNA polymerase from SARS-CoV-2 by remdesivir. Science 2020;368:1499–1504. 422. Young BE, et al. Effects of a major deletion in the SARS-CoV-2 genome on the severity of infection and the inflammatory response: an observational cohort study. Lancet 2020;396:603–611. 423. Yuan S, et al. Nonstructural protein 1 of SARS-CoV-2 is a potent pathogenicity factor redirecting host protein synthesis machinery toward viral RNA. Mol Cell 2020;80:1055–1066.e1056. 424. Yuan S, et al. Metallodrug ranitidine bismuth citrate suppresses SARS-CoV-2 replication and relieves virus-associated pneumonia in Syrian hamsters. Nat Microbiol 2020;5:1439–1448. 425. Yurkovetskiy L, et al. Structural and functional analysis of the D614G SARS-CoV-2 spike protein variant. Cell 2020;183:739–751.e738. 426. Zeng HL, Dichio V, Rodríguez Horta E, et al. Global analysis of more than 50,000 SARS-CoV-2 genomes reveals epistasis between eight viral genes. Proc Natl Acad Sci U S A 2020;117:31519–31526. 427. Zeng LP, et al. Bat severe acute respiratory syndrome-like coronavirus WIV1 encodes an extra accessory protein, ORFX, involved in modulation of the host immune response. J Virol 2016;90:6573–6582. 428. Zhai Y, et al. Insights into SARS-CoV transcription and replication from the structure of the nsp7-nsp8 hexadecamer. Nat Struct Mol Biol 2005;12:980–986. 429. Zhang L, et al. SARS-CoV-2 spike-protein D614G mutation increases virion spike density and infectivity. Nat Commun 2020;11:6013. 430. Zhang L, et al. Crystal structure of SARS-CoV-2 main protease provides a basis for design of improved α-ketoamide inhibitors. Science 2020;368:409–412. 431. Zhang Y, et al. The ORF8 protein of SARS-CoV-2 mediates immune evasion through potently downregulating MHC-I. PNAS 2021;118(23):e2024202118. 432. Zhang J, et al. Structural impact on SARS-CoV-2 spike protein by D614G substitution. Science 2021;372:525–530. 433. Zhang K, et al. Nsp1 protein of SARS-CoV-2 disrupts the mRNA export machinery to inhibit host gene expression. Sci Adv 2021;7. 434. Zhao Z, et al. Description and clinical treatment of an early outbreak of severe acute respiratory syndrome (SARS) in Guangzhou, PR China. J Med Microbiol 2003;52:715–720. 435. Zhao P, et al. Virus-receptor interactions of glycosylated SARS-CoV-2 spike and human ACE2 receptor. Cell Host Microbe 2020;28:586–601.e586. 436. Zhao Y, et al. Crystal structure of SARS-CoV-2 main protease in complex with protease inhibitor PF-07321332. Protein Cell 2021. doi: 10.1007/s13238-021-00883-2. 437. Zheng M, et al. Bat SARS-Like WIV1 coronavirus uses the ACE2 of multiple animal species as receptor and evades IFITM3 restriction via TMPRSS2 activation of membrane fusion. Emerg Microbes Infect 2020;9:1567–1579. 438. Zhou A, et al. Interferon action and apoptosis are defective in mice devoid of 2′,5′-oligoadenylate-dependent RNase L. EMBO J 1997;16:6355–6363. 439. Zhou P, et al. A pneumonia outbreak associated with a new coronavirus of probable bat origin. Nature 2020;579:270–273. 440. Zhou H, et al. A novel bat coronavirus closely related to SARS-CoV-2 contains natural insertions at the S1/S2 cleavage site of the spike protein. Curr Biol 2020;30:2196–2203.e2193. 441. Zhou H, et al. Identification of novel bat coronaviruses sheds light on the evolutionary origins of SARS-CoV-2 and related viruses. Cell 2021;184:4380–4391. 442. Zhu N, et al. Morphogenesis and cytopathic effect of SARS-CoV-2 infection in human airway epithelial cells. Nat Commun 2020;11:3910. 443. Ziebuhr J, Snijder EJ, Gorbalenya AE. Virus-encoded proteinases and proteolytic processing in the Nidovirales. J Gen Virol 2000;81:853–879. 444. Zúñiga S, et al. Coronavirus nucleocapsid protein facilitates template switching and is required for efficient transcription. J Virol 2010;84:2169–2175. 445. Zust R, et al. Ribose 2′-O-methylation provides a molecular signature for the distinction of self and non-self mRNA dependent on the RNA sensor Mda5. Nat Immunol 2011;12:137–143.

829

CHAPTER 22 SARS-CoV-2/COVID-19: Clinical Characteristics, Prevention, and Treatment John H. Beigel • Timothy M. Uyeki Introduction Pathogenesis Pathophysiology Pathology Immune responses to SARS-CoV-2 SARS-CoV-2 variant viruses Epidemiologic parameters Transmission dynamics Transmission modes Duration of infectiousness Clinical disease Severity of disease Spectrum of disease Clinical complications Respiratory complications Extrapulmonary complications Coinfections Laboratory findings Radiographic and imaging findings Risk factors for severe disease Special populations Children Pregnancy Vertical transmission Clinical issues after infection SARS-CoV-2 reinfection Post-COVID conditions Multisystem inflammatory syndrome Diagnostic testing Viral tests Serology Clinical management General Monoclonal antibodies Bamlanivimab/etesevimab Casirivimab/imdevimab 830

Sotrovimab Regdanvimab Variants and monoclonal antibodies Antiviral agents Remdesivir Other antiviral agents Anti-inflammatory Dexamethasone Baricitinib IL-6 pathway inhibitors Other anti-inflammatory agents Convalescent plasma Antithrombotic prophylaxis Ineffective therapies Treatment of COVID-19 Outpatients Hospitalized patients Infection prevention and control Vaccine Adverse events Myocarditis/pericarditis Thrombosis with thrombocytopenia Guillain-Barré syndrome Variant viruses Additional vaccine dose in immunocompromised Vaccine booster Perspective Addendum, April 2022 Epidemiology COVID-19 Vaccines Therapeutics

INTRODUCTION In late 2019, the emergence of a novel respiratory virus caused an unprecedented threat to public health and became a global pandemic. The causative virus was rapidly identified764 (later named severe acute respiratory syndrome coronavirus 2, or SARS-CoV-2) and descriptions of the clinical characteristics of the disease (referred to as COVID-19) and complications were quickly published.137,263,301,690 Within weeks, trials of candidate vaccines and investigational therapeutics began. Within months, there were emergency use authorizations issued in the United States for therapeutics and early approvals in other countries of accurate diagnostics, several efficacious therapeutics were identified in clinical trials, and increased understanding of the pathogenesis and sequelae of the disease were recognized. Within a year, there was authorization or initial approvals of highly effective vaccines worldwide. The emergence of viral variants could reduce the effectiveness of the vaccines and may impact some of the advances noted above in the future. This chapter summarizes the current understanding of clinical characteristics, prevention, and treatment of COVID-19, as of late 2021, primarily based upon findings that preceded emergence of the SARS-CoV-2 Delta (B.1.617.2) variant of concern, while noting the pandemic continues to evolve and the field continues to advance.

PATHOGENESIS 831

Pathophysiology Following host cell binding, the viral and cell membranes fuse, enabling virus entry into the cell444 followed by RNA genome replication and translation.79 SARS-CoV-2 infection induces cellular death via pyroptosis, a highly inflammatory form of programmed cell death commonly seen with cytopathic viruses.217 This process also causes the release of various damage-associated molecular patterns (DAMPs) and pathogenassociated molecular patterns (PAMPs). Pattern Recognition Receptors such as toll-like receptors (TLRs) recognize PAMPs and DAMPs triggering induction of proinflammatory cytokine transcription factors such as NF-κβ, as well as activating interferon regulatory factors that mediate the type I interferon–dependent antiviral response.391 In some patients, SARS-CoV-2 suppresses the early type I and III interferon responses, an immune evasion strategy employed by the virus, leading to early failure to control the virus.74

After SARS-CoV-2 infection, two waves of cellular responses occur. First, a rapid recruitment of monocytes and macrophages into the lungs occurs early after infection.344,373 Then T cells infiltrate into the lungs where they initiate a specific response to clear the virus.135 This results in an increase in proinflammatory cytokines and chemokines and the recruitment of immune cells into affected sites. Several cohort studies in SARS-CoV-2 have observed increased levels of interleukin-6 (IL-6), IL-2, granulocyte colony-stimulating factor (G-CSF), IL-7, IL-10, interferon (IFN)-inducible protein-10 (IP-10), monocyte chemoattractant protein-1 (MCP1), IFN-γ, macrophage inflammatory protein 1α (MIP1α), and tumor necrosis factor (TNF-α).301,650

FIGURE 22.1 Characterization of COVID-19 disease progression. The dark blue shading indicates physiological viral host response over time, and the dark red shading indicates a pathogenic hyperinflammatory host. (Reprinted with permission from Bohn MK, Hall A, Sepiashvili L, et al. Pathophysiology of COVID-19: mechanisms underlying disease severity and progression. Physiology (Bethesda). 2020;35(5):288–301. Copyright © 2020 The American Physiological Society.) In most individuals with SARS-CoV-2 infection, the cytokine release and activation of an antiviral interferon response followed by immune cell recruitment result in successful SARS-CoV-2 clearance. In a subset of patients, there is progression to a hyperinflammatory state that may manifest by organ dysfunction (Fig. 22.1). This hyperinflammatory state is sometimes called a cytokine storm, which is generally described as a collection of clinical manifestations resulting from an overactivated and dysregulated immune response. Cytokine storms are associated with various disorders, such as uncontrolled infectious diseases associated with certain acquired or inherited immunodeficiencies, autoinflammatory diseases, or following therapeutic interventions.135 Distinguishing an appropriate innate immune response triggered by viral infection from a dysregulated abnormal inflammatory response can be difficult, though the clinical benefit in modifying this innate immune response (described below in Treatment) suggests that the cytokine activation may contribute to pathogenesis.

Coagulation abnormalities and a high incidence of thrombotic events occur in COVID-19 patients. Early reports demonstrated prolonged activated partial thromboplastin time (aPTT), prothrombin time, and elevated d-dimer.137,690 A meta-analysis reported that the frequency of disseminated intravascular coagulation (DIC) was 3% in hospitalized COVID-19 patients, DIC was associated with greater severity of illness, and deaths.762 One study reported that 71.4% of nonsurvivors and 0.6% of recovered cases met the criteria for DIC during hospitalization.645 Autopsy data observed fibrin thrombi in pulmonary small arterial vessels in 87% of fatal cases.109 Complement-mediated pulmonary tissue damage and microvascular injury have been observed in severe COVID-19.433 Together, these data suggest that hypercoagulability in addition to hyperinflammation may contribute to the pathogenesis of severe COVID-19.

Pathology The primary pathology of COVID-19 in the lungs is diffuse alveolar damage (DAD), organizing pneumonia, reactive type II pneumocytes, and chronic interstitial pneumonia (Fig. 22.2A).103 There are additional reports of diffuse proteinaceous edema and hyaline membranes.363 Macroscopically, this can appear as very heavy wet lungs593 (Fig. 22.2B). There is also massive capillary congestion often accompanied by microthrombi despite anticoagulation.455 In the proliferative phase of DAD, there is an interstitial infiltrate of lymphocytes and florid, atypical type 2 pneumocyte hyperplasia (within the cytoplasm of which some authors have demonstrated viral inclusions/protein/RNA), sometimes associated with squamous metaplasia.593 Occasionally, the lung consolidation consists of intra-alveolar neutrophilic infiltration, consistent with superimposed bacterial bronchopneumonia,662 though only directly visualized or detected by culture or PCR in 8%.159 Severe tracheobronchitis including aphthous ulcers81 and mononuclear inflammation455 have been described.

Angiotensin converting enzyme 2 (ACE2) is abundantly present in human epithelial cells of the lung, as well as in endothelial cells of the arterial

832

and venous vessels,266 suggesting direct viral infection and injury of the vascular endothelium is possible. Postmortem studies have confirmed venous thromboembolic disease in the majority of patients that died (Fig. 22.2C).709 This has been attributed to increased levels of von Willebrand factor, Toll-like receptor activation, complement deposition, and tissue factor pathway activation.242,640

Histology demonstrated inflammation of the myocardium with predominance of macrophages, and myocyte necrosis has occasionally been reported on endomyocardial biopsy, generally performed after patients present with symptoms of myocarditis or heart failure.588 Autopsy series have demonstrated accumulation of inflammatory cells in the endocardium662 and myocardium.662 The early clinical hypothesis that atherosclerotic plaque rupture results in coronary artery thrombosis has not been supported by postmortem studies.593

FIGURE 22.2 Typical autopsy findings in severe COVID-19. A: Diffuse alveolar damage—congestion/proliferation phases, with lymphocytic infiltration of alveolar septa. B: Typical macroscopic appearance of COVID-19 lung. C: Thrombus in intramyocardial arteriole (arrowhead), with surrounding subacute microinfarct. (Reprinted from Sekhawat V, Green A, Mahadeva U. COVID-19 autopsies: conclusions from international studies. Diagn Histopathol (Oxf). 2021;27(3):103–107. Copyright © 2020 Elsevier. With permission.) The primary pathology findings in the liver are those reflective of underlying diseases such as obesity.593 While elevated liver function tests are seen in many patients, most autopsy series demonstrate nonspecific passive congestion with centrilobular necrosis and collapse. Kupffer cell activation and proliferation has rarely been reported,376 and it is unknown if this is responsible for liver dysfunction or simply reflective of the viremia. The primary findings in the kidneys are also reflective of preexisting diseases: diabetic and hypertensive nephropathy. Acute tubular injury and myoglobin casts have also been commonly reported.630 Noteworthy for its absence in most of the published studies has been glomerular capillary thrombi/evidence of thrombotic microangiopathy (TMA), diverging from the findings in typical DIC.593

In the few published case series examining whole brains, there are infrequent reports of widespread microthrombi, microinfarcts and microhemorrhages, global or watershed anoxic injury, and rarely a focal infiltrate of T lymphocytes and microglia.90,593 White pulp atrophy of the spleen has been commonly described.87,463 Lymphoid depletion has also been documented in lymph nodes, with some studies finding reactive plasmablastic proliferation463 and hemophagocytosis.90 Rarely, skeletal muscle may have mononuclear myositis associated with myocyte necrosis.463

Immune Responses to SARS-CoV-2 T-Cell Immunity Both T- and B-cell responses against SARS-CoV-2 are detected approximately 1 week after the onset of symptoms. The notion that activated T cells are key determinants of protection may explain the increased susceptibility of older individuals to severe COVID-19.282 Aging is associated with thymic involution, which depletes the potential to generate new T-cell repertoires. In contrast, T-cell repertoires are abundant in children which may explain their resistance to severe disease.366

CD8+ T cells are important for directly attacking and killing virus-infected cells, whereas CD4+ T cells are crucial to prime both CD8+ T cells and B cells.650 CD4+ T cells are also responsible for cytokine production to drive immune cell recruitment. After SARS-CoV-1 infection, the CD4+ T-cell response includes production of IFN-γ, TNF, and IL-2, suggesting a TH1 cell response to control the infection.318,610 Spike-reactive CD4+ T cells not only were detected in most infected individuals but also have been detected in up to 1/3 of uninfected unvaccinated controls.88 The SARS-CoV-2-reactive CD4+ T cells from healthy controls also responded to the spike proteins of human endemic coronaviruses 229E and OC43 suggesting a cross reactive response to the seasonal coronaviruses. These findings support the importance of CD4+ T cells in creating effective cross-reactive immune responses. It has been suggested that frequent exposure to endemic coronaviruses in children are part of the reason of decreased severity of disease in this population.

In patients with COVID-19, CD8+ T cells exhibiting activated phenotypes are commonly observed, although the absolute number of CD8+ T cells is decreased.561 SARS-CoV-2–specific CD8+ T-cell responses have been identified in most individuals after recovery from COVID-19.258 These responses are specific to a wide range of SARS-CoV-2 antigens, including spike, nucleocapsid, and membrane proteins, as well as other nonstructural proteins.258,377 In the acute phase of COVID-19, SARS-CoV-2–specific CD8+ T cells express activation markers (CD38 and HLADR), Ki-67, inhibitor checkpoint receptors (PD-1, TIM-3, and LAG-3), and cytotoxic proteins (perforin and granzyme B), indicating that these cells are activated and proliferate with a high cytotoxic capacity.594 However, an exhausted CD8+ T-cell phenotype with an up-regulation of inhibitory receptors such as PD-1, TIM-3, LAG-3, CTLA-4, NKG2A, and CD39, has been described in patients with COVID-19, particularly in those with severe disease.561,618 After recovery from the acute illness, SARS-CoV-2–specific T-cell responses are maintained in convalescent individuals up to 10 months postinfection, indicating that SARS-CoV-2–specific T-cell memory develops successfully and is long lasting.561 Depletion of CD8+ T cells in convalescent macaques partially abrogated the protective efficacy of natural immunity against rechallenge with SARS-CoV-2 suggesting cellular immunity is important for long-term protection.452

B-Cell Immunity B-cell activation and rapid production of antigen-specific antibodies are critical for the control of viral infections. B-cell responses occur concomitantly with T-cell responses, starting around 1 week after symptom onset.650 The seroconversion rate and antibody levels increased 833

rapidly during the first 2 weeks after symptom onset with the cumulative seropositive rate reaching 50% by day 11 and 100% by day 39.282,485,653 Moreover, the median seroconversion time of total antibody, IgM, and IgG were observed at day 11, 12, and 14, respectively.751 Poor antibody responses are associated with ineffective SARS-CoV-2 clearance in some patients.733 Adoptive transfer of purified IgG from convalescent rhesus macaques protects naive recipient macaques against challenge with SARS-CoV-2 in a dose-dependent fashion.452

After infection, there are high-level IgG, IgM, and IgA reactivity to the structural proteins—spike (S), matrix (M), and nucleocapsid (N) of SARSCoV-2, as well as accessory proteins such as ORF3a and ORF7a.106 Antibodies against the N protein typically arise first.643,719 Uninfected unvaccinated individuals can have antibodies that cross-react to SARS-CoV-2, likely from prior seasonal coronavirus, and are mainly targeted to N and the carboxy portion of the S (S2) proteins.222 Antibodies to the amino portion of S (S1) and the receptor binding domain (RBD) are more specific to SARS-CoV-2 infections. With acute infection, substantial populations of endemic human coronavirus reactive antibody-secreting cells expand signifying preexisting immunity though these antibodies were generally nonneutralizing and nonprotective in vivo.199 In a vaccinated population, the risk of symptomatic COVID-19 decreased with increasing levels of anti-spike IgG, anti-RBD IgG, pseudovirus neutralization, and live-virus neutralization titers.213 Correlates of protection have not been established after infection, but likely entail similar immunologic parameters.

Antibody responses tend to be higher in more severe cases, although there is considerable heterogeneity.429 Memory B cells persist or increase even as antibody levels wane.583 There have also been reports of B cells undergoing somatic hypermutation over the course of 6 months, consistent with persistent antigen.228 It is unknown if protection against COVID-19 is durable over years, given that protection against other circulating coronaviruses tends to be short lived, as seen by increased reinfection after 1 year.203,429 Modeling suggests that a loss in protection from SARS-CoV-2 infection may occur, while protection from severe disease including hospitalization and death should be largely retained.349

SARS-COV-2 VARIANT VIRUSES Genetic variants of SARS-CoV-2 have been emerging and circulating around the world throughout the COVID-19 pandemic. Variants of interest are those strains with specific genetic markers that have been associated with changes to receptor binding, reduced neutralization by antibodies generated against previous infection or vaccination, reduced efficacy of treatments, potential diagnostic impact, or predicted increase in transmissibility or disease severity.123 Variants being monitored include variants for which there are data indicating a potential or clear impact on approved or authorized medical countermeasures or that have been associated with more severe disease or increased transmission, but the variant viruses are no longer detected or are circulating at very low levels in the United States. Variants of Concern are those strains with evidence of an increase in transmissibility, more severe disease (e.g., increased hospitalizations or deaths), significant reduction in neutralization by antibodies generated during previous infection or vaccination, reduced effectiveness of treatments or vaccines, or diagnostic detection failures.123 The Delta (B.1.617.2) and Omicron (B.1.1.529) variants are currently classified as Variants of Concern. Alpha (B.1.1.7), Beta (B.1.351), Gamma (P.1), and others are currently considered variants being monitored.

Alpha (B.1.1.7) First identified in September 2020 in the United Kingdom. Includes sublineage with E484K mutation. Most recently has become a minority of isolates.245 Associated with higher viral concentrations in nasopharyngeal swabs compared to the original (wild-type) virus.178 Approximately 30% increased transmission relative to nonvariants.107 In postvaccination sera, exhibited a 1.7-fold reduction in pseudovirus neutralization and 1.3-fold reduction in live virus neutralization compared to wild-type (D614G).518 Risk of death was reported as 55% to 64% higher when compared to other strains.128,178

Beta (B.1.351) First identified in September 2020 in South Africa, but declined in 2021.245 Approximately 25% increased transmission relative to the original wild-type strain.107 In postvaccination sera, exhibited a 6.9-fold reduction in pseudovirus neutralization and 4.6-fold reduction in live virus neutralization compared to wild-type (D614G).518

Gamma (P.1) First identified in October 2020 in Brazil. Most recently has decreased in prevalence in much of the world but continues to widely circulate in South America.245 Approximately 40% increased transmission relative to the original wild-type strain.107 In postvaccination sera, exhibited a 3.2-fold reduction in pseudovirus neutralization compared to wild-type (D614G).518

Delta (B.1.617.2) First identified in September 2020 in India. Includes sublineages AY.1 to AY.12. This strain became widespread and the most common isolate in many countries during 2021.245 Approximately 100% increased transmission relative to the original wild-type strain.107 In postvaccination sera, exhibited a 2.4-fold reduction in pseudovirus neutralization compared to wild-type (D614G).518

Omicron (B.1.1.529) First identified in November 2021 in Botswana and reported to WHO on November 24, 2021. Contains mutations associated with reduced neutralization to some anti-SARS-CoV-2 monoclonal antibodies, increased transmissibility, and immune escape.104,715

Other variants being monitored, but have not widely circulated, include the following: 834

• Epsilon

B.1.427 and B.1.429

• Eta

B.1.525

• Iota

B.1.526

• Kappa

B.1.617.1

• Zeta

P.2

• Mu

B.1.621, B.1.621.1

As variant viruses have the potential to significantly evolve, the data in this chapter including epidemiology, transmission, pathogenesis, and efficacy of vaccines and therapeutics may change; thus, awareness of the circulating variants and their impact on the parameters above is critical to understanding and managing COVID-19.

EPIDEMIOLOGIC PARAMETERS Transmission Dynamics Based upon data from studies conducted when the original Wuhan SARS-CoV-2 strain was predominant, the incubation period for COVID-19 was estimated to be a median of 5.1 days (95% CI 4.5 to 5.8)374 and a mean of 5.2 days after SARS-CoV-2 infection (95% CI 4.1 to 7)399,744 with a range estimated in different populations and countries to be 4 to 7 days.17,205,345,447,700,748 A pooled analysis estimated that 97.5% of symptomatic illness occurs within 11.5 days (CI 8.2 to 15.6) of SARS-CoV-2 infection.374 Meta-analyses have estimated the serial interval (time from illness onset in an index case to illness onset in a secondary case) to be 5.2 to 5.4 days suggesting transmission occurs early after symptom onset.17,541,748 The pooled mean serial interval in China was 4.9 days (range 1.9 to 6.5) after the pandemic peak compared to 6.2 days (range 5.1 to 7.8) before the pandemic peak and may have been affected by differences in time to the isolation of index cases.19 The SARS-CoV-2 incubation period might vary in different variants. For example, an investigation of an outbreak of COVID-19 due to the Delta variant in mid-2021 in southern China reported a median incubation period of 4 days and a median serial interval of 3 days, both shorter than what has been reported for non-Delta SARS-CoV-2 infections.747

Secondary attack rates can vary by transmission settings (e.g., households, workplaces, social settings, health care facilities)660 and extent of prevention and control measures; secondary transmission is highest in households.427,660 One study estimated that susceptibility to SARSCoV-2 infection increased with older age and that most secondary and tertiary transmission occurred in households.299 A meta-analysis reported that the overall estimated secondary household attack rate for SARS-CoV-2 infection was 18.9% but was 24.5% for the Delta variant.427 In one study of SARS-CoV-2–infected index cases aged 7 to 19 years old, of whom 88% were symptomatic, transmission occurred in 18% of households with a secondary attack rate of 45%.157 The basic reproduction number (R0, the average number of secondary cases resulting from exposure to an index case) is typically greater than 2 but has been estimated to vary from 0.48 to 14.8 on a large cruise ship.231

Some SARS-CoV-2 variants, such as those comprising the Alpha lineage, are estimated to have a substantially higher estimated basic reproduction number and are associated with greater transmissibility in community settings than earlier circulating SARS-CoV-2 viruses.177 The Delta variant was estimated to have 55% higher transmissibility than the Alpha variant.107

SARS-CoV-2 can be transmitted by infected persons before they are symptomatic,288,539,720 and by those who never develop symptoms,720 but secondary attack rates are likely lower than from symptomatic individuals. One study estimated that the latent period (time from infection to becoming infectious) was 5.5 days.726 Estimates of secondary attack rates can vary depending on the definition of asymptomatic versus symptomatic or pauci-symptomatic and whether the data were based upon cross-sectional or longitudinal assessments. Two meta-analyses reported that the secondary attack rate was substantially lower in contacts of asymptomatic-infected persons compared with contacts of symptomatic SARS-CoV-2–infected persons.92,428 One meta-analysis estimated that the secondary attack rate for asymptomatic index cases was 1.9% but was 9.3% for presymptomatic and 13.6% for symptomatic index cases.660 Another meta-analysis reported that the relative risk for transmission from asymptomatically infected persons was 42% lower than from symptomatic persons,99 indicating that most transmission of SARS-CoV-2 is from symptomatic persons or infected persons just before symptom onset. A prospective study of U.S. households reported that the incidence of SARS-CoV-2 infections was similar among children and adults, but a greater proportion of infections among children were asymptomatic compared with adults.181

However, depending on the population and the number of asymptomatic individuals, asymptomatic SARS-CoV-2 infections may contribute substantially to transmission. One study that analyzed detailed contact tracing data from China found no difference in transmissibility between symptomatic and asymptomatically infected persons or between age groups and estimated that presymptomatic infected persons accounted for 59% of transmission events.299 A study in university residence halls reported that transmission from SARS-CoV-2–infected persons, most of whom were asymptomatic, to their roommates was more likely to occur from index cases with higher estimated viral RNA levels compared to index cases with lower estimated viral RNA levels.72 One study of a large SARS-CoV-2 outbreak among cruise ship passengers estimated that asymptomatic persons were the source for 69% of SARS-CoV-2 infections.206

Transmission Modes 835

Respiratory transmission through expelled large droplets and small particle droplet nuclei generated through aerosols is the primary mode of SARS-CoV-2 spread. SARS-CoV-2 has been isolated from nasopharyngeal,105,280,302,394,737 oropharyngeal,280,302 saliva,320 sputum,302 and bronchoalveolar lavage fluid764 specimens of COVID-19 patients, indicating the potential for respiratory transmission. SARS-CoV-2 can be isolated from upper respiratory tract specimens of asymptomatic and presymptomatic individuals.35,101,475,477 SARS-CoV-2 RNA is detectable in exhaled breath condensate from patients423,578,760 and experimentally infected rhesus macaques,204 suggesting potential for transmission through breathing or speech to close contacts. However, SARS-CoV-2 RNA may be more frequently detectable in aerosols produced by talking and singing, especially early in the clinical course, than in exhaled breath specimens.162 Experimentally infected ferrets can transmit infectious SARS-CoV-2 through droplets to ferrets in an adjacent cage.562 Air sampling from COVID-19 patients in hospital rooms has occasionally detected SARS-CoV-2 RNA, but the isolation of infectious virus is uncommon.200,381,497,587 Superspreading events have occurred in nightclubs and bars,133,331,472,633 on an aircraft carrier,337 and aboard a cruise ship.208 Epidemiological investigations have documented SARS-CoV-2 transmission in other closed settings including planes,150,347 restaurants,362,401,418,419 fitness centers and exercise facilities,48,155,261,317,389 church services,338 and church choir practice.275 Air sampling has detected SARS-CoV-2 RNA after Cesarean and vaginal deliveries from asymptomatic pregnant women.291 These events suggest that droplet spread facilitated by airflow or airborne transmission of aerosolized small particle SARS-CoV-2 more than 2 m is possible. Generally, these exposures are for periods of time of 30 minutes or more, but transmission can occur with exposures as short as 5 minutes.362

Infectious SARS-CoV-2 has rarely been isolated from nonrespiratory specimens such as urine320,635 or feces.175,192,320,725 Viral RNA has been detected more frequently in feces than other nonrespiratory specimens.140,143,509,571,711 SARS-CoV-2 RNA has been identified by sampling air in toilets and bathrooms of COVID-19 patients,70,410 although infectious virus was not recovered.70 One study suggested that fecal aerosol transmission of SARS-CoV-2 may have contributed to an outbreak of COVID-19 in a high-rise building in southern China.332 Although SARSCoV-2 RNA is detectable in plasma65,136,210 and serum,27,139 particularly in critically ill COVID-19 patients,65,136,652 and is suggestive of viremia, infectious SARS-CoV-2 virus has not been isolated from blood.27

Extrapulmonary dissemination through viremia could explain the detection of viral RNA in cardiac tissues and recovery of infectious virus in urine. Some autopsy studies have reported extrapulmonary SARS-CoV-2 infection of multiple organ tissues based upon electron microscopy findings, but one comprehensive study noted that many of these reported results are based upon a misidentification of nonviral subcellular structures.94 SARS-CoV-2 RNA was detected in the cerebrospinal fluid of an adolescent diagnosed with Guillain-Barré syndrome.33 Detection of SARS-CoV-2 RNA has been reported infrequently in semen in patients with acute illness and recovered COVID-19 patients.227,249,397 Detection of SARS-CoV-2 RNA has very rarely been reported in vaginal fluid40,540,591,621,736 or amniotic fluid.359,537,686 SARS-CoV-2 RNA has been detected infrequently in breast milk samples, but replication-competent virus has not been identified.129,165,260 While respiratory droplets and secretions pose the highest potential of containing infectious SARS-CoV-2, and virus has been isolated sporadically from urine and feces during the acute phase of COVID-19, given the paucity of available data in which viral culture was performed on extrapulmonary specimens that tested positive for SARS-CoV-2 RNA, all extrapulmonary specimens should be considered potentially infectious during the acute phase of COVID-19, especially in patients with severe illness.

One experimental study demonstrated that SARS-CoV-2 can remain viable on different surfaces up to 72 hours and that aerosols may be viable for several hours.677 Although SARS-CoV-2 RNA has been detected on various surfaces for prolonged periods, suggesting potential for fomite transmission, viral culture of patient fomites, and surfaces in households and health care settings and a quarantine hotel has only yielded recovery of infectious virus from one surface sample.64,439,461

Reverse zoonosis of SARS-CoV-2 transmission from people to animals has been reported in domesticated dogs and cats, farmed mink, captive tigers, and lions.273,274,435 Experimental infection models are established in mice, Syrian golden hamsters, Chinese hamsters, cats, dogs, ferrets, raccoon dogs, cattle, tree shrews, and nonhuman primates (cynomolgus and rhesus macaques, and common marmosets),83,435 However, animal-to-human SARS-CoV-2 transmission has only been reported from farmed mink, an event that also had evidence of reverse zoonosis and viral evolution in mink.372,502

Duration of Infectiousness Peak SARS-CoV-2 RNA levels in the upper respiratory tract are typically observed during the first week after symptom onset and in the lower respiratory tract of persons with severe COVID-19 during the 2nd week of illness.126 Reverse transcription polymerase chain reaction (RT-PCR) detection of SARS-CoV-2 RNA was most frequent in nasopharyngeal swab specimens collected within 4 days of symptoms onset in one systematic review.437 SARS-CoV-2 RNA levels may be higher in respiratory specimens infected with the Alpha or Delta variants than viruses not considered variants of concern or interest.326,395 In hospitalized COVID-19 patients, infection with the Delta variant was associated with significantly lower cycle threshold values and longer duration of SARS-CoV-2 RNA detection compared with wild-type SARS-CoV-2 from the first pandemic wave in Singapore.495 Unvaccinated and fully vaccinated persons infected with the SARS-CoV-2 Delta variant may have similar viral RNA levels in respiratory specimens.89,145 One meta-analysis of studies on wild-type Wuhan SARS-CoV-2 infections reported that the mean duration of SARS-CoV-2 RNA detection was 17 days (maximum 83 days) in the upper respiratory tract, 14.6 days in the lower respiratory tract (maximum 59 days), 17.2 days in stool (maximum 126 days), and 16.6 days in serum specimens (maximum 60 days).126 Longer duration of SARS-CoV-2 RNA detection is associated with older age.126,731,758 While SARS-CoV-2 RNA can be detected for prolonged periods in different clinical specimens from persons with asymptomatic infection, mild-to-moderate disease, and severe COVID-19, the presence of viral RNA does not necessarily indicate that infectious virus is present. Therefore, studies to assess evidence of replication–competent SARS-CoV-2, such as isolation in viral culture, are needed to inform the duration of infectiousness.

The duration of detection of infectious virus is shorter than detection of viral RNA. SARS-CoV-2 infectious virus has been isolated from upper respiratory tract specimens of asymptomatic35,294,614 and presymptomatic persons.35,486 In persons with mild-to-moderate COVID-19, SARSCoV-2 RNA can be detected for prolonged periods, but infectious virus identified by viral culture is unlikely to be present in upper respiratory tract specimens beyond 8 to 10 days after illness onset.35,93,504,521,711 However, one case report described isolation of SARS-CoV-2 infectious virus from sputum up to 18 days from illness onset in a patient with mild COVID-19 who had evidence of an endogenous antibody response409 and an asymptomatic, previously healthy immunocompetent adolescent hospitalized for injuries sustained in a motor vehicle accident had 836

SARS-CoV-2 isolated from a throat swab and tracheal aspirate specimens 54 days after hospital admission.582 One study estimated that isolation of infectious virus was highest 2 days before symptom onset to 1 day after onset and declined within 7 days after onset.288 Patients with severe disease may have a longer duration of infectiousness compared to patients with mild-to-moderate COVID-19. In hospitalized patients with severe COVID-19, SARS-CoV-2 has been isolated from respiratory specimens up to 20 days from illness onset, but infectious virus is unlikely to be recovered after 15 days from illness onset or when a neutralizing antibody titer of 1:80 was present.679

Some severely immunocompromised persons can have infectious SARS-CoV-2 detected in the upper respiratory tract for prolonged periods and pose a transmission risk to close contacts. Prolonged detection of the replication–competent infectious virus has been reported in case reports of patients with chronic lymphocytic leukemia and acquired hypogammaglobulinemia, lymphoma, hematopoietic stem cell transplant, kidney transplant, heart transplant, chimeric antigen receptor T-cell therapy, or AIDS, and with shedding often beyond 20 days and as long as 143 days after an initial positive SARS-CoV-2 test results.42,44,46,62,149,188,478,648 During prolonged SARS-CoV-2 replication in immunocompromised patients, especially those who are severely immunocompromised, viral evolution with emergence of mutations can occur with potential implications for treatment.42,46,149,413

CLINICAL DISEASE Severity of Disease Patients with SARS-CoV-2 infection can have clinical manifestations ranging from no symptoms to critical illness (Fig. 22.3). Discussions of severity of disease and treatment require a common understanding on how to define severity of disease. The US National Institutes of Health (NIH) COVID-19 Treatment guidelines define the following groups289:

Asymptomatic Infection: Individuals who test positive for SARS-CoV-2 using a virologic test but who have no symptoms consistent with COVID-19. Mild Illness: Individuals who have any of the various signs and symptoms of COVID-19 but do not have shortness of breath, dyspnea, or abnormal chest imaging. Moderate Illness: Individuals with evidence of lower respiratory disease during clinical assessment or imaging and oxygen saturation (SpO2) ≥94% on room air at sea level. Severe Illness: Individuals who have SpO2