The Mouse in Biomedical Research, 2nd Edition Volume II Diseases
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THE MOUSE IN BIOMEDICAL RESEARCH, 2ND EDITION Volume II Diseases EDITED
BY
James G. Fox
Muriel T. Davisson
Fred W. Quimby
Division of Comparative Medicine, MIT Cambridge, MA
The Jackson Laboratory Bar Harbor, ME
Laboratory Animal Research Center The Rockefeller University New York, NY
Stephen W. Barthold
Christian E. Newcomer
Abigail L. Smith
Center for Comparative Medicine Schools of Medicine and Veterinary Medicine University of California Davis, CA
Research Animal Resources and Department of Molecular and Comparative Pathobiology Johns Hopkins University Baltimore, MD
School of Veterinary Medicine University of Pennsylvania Philadelphia, PA
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Table of Contents Volume I History, Wild Mice, and Genetics List of Reviewers List of Contributors Foreword Preface
1.
Building a Better Mouse: One Hundred Years of Genetics and Biology
x xi xiii xv
10. 1
Herbert C. Morse III
Mouse Embryology: Research Techniques and a Comparison of Embryonic Development between Mouse and Man
165
Matthew H. Kaufman 2.
Systematics of the genus Mus
13
Priscilla K. Tucker
11.
Gamete and Embryo Manipulation
211
K.C. Kent Lloyd 3.
The Secret World of Wild Mice
25
Grant R. Singleton and Charles J. Krebs 12. 4.
Breeding Systems: Considerations, Genetic Fundamentals, Genetic Background, and Strain Types
Mouse Strain and Genetic Nomenclature: an Abbreviated Guide
225
Martin Hrabé de Angelis, Dian Michel, Sibylle Wagner, Sonja Becker, and Johannes Beckers 53
Melissa L. Berry and Carol Cutler Linder 5.
Chemical Mutagenesis in Mice
13.
Gene-Specific Mutagenesis
261
K.C. Kent Lloyd 79
Janan T. Eppig
14.
Gene Transfer Studies Using Mouse Models
267
Robert G. Pergolizzi and Ronald G. Crystal 6.
The Mouse Genome
99
Mark D. Adams 7.
Gene Mapping
15.
Mouse and Human Pluripotent Stem Cells
281
Leslie F. Lock
115
Muriel T. Davisson 16. 8.
Genetic Monitoring
135
Cytogenetics Muriel T. Davisson and Mary Ann Handel
289
Lucia F. Jorge-Nebert, Sandrine Derkenne, and Daniel W. Nebert
Richard R. Fox, Michael V. Wiles, and Petko M. Petkov 9.
Drugs and the Mouse: Pharmacology, Pharmacogenetics, and Pharmacogenomics
145 Index
321
v
vi
TA B L E O F C O N T E N T S
Volume II Diseases
10.
Retroelements in the Mouse
269
Herbert C. Morse III List of Reviewers List of Contributors Foreword Preface
x xi xiii xv
Viral Diseases
11.
Sendai Virus and Pneumonia Virus of Mice (PVM)
281
David G. Brownstein
12.
DNA Viruses
Cardioviruses: Encephalomyocarditis Virus and Theiler’s Mouse Encephalomyelitis Virus
311
Howard L. Lipton, A.S. Manoj Kumar, and Shannon Hertzler 1.
Murine Cytomegalovirus and other Herpesviruses
1 Bacterial Diseases
Geoffrey R. Shellam, Alec J. Redwood, Lee M. Smith, and Shelley Gorman 13. 2.
Mouse Adenoviruses
325
Roger G. Rank
49
Katherine R. Spindler, Martin L. Moore, and Angela N. Cauthen
Chlamydial Diseases
14.
Clostridial Species
349
Kimberly S. Waggie 3.
Mousepox
67
R. Mark L. Buller and Frank Fenner
4.
Parvoviruses
15.
Enterobacteriaceae, Pseudomonas aeruginosa, and Streptobacillus moniliformis
365
Hilda Holcombe and David B. Schauer
93
Robert O. Jacoby and Lisa Ball-Goodrich 16. 5.
Polyoma Viruses
Aerobic Gram-positive Organisms
389
Cynthia Besch-Williford and Craig L. Franklin
105
Thomas L. Benjamin 17.
RNA Viruses
Helicobacter Infections in Mice
407
James G. Fox and Mark T. Whary 6.
Mouse Hepatitis Virus
141 18.
Stephen W. Barthold and Abigail L. Smith
Mycoplasma pulmonis, other Murine Mycoplasmas, and Cilia-Associated Respiratory Bacillus
437
Trenton R. Schoeb 7.
Lymphocytic Choriomeningitis Virus
179
Stephen W. Barthold and Abigail L. Smith
19.
Pasteurellaceae
469
Werner Nicklas 8.
Lactate Dehydrogenase-Elevating Virus
215 Mycotic and Parasitic Diseases
Jean-Paul Coutelier and Margo A. Brinton
9.
Reoviridae Richard L. Ward, Monica M. McNeal, Mary B. Farone, and Anthony L. Farone
235
20.
Fungal Diseases in Laboratory Mice Virginia L. Godfrey
507
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TA B L E O F C O N T E N T S
21.
Protozoa
517
3.
Katherine Wasson
22.
Helminth Parasites of Laboratory Mice
Reproductive Biology of the Laboratory Mouse
91
Kathleen R. Pritchett and Robert Taft
551
4.
Kathleen R. Pritchett
Endocrinology: Bone as a Target Tissue for Hormonal Regulation
123
Krista M. Delahunty and Wesley G. Beamer 23.
Arthropods
565 5.
David G. Baker
Hematology of the Laboratory Mouse
133
Nancy E. Everds Miscellaneous Diseases 6. 24.
The Tumor Pathology of Genetically Engineered Mice: A New Approach to Molecular Pathology 581
Spontaneous Diseases in Commonly Used Mouse Strains
171
Fred W. Quimby and Richard H. Luong
Robert D. Cardiff, Robert J. Munn, and Jose J. Galvez
25.
Clinical Chemistry of the Laboratory Mouse
Management, Techniques, and Husbandry
7. 623
Gnotobiotics
217
Richard J. Rahija
Cory Brayton 8. 26.
Zoonoses and other Human Health Hazards
Management and Design: Breeding Facilities
235
William J. White
719
Christian E. Newcomer and James G. Fox 9. Index
747
Design and Management of Research Facilities for Mice
271
Neil S. Lipman
Volume III Normative Biology, Husbandry, and Models List of Reviewers List of Contributors Foreword Preface
10.
Nutrition
321
Graham Tobin, Karla A. Stevens and Robert J. Russell x xi xiii xv
Normative Biology
11.
Health Delivery and Quality Assurance Programs for Mice
385
Diane J. Gaertner, Glen Otto and Margaret Batchelder
12.
Environmental and Equipment Monitoring
409
J. David Small and Rick Deitrich 1.
Gross Anatomy
1
Vladimír Komárek
2.
Mouse Physiology Robert F. Hoyt, Jr., James V. Hawkins, Mark B. St. Claire, and Mary B. Kennett
13.
23
Biomethodology and Surgical Techniques
437
Alison M. Hayward, Laura B. Lemke, Erin C. Bridgeford, Elizabeth J. Theve, Courtnye N. Jackson, Terrie L. Cunliffe-Beamer, and Robert P. Marini
viii 14.
TA B L E O F C O N T E N T S
In Vivo Whole-Body Imaging of the Laboratory Mouse 489 Simon R. Cherry Use of Mice in Biomedical Research
Foreword Preface
xiii xv
Overview
1
Fred W. Quimby and David D. Chaplin 15.
Behavioral Testing
513
Douglas Wahlsten and John C. Crabbe
16.
Cardiovascular Disease: Mouse Models of Atherosclerosis
1.
The Molecular Basis of Lymphoid Architecture in the Mouse
57
Carola G. Vinuesa and Matthew C. Cook 535
Nobuyo Maeda, Raymond C. Givens, and Robert L. Reddick
2.
The Biology of Toll-like Receptors in Mice
109
Osamu Takeuchi and Shizuo Akira 17.
Convulsive Disorders
565 3.
Mariana T. Todorova and Thomas N. Seyfried
Genomic Organization of the Mouse Major Histocompatibility Complex
119
Attila Kumánovics 18.
Eye Research
595
Richard S. Smith, Patsy M. Nishina, John P. Sundberg, Johann Zwaan, and Simon W.M. John
4.
Some Biological Features of Dendritic Cells in the Mouse
135
Kang Liu, Anna Charalambous, and Ralph M. Steinman 19.
Genetic Analysis of Rodent Obesity and Diabetes
617
Sally Chiu, Janis S. Fisler, and Craig H. Warden 5. 20.
Mouse Models in Aging Research
637
Kevin Flurkey, Joanne M. Currer, and D.E. Harrison
21.
Mouse Models of Inherited Human Neurodegenerative Disease 673
Mouse Skin Ectodermal Organs
155
Maria D. Iglesias-Ussel, Ziqiang Li, and Matthew D. Scharff
6.
Karl Herrup
22.
Mouse Models Revealed the Mechanisms for Somatic Hypermutation and Class Switch Recombination of Immunoglobulin Genes
Mouse Natural Killer Cells: Function and Activation
169
Francesco Colucci 691 7.
Maksim V. Plikus, John P. Sundberg, and Cheng-Ming Chuong
Cytokine-activated JAK-STAT Signaling in the Mouse Immune System 179 Bin Liu and Ke Shuai
23.
Quality Control Testing of Biologics
731 8.
William R. Shek
Signal Transduction Events Regulating Integrin Function and T Cell Migration in the Mouse
195
Lakshmi R. Nagarajan and Yoji Shimizu Index
759 9.
Volume IV Immunology List of Reviewers List of Contributors
x xi
Mouse Models of Negative Selection Troy A. Baldwin, Timothy K. Starr, and Kristin A. Hogquist
207
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TA B L E O F C O N T E N T S
10.
Peripheral Tolerance of T Cells in the Mouse
223
14.
Vigo Heissmeyer, Bogdan Tanasa, and Anjana Rao
Mouse Models to Study the Pathogenesis of Allergic Asthma
291
Chad E. Green, Nicholas J. Kenyon, Scott I. Simon, and Fu-Tong Liu 11.
The Genetics of Mouse Models of Systemic Lupus
243
Srividya Subramanian and Edward K. Wakeland 15. 12.
Inhibitory Receptors and Autoimmunity in the Mouse 261
The Mouse Trap: How Well Do Mice Model Human Immunology?
303
Javier Mestas and Christopher C.W. Hughes
Menna R. Clatworthy and Kenneth G.C. Smith Index 13.
Mouse Models of Immunodeficiency B. Anne Croy, James P. Di Santo, Marcus Manz, and Richard B. Bankert
275
313
List of Reviewers for Chapters in this Volume Baker, David G. Besselsen, David G. Brayton, Cory Brownstein, David Buchmeier, Michael Bunte, Ralph Castleman, William Clark, H. Fred Clifford, Charles B. Compton, Susan R. Conner, Margaret E. Eckhardt, Laurel Fister, Richard Foil, Lane Franklin, Craig Gardner, Murray Jacoby, Robert O. Kaltenboeck, Bernhard Kuo, Cho-chou “Ted” Lindsay, J. Russell Maggio-Price, Lillian Maronpot, Robert Murphy, Frederick A. Ramsey, Kyle Rand, Michael S. Schauer, David B. Shek, William R. Taylor, Nancy Weisbroth, Steven H.
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Louisiana State University, Baton Rouge, LA University of Arizona, Tucson, AZ Johns Hopkins University School of Medicine, Baltimore, MD University of Edinburgh, Edinburgh, UK Scripps Research Institute, La Jolla, CA University of Pennsylvania, Philadelphia, PA College of Veterinary Medicine, University of Florida, Gainesville, FL Children’s Hospital of Philadelphia, PA Charles River Laboratories, Inc., Wilmington, MA Yale University, New Haven, CT Baylor College of Medicine, Houston, TX Hunter College of CUNY, New York City, NY Charles River Laboratories, Inc., Wilmington, MA Louisiana State University, Baton Rouge, LA University of Missouri, Columbia, MO University of California, Davis, CA Yale University, New Haven, CT Auburn University, Auburn, AL University of Washington, Seattle, WA University of Alabama, Birmingham, AL University of Washington, Seattle, WA National Institute of Environmental Health Sciences, Research Triangle Park, NC University of Texas Medical Branch, Galveston, TX Midwestern University, Downer’s Grove, IL University of Arizona, Tucson, AZ Massachusetts Institute of Technology, Cambridge, MA Charles River Laboratories, Inc., Wilmington, MA Massachusetts Institute of Technology, Cambridge, MA McLean, VA
Contributors David G. Baker Division of Laboratory Animal Medicine School of Veterinary Medicine Louisiana State University Baton Rouge, LA 70803 Lisa Ball-Goodrich Section of Comparative Medicine Yale University School of Medicine New Haven, CT 06520-8016 Stephen W. Barthold Center for Comparative Medicine Schools of Medicine and Veterinary Medicine University of California Davis, CA 95616 Thomas L. Benjamin Department of Pathology Harvard Medical School Boston, MA 02115 Cynthia Besch-Williford Research Animal Diagnostic Laboratory University of Missouri, COVM Columbia, MO 65211 Cory Brayton Department of Comparative Medicine Johns Hopkins University School of Medicine Baltimore, MD 21205 Margo A. Brinton Department of Biology Georgia State University Atlanta, GA 30302 David G. Brownstein Research Animal Pathology Core Institute for Medical Cell Biology University of Edinburgh Edinburgh, Scotland EH16 4TJ
R. Mark L. Buller Department of Molecular Microbiology and Immunology St. Louis University Health Sciences Center St. Louis, MO 63104 Robert D. Cardiff Department of Pathology and Laboratory Medicine Center for Comparative Medicine University of California at Davis Davis, CA 95616 Angela N. Cauthen Department of Natural Sciences Clayton College and State University Morrow, GA 30260 Jean-Paul Coutelier Unité de Médecine Expérimentale Université Catholique de Louvain Bruxelles, Belgium Mary B. Farone Biology Department Middle Tennessee State University Murfreesboro, TN 37132 Anthony L. Farone Biology Department Middle Tennessee State University Murfreesboro, TN 37132 Frank Fenner John Curtin School of Medical Research Australian National University Canberra, Australian Capital Territory, Australia James G. Fox Division of Comparative Medicine Massachusetts Institute of Technology Cambridge, MA 02139
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CONTRIBUTORS
Craig L. Franklin Research Animal Diagnostic Laboratory University of Missouri, COVM Columbia, MO 65211
Monica M. McNeal Infectious Diseases Children’s Hospital Medical Center Cincinnati, OH 45229-3039
Jose J. Galvez Department of Pathology and Laboratory Medicine Center for Comparative Medicine University of California at Davis Davis, CA 95616
Martin L. Moore Department of Medicine Vanderbilt University Medical Center Nashville, TN 37232-2650
Virginia L. Godfrey Division of Laboratory Animal Medicine Department of Pathology and Laboratory Medicine University of North Carolina Chapel Hill, NC 27599-7115 Shelley Gorman Telethon Institute for Child Health Research Centre for Child Health Research University of Western Australia Nedlands 6907 Western Australia Shannon Hertzler Department of Neurology Evanston Hospital Northwestern University Evanston, IL 60201 Hilda Holcombe Division of Comparative Medicine Massachusetts Institute of Technology Cambridge, MA 02139 Robert O. Jacoby Section of Comparative Medicine Yale University School of Medicine New Haven, CT 06520-8016 A.S. Manoj Kumar Department of Neurology Microbiology-Immunology, and Biochemistry Molecular and Cell Biology Evanston Hospital Northwestern University, Evanston, IL 60201 Howard L. Lipton Department of Neurology Microbiology-Immunology, and Biochemistry Molecular and Cell Biology Evanston Hospital Northwestern University, Evanston, IL 60201
Herbert C. Morse, III Laboratory of Immunopathy National Institute of Allergy and Infectious Diseases NIH Rockville, MD 20852 Robert J. Munn Center for Comparative Medicine University of California at Davis Davis, CA 95616 Christian E. Newcomer Research Animal Resources and Department of Molecular and Comparative Pathobiology Johns Hopkins University Baltimore, MD 21205 Werner Nicklas Central Animal Laboratories German Cancer Research Centre D-69120 Heidelberg Germany Kathleen R. Pritchett Charles River Laboratories Research Models and Services Domaine des Oncins l′Arbresle, Lyon 69592 France Roger G. Rank Department of Microbiology and Immunology University of Arkansas for Medical Sciences Little Rock, AR 72205 Alec J. Redwood Department of Microbiology University of Western Australia Nedlands 16907 Western Australia David B. Schauer Division of Comparative Medicine Massachusetts Institute of Technology Cambridge, MA 02139
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CONTRIBUTORS
Trenton R. Schoeb Department of Genetics Division of Genomics University of Alabama at Birmingham Birmingham, AL 35294-0024 Geoffrey R. Shellam Department of Microbiology University of Western Australia Nedlands 6907 Western Australia Lee M. Smith Department of Microbiology University of Western Australia Nedlands 6907 Western Australia Abigail L. Smith School of Veterinary Medicine University of Pennsylvania Philadelphia, PA 19104 Katherine R. Spindler Microbiology and Immunology University of Michigan School of Medicine Ann Arbor, MI 49109-0620
Kimberly S. Waggie Preclinical Development ZymoGenetics Inc. Seattle, WA 98102 Richard L. Ward Infectious Diseases Children’s Hospital Medical Center Cincinnati, OH 45229-3039 Katherine Wasson Center for Comparative Medicine University of California at Davis Davis, CA 95616 Mark T. Whary Division of Comparative Medicine Massachusetts Institute of Technology Cambridge, MA 02139
Foreword for Volume II The second edition of The Mouse in Biomedical Research reflects the revolution in mouse biology inaugurated during the past quarter century. It is exemplified by the heavy flavoring of genetics in Volume I and the focus of Volume IV on immunology. These trends re-emphasize the indispensability of mice for biomedical research and the influence of genetic engineering in driving it. Setting mouse genes in motion through molecular wizardry has accelerated understanding of mammalian biology and disease at a stunning pace. It also has provoked domination of mouse populations by novel strains which are scientifically and financially among biomedicine’s most precious assets. Therefore, the health and, implicitly, the diseases of laboratory mice are transcendent issues for investigators and laboratory animal medicine specialists. The roster of health risks for mice also has evolved significantly since the original edition of this text. Improvements in housing, husbandry and health care have reduced the impact of traditional infections, such as mycoplasmosis, viral pneumonias and enteridites, acariasis and endoparasitism. However, they remain, although weakened, incompletely dispatched and, therefore, worthy subjects for this text. Additionally, specific pathogen-free mice, which constitute the lion’s share of contemporary mousedom, are, by definition, highly susceptible to these adventitious infections. Of equal or greater importance, historically notorious agents have been superceded epidemiologically by helicobacters and recently recognized parvoviruses, both of which carry the means for serious disruption of research. Awareness of “new” agents is illustrated further by contemporary investigations of norovirus infection in mice, which are gathering interest as this edition goes to press, and thus too late for formal inclusion. (See Wobus, C.E. et al. 2006, J. Virol., 80, 5104–5112). Lastly, current understanding of agent-host interactions and their
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effects on health and research is based largely on responses of standard stocks and strains. Serendipitous phenotypes resulting from genetic engineering are likely to change these interactions in ways that are currently hard to predict. Fundamental information on pathogenesis and related factors, provided here, are essential to detect and assess divergent outcomes. It is no surprise, in these contexts, that infectious agents continue dominate contemporary murine health concerns and this volume. The potential effects of genetic engineering on the expression and prevalence of non-infectious diseases are illustrated by chapters on tumor pathology and spontaneous diseases. They also highlight the value of phenotyping for murine health. While researchers often seek categorical effects (ie predicted or desired mode of gene expression on a given cell type, tissue or organ), erudite phenotyping is the sentinel for unexpected or undesirable lesions which may compromise potentially novel models. The text also shows that mouse medicine has matured into a highly sophisticated discipline during the past 25 years. Reagents and methods for the early and accurate detection and diagnosis of disease are arguably the most visible vanguards of this advance. Nevertheless, the second edition also illustrates that collaborations with geneticists, embryologists, immunologists, epidemiologists and other scientific colleagues is developing a new and improved generation of health care paradigms and the laboratory animal experts to deliver them.
ROBERT O. JACOBY, DVM, PhD NEW HAVEN, CONNECTICUT
Preface The American College of Laboratory Animal Medicine (ACLAM) was formed in 1957 in response to the need for specialists in laboratory animal medicine. The college has promoted high standards for laboratory animal medicine by providing a structured framework to achieve certification for professional competency and by stressing the need for scientific inquiry and exchange via progressive continuing education programs. The first edition of “The Mouse in Biomedical Research” consisting of four volumes, and published in 19811983 was a part of the College’s effort to fulfill those goals. It is one of a series of comprehensive texts on laboratory animals developed by ACLAM over the past three decades: “The Biology of the Laboratory Rabbit” was published in 1974, “The Biology of The Guinea Pig” in 1976 and a twovolume work “Biology of The Laboratory Rat” in 1979 and 1980. Also, in 1979 the College published a two-volume text on “Spontaneous Animal Models of Human Disease”. In 1984 the first edition of “Laboratory Animal Medicine” appeared in print followed by “Laboratory Hamsters” in 1987. The second edition of The Biology of the Laboratory Rabbit was published in 1994. A two-volume treatise on “Nonhuman Primates in Biomedical Research” was published in 1995 and 1998. A text “Anesthesia and Analgesia in Laboratory Animals” was published in 1997 followed by the second edition of “Laboratory Animal Medicine” in 2002. Most recently, the second edition of “The Laboratory Rat” was published in 2005. The estimated annual use of 100 million-plus mice worldwide attests to the importance of the mouse in experimental research. The introduction of genetically engineered mice has only increased the usefulness of the mouse model in biomedical research. In no other species of animal has such a wealth of experimental data been utilized for scientific pursuits. Knowledge of the mouse that has been accumulated is, for the most part, scattered throughout a multitude of journals, monographs and symposia. It has been 25 years since the publication of the first edition of the “Mouse in Biomedical Research”. The intent of this second edition is to build upon the framework of the first edition, rather than simply to update and duplicate the earlier effort. The intended purpose of this text is to assemble established scientific data emphasizing recent information on the biology and use of the laboratory mouse. Separation of the material into multiple volumes was essential because of the number of
subject areas covered. The four volumes consist of 80 chapters coauthored by 167 scientists. The information in Volume 1 serves as a primer for scientists new to the field of mouse research. It provides information about the history, basic biology and genomics of the laboratory mouse (Mus musculus), as well as basic information on maintenance and use of mouse stocks. Mouse origins and relationships are covered in chapters on history, evolutionary taxonomy and wild mice. Genetics and genomics of the mouse are covered in chapters on genetic nomenclature, gene mapping, cytogenetics and the molecular organization of the mouse genome. Maintenance of laboratory mice is described in chapters on breeding systems for various types of strains and stocks and genetic monitoring. Use of the mouse as a model system for basic biomedical research is described in chapters on chemical mutagenesis, gene trapping, gene therapy, pharmacogenetics and embryo manipulation. Volume 2 entitled Diseases departs from the first edition of the same title by discussing specific disease-causing microorganisms, whereas the first edition discussed infectious diseases affecting specific organs and tissues. This volume consists of 26 chapters subdivided into RNA viruses and DNA viruses, as well as bacterial, mycotic and parasitic infections. These chapters not only provide updates on pathogenesis, epidemiology and prevention of previously recognized murine pathogens, but also include chapters on newly recognized disease-causing organisms: mouse parvovirus, cilia-associated respiratory bacilli and Helicobacter spp. A separate category, consisting of 3 chapters, discusses zoonoses, tumor pathology of genetically engineered mice and spontaneous diseases in commonly used mouse strains. Volume 3 encompasses 23 chapters whose contents provide a broad overview on the laboratory mouse’s normative biology, husbandry and its use as a model in biomedical research. This consists of chapters on behavior, physiology, reproductive physiology, anatomy, endocrinology, hematology and clinical chemistry. Other chapters cover management, as well as nutrition, gnotobiotics and disease surveillance. Individual chapters describe the mouse as a model for the study of aging, eye research, neurodegenerative diseases, convulsive disorders, diabetes and cardiovascular and skin diseases. Chapters on imaging, surgical and other research techniques and the use of the mouse in assays of biological products also are included.
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xvi Volume 4 is a completely new addition to this series, dedicated to mouse immunology. It is based on the vast body of knowledge which has made the mouse the model of choice when studying immunity in human beings. Arguably more is known about the immune system in mice than any other species except human. In large part this is due to the power of genetic engineering to delineate molecular mechanisms. This volume includes an overview of mouse immunology, including both the innate and adaptive immune systems, followed by 15 chapters (mini-reviews), each dealing with a specific area of immunology. The overview addresses broad concepts concerning molecular and cellular immunology and cites both current references and the appropriate chapter, for more detailed information, from the mini-reviews which follow. The 15 chapters illustrate the power of genetic engineering in dissecting each component of the immune response from the development of lymphoid tissues to signal transduction pathways in activated cells. Individual chapters address: The Genomic Organization of the MHC, Toll-like Receptors, The Molecular Basis of Lymphoid Architecture, The Biology of Dendritic Cells, Somatic Hypermutation and Class Switching, Natural Killer Cell Function and Activation, Cytokine Mediated Signaling, Signal Transduction Events Regulating Integrin Function and T-Cell Migration, Central Tolerance in T-Cells, Peripheral Tolerance in T-cells, Inhibitory Receptors and Autoimmunity. The volume also includes the use of mice in studies of Systemic Autoimmunity, Immunodeficiency, Allergic Airway Inflammation and the Differences Between Mouse and Human Immunology. This treatise was conceived with the intent to offer information suitable to a wide cross section of the scientific community. It is hoped that the four volumes will serve as a standard reference source for scientists using mice in biomedical research. Students embarking on scientific careers also will benefit from the broad coverage of material presented in compendium format. Certainly, specialists in laboratory animal
P R E FA C E
science will benefit from these volumes; technicians in both animal care and research will find topics on surgical techniques, management and environmental monitoring of particular value. The editors wish to extend special appreciation to the contributors to these volumes. Authors were selected because of knowledge and expertise in their respective fields. Each individual contributed his or her time, expertise and considerable effort to compile this resource treatise. In addition, the contributors and editors of this book, as with all volumes of the ACLAM series texts, have donated publication royalties to the American College of Laboratory Animal Medicine for the purpose of continuing education in laboratory animal science and comparative medicine. This book could not have been completed without the full support and resources of the editors’ parent institutions which allowed us the time and freedom to assemble this text. A special thanks is also extended to the numerous reviewers of the edited work whose suggestions helped the authors and editors present the material in a meaningful and concise manner. We also thank the editorial staff of Elsevier for their assistance. Finally, we especially acknowledge with deep appreciation the editorial assistance of Lucille Wilhelm, whose dedication and tireless commitment, as well as good humor, throughout this project were of immeasurable benefit to the editors in the completion of this text.
JAMES G. FOX STEPHEN W. BARTHOLD MURIEL T. DAVISSON CHRISTIAN E. NEWCOMER FRED W. QUIMBY ABIGAIL L. SMITH
Chapter 1 Murine Cytomegalovirus and Other Herpesviruses Geoffrey R. Shellam, Alec J. Redwood, Lee M. Smith, and Shelley Gorman
I. Introduction and History . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . II. Properties of the Virus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Classification . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Virion Structure and Replication . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1. Virion Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2. Replication of MCMV . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Virus Strains: Antigenic and Genetic Relationships . . . . . . . . . . . . . . . III. Growth In Vitro and In Vivo . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. In Vitro Propagation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1. Propagation in Permissive Murine Cells . . . . . . . . . . . . . . . . . . . . . 2. Nonpermissive Murine Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3. Non-Murine Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4. Features of MCMV Replication In Vitro . . . . . . . . . . . . . . . . . . . . . B. Infection of Mice . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . IV. Epizootiology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Natural History . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Natural Infection of Laboratory Mice . . . . . . . . . . . . . . . . . . . . . . . . . . C. Mode of Transmission . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. Host Range . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . V. Pathogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Effect of Age, Dose, and Route of Inoculation . . . . . . . . . . . . . . . . . . . 1. Age and Dose . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2. Route of Inoculation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Genetic Control of Host Resistance or Susceptibility to MCMV . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1. Resistance to Acute Infection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2. Mechanisms of Host Resistance to Acute Infection In Vivo . . . . . . 3. Mouse Strain Variation in the Resolution of Chronic MCMV Infection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4. In Vitro Studies of Resistance . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Latency and Reactivation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1. Pathogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
THE MOUSE IN BIOMEDICAL RESEARCH, 2ND EDITION
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Copyright © 2007, 1980, Elsevier Inc. All rights reserved.
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GEOFFREY R. SHELLAM, ALEC J. REDWOOD, LEE M. SMITH, AND SHELLEY GORMAN
D. Suitability of MCMV as a Model of HCMV Infection and Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1. Intrauterine Infection and Congenital Disease . . . . . . . . . . . . . . . . . 2. Interstitial Pneumonitis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3. Hepatitis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4. Cardiovascular Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5. Adrenalitis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6. Infection of the Central Nervous System . . . . . . . . . . . . . . . . . . . . . 7. Retinitis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8. Effect on the Developing Ear . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9. Effects on Hemopoiesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VI. The Immune Response to MCMV Infection . . . . . . . . . . . . . . . . . . . . . . . . 1. Immunosuppression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2. Macrophages and Dendritic Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . 3. Production of Interferon and Other Cytokines . . . . . . . . . . . . . . . . . 4. Natural Killer (NK) Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5. B Cell–Mediated Immune Responses . . . . . . . . . . . . . . . . . . . . . . . 6. T Cell–Mediated Immune Responses . . . . . . . . . . . . . . . . . . . . . . . . 7. Immune Evasion by MCMV . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VII. Diagnosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Serology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Molecular Detection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VIII. Control and Prevention . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1. Development of Vaccines . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . IX. Mouse Thymic Virus: Mouse T Lymphotrophic Virus (MTLV) or Murid Herpesvirus 3 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Introduction and History . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Properties of the Virus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Pathogenesis and Cell Tropism. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. Diagnosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
I.
INTRODUCTION AND HISTORY
Murine cytomegalovirus (MCMV) is a very well studied virus of laboratory mice. Together with the cytomegaloviruses of the rat, guinea pig, human, and other species, MCMV belongs to the b herpesvirinae subfamily of the Herpesviridae. The CMVs exhibit distinctive features that include specificity for their natural host species and the ability to establish persistent and latent infections, which are generally asymptomatic in the immunocompetent host. The discovery of the cytomegaloviruses had its origins in early studies of the etiology of a distinctive cytopathology associated with intranuclear inclusions and cellular enlargement. These cellular changes, which were termed cytomegalia (Goodpasture and Talbot 1921), were observed in the tissues of humans (Jesionek and Kiolemenoglou 1904), guinea pigs (Jackson 1920), rats (Thompson 1932), mice (Findlay 1932; Thompson 1934), and other species. The viral nature of the causative agent was first suggested by experiments in guinea pigs (Cole and Kuttner 1926), in which the inoculation of filtered homogenates of submaxillary glands induced cellular inclusions in the recipients. Murine cytomegalovirus was first isolated in tissue culture in 1954 by Margaret Smith from the salivary gland tissue of
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infected laboratory mice (Smith 1954). This followed earlier demonstrations that the etiological agent of intranuclear inclusion bodies in murine tissues could be transmitted by the inoculation of tissue homogenates into healthy mice (Kuttner and Wang 1934; McCordick and Smith 1936). Human cytomegalovirus (HCMV) was confirmed as the viral agent responsible for human salivary gland disease following its isolation in human cells in vitro (Rowe et al. 1956; Smith 1956; Weller et al. 1957). Subsequently, guinea pig cytomegalovirus (Hartley et al. 1957) and rat cytomegalovirus (RCMV) (Bruggeman et al. 1982) were isolated. The term cytomegalovirus was introduced by Weller and colleagues (Weller et al. 1960). Over the last 20 years, murine cytomegalovirus has become one of the best-studied viral infections of laboratory mice and is the subject of a very extensive literature. There are several reasons for this. First, MCMV research has benefited from the similarities between the diseases caused by human CMV (HCMV) and MCMV in their respective host species. There has been an increased awareness of the importance of HCMV-associated diseases in recipients of solid organ or bone marrow transplants and in HIV/AIDS, where HCMV is a very important cause
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of morbidity and mortality. The role of HCMV in inducing immunopathological diseases such as pneumonitis, retinitis, adrenalitis, and atherosclerosis has received much attention. Because of the strict species-specificity of the cytomegaloviruses, HCMV cannot be studied experimentally in animal models, and increasingly MCMV infection of mice has been used as a model of HCMV-associated diseases in humans. Second, there has been strong interest in how MCMV engages with the immune system. While T cell responses have been shown to be responsible for recovery from MCMV infection, the innate immune system, especially natural killer (NK) cells, has been found to be very important in controlling the extent of MCMV infection in mice. Research in this area has provided fascinating insights into how NK cells interact with viral pathogens. A host gene that controls resistance to MCMV in mice, Cmv1, has been identified and shown to encode a receptor molecule on NK cells. Interestingly, despite the presence of strong innate and adaptive immune responses, MCMV infection persists. This is due in part to virus-encoded proteins that assist the virus to evade the host response, particularly cell-mediated responses. MCMV has become one of the best-studied viruses for understanding the function of these immune evasion proteins. Finally, the strong growth of molecular virology over the last 20 years has significantly influenced the direction of research on MCMV. The genomes of MCMV and several other CMVs have been sequenced, and this information has been enormously valuable for understanding gene function, the relatedness between CMVs, and their evolution. The ability to clone the MCMV genome into a bacterial artificial chromosome has greatly facilitated the production of mutant viruses for the study of gene function, and the ease with which this can be studied in vivo in mice has been of great benefit. In all these areas of progress there has been one common feature that has contributed to the popularity of this virus model. This is that MCMV is a natural pathogen of its host species, the house mouse, infecting both laboratory mice and free-living wild Mus musculus domesticus. As with herpesviruses in general, which are believed to have co-evolved with their particular host species (Karlin et al. 1994; McGeoch and Cook 1994; McGeoch et al. 1995), the infection of mice by MCMV is considered to be a highly evolved host-parasite relationship that is free of the bias introduced by using a virus in its unnatural host. New studies, which are exploring the mutual adaptations made by MCMV and its natural host, will provide an important direction for research over the next few years. Given the size of the literature concerning MCMV, the reader may wish to consult other reviews that deal with the history, immunobiology, or pathogenesis of this virus in more detail than is possible here (Lussier 1975b; Hudson 1979; Osborn 1982; Staczek 1990; Hudson 1994a; Price and Olver 1996; Sweet 1999; Reddehase et al. 2002; Vink et al. 2001;
Gutermann et al. 2002; Scalzo 2002; Lee et al. 2002; Fairweather et al. 2001).
II.
PROPERTIES OF THE VIRUS A.
Classification
The cytomegaloviruses are large, enveloped, doublestranded DNA viruses with an icosahedral capsid that belong to the Betaherpesvirinae subfamily of the family Herpesviridae (van Regenmortel et al. 2000). There are three genera. The genus Cytomegalovirus contains human cytomegalovirus (Human herpesvirus 5). Muromegalovirus contains mouse cytomegalovirus (Murid herpesvirus 1, MuHV-1), which is also the type species, and rat cytomegalovirus (Murid herpesvirus 2, MuHV-2). The genus Roseolovirus contains Human herpesvirus 6 (HHV-6). The general characteristics of the cytomegaloviruses include strict species specificity, an ability to induce cytomegalia in infected cells, cell-associated replication in cell culture, a slow replicative cycle, and the establishment of persistent and latent infection in the natural host. Infection is generally asymptomatic unless the host is immunosuppressed or has an immature immune system. In these situations, infection may result in morbidity or even mortality. The house mouse, Mus domesticus, is considered to be the natural host for MCMV. Because of its common usage, the term murine cytomegalovirus (MCMV) rather than Murid herpesvirus 1 will be used throughout this chapter. Apart from mouse thymic virus (Murid herpesvirus 3), which is described elsewhere in this chapter, other herpesviruses of rodents have been discovered. One of them, mouse herpesvirus strain 68 (MHV-68), was isolated from bank voles in Slovakia (Blaskovic et al. 1980). This virus, which is also known as Murid herpesvirus 4, has been assigned to the Rhadinovirus genus of the Gammaherpesvirinae subfamily of the Herpesviridae (van Regenmortel et al. 2000). However, it appears that MHV-68 is not a natural pathogen of house mice. Although MHV-68 exhibits the features of a herpesvirus infection when it is deliberately inoculated into laboratory mice, including a productive infection in the lungs (Sunil-Chandra et al. 1992) and the establishment of persistence and latency (Stewart et al. 1998; Flano et al. 2000), there have been no reports of natural infection of colonies of laboratory mice with MHV-68. Recent serological evidence also indicates that free-living wild house mice are not naturally infected with MHV-68 (J. Stewart, personal communication). In contrast, MHV-68 appears to be endemic in free-living wood mice (Apodemus sylvaticus) in the United Kingdom although, interestingly, evidence of infection was only rarely found in voles in this study (Blasdell et al. 2003). Other studies from Slovakia
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GEOFFREY R. SHELLAM, ALEC J. REDWOOD, LEE M. SMITH, AND SHELLEY GORMAN
have reported a sero-prevalence of 1%–12% in wild rodents depending on the locality (Kozuch et al. 1993; Mistrikova and Blaskovic 1985), but the prevalence in particular rodent species was not recorded. As a naturally occurring rodent gammaherpesvirus that induces persistent and latent infection when inoculated into inbred laboratory mice, MHV-68 has received considerable attention in recent years for its potential as a model of human gammaherpesvirus infection. The immunobiology of MHV-68 has been widely studied in inbred mice, and there is a large body of literature. The viral genome has been sequenced (Virgin et al. 1997). Nonetheless, MHV-68 will not be discussed further in this chapter because the house mouse does not appear to be the natural host species. MHV-68 has been the subject of a number of comprehensive reviews to which the reader is referred (Nash et al. 2001; Doherty et al. 2001; Blackman et al. 2000; Stevenson et al. 2002; Stewart 1999; Virgin and Speck 1999). The Tupaia herpesvirus (THV) of tree shrews (Mirkovic et al. 1970; McCombs et al. 1971) has been sequenced (Baker and Darai 2001) and has been found to resemble MCMV and other β herpesviruses. However, it is not a virus of wild or laboratory mice and will not be considered further here. Finally, a cytomegalovirus has been isolated from deer mice (Peromyscus maniculatas) in North America (Rizvanov et al. 2003). The virus, which has been designated Peromyscus cytomegalovirus (PCMV), has been characterized as a cytomegalovirus based on physical and biological properties and genetic homology with several genes of other CMVs (Rizvanov et al. 2006). It does not replicate in mouse 3T3 cells, and it is not known whether it infects laboratory mice or free living Mus species.
B. 1.
Virion Structure and Replication
Virion Structure
Herpesvirus virions are spherical and comprise four morphologically distinct elements: the core, capsid, tegument, and envelope. The core encompasses the dsDNA viral genome, which is packaged as a single linear molecule into the protein capsid that exhibits icosahedral symmetry. The tegument is a poorly defined layer of proteinaceous material between the capsid and envelope which may be equivalent to the matrix of other viruses. It contains a number of proteins. The envelope is a lipid bilayer that contains a number of different integral viral glycoproteins (van Regenmortel et al. 2000). The virions of the cytomegaloviruses, including MCMV, share these distinctive features. However, while the diameter of single virions of MCMV is approximately 230 nM, the virions are quite pleiomorphic and may include a high proportion of multicapsid virions containing a number of capsids enclosed within a common membrane (Hudson et al. 1976b). The production of multicapsid virions appears to be an unusual property of MCMV.
The genomes of cytomegaloviruses are significantly larger than those of other herpesviruses, being over 200 kbp in size (Mocarski and Courcelle 2001a). The genome of the Smith strain of MCMV (ATCC VR-194) is 230, 278bp (Rawlinson et al. 1996), which is very similar in size to the genomes of HCMV and RCMV (Chee et al. 1990; Vink et al. 2000). The MCMV genome was originally described as encoding 170 open reading frames (Rawlinson et al. 1996). However, since this time, several new spliced genes have been identified within the genome (Ciocco-Schmitt et al. 2002; Loewendorf et al. 2004; Scalzo et al. 2004). Using comparative genomics techniques and new gene predicting algorithmns, the coding potential of human CMV has been reinterpreted recently (Davison et al. 2003; Murphy, Rigoutsos, et al. 2003 Murphy, Yu, et al. 2003), and using similar approaches, a further 34 open reading frames have been predicted to exist within the MCMV genome (Brocchieri et al. 2005). The MCMV genome comprises a single unique sequence with short terminal direct repeats and several short internal repeats (Ebeling et al. 1983; Mercer et al. 1983; Rawlinson et al. 1996). Unlike HCMV, the linear genome of MCMV does not have an isomeric structure because it lacks the larger terminal or internal repeat sequences (Ebeling et al. 1983; Mercer et al. 1983). The nomenclature devised for naming MCMV genes numbers the genes from the 5′ to the 3′ end of the genome. MCMV genes with homologs in HCMV are assigned the uppercase prefix M, while genes with no sequence identity with HCMV genes are identified by the lowercase prefix m (Rawlinson et al. 1996). The MCMV and HCMV genomes are co-linear over the central 180 kb, and there is significant similarity to the HCMV genome, especially for 78 open reading frames located in the central region of the genome (Rawlinson et al. 1996). MCMV shares with other members of the herpesvirus family a number of evolutionarily conserved proteins that are involved in processes such as DNA replication and virion maturation and structure. However, there are two regions of the genome that contain genes unique to MCMV. These are the m02 gene family at the left-hand end and the m145 gene family at the right-hand end of the genome. Many of the genes in these unique regions encode immune evasion proteins (Rawlinson et al. 1996). The capsid that encloses the viral genome is composed of 162 capsomers that exhibit icosahedral symmetry. Much of the information available on MCMV capsid proteins is inferred from their homology with the proteins of HCMV. The HCMV capsid proteins are also homologous to those found in the alphaherpesvirus, herpes simplex virus (HSV). The capsids of CMVs are larger and incorporate a larger genome than other herpesviruses. The capsid of MCMV is composed of seven proteins. The major capsid protein (MCP), the minor capsid protein, the minor capsid binding protein, and the smallest capsid protein are encoded by the MCMV genes M86, M85, M46, and m48.5, respectively. There are also three distinct assemblin-related proteins, which are encoded by M80, M80a,
1. MURINE CYTOMEGALOVIRUS AND OTHER HERPESVIRUSES
and M80.5 and associate with capsids (Baldick and Shenk 1996; Gibson 1996; Gibson, Baxter, et al. 1996; Gibson, Clopper et al. 1996; Chen et al. 1999). The MCP is the most strongly conserved protein among herpesviruses. The tegument of HCMV is composed of up to of 25 proteins, many of which are phosphorylated and have the prefix pp. By electron microscopy, the tegument is seen to have an ordered structure, particularly proximal to the capsid. The genes encoding the tegument proteins of HCMV have homologues in MCMV. These include M32 (pp150), M48 (large tegument protein) M83 (pp65), M99, and M82 (Rawlinson et al. 1996). Several transcriptional transactivators such as M82 (pp71) have also been localized to the tegument. However, the function of most tegument proteins remains undefined. Individual tegument proteins are conserved in the b herpesvirinae but are not shared with members of the α or γ herpesvirinae. The envelope of MCMV is composed predominantly of lipids obtained from the intracellular membranes of the host cell. The CMV envelope also contains a considerable number of viral encoded glycoproteins, which exceed the number of envelope glycoproteins found in other viruses. It is more pleiomorphic than the envelope of other herpesviruses, and is a distinguishing feature of the virus when observed by electron microscopy. The major glycoprotein is the highly conserved glycoprotein B (gB), which is a dominant B-cell antigen in CMV-infected animals and humans. It has been found in every herpesvirus and is one of the most highly conserved herpesvirus proteins. The amino acid homology between MCMV and HCMV gB is 45% (Rapp et al. 1992). The herpes virus gB is essential for viral entry into the cell (Little et al. 1981). Additional MCMV glycoproteins gL, gH, and gO (encoded by M115, M75, and m74 respectively) closely resemble their HCMV counterparts, which form the heterotrimeric complex gC111 (Huber and Compton 1998). This complex is essential for the entry of HCMV. 2.
Replication of MCMV
The life cycle of MCMV is initiated when virions bind to receptors on susceptible cells and enter the cytoplasm, where the viral envelope is removed and the capsid is transported to the nucleus. VIRAL ENTRY The cellular receptors used by MCMV have not been demonstrated conclusively. However, more than one receptor is likely to be involved. A role for MHC class I molecules (Wykes et al. 1993) and beta-2-microglobulin (Wykes et al. 1992) in facilitating the entry of MCMV has been demonstrated in vitro, although a significant role for MHC class I molecules was not confirmed by others in vitro or in vivo (Polic et al. 1996; Tay et al. 1995). In addition, heparan sulfate proteoglycans contribute to MCMV binding and entry (Price et al. 1995). These molecules also contribute to the binding of HCMV to target cells (Compton et al. 1993). HCMV uses the epidermal growth factor receptor (EGFR) (Wang et al. 2003) and integrin αvβ3 (Wang et al. 2005) as
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co-receptors for viral entry, using HCMV gB and gH, respectively. Integrin αvβ1 is also a co-receptor for HCMV (Feire et al. 2004). The binding of HCMV to these molecules initiates translocation of integrin αvβ3 into lipid rafts in the cell membrane. This results in interaction with EGFR and coordinated signaling, which triggers potent cooperative effects in target cells that appear to be necessary for successful viral infection (Wang et al. 2005). The role of these molecules as possible co-receptors in MCMV infection is yet to be determined. Following viral attachment to cell surface molecules, the HCMV envelope fuses with the cell surface in a pH-independent manner (Compton et al. 1992), involving the HCMV gCIII heterotrimeric complex (Keay and Baldwin et al. 1991; Huber and Compton 1998). It is assumed that MCMV enters cells in a similar manner. In support of this, the deletion of MCMV gL prevents initiation of MCMV infection (J. Allan, personal communication). Viral nucleocapsids then enter the cytoplasm and are transported through nuclear pores to the nucleus. The transcription of MCMV genes, the replication of viral DNA, and the subsequent assembly of capsids occurs in the nucleus of infected cells. REPLICATION OF VIRAL DNA MCMV DNA replication appears to follow the same sequence of events that occurs in the replication of other herpesviruses, via circular and/or concatermeric intermediates (Marks and Spector 1988). The sequences at the termini of the genome fuse via a 3′ nucleotide extension to form the intermediates for MCMV replication, by a rollingcircle model (Marks and Spector 1988). MCMV genomes circularize as soon as 2 hr after infection (Marks and Spector 1988). Host DNA synthesis is inhibited by more than 95% by 10–12 hr post infection (p.i.) (Moon et al. 1976), and the onset of viral DNA synthesis in mouse embryo fibroblasts (MEF) occurs during 8–16 hr p.i. depending on the mitotic phase of the host cells (Misra et al. 1978; Moon et al. 1976; Muller and Hudson 1977; Muller et al. 1978). By 22 hr p.i. the equivalent of 900 viral genomes per cell have been synthesized (Misra et al. 1977). The origin of replication of MCMV is known as the oriLyt replicator gene region, which extends over 1.7 kb (Masse et al. 1997). The oriLyt region in MCMV is extremely rich in repeat sequences that act as binding elements for various transcription factors. MCMV-encoded proteins that are required for origindependent replication include the DNA polymerase (M54), a polymerase accessory protein (M44), the single-stranded DNA binding protein (M57), and a helicase-primase complex of the viral proteins M70, M102, and M105 (Elliott et al. 1991, Rawlinson et al. 1996). The maturation of the MCMV genome involves the processing and cleavage of newly synthesized concatermeric viral DNA into genome-length monomers, prior to packaging into preformed nucleocapsids in the cell nucleus. Herpesvirus-conserved pac1 and pac2 DNA sequence motifs are required for cleavage and packaging of the MCMV genome (McVoy et al. 1998).
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GEOFFREY R. SHELLAM, ALEC J. REDWOOD, LEE M. SMITH, AND SHELLEY GORMAN
REGULATION OF MCMV GENE EXPRESSION Gene expression is temporally controlled in a cascade manner. All betaherpesviruses (Honess and Roizman 1974) including MCMV (Keil et al. 1984; Misra et al. 1978) have three gene families, α, β, and γ, that are temporally regulated. These genes are expressed in the immediate early (IE), early (E), and late (L) phases of viral replication, respectively. The IE phase occurs immediately after viral DNA enters the nucleus and is controlled by the major IE promoter (MIEP) in MCMV (Dorsch-Hasler et al. 1985) or HCMV (Boshart et al. 1985). The IE genes do not require de novo protein synthesis, and their transcription is not inhibited by protein inhibitors such as cyclohexamide (Chantler and Hudson 1978). In MCMV, the MIEP controls expression of the ie1 (m123) and ie3 (M122) genes (Keil et al. 1987a,b). MCMV ie1 (pp89) and ie3 are transcriptional activators (Koszinowski et al. 1986; Messerle et al. 1992). The ie3 gene is essential for MCMV replication (Angulo et al. 2000). A third IE gene, ie2, not present in HCMV (Chee et al. 1990), is transcribed from a different promoter and in the opposite direction (Keil et al. 1987a). The ie2 gene is not essential for MCMV replication either in vitro or in vivo in the situations that have been examined so far (Manning and Mocarski 1988; Cardin et al. 1995). While IE gene expression is confined to discrete regions of the genome, early gene transcripts come from various regions of the genome (Marks et al. 1983; Keil et al. 1984). Entry into the E phase is dependent on de novo protein synthesis and is therefore inhibited by cyclohexamide. There is an absolute requirement for IE gene expression prior to E gene expression (Honess and Roizman 1974), while E gene expression can be regulated by proteins derived from IE genes or from other E genes (Buhler et al. 1990). Genes transcribed during the E phase of viral replication include those required for entry into the L phase of viral replication, as well as other genes, such as the immune evasion genes (Ziegler et al. 1997). Entry into the L phase of viral replication requires DNA synthesis and is consequently inhibited by phosphonoacetic acid (Misra et al. 1977; Chantler and Hudson 1978). MCMV DNA replication requires proteins synthesized during the E phase of replication and occurs in the nucleus. The L phase of viral replication occurs approximately 16 hours after infection (Moon et al. 1976; Keil et al. 1984). However, both viral DNA replication and the kinetics of virus production are likely to be dependent on the cell culture system or cell type studied (Misra and Hudson 1977; Andrews et al. 2001). L phase genes mostly encode structural proteins. MORPHOGENESIS OF MCMV Along with viral gene transcription and DNA replication, the formation of capsids and the packaging of viral DNA occurs in the nucleus of infected cells (reviewed by Gibson 1996). Capsid proteins are produced in the cytoplasm and are transported back to the nucleus across the nuclear membrane. The transport of capsid proteins is due either to their small size, which allows passage across the nuclear pore
complex, or to the presence of a nuclear localization signal. Large capsid proteins such as MCP, that do not contain nuclear localization signals are transported in association with those that do (Wood et al. 1997; Plafker and Gibson 1998). Viral DNA is packaged into complete capsids and transported to the cytoplasm via the nuclear membrane, where the capsids acquire their primary envelope as they bud through the membranes. The two conserved herpesvirus genes UL31 (Chang and Roizman 1993; Reynolds et al. 2001; Fuchs et al. 2002) and UL34 (Purves et al. 1992; Klupp et al. 2000; Reynolds et al. 2001) are involved in this process, and loss of these genes impairs primary envelopment of HSV-1 and pseudorabies virus (PrV) (reviewed in Mettenleiter 2002). In MCMV infection, the virus penetrates the nuclear membrane with the aid of M50/p35 (a homologue of UL34) and a partner MCMV gene M53/p38 (a homologue of UL31). M50/M53 recruit cellular kinases to the inner nuclear membrane, specifically to the nuclear lamina. The nuclear lamina is a filamentous network that prevents budding. Recruitment of cellular kinases results in phosphorylation and degradation of the lamina (Muranyi et al. 2002), and facilitates egress of the virus from the nucleus. The mechanism of egress of herpesviruses from infected cells remains incompletely understood. The envelope and tegument of perinuclear virions differ from those of mature virions (Gershon et al. 1994; Granzow et al. 1997). In addition, the phospholipid content of mature HSV-1 virions differs from that of the nuclear membrane (van Genderen et al. 1994). Finally, the glycoproteins found in the mature virion that are responsible for viral entry into new host cells are not required for primary envelopment (Granzow et al. 2001). The synthesis and processing pathways of the major envelope glycoprotein complex (gp52/105/150) of MCMV have been characterized (Loh 1991). Ultrastructural differences are also apparent between primary enveloped virions and mature virions. These data suggest a secondary envelopment; however, the exact mechanisms by which the virus loses the primary envelope and tegument and acquires mature tegument and envelope has not been fully elucidated. Several models of herpesvirus egress have been proposed (reviewed by Mettenleiter 2002). However, it is suggested that secondary envelopment of virions is preceded by de-envelopment in the cytoplasm. Recent studies with α- and β- herpesviruses suggest that the primary enveloped virus migrates to the rough endoplasmic reticulum that is contiguous with the nuclear membrane, and exits into the cytoplasm, losing its primary envelope (reviewed by Mettenleiter 2002). The virus then migrates to an intracellular compartment, possibly the trans Golgi network, where the virions gain their tegument and final envelope, complete with glycoproteins (Gershon et al. 1994; Mettenleiter 2002; Mettenleiter 2004). The mature enveloped virions are now present in secretory vesicles that are transported to the plasma membrane, where they are released into the extracellular space by exocytosis. Little is known of this process, although UL20 and gK from HSV-1 and PrV have been implicated (reviewed in Mettenleiter 2004).
1. MURINE CYTOMEGALOVIRUS AND OTHER HERPESVIRUSES
The formation of multicapsid virions, which appears to be unique to MCMV infection (see IIIA, 4), is also not well understood. It has been postulated (Weiland et al. 1986) that they are formed when capsids exit the nucleus via nuclear pores and enter cytoplasmic aggregates of capsids. These capsids then receive an envelope via budding into extended cytoplasmic vacuoles derived from the Golgi apparatus. The multicapsid virions are also believed to be released from the cell by exocytosis (Weiland et al. 1986).
C.
Virus Strains: Antigenic and Genetic Relationships
A number of strains of MCMV have been described, but only the Smith and K181 strains have been widely used. The Smith strain was derived from the original isolation of MCMV by Margaret Smith (Smith 1954) and has been the most commonly used strain. The Smith strain has been held by the American Type Culture Collection for a number of years as ATCC number VR-194, recently reissued as VR-1399. However, the strain has been passaged in many different laboratories, resulting in the emergence of variants that exhibit different biological properties (reviewed by Osborn 1982). A more recent report characterizes the variant Vancouver strain, which emerged following the serial passage of the Smith strain in cell culture in Hudson’s laboratory over a number of years (Boname and Chantler 1992). This strain was found to have a 9.4 kb deletion spanning the XbaI I/L junction and a 0.9 kb insertion in the EcoR1 K fragment. It exhibits altered tissue tropism, failing to grow in the salivary gland, while exhibiting enhanced in vitro growth compared to the parental Smith strain. The K181 strain arose in Osborn’s laboratory from the salivary gland passage of the Smith strain in mice, following selection for virulent variants (cited in Misra and Hudson 1980). K181 reached 100-fold higher titers in the salivary glands than Smith (Misra and Hudson 1980), but exhibited different plaque morphology and lower yield in cell culture (Hudson et al. 1988). Differences in restriction endonuclease profiles between Smith and K181 were found for BamH I, Bgl II, EcoRI, HindIII, and XbaI restriction enzymes (Hudson et al. 1988). Where K181 genes such as m133 (Lagenaur et al. 1994), M55 (Elliott et al. 1991; Xu et al. 1996), M75 (Xu et al. 1992), and the termini of the short direct repeats (Marks and Spector 1984) have been sequenced, only minor differences between the DNA and predicted amino acid sequences of Smith and K181 have been found (Rawlinson et al. 1996). However, the exact origin of K181 is unknown. There are three possibilities: it could be a variant of Smith, an endogenous MCMV present in the laboratory mice, or a recombinant between Smith and an endogenous virus. As discussed by Hudson, the use of variants of the Smith strain may explain some of the discrepant results obtained by different investigators (Hudson 1994a). Clearly, there remains a need for reliable reporting of the strain of MCMV used, together
7
with a description of the mode of passage, passage history, and its biological properties. A comparison of the restriction endonuclease profiles of the strain and the ATCC stock of the Smith strain would be valuable. Indeed, restriction enzyme digest analysis of the viral DNA was important in determining that a virulent MCMV strain that was thought to be Smith (Chalmer et al. 1977) had a restriction a pattern identical to that of the K181 strain (Xu et al. 1992) described previously (Hudson et al. 1988). This strain has now been designated K181 (Perth) to identify its laboratory history (Scalzo et al. 1992). Finally, the use of a low-passage master stock would minimize the emergence of variants. Nonetheless, as discussed elsewhere, even a few salivary gland passages of K181 (Perth) in genetically resistant mice were sufficient to mutate the MCMV m157 gene, which is a ligand for the natural killer cell receptor Ly49H controlling MCMV resistance in this mouse strain (Voigt et al. 2003). Following the complete sequencing of the Smith strain (ATCC: VR-194) (Rawlinson et al. 1996), the genetic basis for biological differences between variants of the Smith strain as well as differences between other strains and isolates of MCMV will be readily determined. The construction of bacterial artificial chromosomes of the Smith (Messerle et al. 1997) and K181 (Redwood et al. 2005) strains will facilitate the study of genes associated with viral virulence. Because of its greater virulence in mice (Misra and Hudson 1980), K181 has proved to be useful for in vivo studies of host innate resistance (Chalmer et al. 1977; Grundy (Chalmer) et al. 1981; Allan and Shellam 1984; Scalzo et al. 1992) and the induction of immunopathological diseases (Lenzo et al. 2002; Lawson et al. 1990; Olver et al. 1994; Price et al. 1990), without the need for either immunosuppression or large viral doses. Several viruses that resemble MCMV have been isolated from wild rodents. Raynaud and colleagues isolated an MCMV-like agent from Apodemus sylvaticus (Raynaud and Barreau 1965), and Diosi and colleagues isolated and briefly characterized a similar viral agent from Microtus arvalis (Diosi et al. 1972). These viruses induce characteristic cytopathic effects in mouse embryo fibroblasts, and multicapsid virions are produced (Hudson et al. 1976b) suggesting that they may be related to MCMV. However, as these viruses have not been characterized further, it is not possible to consider them as isolates of MCMV at this stage. Finally, however, it is important to recognize that research on MCMV over the last 50 years has been based on the use of a single strain of MCMV, the Smith strain, and laboratoryderived variants of this strain. These viruses have been maintained by either salivary gland passage in laboratory (usually inbred) mice or by passage in cultured embryo fibroblasts or fibroblastic cell lines, a cell type that is not a primary target of infection in vivo. It is highly likely therefore that the strains or variants of MCMV that are in common use have acquired significant genetic and biological differences from early passages of the Smith isolate.
8
GEOFFREY R. SHELLAM, ALEC J. REDWOOD, LEE M. SMITH, AND SHELLEY GORMAN
This issue has also been recognized in HCMV research (Mocarski and Courcelle 2001b), where the widely used AD169 strain was isolated only 2 years after the isolation of the Smith strain of MCMV (Rowe et al. 1956). AD169 has been maintained by in vitro culture, mainly in fibroblastic cell lines, and multiple variants have arisen in different laboratories. In addition to the use of other strains of HCMV such as Toledo, which was more recently derived from a clinical specimen, and Towne, a strong emphasis in recent HCMV research has been placed on the use of freshly derived clinical isolates. These are now widely used in the in vitro testing of antiviral agents as well as in studies identifying the viral genes associated with particular cell or tissue tropism, disease causation, or immune evasion. A single strain of MCMV that has been strongly adapted by laboratory passage for over 50 years cannot be expected to replicate the range or extent of clinical conditions that are associated with natural HCMV infections in humans. Therefore, if MCMV is to continue to be relevant as a model system for understanding HCMV infections, it is important that more emphasis is placed on the use of recent isolates of MCMV, and care is taken to avoid laboratory adaptation. Free-living wild mice are naturally infected with MCMV (Mannini and Medearis 1961; Rowe et al. 1962; Gardner et al. 1974; Booth et al. 1993; Smith et al. 1993; Singleton et al. 1993, 2000, Gorman et al., 2006) and have been the source of new isolates of MCMV (Booth et al. 1993). Twenty-six isolates of MCMV were obtained from the salivary glands of seropositive wild Mus domesticus that were trapped on subantarctic Kerguelen Island and on the Australian mainland at Geraldton, Nannup, and Walpeup (Booth et al. 1993). The viruses were originally characterized by their induction of cytomegaly in cultured mouse embryo fibroblasts, the reactivity of viral proteins with antibodies to MCMV by Western blotting, and the neutralization of these strains by sera from mice immunized with the K181 strain (Booth et al. 1993). Subsequently, viral genes such as gB and m157 and the H2LLdrestricted cytotoxic T cell epitope within ie1 of many of these strains have been sequenced and compared with the Smith sequence (Xu et al. 1996; Voigt et al. 2003; Lyons et al. 1996). Restriction enzyme analysis of these isolates using EcoRI, XbaI, and HindIII has revealed unique restriction profiles that differ from K181, and demonstrates that natural variation occurs throughout the MCMV genome (Booth et al. 1993). Western blot analysis of viral proteins from these isolates revealed considerable antigenic cross-reactivity among the new isolates when a polyclonal antiserum to K181 was used. However, significant variations in the electrophoretic mobility and intensity of staining of certain proteins were also observed. Interestingly, when 15 of these strains were compared for their ability to replicate in the salivary gland of weanling BALB/c mice, marked differences were observed. Nine of the strains did not replicate to significant titers, while replication was observed with the remaining 6 strains. The failure of 9 strains
that were originally isolated from the salivary gland to replicate there following their isolation and plaque purification is intriguing, especially since all isolates replicated equally well in vitro (Booth et al. 1993; and unpublished observation). This study was also the first report of mixed infection with different MCMV strains in a single mouse. Three mice were found to harbor genetically different strains in their salivary glands. In one mouse, identified as G3, 4 genetically different strains, G3A, G3B, G3C, and G3E, were detected by restriction enzyme analysis. Mixed infection is presumed to have occurred in apparently normal adult mice. This phenomenon appears to be widespread in wild mice (Gorman et al. 2006). For HCMV, mixed infections with different strains have also been described, although most reports relate to immunosuppressed patients. Mixed infections may arise by simultaneous infection with different strains or by sequential infection with individual strains. However, in the latter case, infection would have to be established in the presence of an existing immune response. This eventuality may be favored by the failure of cytomegaloviruses to induce sterilizing immunity. A recent experimental study has established that sequential infection with the G4 and then the K181 strains occurs despite the presence of MCMV-specific antibody and cytotoxic T cells (Gorman et al. 2006). The benefit to the virus of mixed infection is the possibility of complementation between genetically different strains (Cicin-Sain et al. 2005), resulting in persistent infection and enhanced transmission. Studies in laboratory mice offer an excellent means of elucidating this phenomenon.
III.
GROWTH IN VITRO AND IN VIVO A.
In Vitro Propagation
The cytomegaloviruses exhibit species specificity, and infection in vivo is restricted to their natural host species (van Regenmortel et al. 2000). The same is generally true in vitro, although there have been reports of MCMV infecting cell cultures derived from species other than the mouse. 1.
Propagation in Permissive Murine Cells
In permissive cells MCMV replicates to high titer, achieving yields of 10–100 plaque-forming units (pfu) per cell, resulting in the death of infected cells (Hudson 1994a). A feature of MCMV infection is the induction of a characteristic cytoplasmic effect (CPE) in which nuclear and cytoplasmic swelling and chromatin margination occurs (Lussier 1975b) and the formation of both intranuclear and intracytoplasmic inclusions is observed (Ruebner et al. 1966). MCMV replicates in the cell nucleus and buds into the perinuclear cisternae to associate with secondary lysosomes and the Golgi apparatus to form
1. MURINE CYTOMEGALOVIRUS AND OTHER HERPESVIRUSES
intracytoplasmic inclusions (Joseph et al. 1978). These intracellular changes are known as cytomegalia. MCMV can be propagated in a variety of cells or cell lines in vitro (reviewed by Hudson 1994a), and generally replicates most productively in murine embryo fibroblasts (MEF) and in murine fibroblastic cell lines (Osborn 1982). Cell lines that support productive MCMV infection include two fibroblastic lines, NIH 3T3 cells, and the mouse embryo line SC-1, as well as the mouse mammary tumor cell line C127I (Smee et al. 1989). However, for the detection of MCMV by plaque assay, MEF were found to be 10 times more sensitive than these cell lines (Smee et al. 1989). Nonetheless, the use of MEF has some disadvantages. Their preparation is labor intensive, they become resistant to the growth of MCMV after serial passage in culture (Misra and Hudson 1977), and they have a finite life span due to programmed senescence. More recently, the continous murine bone marrow stromal cell line M2-10B4 was found to be as permissive for MCMV replication as MEF (Lutarewych et al. 1997). Other murine cells that are permissive for MCMV include central nervous system stem cells (Kosugi et al. 2000) and microglial cells (Schut et al. 1994). Mast cells can be infected by MCMV, but less than 12% release infectious virus (Gibbons et al. 1990). Similarly, resident peritoneal macrophages can support MCMV infection in culture (Shanley and Pesanti 1983), although reports differ on the extent of infection (Brautigam et al. 1979; Selgrade and Osborn 1974; van Bruggen et al. 1989). As a result of infection, various macrophage functions are down-regulated (Price et al. 1987). Immature dendritic cells also support MCMV replication, although mature dendritic cells are much less permissive (Andrews, Andoniou, et al. 2001; Mathys et al. 2003). In general, the replication of MCMV in these various cells or cell lines results in lower yields than are achieved in MEF (Kosugi et al. 2000; Shanley and Pesanti 1983). 2.
Nonpermissive Murine Cells
In these cells, a low-level infection may occur, resulting in yields of virus of less than 1 pfu per cell (Hudson 1994a). Such cells include the L929 fibroblast line, the Y-1 adrenal cell line, and J774A.1 and other macrophage lines (Hudson 1994a). In other cell types or lines, infection is abortive, and only the transcription of viral genes occurs without the production of viral DNA or infectious virus. Examples include murine T and B cells (Hudson 1994a). 3.
Non-Murine Cells
Several studies have shown that MCMV can replicate in certain primary and continous cell lines derived from other species. These include primary rat embryo fibroblasts (Smith et al. 2005), the rat NRK cell line (Hudson 1994a), African green monkey kidney (BSC-1) cells, primary rabbit kidney cells, and baby hamster kidney (BHK-21) cells (Kim and Carp 1971).
9
However, a number of heterologous cells and cell lines were not found to support MCMV replication, including human cell lines W1-38, HeLa, Hep-2 cells (Kim and Carp 1971), and MRC-5, KB, and the African green monkey cell line Vero (Hudson 1994a). 4.
Features of MCMV Replication In Vitro
The replication of MCMV has been studied extensively in mouse embryo fibroblasts and has been reviewed previously (Osborn 1982; Hudson 1994a). KINETICS OF MCMV REPLICATION In brief, following adsorption of the virus to the fibroblasts, there is a lag period of 16–18 hours before infectious progeny are detected. The lag period is followed by the release of extracellular virus which continues over 18–36 hours p.i. with the maximum yield reached at 30–36 hours p.i.. Infected cells remain viable for 36–48 hours p.i.. As discussed by Hudson (Hudson 1994a), infected cells contain an average of 3000 viral genomes but only release about 100 pfu, which is in part due to the packing of MCMV genomes into multicapsid virions, which contain a number of nucleocapsids but may each produce only a single pfu. These figures may need revision in view of the more sensitive assays that are now available. Two additional factors, centrifugal enhancement of infection and the cell growth cycle, have been shown to influence the replication of MCMV in permissive cells. CENTGRIFUGAL ENHANCEMENT Low-speed centrifugation of the viral inoculum against the cell monolayer at 1000g for 30 minutes results in a 10 to 100-fold increase in the number of pfu that are detected (Osborn and Walker 1968; Hudson et al. 1976a; Hudson 1988, 1994a). The conditions required are precise, and the virus and cells must be centrifuged simultaneously within 60 minutes of inoculation of MCMV. The phenomenon is not dependent on the presence of multicapsid virions and is also applicable to HCMV (Woods et al. 1987) and a variety of other viruses (Pietroboni et al. 1989), although the effect seems to be most pronounced with MCMV. The mechanism is not completely understood, but may involve stabilization of the initial weak interactions between the virus and cellular receptors (Hodgkin et al. 1988). EFFECT OF THE CELL GROWTH CYCLE In permissive cells, the replication of MCMV depends upon the cell cycle (Keil et al. 1984). Production of viral progeny was found to be most effective in cells synchronized at the S phase (Muller and Hudson 1977) and least effective at the G0 phase. This cell cycle dependency is probably due to the occurrence of maximal activity of ribonucleotide reductase (RNR) during the S phase. The IE1 protein of MCMV increases the activity of R2, an enzymatic subunit of RNR, and the M45 protein is a viral homologue of the R1 enzyme subunit of RNR (Brune et al. 2001;
10
GEOFFREY R. SHELLAM, ALEC J. REDWOOD, LEE M. SMITH, AND SHELLEY GORMAN
Lembo et al. 2000). These viral products act to up regulate the catalysis of RNR to convert ribonucleotide diphosphate into deoxyribonucleoside and increase the dNTP pool for viral DNA synthesis and replication. Other cellular proteins utilized by MCMV to regulate the cell cycle and enhance viral DNA synthesis include dihydrofolate reductase, thymidylate synthase (Cavallo et al. 2001), and interferon (IFN)-inducible 204 (Rolle et al. 2001). MULTICAPSID VIRIONS An unusual feature of the replication of MCMV is the production of multicapsid virions, as well as virions containing a single capsid. Multicapsid virions are particles with from 2 to 20 or more capsids enclosed in a common envelope (Hudson et al. 1976b). All size classes of virion appear to be infectious (Hudson et al. 1976b; Chong and Mims 1981), although a multicapsid virion is a single infectious unit. Large multicapsid virions are readily disrupted during purification; free nucleocapsids are noninfectious (Chong and Mims 1981). Mixtures of multicapsid and single virions are produced during in vitro infection of murine embryo cells and fibroblastic 3T3 cells (Hudson et al. 1976b; Chong and Mims 1981; Weiland et al. 1986). Multicapsid virions are also produced during the replication of MCMV in vivo, and have been reported in the liver (Papadimitriou et al. 1984), lung (Reddehase et al. 1985), and spleen (cited in Hudson et al. 1976b). However, multicapsid virions have not been detected in the salivary glands (Hudson et al. 1976b; Chong and Mims 1981), perhaps reflecting the different morphogenesis of MCMV in this organ. The morphogenesis of multicapsid and single virions of MCMV is discussed in Section II, B, 2. The ratio between multicapsid and single virions produced in vivo is not known, but in vitro the multicapsid form accounts for 90%–95% of all virions produced (Hudson et al. 1976b; Chong and Mims 1981). The production of multicapsid virions is not a feature of other herpesvirus infections. Occasional virions encompassing two nucleocapsids have been reported in infections with guinea pig CMV (Fong et al. 1979), rat CMV (Bruggeman et al. 1982), and herpes simplex virus (Nii et al. 1968; Watson 1973), perhaps reflecting the occasional budding of two nucleocapsids into the same vesicle. The production of multicapsid virions has not been detected with HCMV (Hudson et al. 1976b).
B.
Infection of Mice
The replication of MCMV in mice, including the kinetics of the infection, the organs and tissues that support viral growth, the establishment of persistent and latent infection, and host parameters that influence the outcome of the infection (age, mouse strain, route of inoculation) is discussed in Section V. Mice have also been used for virus titration and for virus isolation. Virus can be quantified in vivo by the LD50 assay, in
which adult mice are inoculated intraperitoneally with serial dilutions of viral stock. Interestingly, the effect of varying the dose of virus is pronounced, with mortality changing from 0% to 100% over a 4-fold increase in dose (Selgrade and Osborn 1974; Chalmer et al. 1977; Trgovcich et al. 2000). Hence, serial 1:2 dilutions of virus stock are employed, and ideally 10 mice are used for each virus dilution. Given the well-established age-dependent variation in susceptibility to MCMV (Osborn 1982 and Section V, A, 1), mice of a specified age are used, and the age should be stated. At high virus doses, mice begin to show signs of illness by the second or third day, exhibiting hunching, ruffled fur, reduced activity, and weight loss. Deaths usually occur around day 5. The cause of death is unknown, but as will be discussed in Section V, this assay was very useful in studies of host genetic control of resistance to MCMV in adult inbred mice. The difference in LD50 dose between the most susceptible (BALB/c) and the most resistant mouse strain (B10.BR) was about 30-fold (Grundy (Chalmer) et al. 1981), but even differences of 2- to 4-fold could be measured accurately to distinguish BALB/c from the more resistant C57BL/6 strain (Grundy (Chalmer) et al. 1981). However, this assay raises animal welfare issues, and it is usually not permissible now to use mortality as an endpoint. As an alternative, assessing morbidity by the daily or twice-daily recording of body weight may be useful, since large doses of MCMV cause marked body weight loss, with mice losing 20%–30% of their body weight as the infection progresses (Pomeroy et al. 1998; Bolger et al. 1999). Again, the group size should be sufficiently large to allow firm conclusions to be drawn. ID50 assays involving the inoculation of adult mice with serial dilutions of MCMV, followed by assays for seroconversion, have not been widely used. This may be because MCMV infection does not generally induce high antibody titers in a primary infection. Also, because MCMV is very successful at establishing persistent infection, even rather small viral inocula are often sufficient to allow MCMV to reach the salivary gland, where high viral titers may result. Antibody titers may not therefore adequately reflect the original viral dose. Where the kinetics of infection and the organs involved are known, measuring the viral titer by the plaque assay in a particular organ such as the spleen or liver has been used to discriminate between mouse strains on the basis of innate resistance to MCMV (Allan and Shellam 1984; Mercer and Spector 1986; Scalzo et al. 1992). Mice are commonly used for the preparation of virulent stocks of MCMV by salivary gland passage. Virus harvested from other organs such as the spleen, liver, or kidneys exhibits reduced virulence (Eizuru and Minamishima 1979). Passage of salivary gland–derived MCMV in cell culture results in its attenuation, with effects being observed following even one passage in vitro. However, virulence is restored following salivary gland passage (Osborn and Walker 1971). The mechanism of loss of virulence in cell culture remains to be determined.
11
1. MURINE CYTOMEGALOVIRUS AND OTHER HERPESVIRUSES
Interestingly, even the virulence of salivary gland stocks of MCMV differs according to the time of harvest (Selgrade et al. 1981). Salivary glands also contain factors that inhibit the virulence of MCMV; this property increases after weaning (Mims and Gould 1979). Concern about the presence of cytokines and hormones in salivary gland tissues has convinced Podlech and colleagues (2002) to use tissue culture–prepared virus, despite its reduced virulence. The method of salivary gland passage has been described previously (Osborn and Walker 1970; Osborn 1982; Chalmer et al. 1977). Briefly, inbred female 21-day-old (weanling) mice of a susceptible strain such as BALB/c are inoculated i.p. with a sublethal dose of MCMV. The dose will depend on the strain of MCMV in use. Using the virulent K181 (Perth) strain, we inoculate approximately 5 × 103 pfu per mouse, and harvest salivary glands 17 days later. With this dose, a low level of mortality is observed and high titer stocks are achieved. A 10% w/v stock is prepared from homogenized salivary gland tissue, and after clarification by low-speed centrifugation, the stock is stored in small vials in either liquid nitrogen or at −80°C.
IV. A.
EPIZOOTIOLOGY Natural History
The natural host for MCMV in the wild is considered to be the house Mus musculus domesticus (Mus domesticus). MCMV infection in free-living Mus domesticus is widespread in different parts of the world. In North America, Rowe and colleagues sampled wild mice from a large number of locations and found that most mice were infected with MCMV (Rowe et al. 1962). Similarly, wild mice trapped in 3 separate locations in California showed evidence of infection (Gardner et al. 1974). In Australia, the seroprevalence of MCMV was found to be high in wild Mus domesticus. In mice trapped in wheat fields in 7 separate locations spread across eastern and southeastern Australia, almost 100% had antibodies to MCMV (Smith et al. 1993). Similarly, wild house mice on the Western Australian islands of Thevenard (Moro et al. 1999) and Boullanger (Moro et al. 2003) and the oceanic islands of Kerguelen (Booth et al. 1993), Macquarie (Moro et al. 2003), and Gough (G.R. Singleton, personal communication) exhibited very high rates of MCMV infection. Mus domesticus is a member of the Mus musculus complex of species, which consists of domesticus, musculus, molossinus, castaneus, and bactrianus (Guenet and Bonhomme 2003). A discussion of the taxonomy of the musculus-domesticus complex of species lies outside the scope of this review and is considered in the companion volume of this series. There do not appear to have been studies of the ability of strains of MCMV
obtained from Mus domesticus to infect other members of the Mus musculus complex. Similarly, there are no reports of isolations of cytomegaloviruses from members of this complex other than from Mus domesticus. Thus the question of whether members of this complex other than Mus domesticus can serve as a reservoir of MCMV in the wild remains unanswered. This worldwide distribution of MCMV in wild Mus domesticus reflects not only the very close association between the virus and its host, which is a feature of herpesviruses, but also the successful establishment of populations of house mice in environmentally diverse parts of the world. It is presumed that Mus domesticus reached the Americas, Australasia, and the subantarctic islands during human exploration and colonization (Guenet and Bonhomme 2003). The habitats in which MCMV has been detected in Mus domesticus range from temperate regions in California (Gardner et al. 1973) to arid islands (Moro et al. 1999, 2003) and dry wheatlands with sandy soils (Smith et al. 1993; Singleton et al. 1993, 2000) in Australia to cold subantarctic environments (Booth et al. 1993; Moro et al. 2003). In all environments, MCMV is ubiquitous, with high seroprevalence rates. Interestingly, there was a strong relationship between population size and the extent of MCMV infection. In the mallee wheatlands of southeastern Australia where mouse plagues erupt periodically, the seroprevalence of MCMV was 20%–30% during periods of low mouse abundance but reached approximately 75%–95% at times of high mouse density (Singleton et al. 2000). These trends were supported in other studies (Singleton et al. 1993; Smith et al. 1993).
B.
Natural Infection of Laboratory Mice
Early studies documented natural MCMV infection in laboratory mice. The first isolation of MCMV was made from salivary gland stock that was derived from naturally infected laboratory mice (Smith 1954). Subsequently, studies that employed virus isolation in cell culture or suckling mice found that the rate of infection was 2%–3% (Rowe et al. 1962) or less (Mannini and Medearis 1961), although these are likely to be underestimates. In the 1980s, naturally occurring MCMV infections in laboratory colonies were detected by serological techniques. Using sera provided by different commercial suppliers, 140 of 256 sera contained antibodies to MCMV, and mice from 8 of the 9 suppliers were found to be antibody positive (Anderson et al. 1986). These findings have not been reproduced. Indeed, in a similar study, no anti-MCMV antibodies were detected in sera from mice provided by 4 commercial suppliers (Classen et al. 1987). Naturally occurring MCMV infections in laboratory mice have not been reported in recent years. Nonetheless, MCMV infection may be a problem in laboratory colonies in some parts of the world. A serological survey of 556 mouse sera from 40 laboratory animal facilities in China found a high seroprevalence
12
GEOFFREY R. SHELLAM, ALEC J. REDWOOD, LEE M. SMITH, AND SHELLEY GORMAN
to MCMV, with up to 60% of mice in some institutions being antibody positive (Tang et al. 1992).
C.
Mode of Transmission
A distinctive feature of the cytomegaloviruses is their ability to establish long-term persistence of infectious virus in the salivary gland, as demonstrated in mice (Brodsky and Rowe 1958; Medearis 1964), rats (Bruggeman et al. 1983, 1985), and humans (Lucht et al. 1998). It is presumed that the transfer of infectious virus in saliva to susceptible individuals is the major mode of transmission. In addition, in humans and guinea pigs but not in mice or rats, CMV infection of pregnant mothers may result in transplacental infection of the fetus. The salivary glands in mice and man consist of three paired major glands, which in rank order of salivary output are the submandibular, parotid, and sublinguinal glands. Each gland is connected to the oral cavity by a single excretory duct. In the mouse each gland is limited to one type of secretory cell: serous in the submandibular and parotid glands and mucous in the sublinguinal gland. In addition to these major glands, there are a number of minor glands that in man contribute childhood > adult life]. As expected, a reduction in the load of latent MCMV in mice following therapy with antiviral CD8+ T cells reduced the risk of recurrence (Steffens, Kurz, et al. 1998). These various studies also raised an interesting question. Since the production of infectious MCMV usually results in cell death, the cells that are responsible for producing most of the infectious virus may not be the same cells in which latency occurs, in line with the paradigm established for herpes simplex virus (Reddehase et al. 2002). The recognition of molecular latency in the absence of lowlevel chronic infection as the characteristic latent state in MCMV infection has allowed detailed studies of the mechanism of reactivation from latency and recurrence of infectious MCMV to be undertaken. As already discussed, it has been established that the recurrence of infectious MCMV following reactivation is controlled by the immune system, and CD8+ T cells play the leading role (Polic et al. 1998). However, the molecular reactivation of latent MCMV genomes is regulated by different means. This has been largely determined by analysis of MCMV gene expression in latently infected lungs. It is clear now that latent viral genomes are not necessarily transcriptionally silent. The highly regulated transcriptional program can be interrupted at many checkpoints before assembly and release of infectious virus. The regulation of IE gene expression is an early checkpoint on the way from latency to recurrence. The 1E1/3 transcriptional unit gives rise to 1E1 and 1E3 mRNAs by differential splicing (Keil et al. 1987b; Messerle et al. 1992). This is driven by the P1/3 promoter with a strong upstream enhancer, which serves as a molecular switch, connecting 1E1/3 transcription to the cellular environment. External stimuli, such as the pro-inflammatory cytokine TNF α, act as the first signal in the reactivation pathway, inducing transcription factors that activate the enhancer by binding to defined sequence motifs. 1E3 is believed to be the major transactivator of E gene expression (Messerle et al. 1992; Angulo et al. 2000). MCMV latency is controlled after the initiation of 1E1/3 transcription (Kurz et al. 1999), and this is the second checkpoint in the pathway of molecular reactivation. Reddehase and colleagues have proposed a multistep model of MCMV reactivation and the subsequence recurrence of infectious virus, in which
many checkpoints may be involved (Kurz and Reddehase 1999; Reddehase et al. 2002). Intriguingly, when the early part of this pathway was studied in lung pieces from the lungs of latently infected mice, it was found that latency-associated 1E1 transcription exhibited a random pattern among individual lung pieces. It was calculated that 1 transcriptional focus occurred per 25,000 latent viral genomes, indicating that the expression of ie1 is a rare event (Kurz et al. 1999). Although the reason for the random expression of ie1 in latently infected lungs is not known, it may reflect low-level expression of cytokines such as TNF-α which could activate the sporadic transcription of 1E1. Alternatively, activated CD8+ T cells secreting TNF-α may be responsible (Reddehase et al. 2002). In this concept, TNF-α activates the enhancer, resulting in 1E1 transcription, the synthesis of 1E1 protein, and the presentation of the immunodominant 1E1 nonapeptide to 1E1-specific memory CD8+ T cells. In turn, these cells are activated and release TNF-α, which activates 1E1 expression in another latently infected cell. Although there is more to discover about the latent state of MCMV, it is likely that this knowledge will be generally applicable to HCMV. The mouse model provides the means of studying molecular reactivation in vivo and should continue to provide important insights into how latent CMV infections are regulated. D.
Suitability of MCMV as a Model of HCMV Infection and Disease
Because of the strict species specificity of cytomegaloviruses, it is not possible to undertake experimental studies of HCMV pathogenesis due to the restriction of growth of HCMV in animal models. There is one exception to this: mice with the severe combined immunodeficiency syndrome with engrafted human tissues (SCID-hu mice) support the replication of HCMV in human tissue grafts (Mocarski et al. 1993). A variety of fetal human tissues have been engrafted under the kidney capsule of SCID or SCID beige mice, including thymus, liver, lung, and colon (Mocarski et al.1993; Wang et al. 2005), as well as retinal tissue (Bidanset et al. 2001). HCMV infection is restricted to the grafted human tissues and reaches high titers over 1 month, with persistence for at least 9 months (Mocarski et al. 1993). Since clinical strains of HCMV replicate well in SCID-hu mice while AD169 does not, the model has been very valuable for demonstrating that genes within a 15-kb segment present in clinical strains but not in AD169 are crucial for the replication of HCMV in vivo (Wang et al. 2005). However, the model has significant limitations. It does not replicate the multi-organ infection observed in infected humans, and infection of endothelial and smooth muscle cells of human origin cannot be readily studied. Meaningful studies of HCMV latency or vaccination are also not possible (Wang et al. 2005). Since MCMV infection of mice shares many features with HCMV infection of humans, the mouse model has been used
18
GEOFFREY R. SHELLAM, ALEC J. REDWOOD, LEE M. SMITH, AND SHELLEY GORMAN
extensively for studying the pathogenesis of acute, latent, and recurrent infections. The use of RCMV and guinea pig CMV in their natural hosts has also been valuable for gaining understanding of the pathogenesis of HCMV because these models may reproduce several aspects of HCMV disease (atherosclerosis and congenital disease, respectively) more precisely than MCMV. The merits of all three animal models have been reviewed (Price and Olver 1996). However, the mouse model remains the best studied and has provided a wealth of information about the virus-host relationship. An overview of the usefulness of the mouse model in reproducing HCMV disease is provided in Table 1-1. 1.
Intrauterine Infection and Congenital Disease
HCMV is the leading viral cause of congenital infections in humans in the developed world. With an incidence of 0.2% to 2.2% per live birth in the United States, it is predicted that 40,000 infected infants are born annually in the United States alone, 4000–6000 of whom will experience long-term neurological damage (Britt and Alford 1996). Symptomatic infection is largely restricted to infants born to mothers who experienced a primary rather than a reactivated HCMV infection during pregnancy. Clinical abnormalities in infants with symptomatic
congenital HCMV infection include low body weight, thrombocytopenia, hepatosplenomegaly, microcephaly, and chorioretinitis, and in some, organ dysfunction and death may occur (Britt and Alford 1996). The neurological damage is not reversible and accounts for the long-term morbidity associated with this infection. Hearing loss is the most common neurologic abnormality, and congenital HCMV infection is a leading cause of nonheritable hearing loss in the United States (Hicks et al. 1993). There is thus a clear need for experimental studies to investigate all aspects of intrauterine infection. Unfortunately, there is no conclusive evidence that MCMV can establish intrauterine infection in mice, and most experimental studies use guinea pigs because guinea pig CMV crosses the placenta and initiates fetal infection (Staczek 1990). The early attempts to establish a mouse model of intrauterine infection with MCMV have been reviewed extensively (Osborn 1982; Hudson 1994a) and will not be described in detail. These studies failed to detect in utero infection of mouse embryos following MCMV infection of pregnant female mice, using a variety of inoculation routes, sampling times, and assays for viral detection (Mannini and Medearis 1961; Johnson 1969; Landsdown and Brown 1978). However, several studies using latently infected mothers reported low-level or latent infection of fetuses (Chantler et al.
TABLE 1-1
SUITABILITY OF MCMV AS A MODEL OF HCMV-INDUCED DISEASE Human disease
Features of mouse model
Congenital infection
No transplacental infection, but fetal wastage following maternal infection resembles human condition. MCMV introduced by artificial means affects fetal development of neural tube. No congenital infection. Artificial inoculation results in microphthalmia and cerebral atrophy in fetal mice. Infection of newborns i.c. results in severe necrotizing ependymitis, encephalitis, and cerebral malformation, as in human congenital infection. Infection of adult nude or SCID but not normal mice results in CNS infection and encephalitis. Induction of retinitis in adults requires direct supraciliary intra-ocular infection. Progressive focal necrotizing retinitis observed, enhanced by CD8+ depletion. Mouse not a model for hearing loss due to congenital infection. Infection at birth results in perilabyrinthitis, whereas congenitally infected humans develop endolabyrinthitis. Resembles the human disease in essential features; arises after immunosupression, but viral replication is only indirectly related to pathology. Severity of disease reduced but not prevented by antiviral drugs. Antiviral T cell response necessary for disease development, but the relative importance of CD4 or CD8+ T cells is controversial. Resembles human disease. In adults, virus replicates in hepatocytes. Inflammatory foci are composed mainly of T cells; are initiated by NK cells, IFN-γ, and MIP-α. NK cell–produced IFN-γ important in controlling murine infection. In lethal infection, direct viral cytopathic effects are dominant. High TNF-α and IFN-γ titers may have a toxic rather than beneficial effect. Resembles the human disease observed after heart transplantation. Normal mice experience acute and persistent cardiac inflammation mediated mainly by CD8+ T cells in the presence of very low levels of virus. Viral dose and host genetics influence outcome. MCMV-induced cardiac myosin-autoantibodies may contribute, although no parallels described in humans. HCMV may contribute to disease by enhancing inflammation in blood vessels, augmenting migration of smooth muscle cells, and promoting their proliferation by blocking apoptosis. Mouse model demonstrates similar features but is mainly used for mechanistic studies using gene knockout mice. Adrenal infection in immunodeficient mice resembles adrenal necrosis detected in HIV/AIDS patients. Resembles effect of HCMV. In model of irradiation and auto-reconstitution of bone marrow, infusion of antiviral CD8+ T cells prevents hemopoietic failure and provides basis for human immunocytotherapy. MCMV prevents marrow reconstitution by inhibiting production of hemopoietic cytokines by stromal cells.
Infection of CNS
Retinitis Effect on the developing ear Interstitial pneumonitis
Hepatitis
Myocarditis
Atherosclerosis
Adrenalitis Hemopoietic failure
1. MURINE CYTOMEGALOVIRUS AND OTHER HERPESVIRUSES
1979; Baskar et al. 1985), suggesting that either transplacental infection of a reactivated virus from latently infected mothers or transmission within the germ line had occurred. It has not been established why MCMV does not readily cross the placenta. The murine placenta is hemotrichorial, with multiple cell layers separating the maternal and fetal circulations, in contrast to the hemodichorial structure of the human (and guinea pig) placenta (Pereira et al. 2005). This is considered the most likely reason for the failure of MCMV to cross the placenta. However, there is no evidence to suggest that the murine placenta is intrinsically resistant to the passage of viruses. Polyoma virus (McCance and Mims 1977), ectromelia (Mims 1969), lymphocytic choriomeningitis virus (Traub 1936; Lehmann-Grube 1964), and Coxsackie viruses (Selzer 1969) all cross the murine placenta. If the structure of the murine placenta is a barrier to viral transmission to the fetus, it must be much more effective against MCMV. It is also possible that the Smith strain, which has been used in all studies, has lost its ability to cross the placenta during its adaptation to passage in laboratory mice or cell culture. Further studies of intrauterine infection with MCMV should be undertaken using a variety of viral isolates recently derived from wild mice. Furthermore, the ability of MCMV to infect fetal mice in utero could be investigated in free-living wild mice that are naturally infected with MCMV. Despite the inability of MCMV to cross the placenta and establish productive infection of the fetuses, the infection of pregnant females results in extensive fetal wastage with increased fetal death, fetal resorption, delayed births, and reduced body weight of newborn pups (Mannini and Medearis 1961; Medearis 1964; Johnson 1969; Landsdown and Brown 1978; Huang et al. 1986; Fitzgerald and Shellam 1991; Neighbour 1976). This effect was dose-dependent and varied with the age of the embryo and the timing of the infection (Huang et al. 1986). Time-dependent effects were also observed when pregnant females were infected with MCMV at earlier times (Neighbour 1976). When the infection was timed to coincide with ovulation and pre-implantation development, implantation frequently failed to occur. However, when mice were infected 14 days before or 4 days after mating, the pregnancy rate and preimplantation loss was unaffected. The timing of infection is also important in CMV infections in other species (Griffiths and Hsiung 1980; Boppana et al. 1993; Stagno et al. 1986). A role for genetically determined host resistance in modulating the effect of MCMV infection on fetal outcome was observed in pregnant BALB/c, BALB.K, and CBA mice that were inoculated on day 8 of pregnancy (Fitzgerald and Shellam 1991). In susceptible BALB/c, the percentage of dead or resorbed fetuses was significantly higher in infected compared with control mothers, while in the two genetically resistant strains, no adverse effect of infection was observed. Maternal infection resulted in reduced fetal body weight in all strains, particularly in genetically susceptible BALB/c mice. Infectious virus was not detected in the fetuses either by plaque or co-cultivation assays. Therefore it is likely that the effect of MCMV infection
19
on the fetuses was indirect, reflecting either placental dysfunction following infection or maternal illness affecting maternal metabolism or the immune system. In women who experience a primary HCMV infection during pregnancy, increased fetal loss in the absence of infection of the fetus or the placenta has been reported (Griffiths and Baboonian 1984). Thus, while MCMV does not readily infect fetuses transplacentally, it mimics the indirect effects of maternal HCMV infection on fetal outcome. The study of Fitzgerald and Shellam (1991) also established that genetically determined innate resistance to MCMV influences the level of infection within the fetus itself following direct in utero inoculation of MCMV. Viral titers in fetuses, which were directly inoculated in utero at day 15 of gestation, reflected the resistance status of the mouse strain. These findings suggest that if genetic factors do play a role in modulating HCMV infection in humans, they could influence the outcome of primary HCMV infection during pregnancy. EFFECT OF MCMV ON EMBRYONIC DEVELOPMENT When MCMV was inoculated into the endometrial lumina of pregnant mice 4 days after coitum at the time of embryonic implantation, fetal development was affected. Litter sizes were reduced, and the incidence of abnormal fetuses was significantly increased (Baskar et al. 1987). The fetal damage included maldevelopment of the neural tube and head and ectodermal abnormalities. Infectious MCMV was recovered from the embryos by co-cultivation with MEF. To mimic sperm-mediated MCMV transfer to ova, MCMV DNA was micro-injected into uninfected fertilized murine ova that were cultured and transferred to pseudo-pregnant mice (Baskar et al. 1993). Again, reduction in litter size, fetal growth retardation, maldevelopment, or death as observed. Using PCR and in situ hybridization, MCMV was found in the brains, skin, and salivary glands of the fetuses. However, a recent study in which MCMV was inoculated intratesticularly found no evidence of virus transmission to fertilized oocytes, blastocysts, fetal tissues, or newborn animals following mating of infected males with uninfected females (Tebourbi et al. 2001). Nonetheless, despite the artificial nature of some of these studies, they show that the presence of MCMV in the genital tract at the time of embryonic implantation has the potential not only to initiate fetal infection but also to interfere with morphogenesis. INFECTION OF GONADAL TISSUES MCMV infects the gonadal tissues of both sexes. In infected males, MCMV was recovered from the epididymal sperm, seminal vesicles, and testes (Neighbour and Fraser 1978; Baskar et al. 1986) as already discussed, and was present in the testes during acute infection in athymic nude mice and in the testes of latently infected immunocompetent mice (Dutko and Oldstone 1979). MCMV was found within Leydig cells (Baskar et al. 1983) and in spermatocytes and spermatozoa (Dutko and Oldstone 1979). In females,
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GEOFFREY R. SHELLAM, ALEC J. REDWOOD, LEE M. SMITH, AND SHELLEY GORMAN
MCMV infects the ovaries (Mims and Gould 1979), where it has been located in stromal cells (McCordick and Smith 1936; Brautigam and Oldstone 1980). These studies establish the potential for the sexual transmission of MCMV. EFFECT ON REPRODUCTION The presence of MCMV in gonadal tissues suggests that it may affect reproductive outcomes. While this has not been well studied in laboratory mice, it has been examined in laboratory-reared, wild-derived Mus domesticus housed in secure outdoor enclosures (Farroway et al. 2002). MCMV spread rapidly through founder populations and their offspring. There was little impact on breeding performance. However, the body condition of young mice born in enclosures with MCMV was affected, and where 2 strains of MCMV were present, there was a 20% reduction in the survival of young males. Nonetheless, there was no effect of MCMV on the rate of increase of the populations overall. 2.
Interstitial Pneumonitis
Cytomegalovirus-associated interstitial pneumonitis is a frequent complication in immunosuppressed patients undergoing allogeneic transplantation, especially bone marrow transplantation (BMT). While the incidence of HCMV infection is similar after autologous and allogeneic bone marrow transplantation, HCMV-associated interstitial pneumonitis is more common after allogeneic bone marrow transplantation with concomitant graft versus host disease, where fatalities are often high (Wingard et al. 1988; Meyers et al. 1986; Miller et al. 1986). A diffuse infiltration of mononuclear cells into the lung parenchyma leading to congestion of the septa are features of this disease. The etiology of the disease is complex. Rather than simply reflecting the lack of host control of this opportunistic agent due to immunosuppression, the disease may be immunopathological in nature (Grundy et al. 1987). The replication of HCMV in the lung appears to be only indirectly related to the development of pathological effects, and a host antiviral immune response is an essential component in the development of the disease. MOUSE MODEL The use of MCMV has been very important in understanding the etiology of this disease. Although interstitial pneumonitis has been reported in the absence of immunosuppression in adult BRVS mice following i.n. MCMV infection (Jordan 1978), immunosuppression is usually required. Following the i.n. inoculation of 8 × 104 pfu of MCMV in adult BALB/c mice, followed by a single dose of cyclophosphamide, extensive interstitial pneumonitis developed 10–14 days later (Shanley et al. 1982). However, if the administration of cyclophosphamide was continued, interstitial pneumonitis was not observed, even though the titers of MCMV were increased. The pathology of the disease resembled that seen in humans; mononuclear cells infiltrated the alveolar septa, reducing the alveolar spaces, and accumulation of amorphous eosinophilic material was observed. Lung weights were markedly increased
(Shanley et al. 1982). While the severity of the disease was proportional to virus production, the use of antiviral drugs reduced but did not prevent the development of the disease (Shanley and Pesanti 1985; Shanley et al. 1985, 1988). Other means of immunosuppression such as antilymphocyte serum (Brody and Craighead 1974), graft versus host disease (Shanley et al. 1987, 1988), irradiation (Reddehase et al. 1985), or malnutrition (Price et al. 1990) have also been successful in inducing pneumonitis in infected mice. In these studies, pneumonitis was associated with an influx of T cells into the lungs (Shanley et al. 1987; Shanley and Ballas 1985). In contrast, T cell–deficient athymic nude mice infected with MCMV developed pneumonitis with a progressive focal nodular interstitial pattern, which was quite different from the diffuse pneumonitis seen in the cyclophosphamide model (Shanley et al. 1997). Coalescence of focal areas in the lungs of nude mice suggested that lung pathology was related directly to viral damage in these mice (Shanley et al. 1997). PNEUMONITIS AS AN IMMUNOPATHOLOGICAL DISEASE Using data obtained from the mouse model and from clinical reports, Grundy and colleagues proposed that CMV-pneumonitis is an immunopathological disease that arises following immunosuppression, is triggered by CMV infection in the lung, and is mediated by T cells responding to viral antigens expressed in lung tissue. Since antiviral therapy reduced but did not eliminate pneumonitis, it was suggested that after inducing the response, the continued presence of the virus was not necessary (Grundy et al. 1987, 1985; Grundy 1990). However, this hypothesis has proved to be controversial (Barry et al. 2000; Morris 1993; Podlech et al. 2000). Identifying CD4+ T cells as critical for the development of immunopathology in Grundy’s hypothesis, Barry and colleagues have contended that CD4+ T cells are only present at very low levels in the blood soon after transplantation in humans, when most cases of pneumonitis occur, and that NK cells and CD8+ T cells are the major cell populations present in the lungs (Barry et al. 2000). Podlech and colleagues have emphasized the strongly protective role of antiviral CD8+ T cells in controlling MCMV in the lungs and their major role in controlling HCMV in transplant recipients (Podlech et al. 2000). Instead, it is proposed that early after BMT, when CD8+ T cell responses are low, CMV replicates in the lung without restriction, inducing a cytokine storm involving interferon-γ from NK cells stimulating the release of TNF-α from alveolar macrophages (Barry et al. 2000). Morris takes a somewhat different view, arguing that the immunosuppression associated with transplantation increases the likelihood that CMV viremia will result in pneumonia and that protective CD8+ T cell and NK cell responses will be reduced. The pathological effects in the lung of CMV, irradiation, and cytotoxic drugs act together to enhance the development of pneumonia (Morris 1993). More research is needed to resolve these opposing concepts and to account for the relative lack of interstitial pneumonitis in HCMV-infected AIDS patients. The value of the mouse model
1. MURINE CYTOMEGALOVIRUS AND OTHER HERPESVIRUSES
is clearly evident. It provided the data for the original concept and also the proof that the reconstitution of CD8+ T cells after BMT was responsible for controlling CMV infection (Reddehase et al. 1985). It will also be valuable for elucidating the antigenic targets of the response in the lung, the role of cytokines, and the interplay between the cell types involved. The use of strains of MCMV recently derived from wild mice may also enhance the relevance of the model. 3.
Hepatitis
MCMV-induced hepatitis in mice has been widely studied as a model of hepatitis induced by HCMV. In humans, the liver is frequently infected in cases of disseminated HCMV infection, both in immunocompetent and immunocompromised individuals (Varani and Landini 2002). In vitro, HCMV replicates in hepatocytes, which are the major target cells, producing infectious virus. In vivo, the virus also replicates in bile duct epithelial cells and stromal cells (Sinzger et al. 1999). Early and late HCMV antigens have been detected in the livers of acutely infected patients (Sinzger et al. 1999; Barkholt et al. 1994). Thus, human liver cells can be infected by HCMV in vivo and in vitro. In symptomatic congenital HCMV infection, hepatitis is common. Clinical features include hepatomegaly, which may persist for months, increased levels of serum transaminases, and hyperbilirubinemia with jaundice. Cytomegalic cells are seen most commonly in the bile duct epithelium and infrequently in hepatocytes, indicating that the bile duct epithelium is an important site of HCMV replication in infants (Kosai et al. 1991). In immunocompetent adults, HCMV occasionally induces mononucleosis (Britt and Alford 1996), and in these cases the liver is often involved, with accompanying fever and moderately elevated transaminase activity. A mild, nonspecific hepatitis with a mononuclear cell infiltrate is observed (Ten Napel et al. 1984). Cytotoxic T cells are the predominant cell type (Pape et al. 1983), and may be responsible for rapidly clearing HCMV infected cells in the liver (Pape et al. 1983; Asanuma et al. 1999). Cytomegalovirus infection of the liver is more pronounced in transplant patients. Up to 20% of liver transplant patients have CMV hepatitis (Paya et al. 1989) and, in severe cases, a high HCMV viremia, high levels of serum transaminases, and prolonged fever is common. Histopathologic studies show the presence of typical inclusion bodies in hepatocytes, but with an inflammatory response that is reduced compared to hepatitis in immunocompetent individuals. Early and late HCMV antigens have been detected, providing evidence of productive infection. The hepatic cell damage observed in transplant patients is thought to be caused directly by the virus rather than by the inflammatory response (Varani and Landini 2002). Interestingly, the liver is rarely involved in HCMV infection in HIV-positive patients. Given the morbidity associated with HCMV hepatitis, further research is needed and the study of MCMV-induced hepatitis in mice is amply justified.
21
MCMV HEPATITIS IN MICE The ability of MCMV to infect the liver and induce hepatitis was recognized in the early studies of this virus (McCordick and Smith 1936; Ruebner et al. 1964, 1966; Henson et al. 1967). The liver is infected soon after i.p. inoculation. Hepatocytes become infected by 24 hours p.i., with the extent of infection depending on the dose and route of inoculation. Kupffer cells do not appear to be infected when salivary gland virus is used (Mims and Gould 1978a). At sublethal doses of MCMV, foci of infection develop and increase in size as inflammatory cells are attracted to sites of virus replication. Genetically determined host resistance influences this response; in highly resistant CBA mice, few inflammatory cells were observed compared with the less resistant C57BL strain (Selgrade and Osborn 1974). Viral titers peak in the liver at 2–4 days p.i., with higher titers detected in the livers of susceptible BALB/c than resistant C3H or CBA mice (Allan and Shellam 1984; Mercer and Spector 1986). Infection is more quickly resolved in the livers of resistant mice. The effect of MCMV infection on the liver has been studied in various ways. Foci of infected cells and cells with intranuclear inclusions have been analyzed histologically (Papadimitriou et al. 1982; Olver et al. 1994; Orange et al. 1995). Nuclear DNA has been measured by cytophotometry on liver sections (Papadimitriou and Shellam 1981), and the extent and location of viral antigen expression has been measured by immunocytochemistry (Shanley et al. 1993; Trgovcich et al. 2000; Olver et al. 1994). The measurement of serum transaminase levels has also proved valuable (Papadimitriou et al. 1982; Shanley et al. 1993; Bolger et al. 1999). The levels of alanine transaminase (ALT) and aspartate transaminase (AST) in serum are elevated by MCMV infection, but ALT is more liver specific and appears to be a more sensitive measure of MCMV-induced liver damage (Bolger et al. 1999). The liver is a major target of MCMV infection following i.p. or i.v. inoculation. Several studies have shown that severe hepatitis, which is induced by lethal doses of MCMV in susceptible strains of mice, is likely to be the cause of the early mortality observed (Shanley et al. 1993; Trgovcich et al. 2000). At doses of 1 × 105 pfu and higher, BALB/c mice developed high titers of MCMV in the liver by day 2 that persisted until the deaths of the mice from day 5. Large areas of necrosis were evident in liver sections from day 2 p.i., becoming confluent by day 5 PI with evidence of acute hepatic dystrophy and hypoxic necrosis (Trgovcich et al. 2000). Serum levels of AST were markedly augmented. However, inflammatory infiltrates were modest, suggesting that direct cytopathic effects rather than an immunopathological response were responsible for the liver damage. The very high serum levels of interferon-γ and TNF-α, which developed just before mortalities occurred, were taken as evidence of a toxic rather than a protective role for these cytokines in lethal MCMV infection (Trgovcich et al. 2000). At sublethal doses, titers of MCMV were lower and were reduced by day 5, serum AST titers were not elevated significantly, and, although interferon γ levels were elevated by day 4, they began
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GEOFFREY R. SHELLAM, ALEC J. REDWOOD, LEE M. SMITH, AND SHELLEY GORMAN
to decline by day 5. Genetically resistant B10.BR mice exhibited very low MCMV titers in the liver and little or no elevation of transaminases following i.p. inoculation of a dose of MCMV that was lethal for susceptible BALB/c mice (Shanley et al. 1993). During sublethal infection, however, different mechanisms are involved. Viral titers in the liver declined by day 5 p.i., and foci of inflammatory cells accumulated over the first week p.i., clearing after day 11 with a few foci remaining for 7–8 months in susceptible strains (Olver et al. 1994). From early times p.i. Mac-1+ cells were present in cellular infiltrates, and CD4+ and CD8+ T cells were present after the second day, peaking at day 7 and declining to control levels in resistant CBA and C57BL/6 mice by day 56. However, in susceptible BALB/c and A/J mice, CD8+ T cells persisted for over 30 weeks, dispersed throughout the liver parenchyma. Interestingly, in NK cell–deficient beige mice, the levels of CD8+ T cells were higher at day 7 in both C57BL/6 and CBA mice (Olver et al. 1994). NK cells are important in controlling MCMV in the liver as well as in other tissues. In C57BL/6 beige mice, MCMV replication in the liver was increased (Shellam et al. 1981) and hepatitis was more pronounced (Papadimitriou et al. 1982). NK cells with the distinctive large granular lymphocyte morphology are attracted to the liver during MCMV infection (McIntyre and Welsh 1987). Depletion of NK cells resulted in increased viral titers and hepatitis (Orange et al. 1995; Bukowski et al. 1983). Interestingly, the production of IFN-γ by NK cells was shown to be important in the control of MCMV in the liver (Orange et al. 1995). There is a dichotomy in the effector mechanisms of NK cells, with the production of IFN-γ by NK cells being the major NK effector mechanism in the liver, while in the spleen, cytotoxicity involving perforin is the means by which NK cells control MCMV infection (Tay and Welsh 1997). In the liver, IFN-γ appears to control infection at least in part by the induction of the antiviral molecule nitric oxide (Tay and Welsh 1997). However, very high levels of IFN-γ may also have adverse effects (Pomeroy et al. 1998). A role for TNF-α in the exacerbation of MCMV- induced liver disease has been demonstrated. It caused early hepatic necrosis independent of NK cell or T cell responses, and induced pathology that was largely responsible for liver damage at early times after infection. TNF-α was required for increased levels of serum transaminases but not for the development of inflammatory foci in the liver (Orange et al. 1997). It appears that TNF-α was also responsible for the progressive MCMV-induced liver disease in immunodeficient mice, which resulted in the death of the animals (Orange et al. 1997). 4.
Cardiovascular Diseases
MYOCARDITIS Viral infection has been recognized as one of the etiological agents of myocarditis in humans (Friman et al. 1995; Feldman and McNamara 2000), which may progress to dilated cardiomyopathy. This is a serious and often terminal
condition (Friman and Fohlman 1997; Kawai 1999). Viruses commonly associated with myocarditis include the picornaviruses, particularly Coxsackie viruses, orthomyxoviruses, and herpesviruses (Friman et al. 1995; Huber 1997). HCMV infection has been associated with the development of myocarditis (Wink and Schmitz 1980; Cohen and Corey 1985); cytomegalic cells with intranuclear inclusions are found in the endothelium and myocardial cells in the hearts of infants and adults with generalized CMV disease (Ahvenainen 1952; Myerson et al. 1984). CMV infection is a common major complication in heart transplant recipients and AIDS patients, causing increased rates of allograft rejection and cardiac dysfunction (Fernando et al. 1994; Herskowitz et al. 1994). Furthermore, heart transplant recipients experiencing a primary HCMV infection following transplantation have a 46% incidence of developing myocarditis (Arbustini et al. 1992). Myocardial damage after HCMV infection may persist and lead to dilated cardiomyopathy (Ando et al. 1992; Maisch et al. 1993). However, myocarditis remains difficult to diagnose, and a viral etiology is even more difficult to establish (Feldman and McNamara 2000; Aretz 1987; Lieberman et al. 1993). Hence, an animal model of cytomegalovirus myocarditis is important for establishing the pathogenesis of disease. The induction of myocarditis in mice by MCMV has been well studied and has provided valuable insights into the mechanisms of virusinduced cardiac damage (reviewed in Fairweather et al. 2001). MCMV infection results in the appearance of inflammatory lesions in the heart of both adult (Mims and Gould 1979; Papadimitriou et al. 1982; Bartholomaeus et al. 1988; Leung et al. 1986; Gang et al. 1986; Craighead et al. 1991; Price et al. 1991) and newborn mice (Lussier 1974; Fitzgerald et al. 1990; Price et al. 1991). These lesions persist and may become calcified (Lussier 1974; Gang et al. 1986). The main features of the model are exemplified in a recent study (Lenzo et al. 2002). In adult BALB/c mice, the inoculation of 1 × 104 pfu of salivary gland–derived MCMV [K181 (Perth)] results in myocarditis. At days 3–5 p.i., a few inflammatory foci are seen, without histological evidence of myocyte necrosis. Soon afterward, a mixed cellular infiltrate comprising lymphocytes, macrophages, and polymorphonuclear leucocytes develops around areas of myocyte necrosis. This acute focal myocarditis peaks at day 7 and declines by day 21. A chronic phase of myocarditis, which is characterized by a more dispersed interstitial inflammatory infiltrate with larger areas of necrosis, develops by day 28 p.i., is still prominent at day 56 (Lenzo et al. 2002), and persists until at least day 100 (Price et al. 1991; Lawson et al. 1990). In contrast, C57BL/6 mice inoculated with the same dose develop only a minimal acute response, with minor focal inflammation at days 5–7 p.i. and complete resolution of myocarditis by about 14 days. The size of the inflammatory foci was also generally smaller in C57BL/6. By measuring the number of inflammatory foci per heart section, clear-cut differences in the kinetics of the response and in mouse strain variations in the response could be determined (Lenzo et al. 2002).
1. MURINE CYTOMEGALOVIRUS AND OTHER HERPESVIRUSES
Other studies have also investigated mouse strain variations in the development of myocarditis (Lawson et al. 1990; Price et al. 1991). In order to separate the effects of genetically determined resistance to MCMV infection from any genetic effects that might influence the development of myocarditis per se, adult mice received doses of MCMV that were adjusted in proportion to their sensitivity to lethal infection (Price et al. 1991). Genetically resistant BALB.K and C3H mice bearing the H-2k haplotype developed myocarditis that was as severe as that in susceptible BALB/c mice, indicating that in this kind of study, H-2 genes did not affect resistance to myocarditis other than through modulation of the viral load. The level of MCMV replication in the heart of adult BALB/c mice is, however, very low (Craighead et al. 1991, 1992; Gang et al. 1986; Lenzo et al. 2002; Lawson et al. 1990). Infectious virus generally is cleared from cardiac tissue by 7–10 days p.i., during which time apoptosis of cardiac cells is observed. It was also shown that MCMV replicates in and lyses cardiac myocytes in vitro (Lenzo et al. 2002; Lawson et al. 1990). Despite the absence of infectious virus, the ie1 and gB genes of MCMV can be detected in the heart for up to 100 days in infected adult BALB/c mice (Lenzo et al. 2002). The presence of viral iel and gB RNA transcripts up to day 35 indicates active viral replication, even though this cannot be detected by plaque assay. Beyond this time during chronic myocarditis (days 35–100 PI), viral ie1 RNA transcripts are detected, while viral gB RNA transcripts are not, suggesting that a latent infection had been established in the heart. The ability of MCMV to establish latency in heart tissue has been demonstrated previously (Wilson et al. 1985; Rubin et al. 1984). The cellular infiltrates around infected or necrotic cells consist of Mac-1+ cells and CD4+ T cells, as well as CD8+ T cells, which predominate. Inflammatory T cells persist in the hearts of infected BALB/c mice for long periods (Price et al. 1991; Lenzo et al. 2002). ROLE OF T CELLS AND NK CELLS IN MYOCARDITIS A primary role for T cells in the development of myocarditis has been established. T cell–deficient BALB/c nu/nu mice did not develop myocarditis during acute infection (Lawson et al. 1989) and, similarly, immunocompetent mice in which CD4+ and CD8+ T cells were depleted failed to develop the disease by day 9 p.i., when the acute phase of the disease occurs (Fairweather et al. 2001). However, the relative importance of CD4+ and CD8+ T cells remains unclear. In BALB/c mice, monoclonal antibody depletion or reconstitution of irradiated thymectomized mice with naive CD4+ or CD8+ T cells before MCMV infection demonstrated that CD4+ T cells were responsible for myocarditis at day 21 p.i. (Craighead et al. 1992). However, a similar study using antibody depletion demonstrated a role for both subtypes, but with CD8+ cells playing the major role at day 9 p.i. (Fairweather et al. 2001). Interestingly, in the latter study, a clear protective role for NK1.1+ NK cells was established. The depletion of NK1.1+ NK cells from C57BL/6 or
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BALB.B6–Cmv1r congenic mice resulted in high levels of myocarditis, comparable to the levels in BALB/c mice (Fairweather et al. 2001). Presumably, NK cells and T cells are protective through the reduction of the viral load that reaches the heart, while T cells also accumulate around the sites of myocardial infection and initiate disease. ROLE OF ANTIBODIES Antibodies may also play a role in myocarditis. In response to MCMV infection, both BALB/c and C57BL/6 mice produce antibodies that recognize myosin. However, only susceptible BALB/c mice produce antibodies during chronic myocarditis that interact with the cardiac isoform of myosin (O’Donoghue et al. 1990; Lawson et al. 1992). The passive transfer of affinity-isolated IgG antibodies to cardiac myosin from late immune sera of infected BALB/c mice induces cellular inflammation and myocardial necrosis in naive mice (Lawson et al. 1992). These cardiac myosin–reactive antibodies cross-react with MCMV polypeptides and the S2 region of cardiac myosin (Lawson et al. 1992; Fairweather et al. 1998; Lawson 2000). Furthermore, several monoclonal antibodies that were raised against MCMV and neutralize the virus also react with cardiac myosin (Lawson et al. 1991). Interestingly, cardiac myosin itself is an autoantigen that can induce myocarditis in mice in the absence of viral infection, inducing high titer anti-myosin antibodies (Lawson et al. 1992). Thus, the candidate autoantigen, cardiac myosin, is capable of inducing immunopathological responses in the heart. On the basis of these findings, a possible role for molecular mimicry between MCMV proteins and cardiac myosin in the chronic phase of MCMV myocarditis has been proposed (Lawson 2000). Using evidence obtained from the Coxsackievirus B3 model of myocarditis in mice, which resembles MCMV myocarditis in many ways (Fairweather et al. 2001), the effect of the immunomodulator lipopolysaccharide (LPS) on MCMV myocarditis has been studied (Lenzo, Fairweather, et al. 2001). LPS enhanced myocarditis and serum levels of TNF in BALB/c mice and in the resistant C57BL/6 strain. However, the role of cytokines in MCMV myocarditis in the absence of immunomodulators remains to be determined. RELEVANCE OF THE MOUSE MODEL As well as providing insights into HCMV myocarditis, the relevance of the laboratory model is demonstrated by two additional studies. First, six genetically different isolates of MCMV obtained from wild Mus domesticus induced typical myocarditis in inbred BALB/c mice. Furthermore, myocarditis was detected histologically in 30% of wild mice, all of which were seropositive for MCMV. This does not necessarily imply that the myocarditis was due to MCMV infection. Sera from BALB/c mice infected with wild isolates as well as sera from free-living wild mice, contained antibodies that reacted with MCMV and the S2 region of cardiac myosin (Fairweather et al. 1998). These observations suggest that myocarditis induced in the laboratory model resembles the disease that occurs in free-living wild mice.
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GEOFFREY R. SHELLAM, ALEC J. REDWOOD, LEE M. SMITH, AND SHELLEY GORMAN
Second, the mouse model has been useful for determining the source of primary CMV infection in cardiac transplantation. Hearts from actively or latently infected mice were a source of primary infection in immunosuppressed recipients (Rubin et al. 1984). This study provides valuable information about CMV infection in human transplantation. ATHEROSCLEROSIS A number of infectious agents have been implicated in atherosclerosis (Mattila et al. 1998), and accumulating evidence supports a role for HCMV. The association between HCMV and atherosclerosis has been recently reviewed (Degre 2002). It is based on epidemiological studies and the demonstration of HCMV antigens and DNA in atherosclerotic plaque tissue, as well as on investigations into the mechanisms by which HCMV infection of vascular endothelial and smooth muscle cells could augment the development of atherosclerosis or restenosis. Perhaps because atherosclerosis is a complex multifactorial disease in which infection is only one of several etiologies, evidence for a causal role for HCMV is not yet strong. However, HCMV infection could contribute in several ways, including augmenting inflammation through the activation of NF-κb (Speir et al. 1998) and the release of proinflammatory cytokines and chemokines, enhancing the migration of smooth muscle cells (Zhou et al. 1999) and binding and inhibiting p53, thus blocking apoptosis and promoting the proliferation of smooth muscle cells (Tanaka et al. 1999). HCMV may also act as a prothrombotic agent (Pryzdial and Wright 1994). The ability of RCMV to induce vascular disease in rats has made it the best animal model in which to study the role of cytomegaloviruses in atherosclerosis and transplantation vasculopathy (Span et al. 1992; Kloover et al. 2000; Martelius et al. 2000; Epstein et al. 1996). However, the mouse model has the advantage of the availability of a large number of gene knockout strains and viral mutants for mechanistic studies. As a result, there has been an increase in the use of this model recently. MCMV infection of BALB/c mice results in the expression of MCMV antigens in vascular smooth muscle and endothelial cells, with inflammatory lesions and an increase in low-density lipoprotein cholesterol in the serum (Berencsi et al. 1998; Dangler et al. 1995). In C57BL/6 apolipoprotein-E knockout mice that are genetically susceptible to the development of atherosclerosis, MCMV infection causes a significant increase in the size of atherosclerotic lesions (Hsich et al. 2001), in the expression of pro-atherosclerotic genes in the aorta (Burnett et al. 2004), and in the levels of monocyte chemoattractant protein-1 (MCP-1), which is known to exacerbate atherogenesis (Rott et al. 2001). MCMV infection induced IL-6 from endothelial cells, which enhanced the production of MCP-1 (Rott et al. 2003). In transgenic mice over-expressing MCP-1 in the myocardium and pulmonary arteries, MCMV infection accelerated myocarditis and pulmonary artery inflammation (Froberg et al. 2001). A role for MCMV M33, a G protein–coupled receptor homolog (Davis-Poynter et al. 1997) resembling HCMV US28, in the migration of vascular smooth
muscle cells has been established (Melnychuk et al. 2005). The contribution of MCMV to atherogenesis has been recently reviewed (Froberg 2004). 5.
Adrenalitis
Early studies reported adrenal involvement following MCMV infection (McCordick and Smith 1936; Mims and Gould 1979), but subsequent reports differ about the level of infection achieved. Using the Smith strain, Shanley found no evidence of infection in the adrenal glands of immunocompetent adult BALB/c mice (Shanley and Pesanti 1986). In contrast, the K181 (Perth) strain of MCMV established acute infection in the adrenal glands of adult BALB/c (Price et al. 1996). However, when athymic nude or irradiated BALB/c mice were used, MCMV replicated to high titers in the adrenals (Shanley and Pesanti 1986; Reddehase et al. 1988). This resembled the situation with AIDS patients, in whom extensive adrenal necrosis associated with presumed CMV infection is observed (Tapper et al. 1984), and loss of adrenal function may occur (Pulakhandam and Dincsoy 1990). In MCMV-infected nude mice, the progressive destruction of the adrenals could be arrested by the adoptive transfer of normal spleen cells (Shanley and Pesanti 1986) or CD4+ but not CD8+ T cells from immune mice (Shanley 1987). In irradiated mice, CD8+ but not CD4+ T cells from immune mice protected against adrenal infection (Reddehase et al. 1988). Adrenal infection did not appear to compromise adrenal function in immunocompetent BALB/c mice, as assessed by the levels of circulating ACTH (Price et al. 1996). The levels of corticosterone increased after infection. Also in this strain, an adrenocortical response was important for survival, as adrenalectomized mice died at doses up to 5-fold lower than those they could usually tolerate (Price et al. 1996). A discussion of the physiological role of the adrenal gland in MCMV infection is beyond the scope of this review. A recent review (Silverman et al. 2005) covers this topic comprehensively. 6.
Infection of the Central Nervous System
In humans, HCMV infection of the CNS occurs during severe intrauterine infection. At birth, the majority of these infants show evidence of microcephaly, neurologic abnormalities, periventricular calcification, mental retardation, sensorineural hearing loss, and impaired vision (Bopana et al. 1992; Boppana et al. 1997; Ramsay et al. 1991). CNS involvement also occurs as a complication of HCMV infection of AIDS patients (Cheung and Teich 1999), in whom glial and neuronal cells may be infected (Wiley et al. 1986) and ependymal and subependymal regions of the parenchyma are involved. The retina and cochlea are also often infected (Keithley et al. 1989; Rabb et al. 1992). The mouse does not provide a natural model of congenital infection of the CNS. To circumvent this, Tsutsui (1995) directly
1. MURINE CYTOMEGALOVIRUS AND OTHER HERPESVIRUSES
infected fetuses in utero at various stages of gestation. Microphthalmia and cerebral atrophy developed in fetuses inoculated at 8.5 days of gestation, and viral-infected cells were widely distributed in the mesenchyme, suggesting that mesenchymal infection may be critical in disrupting organogenesis. Infection at day 15.5 led to massive necrosis of the brain. In another investigation, infection of CNS stem cells prepared from fetal brains suppressed growth and inhibited neuronal differentiation, suggesting that these outcomes could be the primary causes of brain disorders in congenital CMV infection (Kosugi et al. 2000). The cerebral infection of newborn mice also gives rise to changes similar to those seen in human congenital infections. Intracerebral inoculation of suckling mice resulted in severe necrotizing ependymitis and encephalitis followed by marked cerebral malformation (Lussier 1973; Lussier 1975a). Clinical signs include mild ataxia, hind leg weakness, and runting. Necrosis and mineralization of the subventricular zone, cerebral cortex, and hippocampus were detected at autopsy. In vitro studies have established that most cell types of the fetal and adult brain support MCMV infection (Schneider et al. 1972; Willson et al. 1974). The ability of MCMV to infect the CNS of adult athymic nude or SCID mice has been studied as a parallel to HCMV infection of the CNS in AIDS patients (Reuter et al. 2004). While immunocompetent mice were resistant to CNS infection following peripheral inoculation, immunodeficient mice showed evidence of CNS infection from 21 days p.i. Many different cell types became infected in the CNS, resulting in meningitis, encephalitis, and choroiditis. Using MCMV tagged with green fluorescent protein, MCMV-infected Mac3+/CD45+ leucocytes were identified in the blood stream and in the brain, suggesting that these cells were the source of infection of the CNS. Despite the ability to establish CNS infection in adult, immunodeficient mice, it appears nonetheless that the cells of the developing brain are intrinsically more susceptible to MCMV infection (van den Pol et al. 2002). 7.
Retinitis
Interest in ocular infections with MCMV has arisen because of the emergence of HCMV-associated retinitis in patients who are immunosuppressed by HIV infection. Prior to the use of highly active antiretiroviral therapy (HAART) to control HIV, HCMV retinitis was the leading cause of vision loss and blindness among patients with AIDS (Cunningham and Margolis 1998), and it remains a chronic, sight-threatening ophthalmic problem among HIV patients who do not respond to HAART (Jabs and Bartlett 1997). Ocular abnormalities including chorioretinitis also affect 20%–25% of symptomatically congenitally infected infants (Pass et al. 1980). The features of HCMV retinitis include typical inclusions and cytomegaly of retinal neurons and retinal pigment epithelium, inflammation in the retina consisting of neutrophils or mononuclear cells, and full-thickness
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retinal necrosis and necrosis of the retinal pigment epithelium (de Venecia et al. 1971; Egbert et al. 1980; Pepose et al. 1985). In AIDS patients with HCMV retinitis, choroiditis is also frequently observed and, less commonly, HCMV infection of cells of the optic nerve (Pepose et al. 1985; Grossniklaus et al. 1987). In immunocompetent mice infected i.p. with MCMV, the virus was recovered from homogenates of eye tissue over the first week and from intraocular fluids most frequently on days 11–21 and from explant cultures of the eye and optic nerve as long as 120 days after infection. However, ocular abnormalities were not detected, and the focal retinal necrosis that is common in HCMV retinitis in humans was not observed (Bale et al. 1984). Similarly, MCMV inoculation by the intraocular route (anterior chamber or intravitreal) results in only minimal disease (Hayashi et al. 1985; Bale et al. 1990), although latent MCMV was detected in ocular tissue (Bale et al. 1990). The use of immunosuppression by cyclophosphamide following inoculation of MCMV behind the lens resulted in uveal infection and focal retinal necrosis involving the outer retinal layers, but lacked the features of HCMV retinitis (Holland et al. 1990). However, the use of the supraciliary route for intraocular inoculation was successful at inducing progressive focal necrotizing MCMV retinitis in adult, immunocomponent BALB/c mice (Atherton et al. 1991). Depletion of CD8+ and to a lesser extent CD4+ T cells induced acute retinitis that progressed to retinal necrosis (Atherton et al. 1992). The depletion of NK cells also promoted the development of retinitis in this model (Bigger et al. 1998). The adoptive transfer of CD8+ T cells from MCMV-immune mice protected against the development of retinitis induced by MCMV in T cell–depleted mice (Bigger et al. 1999). The perforin cytotoxic pathway was recently found to be more important than the Fas/FasL cytotoxoic pathway in protecting against MCMV retinitis in C57BL/6 mice (Dix et al. 2003). Mice with the retrovirus immunodeficiency syndrome MAIDS exhibited enhanced frequency and severity of MCMV retinitis. This was found to be directly related to a decrease in the perforin cytotoxic pathway in these mice. In conclusion, MCMV does not induce retinitis unless it is administered intraocularly by the supraciliary route. Nonetheless, valuable information about protective immune mechanisms has been obtained using the mouse model. 8.
Effect on the Developing Ear
Congenital HCMV infection in humans, whether symptomatic or asymptomatic, is considered to be the leading cause of sensorineural deafness (Fowler et al. 1997; Hicks et al. 1993). Hearing loss is the most common neurologic abnormality in these infants, and may range from mild to profound (Britt and Alford 1996). Of concern is the high rate of hearing loss in infants with no clinical evidence of infection. Hearing loss may progress during the first years of life in congenitally infected children, suggesting the presence of ongoing CNS infection (Britt and Alford 1996).
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GEOFFREY R. SHELLAM, ALEC J. REDWOOD, LEE M. SMITH, AND SHELLEY GORMAN
The mouse does not provide a model of congenital CMV infection leading to hearing loss. However, the newborn mouse ear appears to be equivalent in its embryological development to the ear of the human fetus at 15 weeks’ gestation. The intracranial inoculation of newborn mice with MCMV resulted in infection of the inner ear, inducing perilabyrinthitis in the cochlea, although this was not achieved by i.p. or i.n. inoculation (Davis and Strauss 1973; Davis and Hawrisiak 1977). Infection of cranial nerve ganglion cells and Schwann cells was observed, and in explants of trigeminal ganglia from infected newborn mice, Schwann cells, satellite cells, and neurons showed evidence of infection (Davis et al. 1979). In vitro organ culture of the inner ear from mice infected as newborns established that MCMV replicates in mesenchymal cells (Davis 1981). These observations contrast with the effect of HCMV on the inner ear following congenital infection in humans, where endolabyrinthitis involving the replication of HCMV in epithelial rather than mesenchymal cells is observed (Davis and Hawrisiak 1977). Thus, MCMV infection of newborn mice does not appear to replicate the pathological effects on the developing ear observed following congenital HCMV infection. The role of viruses including CMVs in vestibular neuritis and viral infections of the inner ear has been further reviewed (Davis and Johnsson 1983; Davis 1993). 9.
Effects on Hemopoiesis
The effects of MCMV on the bone marrow have been widely studied to find explanations for the defects in hemopoiesis that accompany HCMV infections in human BMT. In the absence of an adequately reconstituted immune system, BMT patients are prone to debilitating HCMV-induced diseases, particularly interstitial pneumonitis. Although rates of HCMV disease in BMT have declined markedly in recent years due to improved clinical management (reviewed in Pass 2001), HCMV infection remains a significant clinical concern. The mouse model has proved to be ideal for studying this problem. In normal adult mice, sublethal MCMV infection causes a marked atrophy of the bone marrow over the first week p.i., without establishing significant infection in bone marrow cells. Recovery occurs from day 7 p.i. (Gibbons et al. 1994). MCMV interferes with hemapoiesis by reducing the colony forming units–spleen (CFU-S) and CFU-granulocyte/macrophage (CFU-GM) (Gibbons et al. 1995). In persistently infected mice, no effects on the bone marrow were observed, but after 5-fluorouracil treatment, marrow recovery was delayed in infected mice. Stromal cells from infected mice showed a reduced capacity to support hemopoiesis and harbored latent infection (Mori et al. 1999). In experiments employing sublethal irradiation and autoreconstitution of bone marrow from surviving stem cells, MCMV infection prevented marrow reconstitution and interfered with the earliest step in hemopoiesis, the generation of CFU-S-1, without establishing marrow infection (Mutter et al. 1988).
However, the adoptive transfer of antiviral CD8+ T cells prevented hemopoietic failure (Mutter et al. 1988). This study reached the important conclusion that CD8+ T cells normally play a vital role in preventing the anti-hemopoietic effect of MCMV infection. In human allogeneic BMT, the high incidence of lethal HCMV disease therefore probably results from incomplete reconstitution by histoincompatible hemopoietic cells. Consequently, a delay in the generation of immune effector cells results in a failure to control HCMV disease (Mutter et al. 1988). Further studies in the mouse model of auto-reconstitution of bone marrow have established that MCMV-induced bone marrow failure is associated with a greatly reduced capacity of bone marrow stromal cells to produce the essential hemopoietic cytokines, stem cell factor, granulocyte colony stimulating factor, and IL-6 (Mayer et al. 1997). MCMV was shown to impair the engraftment of bone marrow cells in the stroma, which was associated with a lack of hemopoietic progenitor cells expressing SCF receptors and a reduced level of SCF gene expression in stromal cells (Steffens et al. 1998). A recent study suggests a useful therapeutic approach to protect against lethal MCMV infection in either congenic or allogeneic BMT. The inclusion of small numbers of common lymphoid progenitors accelerates immune reconstitution and protects against MCMV infection (Arber et al. 2003). Thus, the mouse model has provided valuable insights into the mechanisms of CMV-induced failure of bone marrow transplantation. It has also established the crucial protective role of CD8+ T cells and the possible use of these cells or common lymphoid progenitors in immunocytotherapy.
VI.
THE IMMUNE RESPONSE TO MCMV INFECTION
Immune responses in mice infected with MCMV involve both the innate and adaptive arms of the immune system. To establish persistent infection, MCMV has evolved mechanisms to modulate immune responses. MCMV also replicates in cells regulating immune responses, including macrophages and dendritic cells. This topic is the subject of a large literature, and the interested reader is directed to more comprehensive reviews. 1.
Immunosuppression
MCMV infection results in significant but temporary suppression of the immune system. A range of responses are affected, including responses to mitogens, antigens, allograft rejection, delayed type hypersensitivity responses, and the clearance of other pathogens (reviewed in Osborn 1982). While MCMV infection may cause splenic necrosis (Mims and Gould 1978b) and the loss of T and B cells (Trgovcich et al. 2000), early evidence suggested that the major effect of MCMV was on
1. MURINE CYTOMEGALOVIRUS AND OTHER HERPESVIRUSES
regulatory cells (Kelsey et al. 1977; Osborn 1982). In support of this, recent evidence shows that MCMV infection of dendritic cells interferes with their key coordinating role in the immune system (Andrews, Andoniou, et al. 2001). Furthermore, infection of another important accessory cell, the macrophage, results in its functional impairment (see below) and loss of response to cytokines, which enhance its response and defense against pathogens (Popkin et al. 2003). Other effects of MCMV on the immune response are discussed in Section 7 below. 2.
Macrophages and Dendritic Cells
Blood monocytes and tissue macrophages play a pivotal role in the pathogenesis of MCMV infection (Hanson et al. 1999). Macrophages are major target cells in many tissues following infection with MCMV (Hudson et al. 1978; Mercer et al. 1988; Mims and Gould 1978a; Selgrade and Osborn 1974; Stoddart et al. 1994) and may harbor latent viral DNA (Brautigam et al. 1979; Koffron et al. 1998; Pollock et al. 1997). Circulating blood monocytes disseminate virus during acute infection and differentiate into mature macrophages, an event that favors MCMV replication (Cavanaugh et al. 1996; Collins et al. 1994; Heise, Connick, et al. 1998; Hesie, Pollock, et al. 1998b; Heise and Virgin 1995; Mitchell et al. 1996). Macrophages are also important mediators of inflammatory and innate responses to infection and, when activated, infiltrate sites of infection (Heise and Virgin 1995) to initiate early antiviral responses by producing tumor necrosis factor-α (TNF-α) or interferon-γ (IFN-γ) (Hanson et al. 1999). However, IFN-γ inhibits the replication of MCMV in macrophages (Presti et al. 2001). The phagocytic function of macrophages is altered following infection with MCMV (Katzenstein et al. 1983; Shanley and Pesanti 1983; van Bruggen et al. 1989). Macrophages contribute to protection by decreasing the viral load and acting as antigen-presenting cells to activate T cell responses (Hamano et al. 1998). Macrophages may protect other highly permissive cell types from MCMV infection (Hanson et al. 1999). In vitro replication of MCMV is slower in macrophages than in fibroblasts (Heise and Virgin 1995; Shanley and Pesanti 1983; van Bruggen et al. 1989). As initiators of immune responses, dendritic cells (DC) contribute to the control of MCMV infection. Immature DC process antigen and, once activated, migrate to lymphoid organs, where they initiate primary immune responses and activate naive and activated T cells. Through the release of cytokines and chemokines, activated DC also play a role in the recruitment and activation of NK cells, NKT cells, macrophages, and B cells. Various subsets of DC limit MCMV replication during infection. Plasmacytoid DC infected with MCMV in vitro produce IFN-α/β, which induces other DC to produce IL-12, in particular the CD11b+ subset (Dalod et al. 2002, 2003). IFN-α/β also promotes the accumulation of the plasmacytoid DC and
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the maturation of CD8α+ DC (Dalod et al. 2003). CD8α+ DC are also essential for the expansion of Ly49H+ NK cells via a mechanism involving the production of IL-12 and IL-18 (Andrews et al. 2003). Recent studies have identified two phases in the response of DCs to MCMV. Upon initial infection, immature DC transiently up regulate MHC and costimulatory molecules (Andrews et al. 2001; Mathys et al. 2003). However, within 4 days, when most immature DC are productively infected, the cell surface expression of MHC and costimulatory molecules is reduced, and the DC lose their ability to prime T cells or respond to maturation stimuli. Mature DC are either infected at a lower level (Andrews et al. 2001) or are not productively infected with MCMV (Mathys et al. 2003). 3.
Production of Interferon and Other Cytokines
IFN is an important inducer of nonspecific host defenses during acute infection with MCMV. The protective role of IFN was first shown by the administration of antiserum specific for IFN-α/β, which significantly reduced the resistance of mice to MCMV infection (Grundy et al. 1982). Many other studies have demonstrated the importance of IFNs for controlling MCMV infection by administering neutralizing antibodies or exogenous cytokines to infected mice (Allan and Shellam 1985; Chong et al. 1983; Cousens et al. 1997; Heise and Virgin 1995; Lucin et al. 1992; Martinotti et al. 1990, 1992, 1993; Orange and Biron 1996b; Orange et al. 1995; Quinnan and Manischewitz 1987). During MCMV infection, IFN α/β peaks were detected in plasma at 6 hr (Shellam et al. 1981) and 48 hr (Allan and Shellam 1985) and again at 10 days PI (Tarr et al. 1978). IFN-γ levels in sera peaked at 40 hr PI (Cousens et al. 1997; Orange and Biron 1996b; Ruzek et al. 1997). IFN-α/β induces an antiviral state during early times of infection, prior to the development of adaptive host immune responses. The kinetics of the IFN-α/β response mimic NK cell activation and proliferation (Bukowski et al. 1984; Shellam et al. 1983). NK cell activity is enhanced by IFN within the first few days of infection (Welsh 1986). IFN has effects on other antiviral mechanisms via NK-cell independent pathways, as shown by prophylactic treatment with IFN-β of NK-cell deficient mice (Bukowski et al. 1987). IFN-α inhibits MCMV replication by impairing viral IE gene transcription (Martinotti et al. 1993). This is achieved by down-regulating the activity of transcription factors such as NF-κB that activate the IE enhancer in the nuclei of infected cells (Gribaudo et al. 1993). IFN-α/β also has regulatory effects on the subsequent expression of endogenous cytokines including 1L-12 and IFN-γ, which induce protective immune responses during MCMV infection (Cousens et al. 1997). IFN-γ is a key regulator during all phases of MCMV infection (Presti et al. 1998). Control of acute infection by MCMV is mediated by IFN-γ secretion by NK cells (Heise and Virgin 1995; Orange et al. 1995) and T cells (Karupiah et al. 1998;
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GEOFFREY R. SHELLAM, ALEC J. REDWOOD, LEE M. SMITH, AND SHELLEY GORMAN
Shanley et al. 2001). Functions of IFN-γ during MCMV infection include macrophage activation (Heise and Virgin 1995), enhancement of antigen presentation by infected cells to CD8+ T cells via MHC class I–dependent pathways (Hengel et al. 1994), and inhibition of lytic MCMV replication and gene expression at the cellular level (Heise and Virgin 1995; Lucin et al. 1994; Presti et al. 1998). IFN-γ may also regulate chronic MCMV replication in the salivary glands (Jonjic et al. 1989; Koszinowski et al. 1990; Lucin et al. 1992; Presti et al. 1998). NK cells are the dominant source of IFN-γ soon after infection. MCMV has evolved immune modulation strategies to reduce the effectiveness of IFN-γ in infected cells (Popkin et al. 2003). MCMV induces the production of other cytokines that are detectable in sera during the acute stage of infection, including IL-18, IL-12, TNF-α, IL-1α, and IL-6 (Pien et al. 2000; Ruzek et al. 1997). Using mice with cytokine deficiencies or neutralized cytokine functions, IL-6 was a pivotal mediator of the glucocorticoid response, and IL-1 contributed to IL-6 production (Ruzek et al. 1997). There may be organ-specific cytokine responses to MCMV infection. There were higher concentrations of IL-10 in the lungs of BALB/c than C57BL/6 mice following infection (Geist and Hinde 2001). 4.
Natural Killer (NK) Cells
The protective role of NK cells in MCMV infection was identified over 20 years ago and remains a very active area of research (see also Section V, B, 2). The NK cell response is a preformed defense mechanism that is active before the development of acquired immunity. NK cells are large granular lymphocytes that lyse a restricted variety of target cells upon contact. NK cells control MCMV replication by direct lysis of infected cells (Bukowski et al. 1985; Shellam et al. 1981) and by producing cytokines (Orange et al. 1995). Direct killing requires NK cells to be localized in close proximity to virusinfected target cells. However, NK cells can recycle after direct lysis to kill other infected cells. NK cells also regulate downstream T cell responses (Su et al. 2001). NK cells respond rapidly to MCMV infection (Bancroft et al. 1981), and limit the severity, extent, and duration of acute infection (Shellam et al. 1981; Bukowski et al. 1983, 1984), and may also regulate viral persistence in the salivary gland (Bukowski et al. 1984). The importance of NK cells in controlling MCMV infection has been shown by depletion or adoptive transfer of NK cells. Depletion of NK cell activity in adult mice using antibodies to asialo-GM1, or NK1.1 monoclonal antibody rendered C5BL/6 mice susceptible to MCMV (Bukowski et al. 1984; Scalzo et al. 1992; Shanley 1990; Welsh et al. 1990, 1991, 1994). Infection of newborn mice with MCMV is usually lethal. However, the adoptive transfer of cloned NK cells or NK cell–enriched fractions from naive adults to neonatal mice prevented mortalities (Bukowski et al. 1985, 1988) by reducing splenic viral titers (Welsh 1986). Genetic susceptibility to infection by MCMV correlates with the inability to mount a sufficient
NK cell response (Bancroft et al. 1981; Shellam et al. 1981). The interaction between NK cell receptors and viral ligands is discussed in Section V, B. Cytokines produced during acute MCMV infection have specific NK cell activation roles. The production of IFN-α/β is required for the accumulation, blastogenesis, and cytotoxicity of NK cells (Biron 1997; Orange and Biron 1996b; SalazerMather et al. 2002). The cytotoxic activity of NK cells is greatly augmented by the presence of IFNs, although they also protect the target cells from lysis (Welsh 1986). MCMV-induced IL-12 is responsible for early NK cell IFN-γ production and viral control, as in vivo IL-12 neutralization by antibody treatment blocks these events. However, treatment with neutralizing IL-12 failed to alter these NK cell responses at 7–9 days p.i. (Orange and Biron 1996a). IFN-α stimulated IFN-γ production by NK cells, but also inhibited NK cell cytotoxicity (Orange and Biron 1996a). NK cells may utilize different mechanisms to regulate viral infection in different organs. NK cells accumulate in the spleen and liver at the sites of viral replication in an IL-12, IFN-γ, and TNF-α dependent manner (Dokun et al. 2001). In the spleen of C57BL/6 mice, NK cells control MCMV via a perforindependent (IFN-γ independent) cytotoxic mechanism (Tay and Welsh 1997). In contrast, IFN-γ produced by NK cells is a major immune regulator of infection in the liver (Orange et al. 1995; Orange and Biron 1996 a, 1996b; Tay and Welsh 1997). The formation of early inflammatory foci in the liver (Bukowski et al. 1983; Orange et al. 1997; Olver et al. 1994) is dependent upon the rapid accumulation of NK cells, IFN-γ, and macrophage inflammatory protein-1α (MIP-1α) (Salazer-Mather et al. 2002). In C57BL/6 mice, liver pathology occurred independently of NK and T cells. However, NK cells may limit the damage elicited by TNF-α (Orange et al. 1997). NK1.1+T (NKT) cells also regulate the innate immune response, especially in tumor surveillance. However, NKT cells are not directly involved in restricting MCMV infection, but when appropriately activated, produce IFN-γ, which activates NK cells to control MCMV infection (van Dommelen et al. 2003). 5.
B Cell–Mediated Immune Responses
The humoral immune response is stimulated during acute MCMV infection. Neutralizing antibody was detected by 3 days p.i. (Araullo-Cruz et al. 1978), while non-neutralizing antibody was first detected in sera at 8–10 days p.i., when titers were 10-fold higher than neutralizing titers (Manischewitz and Quinnan 1980). Antibody titers to MCMV persisted until as late as 6 months p.i. in BALB/c mice (Classen et al. 1987). Longterm persistence of CMV in the salivary glands may contribute to serum titers of antibody, as it does in the rat model (Kloover et al. 2002). Serum IgM antibodies were detected in mice as early as 3–5 days, while IgG antibody was detected between 5–7 days, reaching peak levels at 20 days p.i. (Lawson et al. 1988). The IgG isotype response to MCMV antigens is predominantly
1. MURINE CYTOMEGALOVIRUS AND OTHER HERPESVIRUSES
IgG2a (Price et al. 1993). IgA antibodies were not detected in the sera of MCMV-infected mice (Lawson et al. 1988). Antibody is not critical for controlling primary MCMV infection, as high antibody titers occur in mice with significant viral titers in organs (Lawson et al. 1988). Antibody also is not essential for recovery from a primary MCMV infection, as mutant mice devoid of B cells recovered from infection (Jonjic et al. 1994). Nonetheless, the role of antibody is important, as mice depleted of immunoglobulin from birth were 10-fold more susceptible to MCMV than normal mice (Selgrade et al. 1976). Passive administration of neutralizing antibody to MCMV can restrict virus replication but not the establishment of viral latency (Araullo-Cruz et al. 1978; Farrell and Shellam 1991; Shanley et al. 1981). Antibodies may also limit the reactivation of latent virus and its subsequent dissemination. Factors that influence the production of antibody include the MCMV strain, viral dose, genetic resistance, and secondary infections. Antiviral antibody was detected after the administration of 10–100 but not 1 pfu of MCMV (K181) (van Dommelen and Shellam, unpublished observation). There was no correlation between antibody titers and mouse resistance status (Lawson et al. 1988; Price et al. 1993). However, the specific viral proteins recognized vary between mouse strains (Farrell and Shellam 1989). A large number of viral proteins are recognized by antiMCMV antibody. More than 50 viral antigens were recognized by rabbit sera raised against extracts of MCMV-infected mouse cells (Chantler and Hudson 1978). These proteins included both structural and nonstructural components of the virion (Selgrade et al. 1983). Several structural glycoproteins of MCMV stimulate production of neutralizing antibodies, including gp52, gp105, gp87, and the gp150 complex (Loh 1991; Loh et al. 1988; Loh and Qualtiere 1988). MCMV-induced increases in serum immunoglobulin levels are driven by cytokines such as IL-6 (Karupiah et al. 1998). IFN-γ is important for the predominance of IgG2a antibody isotype compared to the IgG1 isotype in MCMV infection (Karupiah et al. 1998). The appearance of autoantibodies in the serum of MCMV-infected mice (Bartholomaeus et al. 1988) may be due to polyclonal activation of B cells (Price et al. 1993) and may contribute to autoimmune diseases that accompany MCMV infection in mice (Bartholomeaus et al. 1988). Such antibodies have broad cross-reactivity with auto-antigens, conventional antigens, and unrelated viral antigens (Karupiah et al. 1998; Price et al. 1993). 6.
T Cell–Mediated Immune Responses
Both CD4+ and CD8+ T cells play critical roles in the immune response to MCMV. The role of T cells in protective immunity was initially observed in T cell–deficient nude mice, which were very susceptible to MCMV (Grundy and Melief 1982; Starr and Allison 1977). CD8+ T LYMPHOCYTES Adoptive transfer of the CD8+ subset of sensitized T lymphocytes limited MCMV dissemination,
29
prevented tissue destruction, and protected mice against lethal disease (Reddehase et al. 1984a, 1988; Reddehase, Mutter, et al. 1987; Reddhase, Mutter, Munch, et al. 1987). Cytotoxic T lymphocytes (CTL) are crucial for the clearance of acute MCMV infection in BALB/c mice. CTL lysed MCMV-infected fibroblasts in vitro and were found in the spleens of mice from day 3, peaking on day 7–8 (Ho 1980; Quinnan et al. 1978; Sinickas et al. 1985) and were active until 14 days p.i. (Ho 1980). Sensitized CTL recognize both structural and nonstructural viral antigens. However, almost 50% of CTL in BALB/c mice are primarily directed against the nonstructural pp89 (IE1) protein of the Smith strain (Koszinowski, Reddehase, et al. 1987; Reddehase et al. 1984b; Reddehase and Koszinowski 1984). The pp89 protein is recognized by specific CTL in conjunction with the MHC class I H-2d locus (Alexander-Miller et al. 1993; Koszinowski, Keil, et al. 1987). BALB/c (H-2d) mice were protected against lethal MCMV challenge by vaccination with a recombinant vaccinia virus expressing pp89 (Del Val et al. 1988; Koszinowski, Keil, et al. 1987; Volkmer et al. 1987), which, however, did not prevent infection and morbidity (Jonjic et al. 1988). Epitope mapping studies, using vaccinia virus recombinants expressing pp89 synthetic peptides, have shown that pp89 contains an immunodominant CD8+ T cell epitope, YPHFMPTNL (Del Val et al. 1988; Reddehase et al. 1989), of which only 5 amino acids are essential for CTL recognition (Koszinowski et al. 1991). IE1-specific (pp89) CD8+ T cells dominate during the acute phase of infection (Holtappels, PahlSeibert, et al. 2000). Most IE1-specific CD8+ T cells belong to the CD62lo subset of resensitized memory effector cells. IE1specific CTL are likely to be frequently resensitized during latent infection of the lungs and may be involved in maintaining latency (Holtappels, Pahl-Seibert, et al. 2000). Nucleotide sequencing of MCMV isolates derived from wild mice identified variations between amino acids 147 and 192 of pp89, which included the region encompassing the CTL epitope (amino acid residues 168–176) (Lyons et al. 1996). Four groups of variants at this locus were defined. Some wild isolates and the laboratory strains K181 and Vancouver had complete identity with the Smith strain (Lyons et al. 1996). Polyclonal pp89 (Smith)-specific CTL only weakly recognized target cells infected with MCMV from most variant groups (Lyons et al. 1996). Immunization of mice with YPHFMPTNL conferred significant protection against the laboratory isolate K181, but there was no protection of mice challenged with G4 or N1 isolates (Lyons et al. 1996). Other CTL epitopes identified within the MCMV genome include H-2d restricted CTL epitopes from the m04, M45, and m164 gene products (Gold et al. 2002; Holtappels, Thomas, et al. 2000; Holtappels et al. 2002). Eighty percent of memory CD8+ T cells in spleen and pulmonary infiltrates were specific for the ie1 and m164 peptides (Holtappels et al. 2002). M83 and M84 may also encode epitopes that are recognized by CD8+ T cells (Holtappels et al. 2001). M84 encodes a protein that shares significant amino-acid homology with the HCMV pp65
30
GEOFFREY R. SHELLAM, ALEC J. REDWOOD, LEE M. SMITH, AND SHELLEY GORMAN
tegument protein, a major target of protective CTL in humans (Morello et al. 2000). CD8+ T cell responses are not essential for the early control of MCMV replication in the presence of effective NK cell responses, such as those present in genetically resistant CBA and C57BL/6 mice (Lathbury et al. 1996). Susceptible mouse strains (BALB/c, A/J) are dependent upon CD8+ T cells. However, congenic BALB/c mice containing the resistance Cmv1r allele did not require a CD8+ T cell response for viral clearance (Lathbury et al. 1996). Thus, NK cell and CD8+ T cell responses are independently responsible for viral clearance, although CD8+ T cells are required for recovery from infection. Karrer and colleagues (2003) determined that the kinetics of the acute CD8+ T cell response after MCMV infection were characterized by rapid expansion of activated T cells followed by a contraction phase. Thereafter, MCMV-specific memory CD8+ T cells steadily accumulated over time in the spleen, lymph nodes, liver, lungs, and blood. At 1 year p.i., 20% of all CD8+ T cells were specific for the pp89 epitope, and there was also a gradual restriction in the use of the variable region of the TCR β-chain (Karrer et al. 2003). Continuous or repetitive exposure to antigen during latency may enlarge these memory T cell populations over time, as observed in HCMV infection of human populations (Gillespie et al. 2000; Weekes et al. 1999). CD4+ T LYMPHOCYTES CD4+ helper T cells are not absolutely required for initiating CD8+ T cell–mediated immune responses (Ahmed et al. 1988; Jonjic et al. 1989; Reddehase et al. 1988) but are required for viral clearance in certain organs and may enhance CD8+ T cell responses. In mice depleted of the CD4+ subset, clearance of replicating virus occured in infected tissues except for the salivary glands (Jonjic et al. 1989; Lucin et al. 1992). CD4+ lymphocytes can compensate for the absence of CD8+ lymphocytes (Jonjic et al. 1990; Koszinowski et al. 1991). IFN-γ is required for CD4+ T cell activity in the salivary glands (Lucin et al. 1992). CD4+ T cells are also required for the MCMV-specific DTH and IgG antibody responses (Lathbury et al. 1996). The response to MCMV in the salivary gland is intriguing. MCMV continues to persist for long periods despite the presence of CD8α+ DC, γ/δ T cells, NK cells, CD4+ and CD8+ T cells, and the expression of 1FNγ, IL-10, and CC chemokines (Cavanaugh et al. 2003). 7.
Immune Evasion by MCMV
A number of the genes of MCMV and other herpesviruses are responsible for establishing and maintaining the virus in its host. They control a variety of processes, including moderating the immune response, or immune evasion, the inhibition of apoptosis, and specific cell tropism. These genes are not
required for viral replication and can often be deleted without in vitro consequence. A list of the known immune invasion genes is shown in Table 1-2. Detailed discussion of these genes can be found in various reviews (Alcami and Koszinowski 2000; Scalzo 2002; Yewdell and Hill 2002; Krmpotic et al. 2003; Mocarski, 2004). The products of immune evasion genes of MCMV affect various aspects of the immune system. Some examples of the genes involved are m129/131 and M33, which encode a chemokine homolog MCK-2 and a receptor respectively, M27, whose product inhibits the interferon response, and m147.5, which inhibits antigen presentation by DC (Table 1-2). Others, such as those that down-regulate MHC class II expression (Alcami and Koszinowski 2000), antigen presentation by macrophages (Popkin et al. 2003), or DC (Andrews et al. 2001), are yet to be characterized. The best-studied of the immune evasion genes of MCMV are those that subvert NK or T cell responses. Given the importance of NK cells in combating MCMV infection, it is not surprising that MCMV encodes a number of proteins that inhibit NK cell function (Table 1-2). A homolog of cellular class I molecules that inhibits the NK cell–mediated clearance of MCMV in vivo is encoded by m144 (Farrell et al. 1997). It has been suggested that the m144 protein acts as a decoy for NK cells by engaging inhibitory NK cell receptors, although the receptor has not been identified (Farrell et al. 1997; Cretney et al. 1999; Kubota et al. 1999). As will be discussed below, the viral m152 gene inhibits the clearance of MCMV by CD8+ T cells. However, the m152 product also inhibits NK cells by down-regulating the expression of the MHC class 1–like molecule RAE-1, which is a ligand for the NK cell–activating receptor NKG2D on NK cells (Cerwenka et al. 2000; Lodoen et al. 2003). NKG2D has two additional activating ligands, MULT-1 (Carayannopoulos et al. 2002) and H-60 (Malarkannan et al. 1998; Diefenbach et al. 2000). MCMV also interferes with their expression on infected cells. The products of m145 and m155 down-regulate the expression of MULT-1 and H-60, respectively, and the deletion of these genes from MCMV enhances clearance of the virus due to NK cell–specific effects (Krmpotic et al. 2002; Lodoen et al. 2003; Ogasawara et al. 2003; Hasan et al. 2005; Krmpotic et al. 2005). Finally, as already discussed, MCMV can evade Ly49H+ NK cells by mutations in the viral ligand m157 (Voigt et al. 2003). Since mutations in m157 affecting Ly49H binding occur in wild isolates of MCMV, this interaction may not occur frequently in nature. It is likely that m157 normally binds inhibitory NK cell receptors such as Ly49I (Arase et al. 2002) in wild mice, which show significant variability in the NK gene complex (Scalzo et al. 2005). The viral genes affecting NK cell function have all been found within the m145 gene family, but it is likely that additional NK cell inhibitory genes exist within the m02 gene family (Oliveira et al. 2002).
31
1. MURINE CYTOMEGALOVIRUS AND OTHER HERPESVIRUSES
TABLE 1-2
IMMUNE EVASION BY MCMV Viral gene
Function
Mechanism
Reference*
m04 m06
Blocks CTL function Blocks CTL function
[1–4] [2,3,5]
M27 M33
Type I and type II IFN resistance Migration of cells, including smooth muscle cells Promotes inflammation and dissemination to salivary gland Unknown Inhibits NK cell function Inhibits NK cell function
Binds MHC I and remains associated on cell surface Down-regulates MHC I by targeting molecule for lysosome degradation Down-regulates STAT-2 Receptor homologue binds RANTES, functional homologue of HCMV US28 Chemokine homologue, macrophage chemoattractant
m129/131 m138 (fcr-1) m144 m145 m147.5 m152
Inhibits Ag presentation? Blocks CTL function Inhibits NK cell function
m155
Inhibits NK cell function
m157
Activates NK cells Inhibits NK cells?
Unknown
Complement resistance
Binds antibody MHC class I homology. Mechanism unknown Down-regulates MULT-1, a ligand for the NK cell activating receptor NKG2D on infected cells Specifically down-regulates CD86 on dendritic cells Retains MHC I in ERGIC Down-regulates RAE-1, a ligand for the NK cell activating receptor NKG2D on infected cells Down-regulates H-60, a ligand for the NK cell activating receptor NKG2D on infected cells Activates NK cells via Ly49H in C57BL/6 mice Presumed function is to bind NK cell inhibitory receptors such as Ly49I Up regulation of CD46 via MCMV responsive element in CD46 gene promoter
* References are given in full in the bibliography. 1. Kleijnen et al. 1997 7. Melnychuk et al. 2005 2. Wagner et al. 2002 8. Davis-Poynter et al. 1997 3. LoPiccolo et al. 2003 9. Waldhoer et al. 2002 4. Kavanagh et al. 2001 10. Fleming et al. 1999 5. Reusch et al. 1999 11. Saederup et al. 2001 6. Zimmermann et al. 2005 12. Saederup et al. 1999 13. Crnkovic-Mertens 1998
T cells, particularly CD8+ T cells, play a vital role in viral clearance and in the control of chronic infection and recurrence from latency (Krmpotic et al. 2003). The importance of CD8+ T cells is reflected in the number of mechanisms employed by MCMV to circumvent their effect. MCMV acts on both the antigen presentation and effector stages of the immune response. The virus inhibits the capacity of DC to present antigen to naive T cells (Andrews et al. 2001), as well as inhibiting the effector phase of the response by down-regulating the expression of MHC class I molecules, thus reducing the recognition and killing of infected cells by CD8+ T cells. Three MCMV genes, m04, m06, and m152, appear to account for all the inhibitory effects of MCMV on MHC class I function, at least in vitro (Wagner et al. 2002). The product of m04, glycoprotein (gp) 34, binds MHC class I in the ER but does not inhibit its expression on the cell surface. Rather, it is transported to the cell surface as a complex with class I molecules where CD8-mediated killing of infected cells is inhibited (LoPiccolo et al. 2003). The m06
14. 15. 16. 17. 18. 19. 20.
Thale et al. 1994 Farrell et al. 1997 Cretney et al. 1999 Krmpotic et al. 2005 Loewendorf et al. 2004 Ziegler et al. 1997 Krmpotic et al. 2002
21. 22. 23. 24. 25. 26. 27.
[6] [7–9] [10–12] [13,14] [15,16] [17] [18] [2–4,19,20] [20,21] [22,23] [24–26]
[27]
Lodoen et al. 2003 Lodeon et al. 2004 Hasan et al. 2005 Scalzo et al. 1992 Arase et al. 2002 Smith et al. 2002 Nomura et al. 2002
gene encodes gp48, which interferes with the MHC class I pathway of antigen presentation by binding to complexes of MHC class I molecules and antigenic peptides, targeting them for degradation in the lysosome (Reusch et al. 1999). The reduced cell surface expression of these complexes results in reduced recognition and killing of these cells by CD8+ T cells (Reusch et al. 1999; Wagner et al. 2002; LoPiccolo et al. 2003). The m152 gene product, gp40, prevents the transport of class I molecules to the cell surface, causing them to be retained in the ERGIC (Del Val et al. 1992; Ziegler et al. 1997). Since m152 also down-regulates the NK cell ligand RAE-1, this viral gene exerts an inhibitory effect on both CD8+ T cells and NK cells. While these in vitro studies have established a role for m04, m06, and m152, their role in vivo is less clear. Deletion of m152 from MCMV results in enhanced clearance of MCMV in vivo by CD8+ T cells and NK cells (Krmpotic et al. 2002). However, a virus constructed by the deletion of m04, m06, and
32
GEOFFREY R. SHELLAM, ALEC J. REDWOOD, LEE M. SMITH, AND SHELLEY GORMAN
m152 was cleared no more effectively than wild-type virus in vivo (Gold et al. 2004). Understanding the role of immune evasion in vivo is complicated by a number of factors. For example, the m04, m06, and m152 products exhibit distinct preferences for certain MHC class I allelic forms (Wagner et al. 2002), and their downregulation of MHC class I molecules may therefore vary among inbred mouse strains. Furthermore, the m152 product gp40 exhibits dual functions; gp40 also down-regulates H-60, which is a ligand for the NKG2D receptor on NK cells (Krmpotic et al. 2002). Thus, the effect of m152 varies among inbred mouse strains according to the interactions with particular ligands. Finally, isolates of MCMV exhibit polymorphisms in a number of immune evasion genes (Smith et al. 2006). The G4 isolate encodes a variant of m157 that does not activate Ly49H (Voigt et al. 2003). Hence, Ly49H+ mouse strains are no more resistant to G4 than are Ly49H- mouse strains. Given the complexity of the effects of MCMV immune evasion proteins, understanding how they influence the dynamic interaction between MCMV and its host poses a significant challenge.
VII. A.
DIAGNOSIS Serology
Antibody assays have been the standard means of screening for evidence of MCMV infection. Serum neutralization assays were originally used (Mannini and Medearis 1961), and their sensitivity has been improved by the addition of complement (Kim and Carp 1973; Lawson et al. 1988) and the use of MCMV antigen derived from virus passaged in cell culture rather than the salivary gland (Chong et al. 1981; Lawson et al. 1988). Since the 1980s, the use of the enzyme-linked immunosorbent assay (ELISA) has been widespread (Anderson et al. 1983, 1986; Classen et al. 1987; Lawson et al. 1988). ELISA was found to be more sensitive than nuclear anticomplement immuofluorescence (NAIF) or complement fixation assays (Anderson et al. 1986), although NAIF was reported to be more sensitive for the detection of antibodies in the acute stage of the infection (Anderson et al. 1983). Another study showed that ELISA and indirect immunofluorescence were of comparable sensitivity (Classen et al. 1987). ELISA is the method of choice for routine screening of sera from mouse colonies, and it is used by most diagnostic laboratories. However, there are situations in which serological assays are not suitable. Immunodeficient, genetically modified mice such as nude, SCID, RAG gene, or immunoglobulin gene knockout mice cannot produce anti-MCMV IgG antibodies, so sensitive molecular techniques such as the polymerase chain reaction (PCR) are employed (see below). PCR may also be used to confirm MCMV infection in mice whose antibody titers are borderline positive.
B. Molecular Detection Real-time quantitative PCR has recently been developed for the detection of MCMV (Wheat et al. 2003; Farroway et al. 2005) and, as would be predicted, it is much more sensitive than the plaque assay or ELISA for detecting infection. However, given that viral genomes are likely to be restricted to particular tissues in latently infected mice (see Section V, C), it is more feasible to use the ELISA for routine screening. Real-time qPCR has recently been adapted for the detection of mixed infection with different strains of MCMV in individual mice based on detection of different ie1 genotypes (Gorman et al. 2006).
VIII. CONTROL AND PREVENTION With the introduction of regular screening programs for murine pathogens by commercial suppliers of laboratory mice and the widespread use of specific pathogen-free mice in research, naturally occurring MCMV infections in mouse colonies no longer occur. Since MCMV does not infect the fetus transplacentally, the use of cesarean derivation to maintain the disease-free status of laboratory animals ensures that mouse colonies will remain free of MCMV. However, the potential for cross-infection with MCMV derived from experimentally infected mice remains. MCMV appears not to be transmitted from cage to cage (Mannini and Medearis 1961), although a conflicting result has been reported (Anderson et al. 1983). The physical separation of cages of MCMV-infected mice from other mice and the use of filter tops would therefore be prudent. An even greater risk attends the housing of wild-caught mice in the same facility as laboratory colonies. Because wild mice are naturally infected with MCMV (see Section IV) and other viruses, as well as bacterial and parasitic pathogens, these animals should only be housed in a separate quarantine facility with stringent barrier procedures and comprehensive monitoring. There is no need to consider the use of vaccines or other measures to control MCMV in mouse colonies. However, antiviral compounds have been used widely in research, particularly when the control of CMV infections in human patients is modeled in murine studies (Bolger et al. 1999; Lenzo, Shellam et al. 2001; Shanley et al. 1985, 1988). Ganciclovir and cidofovir are very effective against MCMV, and the 50% inhibitory concentrations in vitro are about 5 and 0.24 µM respectively (Okleberry et al. 1997; Lenzo, Shellam, et al. 2001). The use of foscarnet (Smee et al. 1995; Lenzo, Shellam, et al. 2001), acyclovir (Shanley et al. 1985), and HPMPC (Bolger et al. 1999) has also been reported. A number of antiviral drugs have been tested against MCMV and show promise (Rybak et al. 2000). Natural products with anti-MCMV properties also have been described (Hudson 1994a).
33
1. MURINE CYTOMEGALOVIRUS AND OTHER HERPESVIRUSES
1.
Development of Vaccines
While there is no necessity to develop vaccines to control MCMV infection in colonies of laboratory mice, the mouse model has been used to test strategies for vaccination against HCMV and for evaluating the usefulness of MCMV as a persistent viral vector encoding heterologous antigens. VACCINES AGAINST MCMV OR HETEROLOGOUS ANTIGENS The ability of vaccines based on a single MCMV epitope to induce protection against MCMV challenge has been established using the viral peptide in an adjuvant formulation (Scalzo et al. 1995), a recombinant vaccinia vector expressing the peptide epitope (Del Val et al. 1991), or plasmids expressing the viral epitope (Morello et al. 2000). Alternatively, a vaccine based on MCMV that has been genetically disabled through the deletion of gL shows promise (J. Allan, personal communication). A different approach has been to use the property of viral persistence and the induction of long-term memory (Karrer et al. 2003) exhibited by MCMV in developing the virus as a vector for vaccinating against heterologous antigens. A recombinant MCMV vaccine expressing epitopes from influenza A or lymphocytic chorio-meningitis viruses induced CD8+ memory T-cell populations that expanded with time and responded rapidly to challenge with either of these viruses without the need for boosting (Karrer et al. 2004). VIRALLY VECTORED IMMUNOCONTRACEPTION A novel use of MCMV as a vaccine vector is the development of an immunocontraceptive vaccine for possible use as a disseminating agent to control population explosions of house mice in rural Australia (Shellam 1994). A recombinant MCMV vaccine was constructed encoding the murine zona pellucida 3 (ZP3) protein, which coats the egg and is involved in the binding of sperm in the fertilization reaction. Complete and long-lasting sterility was induced following a single vaccination of female BALB/c mice (Lloyd et al. 2003; Redwood et al. 2005) or wild mice (Lloyd et al. 2006). Vaccination was accompanied by the production of antibodies to ZP3 and a profound reduction in ovarian follicles.
IX.
MOUSE THYMIC VIRUS: MOUSE
T LYMPHOTROPHIC VIRUS (MTLV) OR MURID HERPESVIRUS 3 A.
Introduction and History
Mouse thymic virus (MTV) was first described in 1961, due to the occurrence of thymic necrosis during serial passage of organ homogenates in newborn Swiss mice (Rowe and Capps 1961). Inoculation of newborn mice with homogenized necrotic thymus
tissue resulted in necrosis of the thymus in recipient mice. Sera from inoculated mice did not contain antibodies against a range of mouse viruses (Rowe and Capps 1961). The properties of the infectious agent were compatible with those of a virus, being filterable, not affected by antibiotics, and not culturable on bacteriological media. Based on morphology and lability, mouse thymic virus was classified as a herpesvirus (Parker et al. 1973). Despite efforts, a permissive tissue culture system has not been established for mouse thymic agent, limiting subsequent research. The genome of MTV has not been studied.
B.
Properties of the Virus
Mouse thymic virus has typical herpesvirus morphology. Within infected thymus tissues, MTV capsids are icosahedral, ranging from 95 nm to 110 nm in diameter. Enveloped capsids are spherical, ranging from 125 nm to 165 nm in diameter (Parker et al. 1973; Athanassious et al. 1990). The virus displays the same properties of heat (50°C, 30 minutes) and ether (diethyl ether, 2 hours, 20 minutes) lability of other herpesviruses. Infection of newborn mice with MTV results in thymic necrosis and acute immunosuppression over 2–3 weeks. The thymus regenerates in most animals within 1–2 months; however, mice continue to shed virus asymptomatically into the saliva (Rowe and Capps 1961; Parker et al. 1973; Cross et al. 1979; Cohen et al. 1975). All mouse strains tested were found to be susceptible to MTV, but the rate at which thymus necrosis occurred varied between strains (Cross et. al. 1979). Neonatal infection of selected strains of mice results in histologically and serologically evident autoimmune disease, similar to organspecific autoimmune diseases in humans (Morse et al. 1999), possibly due to altered control of self-reactive T cells. Initial attempts were made to culture the virus on primary mouse embryo and kidney cultures, spleen and thymus explants, and several tumor cell lines, all without success (Rowe and Capps 1961). Similarly, several other murine and non-murine cells and cell lines have been used with no apparent virus growth or cytopathic effect (Morse 1988; Hudson 1994b). One unpublished observation (cited in Morse and Valinsky 1989) states that MTV is able to grow in “several T lymphoblastoid lines,” including three CD4+ human lines. However, this has not been repeated elsewhere. When housed with mice i.p. inoculated with MTV, test animals are seropositive by IFA, CF, and ELISA 21–30 days after cohousing, and MTV was identified in the salivary glands of contact-infected mice after 90 days of cohousing (St-Pierre et al. 1987; Lussier et al. 1988a, 1988b). Transmission of MTV from mothers to offspring was observed following inoculation with MTV 24 hours after giving birth. In this instance, virus was identified in the salivary glands of the offspring 21–27 days after inoculation, although no anti-MTV antibody could be detected in these mice (St-Pierre et al. 1987). However, it is unclear if this transmission of MTV was via milk or simply due
34
GEOFFREY R. SHELLAM, ALEC J. REDWOOD, LEE M. SMITH, AND SHELLEY GORMAN
to contact transmission from the mother. Inoculation of pregnant females at various times of gestation yielded no evidence of transplacental transfer of MTV, as all fetuses were MTV-negative and no abortion was recorded (St-Pierre et al. 1987). MTV was originally isolated from a laboratory strain of mouse (Rowe and Capps 1961) and was subsequently identified in other mouse colonies (Cross et al. 1979). Little is known about naturally occurring MTV infection in wild mice, although 4 out of 15 wild-mouse serum samples were positive for MTV by IFA. MTV was identified by infectivity assay in a small number of mainly seronegative wild mice in this study (Cross et al. 1979). Interestingly, MTV and MCMV were frequently isolated from the same populations of mice, and co-infection of MTV with MCMV results in increased MCMV-related mortality, perhaps due to the immunosuppressive effect of MTV (Cross et al. 1979).
i.p. injection. Orally inoculated animals seroconverted to the virus (Morse 1989). ADULT MICE Mice inoculated with MTV as adults (or after 7 days of age) do not develop either autoimmune conditions or thymic necrosis. Within the thymus there is no change in populations of CD4+ cells. In these animals, MTV is detectable in the salivary gland from 7 days p.i. onwards, with virus shedding into the saliva. No virus is detectable in the visceral organs. From day 14 p.i., mice are seropositive for anti-MTV antibodies by IFA, with these antibodies persisting over 70 days (Cross et al. 1979). Nude mice can be infected with MTV, but there is a low level of virus shedding from infected animals (Morse 1988), indicating the requirement of T cells for viral persistence.
D. C.
Pathogenesis and Cell Tropism
NEWBORN MICE In mice infected as newborns, MTV typically induces thymic necrosis. Three days after infection, nuclear inclusions are visible in thymocytes (mostly CD4+ 8+ and CD4+ 8-), which are almost completely destroyed within 1–2 weeks. At this point, the thymus may only be 20% of normal weight. Thymocyte numbers then begin to recover, and within 1–2 weeks the thymus appears macroscopically and histologically normal, with the composition of CD4+ cell subsets returning to normal (Cross et al. 1979; Morse et al. 1999; Morse 1989). As the age of the mouse increases prior to infection, the effect of the virus on the thymus is decreased until from 6 days of age the thymus is not susceptible to MTV-related damage (Cross et al. 1979). In BALB/c and A strain mice, but not in C57BL/6, C3H, or DBA/2 mice, 30%–40% of infected newborns develop autoimmune gastritis, which is characterized by the infiltration of mononuclear cells into the gastric mucosa and destruction of parietal and chief cells (Morse et al. 1999). Smaller numbers of mice from other strains develop oophoritis or antibodies to thyroglobin. In no case was MTV found in the affected tissues. Following i.p. inoculation in newborns, virus can be detected in the thymus between 3 and 10 days after infection, with a peak titer at day 7. Viremia is seen between 3 and 7 days p.i., with MTV identifiable in the visceral organs between days 3 and 14. Virus can be detected in the salivary glands from 5 days p.i., and persists more than 200 days later. At approximately 300 days p.i., virus is still shed into the saliva (Cross et al. 1979). No serum anti-MTV antibodies can be detected by IFA in animals infected as newborns. Interestingly, in newborn animals inoculated orally with MTV, thymic necrosis also occurs, with thymic appearance at 9 days p.i. indistinguishable from that of animals inoculated by
Diagnosis
As there is no information available on the genome of MTV (although it is presumably a ds-DNA virus), there are no methods such as PCR described for the detection of viral genomes within infected animals. Newborn mice infected with MTV produce very little antibody. However, mice infected as adults produce significant amounts of serum anti-MTV antibodies, which persist for long periods. These antibodies can be detected by indirect immunofluorescence (IFA), complement fixation (CF), or ELISA. In the IFA test, cells from thymuses of MTV-infected mice are dispersed and fixed onto slides and subsequently incubated with serial dilutions of serum from test animals. Anti-MTV antibodies bound to infected cells are identified using labeled anti-mouse conjugates (Lussier et al. 1988b). Homogenates of thymuses from 7-day-old mice inoculated with MTV as newborns are used as sources of antigen for both CF and ELISA. In the CF test, serial dilutions of heat-inactivated test sera are tested against a standard antigen dilution, and in the ELISA, a standard dilution of MTV antigen is coated onto microtiter plates prior to incubation with serial dilutions of test serum. Bound anti-MTV antibodies are detected using labeled anti-mouse antibodies (Lussier et al. 1988a, 1988b). In all tests, antibody titer is determined as the reciprocal of the highest dilution giving a positive result. During experimental infection of mice with MTV (by i.p. inoculation or by contact with infected animals), all 3 tests identified seropositive animals with similar sensitivities, although ELISA testing resulted in higher serum antibody titers (Lussier et al. 1988b). Mice were seropositive for MTV up to 230 days after infection. Serum antibody may also be titrated using a neutralization assay in which serum is incubated with a known titer of virus, that is subsequently inoculated into newborn mice. Alternatively, virus can be titrated and the ID50 calculated using an infectivity assay, where thymus homogenates are inoculated into newborn mice. In both cases, thymic necrosis is scored macroscopically after 10–14 days (Cross et al. 1979; Morse 1990).
1. MURINE CYTOMEGALOVIRUS AND OTHER HERPESVIRUSES
In none of the serological tests is there any cross-reaction between MTV and MCMV (Cross et al. 1979).
ACKNOWLEDGMENTS We acknowledge the support of the National Health and Medical Research Council of Australia through Project Grant 254636, the Pest Animal Control Co-operative Research Centre, the Grains Research and Development Corporation, and the University of Western Australia. We are very grateful to Sandra Jones, Catherine Gangell, and Fiona Stanley for assistance with the preparation of this review.
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Chapter 2 Mouse Adenoviruses Katherine R. Spindler, Martin L. Moore, and Angela N. Cauthen
I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . II. History and Isolations of Mouse Adenoviruses, Antigenic Relationships, Virus Strains, and Virus Mutants . . . . . . . . . . . . . . . . . . . . . III. Physical Properties . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . IV. Molecular Genetics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. MAV-1 Genome Features . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. MAV-1 E1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. MAV-1 E3 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. MAV-1 E4 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . E. MAV-1 Major Late Promoter . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . V. Growth of Mouse Adenoviruses In Vitro . . . . . . . . . . . . . . . . . . . . . . . . . . A. In Vitro Infection, wt MAV-1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. In Vitro Infection, MAV-1 E1A Mutants . . . . . . . . . . . . . . . . . . . . . . . C. In Vitro Infection, MAV-1 E3 Mutants . . . . . . . . . . . . . . . . . . . . . . . . . VI. Clinical Disease and Pathogenesis of Mouse Adenoviruses . . . . . . . . . . . . A. Wild-Type MAV-1 Infection In Vivo . . . . . . . . . . . . . . . . . . . . . . . . . . B. MAV-1 E1A Mutant Infection In Vivo . . . . . . . . . . . . . . . . . . . . . . . . . C. MAV-1 E3 Mutant Infection In Vivo . . . . . . . . . . . . . . . . . . . . . . . . . . D. Immune Response to MAV-1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1. Innate Immune Response to MAV-1 . . . . . . . . . . . . . . . . . . . . . . . . 2. Cell-Mediated Immune Response to MAV-1 . . . . . . . . . . . . . . . . . 3. Humoral Immune Response to MAV-1 . . . . . . . . . . . . . . . . . . . . . 4. Model of MAV-1 Immunopathogenesis . . . . . . . . . . . . . . . . . . . . . E. MAV-2 Infection In Vivo . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VII. Host Genetics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VIII. Host Range and Prevalence . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . IX. Diagnosis, Control, and Prevention . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
I.
INTRODUCTION
The first mouse adenovirus was isolated by Hartley and Rowe while trying to establish the Friend leukemia virus in tissue culture from mice (Hartley and Rowe 1960). Mouse adenoviruses THE MOUSE IN BIOMEDICAL RESEARCH, 2ND EDITION
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are useful for study of adenovirus pathogenesis in the natural host, in which they cause acute and persistent infections (Smith and Spindler 1999). Such studies are not possible with the species-specific human adenoviruses (hAds). The availability of immunocompetent and immunodeficient inbred mouse Copyright © 2007, 1980, Elsevier Inc. All rights reserved.
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strains, immunological reagents for mice, and tools for genetic mapping studies combine to make mouse adenovirus studies ideal for understanding virus-host interactions. Commercial mouse suppliers standardly monitor for mouse adenoviruses, and the viruses have been eliminated in commercial mouse colonies and are rare if not absent in institutional colonies (Richter 1986). Methods for propagating and titrating the virus have been described (Cauthen and Spindler 1999a). The hAds were isolated from patients with respiratory illness independently by Rowe et al. (1953) and Hilleman and Werner (1954). The molecular biology of the hAds has been extensively studied in the years since their discovery (Shenk 2001). In addition to their use as models for studying DNA replication and mRNA transcription and processing, hAds are being widely used to develop gene therapy and vaccine vectors (GomezRoman and Robert-Guroff 2003; Hart 2003; Imperiale and Kochanek 2004; Thomas et al. 2003). Far less is known about the pathogenesis of hAds, in part due to the strict species specificity of the adenoviruses. There are currently more than 51 distinct hAd serotypes, some of which cause respiratory disease; others are associated with diseases including conjunctivitis and gastroenteritis (reviewed in Horwitz 2001). Adenoviruses are associated with acute pneumonia in children in developing countries, where they are a major cause of illness and death (Kajon et al. 1996). Severe adenovirus infections occur in immunocompromised people (Kojaoghlanian et al. 2003), including AIDS patients or those undergoing bone marrow or solid organ transplantation (Blanke et al. 1995; Carrigan 1997; Flomenberg et al. 1994). Pediatric bone marrow transplant patients are particularly at risk of hAd infection and mortality (Gavin and Katz 2002; Hale et al. 1999; Walls et al. 2003). In productive infections, the mouth, nasopharynx, or ocular conjunctiva are the initial site of hAd entry, with replication in epithelial cell types (Horwitz 2001). Like the mouse adenoviruses, hAds also cause persistent infections (Lukashok and Horwitz 1999). A study using sensitive real-time PCR coupled with lymphocyte purification suggests that human mucosal T lymphocytes are the site of hAd persistence (Garnett et al. 2002).
II.
HISTORY AND ISOLATIONS OF MOUSE
ADENOVIRUSES, ANTIGENIC RELATIONSHIPS, VIRUS STRAINS, AND VIRUS MUTANTS Although mouse adenoviruses have been isolated numerous times, there are only two serotypes, murine adenovirus 1 and murine adenovirus 2, as classified by the International Committee on Taxonomy of Viruses (2000). These are currently categorized as belonging to two different species, murine adenovirus A and murine adenovirus B, respectively. This is supported by data on serum neutralization (Lussier et al. 1987), growth characteristics (Smith et al. 1986), and genome restric-
tion analysis (Hamelin et al. 1988; Hamelin and Lussier 1988; Jacques, Cousineau, et al. 1994). Because these viruses are not infectious for infant rats (Smith and Barthold 1987) and because of the host species specificity of adenoviruses, we favor nomenclature that uses “mouse” instead of “murine,” and virus abbreviations as suggested by Ishibashi and Yasue (1984). Mouse adenovirus type 1 (MAV-1) was the first mouse adenovirus isolated (Hartley and Rowe 1960) and has also been designated in the literature as FL, MAdV-1, MAdV-FL, and MAd-1. Mouse adenovirus type 2 (MAV-2), also known as strain K87, was isolated from the feces of healthy mice by Hashimoto et al. (1966). In the work describing the isolation of MAV-1 and its classification as an adenovirus, Hartley and Rowe (1960) demonstrated a serologic relationship between MAV-1 and the hAds. Guinea pig serum from hAd-inoculated animals reacted against MAV-1 antigens. Similar results were obtained by Larsen and Nathans (1977) using serum against the hAd group antigen, hexon (a capsid protein). Hashimoto et al. (1966) used serum against hAd3 raised in guinea pigs to demonstrate a positive complement fixation reaction of MAV-2 antigen. For both MAV-1 and MAV-2, anti-MAV sera raised in mice are poorly reactive against hAd antigens (Hartley and Rowe 1960; Hashimoto et al. 1966). The antigenic relationships between MAV-1 and MAV-2 have been examined in several studies. In cross-neutralization tests between the two serotypes, there is a one-sided relationship between them (Wigand et al. 1977). MAV-2 antiserum neutralizes both MAV-1 and MAV-2, whereas MAV-1 antiserum neutralizes MAV-1 but only weakly neutralizes MAV-2. A similar partial serological relationship was identified by Lussier et al. (1987). Smith et al. (1986) found that MAV-1 antiserum neutralizing antibody titer is fourfold higher with MAV-1 than with MAV-2. MAV-1 has been more extensively studied than MAV-2. MAV-1 is available from the American Type Culture Collection (cat. no. VR550), but it should be noted that genome sequence differences and slight pathogenesis differences were found between the “ATCC” strain and the strain that we and others obtained directly from Steven Larsen (referred to as “standard”) (Ball et al. 1991). This was somewhat surprising, since the ATCC strain was deposited by Dr. Larsen. Evidently the ATCC and standard viruses have different passage histories. As discussed below (Section IV) the complete MAV-1 DNA sequence has been compiled, but there have been no sequences reported for MAV-2. A restriction map of MAV-2 has been determined (Jacques, D’Amours, et al. 1994). Mutant strains of MAV-1 have been constructed by sitedirected mutagenesis techniques using the standard strain as the starting virus. Because the standard MAV-1 genome has two EcoRI sites, these were mutated singly to obtain viruses that have a single EcoRI site in either early region 1 (E1) (Smith et al. 1996) or early region 3 (E3) (Beard and Spindler 1996). The resulting viruses, pmE301 and pmE101, have been used to
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2. MOUSE ADENOVIRUSES
construct mutants in E1A and E3, respectively (Beard and Spindler 1996; Cauthen et al. 1999; Smith et al. 1996). These virus mutants were named based on the types of mutations that were introduced (pm, point mutation; dl, deletion) and the gene where the mutation lies, E1 or E3, followed by an isolation number. The in vitro and in vivo growth characteristics of these viruses are discussed below (Sections V, B and C and VI, B and C). A naturally occurring MAV-1 mutant was isolated by Winters et al. (1981). This variant exhibits a large-plaque phenotype in cell culture and causes clinical disease with a distinct pulmonary tropism in adult C3H/HeJ mice. The mutation(s) responsible for this phenotype have not been mapped.
III.
PHYSICAL PROPERTIES
MAV-1 and MAV-2 share many physical properties, including ether resistance, thermal inactivation at 56°C, and a size of ∼80–90 nm (Hartley and Rowe 1960; Hashimoto et al. 1966). Both mouse adenoviruses lack hemagglutinating activity, unlike the hAds. MAV-1 has a buoyant density in CsCl like that of the hAds, 1.34 g/ml (Larsen and Nathans 1977; Wigand et al. 1977). MAV-1 is inactivated by 50% ethanol (Larsen and Nathans 1977).
IV. A.
MOLECULAR GENETICS
mapping of E1, E3, early region 4 (E4), and identification of the major late promoter (MLP) (Ball et al. 1989, 1991, 1988; Beard et al. 1990; Cai et al. 1992; Cauthen and Spindler 1996; Kring et al. 1992; Kring and Spindler 1990; Raviprakash et al. 1989; Song et al. 1996, 1995; Weber et al. 1994). Davison et al. (2003) have analyzed the genetic content, phylogeny, and evolution of the family Adenoviridae, and they have submitted a third-party annotation for MAV-1 (AC_000012). It should be noted that their annotation includes some predicted genes for MAV-1 that are not in agreement with published experimental evidence. For example, their MAV-1 E1A annotation does not take into account E1A transcription mapping and cDNA sequencing data (Ball et al. 1989). Other differences are indicated for IVa2, the DNA polymerase, and pTP genes, which could be correct; these regions have not been transcription mapped. MAV-1 has inverted terminal repeats (ITRs) that are 93 nucleotides (nt) long (Ball et al. 1991, 1988; Temple et al. 1981). These ITRs have the first 18 nt that are highly conserved among the adenoviruses and that are essential for replication of the viral DNA (Challberg and Rawlins 1984; Lally et al. 1984; Tamanoi and Stillman 1983). Unlike hAds, MAV-1 does not encode virus-associated (VA) RNAs (Meissner et al. 1997). Although MAV-1 polypeptide III (penton base) shares high amino acid identity with the hAds, it does not have the arginineglycine-aspartic acid (RGD) motif found in many hAds and thought to be important for viral internalization via cellular integrins (Meissner et al. 1997). However, it has a leucine-aspartic acid-valine (LDV) motif recognized by other integrins, present as LDL and, like porcine adenoviruses, MAV-1 has an RGD sequence in the C-terminus of the fiber protein.
MAV-1 Genome Features
The complete sequence (30,944 base pairs) of the doublestranded DNA genome of MAV-1 has been determined (Meissner et al. 1997) (accession number NC_000942). The ordering of genes in the MAV-1 genome, shown in Fig. 2-1, has been accomplished through sequence comparison with hAds, transcription
B.
MAV-1 E1
Genes expressed prior to early Ad viral replication are designated as “early,” while those expressed at or after the time of DNA replication are “late.” The E1 region in MAV-1
Fig. 2-1 Genomic organization of MAV-1. The genome is indicated by the horizontal lines in the center, and map units are indicated below the line. Circles on the end of the genome indicate the terminal protein. Genes whose transcription has been mapped are indicated by arrows; for E3 and E4, multiple transcripts are indicated by only a single arrow. Open reading frames with similarity to proteins of hAds are indicated by the boxes. Genes transcribed in the rightward direction are indicated above the genome, and genes transcribed in the leftward direction are indicated below. The major late promoter (MLP) is indicated by the arrowhead. DNApol, DNA polymerase; pTP, terminal protein precursor; DBP, DNA binding protein. (Adapted from Smith and Spindler, 1999, with permission.)
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corresponds in location to the E1 region of hAds, at the left end of the genome (Ball et al. 1988), and the mRNAs are transcribed in a rightward direction (Ball et al. 1989). E1 encodes three mRNAs that overlap; one is designated E1A, and the other two correspond to hAd mRNAs that encode E1B 19K and E1B 55K proteins (Ball et al. 1989). The three mRNAs are 3′ coterminal, and the three proteins share a common C-terminal sequence. The MAV-1 E1A protein sequence has little overall sequence similarity to the hAd E1A 289 aa protein (encoded by the 13S E1A mRNA). However, Ball et al. (1989) showed that MAV-1 E1A has approximately 40% sequence similarity in conserved regions 1 (CR1), CR2, and CR3 compared to CR1, CR2, and CR3, respectively, of the hAd 289 aa protein (Moran and Mathews 1987). MAV-1 CR2 is the most similar to that of hAds, with approximately 50% sequence similarity (Ball et al. 1988). The predicted amino acid sequence similarity of the MAV-1 E1B coding regions compared to that of the hAd 19K and 55K proteins is 37% and 42%, respectively (Ball et al. 1988). No further studies involving the E1B proteins have been reported. The E1A protein is detected in MAV-1 infections in cell culture at both early and late times by immunoprecipitation with polyclonal antiserum raised against amino acids 27–200 of the E1A protein (Smith et al. 1996; Ying et al. 1998). The E1A protein is predicted to have a molecular weight of 22 kDa, but migrates slightly larger than 30 kDa, likely due to its phosphorylation (Smith et al. 1996). hAd E1A proteins are also phosphorylated (Gaynor et al. 1982; Yee and Branton 1985a; Yee et al. 1983). However, the significance of the phosphorylation of E1A in MAV-1 infections is not known. MAV-1 viruses with mutations in E1A are described in Table 2-1. The E1A protein expression patterns were evaluated by Western blot analysis and immunoprecipitation (Smith et al. 1996; Ying et al. 1998). The dlE102 (CR2 deletion) and dlE106 (CR3 deletion) E1A proteins migrate faster than the wild-type (wt) E1A, as expected. pmE109 and pmE112, initiator methionine mutants, do not synthesize detectable levels of E1A protein.
TABLE 2-1
EFFECTS OF MAV-1 E1A MUTANTS IN CELL CULTURE AND OUTBRED SWISS MICE E1A mutant
Effect of the mutationa
dlE105 dlE102 dlE106 pmE109 pmE112
Deletion of CR1 region (amino acids 35–78) Deletion of CR2 region (amino acids 111–129) Deletion of CR3 region (amino acids 135–154) Mutation of initiator ATG to TTG Mutation of initiator ATG to CAC
aSmith
Average LD50 (log PFU)b 103.5 100.9 102.6 103.5 103.9
et al. 1996 et al. 1998; the average LD50 for wt virus in these experiments was 10-1.5. bSmith
Although dlE105 (CR1 deletion) has a 43 amino acid deletion, it produces protein that migrates slower than the wt protein; a similar phenomenon is seen in hAd 5 E1A CR1 deletion proteins (Egan et al. 1988). hAd E1A is a transcriptional regulator (reviewed in Gallimore and Turnell 2001), and it is required for activation of early viral transcription (Berk et al. 1979; Jones and Shenk 1979). A plasmid encoding the left end of the MAV-1 genome (including the E1A coding region) transactivates the hAd5 E3 promoter in both HeLa cells and mouse L929 cells, albeit at a level lower than that of hAd E1A (Ball et al. 1988). Although MAV-1 E1A is able to transactivate the hAd5 E3 promoter, unlike hAd E1A, it does not appear to stimulate the expression of the other early mRNAs (E1A, E1B, E2, E3, and E4), at least in 3T6-infected cells at a multiplicity of infection (MOI) of 5 (Ying et al. 1998). Thus, the importance of the transactivation by MAV-1 E1A in reporter assays of transfected cells (Ball et al. 1988) is not understood. hAd E1A CR2 encodes a retinoblastoma protein (pRB) binding motif, (D)-L-X-C-X-E, that is conserved as (D)-L-R-C-Y-E in MAV-1 E1A CR2. MAV-1 E1A protein binds to mouse pRb and related proteins (Smith et al. 1996). This was shown both for in vitro transcribed and translated pRb protein (Smith et al. 1996) and for the endogenous pRb in virus-infected cells (L. Fang et al. 2004). This binding is primarily dependent on the presence of E1A CR2 (Smith et al. 1996). Similar experiments also showed that related pRb family proteins p107 and p130 bind to MAV-1 E1A (Smith et al. 1996; L. Fang et al. 2004). For the in vitro–produced p107, CR2 is necessary and sufficient for binding (Smith et al. 1996); p107 binds to hAd E1A in the CR2 region (Dyson, Guida, Münger, et al. 1992; Harlow et al. 1986; Whyte et al. 1989; Yee and Branton 1985b). In both the pRb and the p107 experiments in which infected cell lysates were mixed with in vitro translated pRb or p107, the E1A CR1 deletion mutant protein bound to pRb and p107 at greatly reduced levels compared to wt E1A (Smith et al. 1996). This suggests that CR1 may play a cooperative or stabilizing role in pRb and p107 binding to CR2 of E1A, as has been shown for hAd E1A (Dyson, Guida, McCall, et al. 1992; Ikeda and Nevins 1993). The functional relevance for the interaction between MAV-1 E1A and pRB was shown by experiments in SAOS-2 cells, which lack pRb (Smith et al. 1996). SAOS-2 cells transfected with mouse pRb alone exhibit an arrested growth phenotype that is reversed when MAV-1 E1A and pRb are introduced into the cells together.
C.
MAV-1 E3
Transcription mapping of the E3 region of MAV-1 indicated that this region produces three early mRNAs that are 5′ and 3′ coterminal (Beard et al. 1990). Each E3 mRNA has three exons; the first and second are identical among the three mRNAs, and the last intron of each is different due to differential splicing.
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Thus, the predicted E3 proteins share a 72 amino acid N-terminus with unique C-terminal domains. The three mRNAs transcribed at early times are referred to as class 1 mRNA, which gives rise to the E3 gp11K protein, class 2, and class 3 mRNAs (Beard et al. 1990). Only the E3 gp11K protein has been detected in wt virus infection of cultured cells (Beard and Spindler 1995). A fourth type of mRNA transcribed at late times (after viral DNA replication) is also detected that encodes E3 gp11K (Beard et al. 1990; Cauthen and Spindler 1999b). A polyclonal antiserum that specifically recognizes the unique portion of the E3 gp11K protein was used to demonstrate that E3 gp11K is detected in infected 3T6 cells beginning at 16 hours postinfection (h PI) and is detected until at least 48 h PI (Beard and Spindler 1995). The E3 gp11K protein can also be immunoprecipitated from radiolabeled infected cell lysates at both early and late times in infection. Additionally, the gp11K protein is recognized by polyclonal antiserum that is specific for the common portion of the E3 proteins (Beard and Spindler 1996). This antiserum fails to detect the other E3 proteins in MAV-1 infection of 3T6 cells, presumably because the proteins produced by the class 2 or class 3 mRNA are absent from infections in cell culture or are produced at levels too low for detection (Beard and Spindler 1996; Cauthen et al. 1999; Cauthen and Spindler 1999b). The E3 gp11K protein appears to be approximately 14K in infected cell lysates, but this is larger than the size of the product produced during in vitro transcription and translation of a plasmid containing the E3 gp11K gene (Beard and Spindler 1995). This size difference can be attributed to modifications made to the protein in infected cells: the cleavage of the signal sequence (predicted at amino acids 1 to 37) at the N-terminus of the E3 gp11K protein, and glycosylation at the predicted N-glycosylation consensus site (N-X-S/T) located at amino acid 56. The E3 gp11K protein localizes to the ER of the cell and is a peripheral membrane protein, as determined by alkaline extraction and phase separation experiments (Beard and Spindler 1995). E3 gp11K is transcribed and translated as an early mRNA and protein. However, a large mRNA expressed at late times can also encode E3 gp11K (Cauthen and Spindler 1999b). This expression of E3 gp11K is due to alternative splicing of a late transcript encoding a capsid protein, pVIII. pVIII is predicted to be translated from an unspliced late mRNA. However, if the primary pVIII transcript is spliced at the “E3” sites, a fusion protein consisting of sequences of pVIII and E3 gp11K is predicted to occur (Cauthen and Spindler 1999b), since they are translated in the same reading frame and their coding regions partially overlap (Raviprakash et al. 1989). It is thought that the signal sequence of the E3 gp11K coding region allows the pVIII-E3 gp11K fusion protein to enter the ER and get processed, producing a mature E3 gp11K (Cauthen and Spindler 1999b). The relevance of transcription and translation of E3 gp11K at late times is not known. The E3 mRNAs and their putative proteins have been identified or described, but the functions of these proteins have not
yet been discovered. hAd E3 proteins are involved in immune evasion (Fessler et al. 2004). For example, the hAd2/5 E3 gp19K prevents the display of class I major histocompatibility complex (MHC) antigens on the surface of infected cells, and this has been proposed as a mechanism enabling persistence of hAds (Levine 1984; Wold and Gooding 1991). However, unlike hAds, MAV-1 does not prevent the display of class I MHC antigens on the surface of infected mouse cells in culture (Kring and Spindler 1996). D. MAV-1 E4 The transcription map of MAV-1 E4 was determined and indicates that E4 encodes seven classes of mRNAs that are 3′ coterminal (Kring et al. 1992). The predicted coding regions of three of these mRNAs have some sequence similarity to hAd E4 proteins. The MAV-1 E4 orf a/b has 48% sequence similarity to the hAd2 E4 34K (orf 6) protein (Ball et al. 1991; Kring et al. 1992). MAV-1 orf a/c has 69% sequence similarity to the hAd2 E4 11K (orf 3) protein. The N-terminus of MAV-1 orf d has 55% sequence similarity to hAd2 E4 orf 2 (Kring et al. 1992), and the entire MAV-1 orf d has 60% sequence similarity to hAdE4-orf6/7 (L. Fang and K. Spindler, unpublished data). Further studies involving the E4 region of MAV-1 have not been reported. E. MAV-1 Major Late Promoter The MLP in hAds directs the synthesis of the late messages, and the MLP in MAV-1 was mapped to a similar region in the genome using ribonuclease protection assays and primer extension analysis (Song et al. 1996). The MAV-1 MLP has a TATA box, an inverted CAAT box, an SP1 binding site, and a DE1 element (Song et al. 1996). Notably, there is an absence of a USF-binding site, an initiator element (INR), and a DE2 element found in hAds and other mammalian viruses (Song and Young 1997). The lack of an INR may explain the finding of more than one start site of the MAV-1 MLP. Song and Young (1997) used a mutational analysis to demonstrate that the TATA box, SP1 site, and CAAT box elements are important for the MAV-1 MLP to function at normal levels in cells. Additionally, gel mobility shift assays were employed to show that the SP1 protein binds to the MAV-1 MLP with high affinity.
V.
GROWTH OF MOUSE ADENOVIRUSES IN VITRO A.
In Vitro Infection, wt MAV-1
Temple et al. (1981) reported that MAV-1 viral DNA synthesis is first observed at 35 h PI, even at MOIs of up to 800 PFU/cell
K AT H E R I N E R . S P I N D L E R , M A RT I N L . M O O R E , A N D A N G E L A N . C A U T H E N
when using [3H]thymidine labeling. However, using 32PO4 labeling and analysis of viral DNA prepared by the Hirt method (1967), we observed MAV-1 DNA synthesis as early as 20 h PI in L929 cells infected at a MOI of 10 PFU/cell (Fig. 2-2). Onestep growth curves in 3T6 cells and mouse brain microvascular endothelial cells (MBMEC) show that MAV-1 exhibits a typical eclipse period, with an increase in virus titer relative to input virus at 36–48 h PI (Cauthen et al. 1999; Ying et al. 1998). Similar results were seen in a two-step growth curve in mouse L929 cells (Fig. 2-3). Cytopathic effects are first detected at 36–48 h and are similar to other adenoviruses: Cells infected with wild-type MAV-1 round up, become refractile, and eventually detach from the substrate (Fig. 2-4). Analysis of [35S]-labeled infected cell proteins by electrophoresis indicates that unlike hAds, MAV-1 does not efficiently shut off host cell protein synthesis (Antoine et al. 1982; Ying et al. 1998). At least two mechanisms are proposed for the selective translation of viral mRNAs in hAd-infected cells resulting in shutoff of
107
106
Titer (PFU)
54
105
104
103 0
24
48
72
96
120
144
168
h PI
Fig. 2-3 Two-step growth curve of MAV-1 on L929 cells. Monolayers were infected with MAV-1 at a MOI of 0.1, harvested at the indicated times, and titrated by plaque assay on L929 cells.
Fig. 2-2 Time course of MAV-1 DNA replication. L929 cells were mock infected or infected at a MOI of 10. The cultures were labeled for one hour with 32PO and harvested at the indicated times PI. Hirt supernatants containing viral 4 DNA were isolated (Hirt 1967) and digested with HindIII and RNase A and electrophoresed on 0.7% agarose gels, dried and autoradiographed. The sizes of the restriction fragments are indicated on the right. In a parallel experiment performed with hAd5 in HeLa cells, DNA replication was first detected at 16 h PI (data not shown).
host protein synthesis (reviewed in Shenk 2001). One of these is via the VA RNAs; the lack of VA RNAs in MAV-1 infected cells (Meissner et al. 1997) could be responsible for defective host protein synthesis shutoff in MAV-1 infection. The other proposed mechanism of host protein synthesis shutoff in hAd-infected cells is via an inactivation of eIF-4F by dephosphorylation observed late after hAd infection; whether this occurs in MAV-1 infection has not been reported. The receptor for MAV-1 has not been described. hAds use a two-step mechanism for entry. The protruding hAd fiber protein interacts with the cellular receptor (Philipson et al. 1968), which has been shown for a majority of hAd serotypes to be an immunoglobulin superfamily member, the Coxsackie-adenovirus receptor (CAR) (Bergelson et al. 1997; Tomko et al. 1997). In the second step, the penton base interacts with host integrin molecules to promote entry (Wickham et al. 1993). The murine homolog of CAR (mCAR) has been identified, and it can serve as a receptor for hAds (Bergelson et al. 1998; Tomko et al. 1997). However, it is not known whether mCAR is a receptor for MAV-1 or whether MAV-1 uses integrins for entry. Productive MAV-1 infections occur in cell lines that do not express mCAR, such as mouse L929 cells and NIH3T3 fibroblasts (Tomko et al. 1997), suggesting that mCAR is not required for entry. It is believed that MAV-1 does not replicate productively in cultured human cells (Antoine et al. 1982; Larsen and Nathans 1977; K. Spindler, unpublished data), although there is one early conflicting report (Sharon and Pollard 1964). Infection of human cells does not yield infectious virus (Larsen and Nathans 1977); viral DNA synthesis occurs, but there appears to be a defect at
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2. MOUSE ADENOVIRUSES
mock
wt
pmE312
Fig. 2-4 Cytopathic effects of MAV-1 infection. Mouse 3T6 fibroblasts were mock-infected or infected with wt or pmE312 mutant virus at a MOI of 1 and photographed at 2.5 days PI. Wild-type virus consistently caused cells to release from the culture dish, whereas pmE312 did not, even upon prolonged incubation (data not shown). (Adapted from Cauthen 1998, with permission.)
the level of formation of virus particles due to defects in virus structural proteins, particularly hexon (Antoine et al. 1982).
B. In Vitro Infection, MAV-1 E1A Mutants Mouse 3T6 cells and 37.1 cells (a 3T6-derived cell line that expresses MAV-1 E1A protein) infected at a MOI of 5 with each of the E1A mutant viruses (Section IV, B) give yields of virus approximately like those of wt virus (Ying et al. 1998). Similarly, serum-starved 3T6 cells and transgenic mouse embryo fibroblasts (MEFs) from pRb+/+, pRb+/-, and pRb−/− embryos infected at a MOI of 5 give yields of E1A mutant viruses similar to that of wt virus, indicating that at high MOIs, E1A is not required to stimulate progression of the cell cycle and DNA replication (Ying et al. 1998). However, infection of 3T6 cells, mouse brain microvascular endothelial cells (MBMECs), and MEFs at MOIs of 0.05 resulted in a two log unit reduction in viral yield for E1A null mutant pmE109 (Fang and Spindler 2005; M. Moore and K. Spindler, unpublished data). Multiplicity-dependent growth of mutant viruses has been observed previously for hAds (Gaynor and Berk 1983; Imperiale et al. 1984; Nevins 1981), human cytomegalovirus (Bresnahan and Shenk 2000; Oliveira and Shenk 2001), and
herpes simplex virus 1 (Cai and Schaffer 1992; Chen and Silverstein 1992; Everett et al. 2004), but its significance is unknown. The hAd E1A protein protects cells from the interferon (IFN) response (IFN-α/β and -γ) by inhibiting the ISGF3 transcription factor, thereby reducing expression of IFN-stimulated genes (ISGs) (Ackrill et al. 1991; Gutch and Reich 1991; Leonard and Sen 1996). Similarly, MAV-1 E1A protects mouse 3T6 cells from IFN-α/β and IFN-γ (Kajon and Spindler 2000). E1A can provide this protection from IFN-α/β in trans to vesicular stomatitis virus (VSV). The presence of MAV-1 E1A (either in 37.1 cells or virus-infected cells) correlates with a reduction in steady-state levels of ISGs, suggesting that the mechanism of protection from IFN effects is like that of the hAds. We have preliminary evidence that MAV-1 E1A is able to bind to STAT-1 (L. Fang and K. Spindler, unpublished data), as has been suggested for hAd5 E1A (Look et al. 1998).
C.
In Vitro Infection, MAV-1 E3 Mutants
A series of mutations were made in E3 of MAV-1 to determine the individual effects of the three putative E3 proteins (Section IV, C) on virus replication in cell culture and on
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TABLE 2-2
EFFECTS OF MAV-1 E3 MUTANTS IN CELL CULTURE AND MICE E3 mutant virus
Effect of the mutation
pmE310 pmE312 pmE314 dlE303 dlE307 dlE309
No early expression of E3 protein; late expression of E3 gp11K No early expression of E3 protein; late expression of E3 gp11K No expression of E3 proteins at early or late times in infection Produces E3 gp11K mRNA only; E3 gp11K detected in cell culture Produces E3 class 2 mRNA only; no protein detected in cell culture Produces E3 class 3 mRNA only; no protein detected in cell culture
a
Average LD50 (log PFU) 101.5 102.9 a 105.2 103.7 103.6 104.7 a
Reference Beard and Spindler 1996 Cauthen and Spindler 1999b Cauthen et al. 1999 Beard and Spindler 1996 Beard and Spindler 1996 Beard and Spindler 1996
This value is from a single experiment.
growth and pathogenesis in mice. The E3 mutant viruses and the effects of their mutations on the synthesis of the putative E3 proteins are shown in Table 2-2. The E3 mutant viruses have growth patterns similar to those of wt virus in mouse 3T6 cells at a high multiplicity (MOI of 5–10). pmE310, pmE314, and dlE309 grow to titers similar to that of wt virus (Beard and Spindler 1996; Cauthen and Spindler 1999b). In addition, pmE314 virus grew as well as wt virus in MBMECs (Cauthen and Spindler 1999b), which were tested since endothelial cells are one of the target cells of MAV-1 in vivo (Charles et al. 1998; Kajon et al. 1998). dlE303 and dlE307 grow to titers approximately 10- to 50-fold lower than that of wt virus (Beard and Spindler 1996). pmE312 grows to titers approximately 10-fold lower than that of wt virus (Cauthen and Spindler 1999a). dlE303, dlE307, and pmE312 all exhibited slightly smaller plaques than wt virus (N. Cauthen and K. Spindler, unpublished data), and pmE312 showed a unique cytopathic effect in 3T6 cells (Fig. 2-4). The multiplicity dependence seen for MAV-1 E1A (Section V, B) is not seen for E3. E3 mutants has yields like wt virus, even at a MOI of 0.05 (M. Moore and K. Spindler, unpublished data).
VI.
CLINICAL DISEASE AND PATHOGENESIS OF MOUSE ADENOVIRUSES A.
Wild-Type MAV-1 Infection In Vivo
MAV-1 permits the study of a replicating Ad in vivo and provides a good model of Ad pathogenesis. hAds do not replicate in rodents, but high dose (108 to 1010 PFU per animal) intranasal (i.n.) infection of cotton rats and mice induces pulmonary disease (Ginsberg et al. 1991; Pacini et al. 1984). In contrast to hAds, MAV-1 doses as low as 1–100 PFU cause fatal disease in newborn and adult mice (Spindler et al. 2001; van der Veen and Mes 1973). The outcome of MAV-1 infection depends on the virus dose, the mouse strain and age, the inoculation route, and the strain of virus.
When mice survive an acute MAV-1 infection, they can become persistently infected (reviewed in Smith and Spindler 1999). Infectious MAV-1 was found at high titers in urine of infected mice at 11 months PI (Rowe and Hartley 1962) and 24 months PI (van der Veen and Mes 1973). Infectious virus was found in kidney 70 days PI (Ginder 1964) and in liver 52 days PI (Wigand 1980). Virus particles were detected in urine, and viral DNA was detected in brains, spleens, and kidneys of mice 55 weeks PI (Smith et al. 1998). E1A mutants of MAV-1 persist for up to 55 weeks PI (Smith et al. 1998) (Section VI, B). The mechanism by which MAV-1 persists is not understood (Smith and Spindler 1999). This long-term shedding of MAV-1 is paralleled in human adenoviruses, which have been found shed in feces up to 2 years after infection (Fox et al. 1969). Human adenoviruses have also been isolated from urine of patients with AIDS (de Jong et al. 1983). Mouse adenoviruses have not been reported to be oncogenic, and MAV-1 virions and viral DNA do not cause transformation of cloned rat embryo fibroblast (CREF) cells, unlike hAd virions and viral DNA (K. Spindler, unpublished data). The effects of mouse age on infection are as follows. In adult mice MAV-1 induces clinical signs of disease and dose-dependent acute encephalomyelitis in outbred (Kring et al. 1995), C57BL/6 (B6) (Guida et al. 1995), 129 Sv/Ev (M. Moore and K. Spindler, unpublished data), and SJL/J (Spindler et al. 2001) mice. Adult BALB/c mice are resistant to MAV-1-induced disease, but high dose infection results in ruffled fur, hyperpnea, and conjunctivitis (Charles et al. 1998; Guida et al. 1995; Moore et al. 2004). In suckling mice MAV-1 produces fatal, disseminated disease and myocarditis (Blailock et al. 1967; Hartley and Rowe 1960), and intranuclear inclusions typical of adenovirus infections are seen in endothelial cells of the brain (Heck et al. 1972). Similar widely disseminated disease is seen in suckling mice infected with both the “standard” and “ATCC” strains of MAV-1 (Ball et al. 1991) (Section II). In both adult and suckling mice, regardless of dose, infection results in viremia as early as 1 day PI and is detected in tissues at 3 days PI (Heck et al. 1972; Spindler et al. 2001); viral DNA is found in tissues as early as 2 days PI (Ball et al. 1991). The effects of mouse strain differences in the disease outcome are discussed below (Section VII).
2. MOUSE ADENOVIRUSES
Inoculation of MAV-1 by different routes does not result in major differences in pathogenesis. Outbred mice infected i.n. with MAV-1 exhibit slightly fewer disease signs and have protracted infection kinetics compared to mice infected intraperitoneally (i.p.) (Kajon et al. 1998; Wigand 1980). High dose (106 PFU) i.n. infection of newborn mice results in a robust macrophage infiltrate in the lung at 3 days PI (Gottlieb and Villarreal 2000). Intravenous (i.v.) infection of inbred mice results in levels of virus in brain and spleen, early antibody responses, and survival in B6 and B cell–deficient mice similar to what is seen in i.p. infection (Moore et al. 2004). Intracerebral (i.c.) inoculation of outbred mice results in significant infection of the adrenal gland (Margolis et al. 1974), an organ which also has significant viral DNA levels when infected i.p. (Kring et al. 1995). Similar to mice infected i.p. with MAV-1 (Spindler et al. 2001), susceptible and resistant adult inbred mice infected i.c. showed replication of the virus in brain and spleen (Fig. 2-5A). Interestingly, in the resistant BALB/c mice, virus was detected at high levels in the spleen as early as 2 days PI and was even detected in one susceptible SJL spleen at 2 days PI. The high levels of virus found earlier in BALB/c spleens were surprising, since BALB/c mice are resistant to the virus (Guida et al. 1995; Spindler et al. 2001). One interpretation of the data is that earlier high levels of virus in spleen correlate with or stimulate a stronger innate immune response in BALB/c mice that is able to control subsequent replication in resistant mice. To determine whether
57 the i.c. injections would result in a difference in viral titers at 8 days PI, mice were infected with three low doses of MAV-1 (Fig. 2-5B). Higher levels of virus were found in the brains and spleens of SJL mice than BALB/c mice at every dose (except spleens at the lowest dose), similar to results of infection of SJL mice by the i.p. route (Spindler et al. 2001; Welton et al. 2005). Taken together, the data suggest a model where MAV-1 replicates equally well in susceptible SJL and resistant BALB/c mouse brains, but as the infection proceeds, viral replication is controlled in resistant mice but not in susceptible mice, resulting in higher viral loads at later times after infection. Pregnant mice infected with MAV-1 show histological signs of infection, but the virus does not cross the placenta (Lipps and Mayor 1980; Margolis et al. 1974). Maternal antibodies to MAV-1 are protective for suckling mice (Hartley and Rowe 1960; Lipps and Mayor 1982). In most strains of MAV-1-infected adult mice, MAV-1 infects cells of the monocyte/macrophage lineage and endothelial cells of the vasculature throughout the mouse; highest levels of virus are found in the spleen and central nervous system (CNS) (Charles et al. 1998; Guida et al. 1995; Kajon et al. 1998; Kring and Spindler 1990). MAV-1 nucleic acid is also detected by in situ hybridization (ISH) in the renal tubular epithelium of adult outbred mice infected i.p. or i.n. (Kajon et al. 1998; Smith et al. 1998). Viral DNA is detected in organ homogenates of bowel, pancreas, spleen, adrenal gland, kidney, liver, lung, heart, brain,
Fig. 2-5 Intracerebral MAV-1 infections of mice. A. Anesthetized susceptible SJL/J and resistant BALB/c mice were injected i.c. with 104 PFU MAV-1 in a volume of 30 µl. Mice were euthanized at the indicated days PI (dpi), and brain and spleen titers were determined by plaque assay. Each symbol represents an individual mouse; the short horizontal lines indicate the mean values. The arrow indicates the input dose in PFU/g, assuming brains have a mass of 0.3 g. The asterisk and dotted line indicate the level of detection. B. Mice were infected i.c. with the indicated dose of virus, and brain and spleen titers were determined at 8 days PI. P < 0.03 for the 3 PFU brain and spleen titers between SJL and BALB/c mice; P = 0.002 for the 30-PFU brain and spleen titers.
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and spinal cord of i.p. infected outbred mice (Kring et al. 1995); most likely this is due to infection of endothelium throughout the mice. MAV-1 has not been observed in the brain parenchyma; it is restricted to the vascular endothelium (Charles et al. 1998; Guida et al. 1995; Kajon et al. 1998). Mice with MAV-1-induced encephalomyelitis exhibit histological evidence of perivascular edema in the CNS, and moribund infected mice also exhibit endothelial cell reactivity, vasculitis, vascular wall degeneration, and viral inclusion bodies in microvascular endothelial cells (Charles et al. 1998; Guida et al. 1995; Kajon et al. 1998; Kring et al. 1995; Spindler et al. 2001).
B.
Sublethal doses of γ-irradiation result in increased levels of MAV-1 in infected mice 42–55 weeks PI (Smith et al. 1998). The percentage of mice with viral DNA–positive organs (brain, kidney, and spleen) increases in post-irradiation mice previously infected with wt or E1A mutant viruses, peaks in the first week following irradiation, and returns to pre-irradiation percentages by 3 weeks PI. Thus MAV-1 can persist in the brains, spleens, and kidneys of infected mice for up to 55 weeks PI, the virus can be shed in the urine, and E1A is not required for the persistence of the virus. It is not known whether the virus produced is infectious virus, nor is it known whether persistence is due to a chronic or latent infection.
MAV-1 E1A Mutant Infection In Vivo
The 50% lethal dose (LD50) of each E1A mutant virus (Section IV, B) was determined and is shown in Table 2-1. The LD50 values for the E1A mutants are 1.5–5 log units higher than that of wt virus, indicating that the E1A protein is important in MAV-1 pathogenesis. pmE112, the virus that does not synthesize the E1A protein, is the least virulent (Smith et al. 1998). The disease signs observed in the mutant virus–infected mice are identical to those of wt-infected mice, and the onset of disease is dose-dependent. Studies of the E1A null mutants, pmE109 and pmE112, were carried out in mice to evaluate histopathology and the presence and quantity of viral DNA in mice (Smith et al. 1998). Neither wt nor E1A null mutant viruses produce detectable levels of virus, as measured by dot blot analysis of DNA, in the spleens at 5 d PI in outbred mice infected with 104 PFU. Brains of wt and pmE109-infected mice have similar levels of viral DNA, but the other E1A null mutant, pmE112, produces significantly less viral DNA in the mouse. At lower dose infections, near the LD50 for the wt virus, no viral DNA is detected in brains or spleens of the pmE109- or pmE112-infected mice at 5 or 14 d PI. Mutations in the E1A null mutants did not revert in vivo, because PCR amplification and sequencing of viral DNAs recovered from infected mice showed they were identical to the starting mutant viruses (K. Smith and K. Spindler, unpublished data). The E1A mutant viruses exhibit histopathology similar to that of wt virus during the acute phase of disease (Smith et al. 1998). When doses of 104 PFU are used, the tropism of pmE109 and pmE112 is similar to that of wt virus, with the exception that pmE112 is also found in thymus of infected mice. Persistence of wt and E1A mutant MAV-1 was evaluated using an immunocapture assay of urine from virus-infected mice (Smith et al. 1998). During the 12–22 week period PI, mice infected with wt and E1A mutant viruses all shed MAV-1. From 42–55 weeks PI, only wt-infected mice shed detectable levels of virus. At 42 weeks PI viral DNA is detected in brains, spleens, and kidneys of mice infected with wt and E1A mutant viruses by PCR amplification and ISH, indicating that a persistent MAV-1 infection can be established in the absence of E1A.
C.
MAV-1 E3 Mutant Infection In Vivo
Table 2-2 shows the LD50 values for infection of outbred mice by E3 mutant viruses (Section V, C), which are all less virulent than wt virus; the E3 null mutant pmE314 has the most severe defect in virulence. The elevated LD50 levels for each of the E3 mutant viruses suggest that each of the three E3 gene products plays a role in the pathogenesis of MAV-1 in vivo. Infections with E3 mutant viruses result in the same clinical signs of disease in mice as wt virus, and the onset of disease is dependent on the dose of virus (Cauthen et al. 1999; and C. Beard, N. Cauthen, and K. Spindler, unpublished data). Similar to wt virus, pmE314 is found primarily in endothelial cells of the brain and spinal cord and in endothelial cells and stationary macrophages in the spleen (Cauthen et al. 1999). Outbred mice given 105 or 106 PFU of pmE314 die 3 or 4 d PI with large numbers of viral inclusion bodies and ISH evidence of virus; it is thought that at this dose the mice die of an overwhelming infection of endothelial cells. Lower doses (103 or 104 PFU) of pmE314 given to outbred mice result in fewer and less severe histopathological changes than wt virus, particularly with respect to inflammation and endothelial damage. Viral nucleic acid is distributed in a similar pattern in the brains and spinal cords of both pmE314- and wt-infected mice, albeit at slightly lower levels in pmE314-infected brains and spinal cords. The functions of the E3 gene products have not yet been elucidated, thus the mechanism by which the lack of E3 results in reduced inflammation in pmE314 infection is unknown. D. 1.
Immune Response to MAV-1
Innate Immune Response to MAV-1
The innate immune response to MAV-1 includes early increases in the steady-state levels of the mRNAs of cytokines and chemokines (Charles et al. 1999; Charles et al. 1998). Charles and coworkers quantitated cytokine and chemokine mRNAs in mock-infected and MAV-1-infected B6 and BALB/c brains because BALB/c mice are more resistant to MAV-1-induced encephalomyelitis than B6 mice (Guida et al. 1995). MAV-1
2. MOUSE ADENOVIRUSES
increases the mRNA steady-state levels of the cytokines IFN-γ, tumor necrosis factor alpha (TNF-α), interleukin-1 (IL-1), IL-6, and lymphotoxin (LT) in B6 and BALB/c brains 4 days PI (Charles et al. 1998) and transiently increases expression of IL-12 mRNA by macrophages in CBA/Ht mice (Coutelier et al. 1995). MAV-1 also increases the mRNA steady-state levels of the chemokine receptors CCR1–CCR5 in B6 and BALB/c brains 4 days PI (Charles et al. 1999). Steady-state mRNA levels of the chemokines IFN-γ-induced protein 10 (IP-10), monocyte chemoattractant protein 1 (MCP-1), and T cell activation gene 3 (TCA-3) are increased by MAV-1 infection in B6 but not BALB/c brains 4 days PI, suggesting that innate immune responses contribute to the MAV-1-induced encephalomyelitis in B6 mice. IFN-α/β signaling plays a role in MAV-1 pathogenesis and is a determinant of MAV-1 organ tropism. In single-cycle infectious yield reduction assays in vitro, MAV-1 is more resistant than VSV to pretreatment with mouse IFN-α/β (Kajon and Spindler 2000). However, MAV-1 replication is not completely resistant to IFN-α/β, since virus yields are reduced about 10-fold in high concentrations of IFN. E1A mutants of MAV-1 are more sensitive to IFN-α/β in vitro than wt MAV-1. These data indicate that MAV-1 E1A counteracts the IFN-α/β antiviral response in vitro. The role of IFN-α/β in MAV-1-induced encephalomyelitis was examined by comparing the pathogenesis of MAV-1 in 129 Sv/Ev and IFN-α/βR−/− mice (M. Moore and K. Spindler, unpublished data), which are defective for type I IFN signaling (M¨uller et al. 1994). IFN-α/βR−/− mice had higher levels of infectious MAV-1 in spleens at 4 and 7 days PI and exhibited a more disseminated MAV-1 infection at 7 days PI than control mice (M. Moore and K. Spindler, unpublished data). However, this disseminated MAV-1 infection in IFN-α/βR−/− mice did not show clinical or histopathological differences from infection of control mice, and survival was not different between IFN-α/βR−/− mice and controls given a 700 PFU dose of virus. Virus levels in brain were high in both 129 Sv/Ev (control) and IFN-α/βR−/− mice. These results suggest that IFN-α/β signaling is correlated with reduced MAV-1 replication in the spleen and prevention of widespread infection of vascular endothelial cells. However, IFN-α/βR signaling does not prevent MAV-1-induced encephalomyelitis or limit MAV-1 replication in the brain. ISGs whose steady-state RNA levels were increased by MAV-1 infection in vitro and in vivo were identified first by cDNA arrays and confirmed by Northern analysis (M. Moore and K. Spindler, unpublished data). These ISGs included the transcription factors interferon regulatory factor 7 (IRF-7), interferon regulatory factor 1 (IRF-1), and signal transducer and activator of transcription 1 (STAT-1). 2. Cell-Mediated Immune Response to MAV-1
Outbred mice infected i.p. with a sublethal dose develop MAV-1-specific cytotoxic T cells (CTL) that are detectable 4 days PI, peak at 10 days PI, then rapidly decline (Inada and Uetake 1978a, 1978c). These kinetics are paralleled by
59 cell-mediated immunity measured by induction of macrophage migration inhibitory factor (Inada and Uetake 1978b) and are typical of acute viral infections in mice. Several studies implicate a protective role for adaptive immunity in MAV-1-induced disease. MAV-1-infected athymic nu/nu (T cell–deficient) mice on a mixed NIH Swiss and C3H/HeN background succumb to a wasting disease with characteristic duodenal hemorrhage and intranuclear adenovirus particles in endothelial cells (Winters and Brown 1980). Mice that are homozygous for the severe combined immunodeficiency (SCID) mutation (T cell– and B cell–deficient) on a CB.17 or BALB/c background succumb to MAV-1 infection with diffuse hepatic injury that resembles Reye syndrome pathology (Charles et al. 1998; Pirofski et al. 1991). RAG-1−/− (T cell– and B cell–deficient) mice are more susceptible to MAV-1 infection than B6 controls (Moore et al. 2004). These studies suggest that adaptive immunity protects adult mice from MAV-1-induced disease. Furthermore, sublethal irradiation of inbred mice that are resistant to MAV-1 infection renders them susceptible, suggesting that resistance to MAV-1 infection has an immunological basis (Spindler et al. 2001). Infection of mice deficient for T cells, T cell subsets, and T cell–related functions revealed that T cells cause acute immunopathology and are required for long-term survival in MAV-1-induced encephalomyelitis (Moore et al. 2003). Brains harvested from MAV-1-infected mice lacking α/β T cells or perforin have less histological evidence of MAV-1 encephalomyelitis and less cellular inflammation than brains harvested from control mice. Mice lacking α/β T cells, MHC class I (β2m−/−), or perforin have fewer disease signs at 8 days PI than control B6 mice, whereas mice lacking MHC class II have acute disease signs like B6 controls, such as hunched posture, ataxia, and ruffled fur. Thus, CD8+ CTL contribute to disease severity in the acute phase of MAV-1 infection. Similar to virus-induced disease in other virus infections, MAV-1-induced disease in B6 mice depends on virus dose and cell-mediated immunity (Moore et al. 2003), supporting the view that antigen quantity controls T cell–mediated immunity (Zinkernagel and Hengartner 2001). Mice lacking α/β T cells succumb to MAV-1 infection 9 to 16 weeks PI. These mice have detectable viral loads in spleens and brains at 3 weeks PI and high viral loads in spleen and brains when moribund (Moore et al. 2003). In contrast, control B6 mice clear MAV-1 to a level below the plaque assay detection limit by approximately 12 days PI, and no infectious virus is recovered 12 weeks PI from B6 mice (Moore et al. 2003; M. Moore and K. Spindler, unpublished data). Somewhat surprisingly, neither MHC class-I deficient, MHC class II-deficient, CD8−/−, CD4−/−, perforin-deficient, nor IFN-γ-deficient mice have any detectable infectious virus in spleens or brains at 12 weeks PI. Since almost all α/β T cells are either CD8+ or CD4+ (Mombaerts et al. 1992), these results suggest that having either CD8+ or CD4+ effector α/β T cells is sufficient for α/β T cell–mediated clearance of MAV-1.
60 3.
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Humoral Immune Response to MAV-1
Early studies showed that MAV-1, like hAds, induces a strong humoral immune response. Outbred mice infected i.p. with a sublethal dose of MAV-1 develop high neutralizing antibody (nAb) titers 2 weeks PI, and nAb titers increase for one year then decline (van der Veen and Mes 1973). MAV-1 infection of CBA/Ht mice with 107 50% infectious doses (ID50) results in significant splenic B cell proliferation at 10 days PI (Coutelier et al. 1990) and, like other viral infections of mice, stimulates predominantly antiviral IgG of the IgG2a subtype (Coutelier et al. 1990, 1988, 1987). MAV-1 acts as a T cell–independent (TI) Ag of the TI-2 type (Moore et al. 2004); that is, like polyomavirus, it induces TI antibodies in immunocompetent but not Xid mice (SzomolanyiTsuda and Welsh 1998). Early TI antiviral IgM plays a crucial role in protection against disseminated MAV-1 infection (Moore et al. 2004). In contrast to mice lacking T cells, mice lacking B cells are more susceptible to acute MAV-1-induced disease than B6 controls. B cell–deficient mice die early (7 to 10 days PI), and T cells are not required for survival of acute infection (Moore et al. 2003). These findings are consistent with TI B cell responses being critical for protection against MAV-1-induced encephalomyelitis. Mice lacking Bruton’s tyrosine kinase (Btk) have reductions in serum immunoglobulin, conventional B cells, and peritoneal B-1 cells (Khan et al. 1995). Btk is required for survival of acute MAV-1 infection, since Btk-deficient mice succumb to MAV-1 infection, (Moore et al. 2004). This was the first demonstration that Btk plays a role in protection from virus-induced disease in mice. Btk-deficient and µMT mice, deficient for B cells and on B6.129 and B6 strain backgrounds, respectively, succumb to acute MAV-1 infection with systemically high viral loads and histological evidence of hepatitis in addition to the histological evidence of MAV-1-induced encephalomyelitis (Moore et al. 2004). B cell–deficient mice on a BALB/c background (Jh mice) are more susceptible to acute MAV-1-induced disease than BALB/c controls, and succumb with systemically high viral loads and evidence of significant hepatitis. However, moribund Jh mice do not exhibit encephalomyelitis; MAV-1-infected Jh mice likely die of hemorrhagic enteritis. SCID (T cell– and B cell–deficient) mice on a BALB/c background succumb to acute MAV-1 infection with evidence of liver infection but no histological evidence of hepatitis (Charles et al. 1998). One explanation for this strain-specific pathology is as follows. In the absence of B cells, MAV-1 replicates to high titers throughout the mouse (Moore et al. 2004). In the presence of T cells, acute MAV-1 infection elicits dose-dependent immunopathology (Moore et al. 2003). T cell–mediated immunopathology may be directed to various organs in different mouse strains by strain-specific innate immune responses, for example, differential chemokine expression in the brains of MAV-1-infected B6 and BALB/c brains (Charles et al. 1999). Data showing that MAV-1 replicates to high levels in the brains of Jh mice without inducing encephalomyelitis (Moore et al. 2004) support this
model rather than a model in which receptor differences account for differential pathology in MAV-1-infected B6 and BALB/c mice (Charles et al. 1998). 4.
Model of MAV-1 Immunopathogenesis
T cells, B cells, and type I IFN each have a distinct protective role in MAV-1 pathogenesis, and the data suggest their roles are interrelated. B cell–deficient mice succumb to disseminated infection with high virus loads throughout the mouse (Moore et al. 2004). T cell–mediated immunopathology is implicated in exacerbating disease in B cell–deficient mice (Moore et al. 2003, 2004). Similar to B cell–deficient mice, type I IFN–deficient mice also had a more disseminated infection with higher virus loads than control mice (M. Moore and K. Spindler, unpublished data). However, unlike B cell–deficient mice, type I IFN–deficient mice did not exhibit more clinical disease or more histopathological evidence of disease than control mice, even in organs with high virus loads, such as the liver (M. Moore and K. Spindler, unpublished data). This suggests that type I IFN controls MAV-1 replication but also contributes to T cell–mediated immunopathology.
E. MAV-2 Infection In Vivo Hashimoto and colleagues studied the pathogenesis of MAV-2 in mice infected perorally (Hashimoto et al. 1970). The virus grows in the intestinal tract and is shed in feces for 3 weeks after infection, but there are no clinical signs of disease. Inbred DK1 mice given 2 × 105 TCID50 of virus show virus replication from 3–14 days PI, with a peak of virus yield from 7–14 days PI. At the peak times of infection, virus is seen by immunofluorescence and electron microscopy in epithelial (columnar, goblet, and Paneth) cells of the ileum (Takeuchi and Hashimoto 1976). Mesenchymal cells are not infected; the virus has a specific tropism for the villus epithelium. The infection results in little cytopathic effect, but infected epithelial cells are shed at a high rate into the gut lumen. Interestingly, MAV-2 peroral infection of BALB/c nude (nu/nu) mice, which lack T cells, results in prolonged viral proliferation in the gut, but viral replication is suppressed 6 weeks PI (Umehara et al. 1984). MAV-2 is not found in organs other than the gut. Antiviral resistance in BALB/c nu/nu mice 6 weeks PI does not correlate with interferon levels or NK cell activity, and the resistance is not affected by administration of anti-asialo GM1 antibody or carrageenan (Umehara et al. 1987). However, antiviral resistance is completely abolished by cyclophosphamide treatment. Cyclophosphamide is a carcinogen and mutagen that is toxic to actively cycling cells, reduces peripheral lymphocytes, and reduces serum IgG levels in nu/nu mice. Mice rechallenged with MAV-2 28 days after initial infection are resistant to virus growth (Hashimoto et al. 1970). The data suggest that antibody
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responses are responsible for the antiviral resistance of nu/nu mice to MAV-2.
VII.
HOST GENETICS
Different strains of outbred and inbred mice have been shown to differ in their susceptibility to MAV-1 (Guida et al. 1995; Kring et al. 1995; Spindler et al. 2001). Adult B6 mice infected with MAV-1 succumb to a fatal hemorrhagic encephalitis whereas BALB/c mice do not (Guida et al. 1995). B6 mice show clinical signs of acute CNS disease accompanied by histological evidence of hemorrhage and inflammation, and high levels of virus in brain and spinal cord. In contrast, infected BALB/c mice do not have detectable levels of viral mRNA in brain or spleen, and they lack histological and clinical signs of disease. Enteritis has been seen in BALB/SCID (T and B lymphocyte–deficient) mice (Charles et al. 1998), and B lymphocyte–deficient mice on a BALB background (Jh) (Moore et al. 2004) likely succumb due to a hemorrhagic enteritis. These results suggest that there are mouse strain differences in tropism and cause of death. SJL/J mice are more than four log units more susceptible to MAV-1 than most inbred strains of mice (Spindler et al. 2001). A sublethal dose of gamma irradiation renders resistant mice susceptible, and there are no differences in viral yield in ex vivo MAV-1 infection of primary cells from susceptible and resistant mice. Susceptible mice have higher virus loads in brain and spleen than resistant mice but only modest differences in histopathology. The results suggest that immune response differences may account for differences in susceptibility. A genetic mapping approach (positional cloning) is being used to identify the host gene(s) for susceptibility to MAV-1 (Welton et al. 2005).
VIII.
HOST RANGE AND PREVALENCE
The mouse strain used when MAV-1 was isolated was not specified by Hartley and Rowe (1960), but MAV-1 infects inbred and outbred strains of mice in the Mus genus. There is a report of adenovirus isolation from Peromyscus (Reeves et al. 1967) and a report of a seropositive wild rodent (Kaplan et al. 1980), but these do not provide strong support for infections of non-Mus mice. The host range of MAV-1 in cells in culture is limited to mouse cells; infection of a variety of human and monkey cells does not result in infectious virus (Antoine et al. 1982; Larsen and Nathans 1977). A survey reported in 1966 testing mice in U.S. laboratory colonies indicated that evidence of MAV-1 infection was only found in 4 of 34 colonies, much less frequently than other viruses tested (Parker et al. 1966). A survey from 1984–1988 of
laboratory colonies in 10 European countries showed no mouse adenovirus infections (Kraft and Meyer 1990). MAV-1 is virtually absent from commercial colonies, which are now monitored routinely for MAV-1 (Otten and Tennant 1982). Commercial and laboratory colonies are not usually tested for MAV-2 (A. Smith, personal communication). It has been postulated that MAV-1 occurred enzootically in infected laboratory colonies as a silent infection that was transmitted orally to cage mates (Richter 1986). Although infection can be transmitted to cage mates, there is no seroconversion for animals held in the same animal room but in separate cages (Hartley and Rowe 1960). Mice kept in close contact with MAV-1-infected mice seroconvert by day 21 PI (Lussier et al. 1987). Animals placed on bedding obtained from cages of MAV-1-infected animals do not show signs of disease, seroconversion, or shedding of virus in urine (Smith et al. 1998). The prevalence of mouse adenoviruses in the wild has not been systematically studied. However, a serological survey of wild M. domesticus in southeastern Australia indicated that of mice isolated from 14 sites, 37% were seropositive for MAV-2 and 0% were seropositive for MAV-1 (Smith et al. 1993). There was significant variation in MAV-2 seroprevalence at two study sites in southeastern Australia investigated over 13 months, and again, no evidence of MAV-1 was seen (Singleton et al. 1993). M. domesticus was introduced to an island off the northwest coast of western Australia and first identified in 1986 (Moro et al. 1999). A study of mice from this island in 1994–1996 indicated no serological evidence of MAV-1 or MAV-2 in M. domesticus or in an indigenous mammal, the short-tailed mouse Leggadina lakedownensis.
IX.
DIAGNOSIS, CONTROL, AND PREVENTION
MAV-1 infection can be diagnosed by serological testing by ELISA; test kits are available commercially (Charles River Laboratories). Complement-fixing antibodies are detected in mice inoculated i.p. with 104 TCID50 as early as 14 days PI (Lussier et al. 1987). Neutralizing Ab is detected in mice infected with 1 PFU at 12 days PI (Moore et al. 2004). Because commercial colonies are free of mouse adenoviruses (Section VIII), control is unlikely to be needed. If necessary, control in an infected colony can be achieved by embryo rederivation (Richter 1986; Trentin et al. 1966). Mice to be infected with MAV-1 should be maintained in a room where their cages and bedding can be autoclaved after use. Because the virus appears to require close mouse-mouse contact for transmission (Hartley and Rowe 1960; Lussier et al. 1987), it is straightforward to work with these mice using standard microisolator techniques. In 20 years of performing MAV-1 infections in animal facilities in four different buildings at two institutions, we have never had mock-infected experimental mice (in separate cages) or sentinel mice (even in open cages) in the same
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room as infected mice show disease signs or become seropositive for MAV-1 (K. Spindler, unpublished data). ACKNOWLEDGMENTS We thank Lei Fang for critical reading of the manuscript. This work was supported by NIH R01 AI023762.
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65 Winters, A. L., Brown, H. K., and Carlson, J. K. (1981). Interstitial pneumonia induced by a plaque-type variant of mouse adenovirus. Proc. Soc. Exp. Biol. Med. 167, 359–364. Wold, W. S. M., and Gooding, L. R. (1991). Region E3 of adenovirus: a cassette of genes involved in host immunosurveillance and virus-cell interactions. Virology 184, 1–8. Yee, S.-P., and Branton, P. E. (1985a). Analysis of multiple forms of human adenovirus type 5 E1A polypeptides using an antipeptide antiserum specific for the amino terminus. Virology 146, 315–322. –– –– ––. (1985b). Detection of cellular proteins associated with human adenovirus type 5 early region 1A polypeptides. Virology 147, 142–153. Yee, S.-P., Rowe, D. T., Tremblay, M. L., McDermott, M., and Branton, P. E. (1983). Identification of human adenovirus early region 1 products using antisera against synthetic peptides corresponding to the predicted carboxy termini. J. Virol. 46, 1033–1013. Ying, B., Smith, K., and Spindler, K. R. (1998). Mouse adenovirus type 1 early region 1A is dispensable for growth in cultured fibroblasts. J. Virol. 72, 6325–6331. Zinkernagel, R. M., and Hengartner, H. (2001). Regulation of the immune response by antigen. Science 293, 251–253.
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Chapter 3 Mousepox R. Mark, L. Buller, and Frank Fenner
I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . II. History . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . III. Properties of the Agent . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Classification . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Virion Morphology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Physical Properties . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1. Virion Structure and Composition . . . . . . . . . . . . . . . . . . . . . . . . . . 2. Virion Stability . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . IV. Strains of Virus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Hampstead Strain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Moscow Strain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. NIH-79 Strain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. Other strains . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . V. Growth In Vitro and In Vivo . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Replication . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Growth in Tissue Culture and Eggs . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1. Chick Embryo . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2. Tissue Culture . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VI. Clinical Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Clinical Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Factors Affecting Clinical Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1. Age . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2. Gender . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VII. Pathogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. The Mechanism of Infection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1. Intradermal Inoculation and Scarification . . . . . . . . . . . . . . . . . . . . 2. Feeding . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3. Arthropod Vectors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4. Intraperitoneal Inoculation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5. Intranasal Inoculation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6. Lower Respiratory Tract Inoculation . . . . . . . . . . . . . . . . . . . . . . . . 7. Intracerebral Inoculation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8. Intrauterine Infection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Virus Spread in the Animal Following Footpad Infection . . . . . . . . . . .
THE MOUSE IN BIOMEDICAL RESEARCH, 2ND EDITION
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Copyright © 2007, 1980, Elsevier Inc. All rights reserved.
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C. Gross and Microscopic Pathology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1. Susceptible Mouse Strains . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2. Resistant Mouse Strains . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VIII. Innate and Adaptive Immune Responses . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Innate Resistance . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Adaptive Immunity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Genetics of Resistance to Lethal Mousepox . . . . . . . . . . . . . . . . . . . . . IX. Epizootiology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Host Range . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1. Species . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2. Strains of Mice . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Prevalence and Distribution . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1. Europe . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2. United States . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3. Other Regions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Epizootic Mousepox . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. Enzootic Mousepox . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . X. Diagnosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Clinical Signs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Serology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Pathology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. Virus Isolation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . E. Molecular and Antigen Detection Techniques . . . . . . . . . . . . . . . . . . . . XI. Control and Prevention . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Control . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1. Depopulation and Disinfection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2. Vaccination . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3. Serological Screening . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4. Rederivation of Mouse Strains . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Prevention . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1. Sentinel Surveillance . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2. Quarantine . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3. Mouse Antibody Production . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References .....................................................
I.
INTRODUCTION
First recognized in 1930 (Marchal 1930), when the use of mice as experimental animals in virology was just beginning (see Burnet 1960, Table 3-1), infectious ectromelia, or mousepox, has had a rather different history in the four continents where laboratory mice have been extensively used: Europe, North America, Australia, and Japan. In Europe and Japan it was soon found to be present, usually as an unrecognized enzootic infection, in many breeding colonies. As well as threatening potentially valuable mouse stocks, this infection complicated much virological research involving serial passages of viruses or tumors in mice, and the presence of enzootic mousepox has also rendered suspect several studies of the nature of the disease itself. By chance, most mouse colonies established in North America were free from the disease. When the virus was inadvertently imported into the United States from Europe with mouse strains or mouse tissues (cell lines or tumors), there were sometimes disastrous outbreaks in colonies (see Briody 1955; Whitney 1974; Lipman et al. 2000); hence, quarantine precautions were instituted, and research with the
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virus in the United States was carried out with great care. Enzootic mousepox does not occur in Australia. Following the recognition of the relation of the virus to vaccinia virus by Burnet and Boake (1946), mousepox has been used as a model for research in several fields, notably the pathogenesis of generalized infections, experimental epidemiology, and the cellular immune response. In this age of concern over the potential use of pathogens as weapons of mass destruction, there is a renewed interest in development of prophylactics and therapeutics against smallpox. The mousepox model is arguably the best mouse model available for this purpose, as it provides a much greater dynamic range for evaluating antivirals and vaccines and shares a number of important similarities with smallpox (Buller 2004; Schriewer et al. 2004). Also, studies with ectromelia have suggested methods for generating more virulent orthopoxviruses that possibly could be used as bioweapons. For example, by accident, Jackson et al. (2001) using ectromelia virus (ECTV) as a vector for immunocontraception of feral mouse populations discovered that ECTV expressing IL-4 was able to break through vaccine-induced immunity.
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TABLE 3-1
TAXONOMIC POSITION OF ECTROMELIA VIRUS Virus
Animals found naturally infected
Camelpox Cowpox Ectromelia Monkeypox Raccoonpox Skunkpox Taterapox Uasin Gishu pox Vaccinia Variola Volepox
Camel Cow, man, rodents, carnivores, elephant, okapi Mice, ? vole African monkeys, anteater, great apes, man, squirrels Raccoon Skunk Tatera kempi (an African gerbil) Horse Man, cow, buffalo, rabbit Man Vole
II.
histocompatibility complex (MHC) antigens in viral infections. In the 1980s and 1990s, Brownstein (1997) provided insights into the genetic basis of resistance to mousepox; most recently, Karupiah and colleagues explored the role of innate/immune host responses for recovery from infection, and Buller and coworkers investigated the genetic basis for ECTV virulence (reviewed by Esteban et al. 2005). In 2003, the annotated genomic sequence of the Moscow strain of ECTV was published (Chen et al. 2003). Mousepox is a troublesome and often devastating disease that interferes with experiments involving mice. This chapter will examine the natural history of mousepox in laboratory mice and the measures that can be used in prevention and control. In a less detailed manner, we will explore some results that have emerged from the use of mousepox as a model infection by virologists and immunologists.
HISTORY
Mousepox was first recognized by Marchal (1930) as an epizootic disease of laboratory mice in England, following investigation of unusually high mortality in mice received from commercial breeders by the National Institute of Medical Research at Hampstead. She named it infectious ectromelia because of frequent amputation of the extremities during the outbreak that she investigated. Soon after, Barnard and Elford (1931) demonstrated by ultraviolet microscopy that the virion (“elementary particle”) was similar in size and shape to that of vaccinia virus. However, no serological comparisons with other poxviruses were made until 1946, when Burnet and Boake (1946) demonstrated by hemagglutination inhibition (HI) (Nagler 1944) that ectromelia and vaccinia viruses were closely related. Fundamental studies with mousepox have been carried out in Australia, first at the Walter and Eliza Hall Institute in Melbourne and then at the John Curtin School of Medical Research in Canberra, in Spain at Madrid, and in the United States at the National Institutes of Health, Yale University, and Saint Louis University. During the 1930s, Greenwood and collaborators (1936) used ECTV as a model pathogen in experimental epidemics in mice, and in 1946 Fenner (for review, see Fenner 1949a) revived this work with the added knowledge of the taxonomic status of the virus. He soon found (Fenner 1948b) that infection produced a rash and revealed the pathogenesis of mousepox (Fenner 1948a) as a useful model for generalized viral exanthemata (Fenner 1948d). He suggested that the disease be called mousepox and the virus ectromelia virus (cf. smallpox and variola virus). During the 1950s and early 1960s, Mims and colleagues expanded Fenner’s work on pathogenesis using fluorescent antibody staining to probe events at the cellular level (for reviews, see Mims 1964, 1966). Blanden and colleagues exploited the mousepox model to explore the role of cell-mediated immunity in poxvirus infections and demonstrate the role of major
III.
PROPERTIES OF THE AGENT A.
Classification
Three distinct species of poxviruses produce natural infections of rodents: infectious ectromelia or mousepox virus (Marchal 1930), cowpox virus (Chantrey et al. 1999), and Turkmenia rodent virus (Marennikova et al. 1978), which is a close relative of cowpox virus. All three are orthopoxviruses with strong serological cross-reactivity to other orthopoxviruses (see Table 3-1). Comparative molecular studies have shown that the DNAs of orthopoxviruses are closely related, with greater than 90% nucleotide identity exhibited over the central ~100 kbp of their genomes (Bellett and Fenner 1968; Müller et al. 1978; Gubser et al. 2004). Serological comparisons of ECTV with other orthopoxviruses have been made mainly by neutralization and HI tests. Cross-neutralization by vaccinia virus–immune sera is readily demonstrable in pock-reduction tests on the chorioallantoic membrane (McCarthy and Downie 1948), and ECTV can be distinguished from vaccinia and cowpox viruses in plaque-reduction assays (McNeill 1968). Although they cross-react, the HI titers of ECTV and vaccinia virus antiserum are substantially higher with homologous antigen (Fenner 1947a).
B.
Virion Morphology
Poxviruses are the largest of all animal viruses and can be visualized by light microscopy, although details of virion structure remain obscure. With the advent of high-resolution electron microscopy, the morphology of the virion structure began to be revealed during the 1940s and 1950s. Ectromelia virus is a
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2.
Immature particles
IMV
Virion Stability
Like other orthopoxviruses, ECTV is relatively resistant to heat and to many disinfectants (see Fenner 1949a; Bhatt and Jacoby 1987c). For example, virus infectivity was still detectable following 50 days at room temperature or treatments for 24 hr with 10% ether, 0.5% chloroform, 1% phenol, or 0.01% formalin. Virus infectivity was not detectable following a 10-minute incubation with 2% phenol or 40% alcohol.
IV.
Fig. 3-1 An electron micrograph of a thin section of orthopoxvirusinfected cells. The structure of the virion reveals a condensed nucleoprotein organization of DNA. The core assumes a dumbbell shape surrounded by a core wall invaginated by large lateral bodies, which are in turn enclosed within a membrane to form intracellular mature virus (IMV). The circular and partially circular shapes are intermediate stages in virus assembly (immature particles).
typical orthopoxvirus, morphologically indistinguishable from the prototype species, vaccinia virus. The intracellular mature virus (IMV) appears to be an oval or brick-shaped structure of about 200 to 400 nm in length, with axial ratios of 1.2 to 1.7 (see Fig. 3-1) (Peters 1956).
C. 1.
Physical Properties
Virion Structure and Composition
The virion contains a noninfectious, linear, 67% A+T-rich, double-stranded DNA genome of 209,771 base-pairs in length (Chen et al. 2003). The virion has more than 100 polypeptides, arranged in four distinct structures (core, lateral body, membrane, and envelop), as determined by electron microscopy of thin sections and negatively stained preparations of purified virions. The membrane contains more than eight proteins, of which 129, 72, and 97 are targets of neutralizing antibody (Moss 2003). The extracellular enveloped virus (EEV) is an IMV virion with a second membrane (see below). The unique EEV membrane has six major proteins, with the 155 and 135 proteins implicated as important targets of protective antibody. Like most other orthopoxviruses, ECTV produces a hemagglutinin (151) that by analogy with vaccinia virus (Payne and Norrby 1976) is part of the viral envelope, the presence of which is not necessary for infectivity.
STRAINS OF VIRUS
Isolates from Manchester, England (McGaughey and Whitehead 1933), Paris (Schoen 1938), Germany (Kikuth and Gönnert 1940), Moscow (Andrewes and Elford 1947), Japan (Ichihashi and Matsumoto 1966), and the United States (New 1981; Dick et al. 1996) were found to be almost indistinguishable from the original Hampstead strain isolated by Marchal (1930) by serologic or genetic means. For example, restriction endonuclease analysis of genomic DNA from Hampstead and Moscow strains revealed an almost identical pattern of fragments following digestion with restriction endonucleases Hind III and Xho I (Mackett and Archard 1979). Similarly, DNA sequence analysis detected nucleotide identity of 99.5% over the entire length of the genomes of the Moscow strain and an isolate from an outbreak at the Naval Medical Research Institute in the United States (referred to as the NAV isolate), even though these isolates were obtained roughly 50 years apart (Chen et al. 2003). Taken together, the serologic and genomic nucleotide sequence similarity suggests that there is little genetic diversity among the strains; however, there are distinct biologic differences. Fenner (1949c) found that virulence and transmissibility varied independently and differed between strains. The Moscow strain was highly virulent and highly infectious; the Hampstead (mouse-passaged) strain had similar virulence but was less infectious. The Hampstead (egg) strain had low virulence and low infectivity. Table 3-2 illustrates differences in the LD50 of several strains of ECTV in BALB/c mice. The Hampstead, Moscow, and NIH-79 strains have been extensively characterized, as summarized below.
A.
Hampstead Strain
The original isolation was made by Marchal (1930). This strain was used in most of the early studies characterizing mousepox, and was used in Greenwood’s epidemiological experiments (Greenwood et al. 1936). A mouse-passaged Hampstead strain retained its high virulence, but egg passage led to a substantial reduction in its virulence for mice (Fenner 1949c), although it
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was then more readily adapted to growth on the rabbit cornea (Paschen 1936).
V.
GROWTH IN VITRO AND IN VIVO A.
B.
The Moscow strain of virus was isolated by Professor V. D. Soloviev and coworkers and is highly virulent and highly infectious. It was used extensively in experiments by a succession of Australian workers (Fenner, Mims, Roberts, Blanden, and Karupiah), by Andrewes and Elford, and more recently by Buller. The Moscow strain continues to be the most frequently studied ECTV strain.
C.
NIH-79 Strain
The NIH-79 strain of virus was isolated by Dr. Anthony Allen, Veterinary Resources Branch, National Institutes of Health (Allen et al. 1981). It was used by both Buller and Wallace and Bhatt and Jacoby to study the clinical response, pathogenesis, and transmissibility of ECTV in inbred strains of mice. The NIH-79 strain is less virulent than the Moscow strain, causing less clinical illness and fewer deaths among genetically resistant C57BL/6 mice and susceptible BALB/c mice (Bhatt et al. 1988; see Table 3-2).
D.
Other Strains
Japanese workers have made considerable use of strains recovered in Japan, as well as the Hampstead strain. Of these, the Ishibashi strain was the most thoroughly studied and was shown to have a larger plaque size than the Hampstead strain (Ichihashi and Matsumoto 1966). The NAV strain was isolated from commercial, pooled mouse sera, which was the source of a 1995 outbreak of mousepox in a laboratory mouse colony at the Naval Medical Research Institute in the United States (Dick et al. 1996; see Section IX, C).
TABLE 3-2
LETHAL DOSE50 OF ECTROMELIA VIRUS STRAINS AFTER FOOTPAD ROUTE INOCULATION IN BALB/CBYJ MICEa Virus strain Washington Univ. St. Louis 69 Moscow Beijing 70 NIH-79 Ishibashi I-III aR.
Replication
Moscow Strain
M. L. Buller, unpublished results
LD50 in PFU 1.0 × 101 2.8 × 101 3.9 × 101 9.3 × 101 7.9 × 102 4.3 × 104
Key features of the intracellular replication cycle of orthopoxviuses are shown in Fig. 3-2 (Moss 2003). The virion containing early RNA transcription machinery attaches to and fuses with the plasma membrane. Within 15 minutes, transcription machinery is activated (uncoating I). Early genes are expressed that code for a variety of functions that modify the host cell for optimal virus replication, attentuate the host response to infection, and mediate virus synthetic processes. After further uncoating (II), and between 1.5 to 6 hr PI, the virus genome is replicated via concatamers. Virus DNA is replicated in a region of the cytoplasm that is “cleared” of organelles and shows characteristic staining patterns by light and electron microscopy. Kato et al. (1955) called these structures B-type inclusion bodies and Cairns (1960) referred to them as viral factories. From progeny DNA templates, intermediate genes encoding late transcription factors are expressed, leading to the sequential synthesis of late gene RNA and proteins. Late genes encode the early transcription system, enzymes, and structural proteins necessary for virion assembly. By 4 hr PI, virion morphogenesis commences with the formation of membrane structures in the intermediate compartment of the cell and the packaging of resolved unit length genomic DNA. The IMV has one membrane derived from the intermediate compartment. Some IMV acquire an additional double layer of intracellular membrane derived from the trans Golgi network that contains unique virus proteins (intracellular enveloped virus, IEV). These IEVs are transported to the periphery of the cell, where fusion with the plasma membrane ultimately results in release of EEV or, if attached to the exterior surface of the plasma membrane, remain as cell-associated enveloped virus (CEV). While IMV and CEV/EEV are infectious, the external antigens on the two virion forms are different. EEV are thought to be most important in cell-to-cell spread and systemic disease. In addition to these forms, ECTV also encodes a major late protein of 130 kDa that forms characteristic spherical, acidophilic, cytoplasmic masses in infected epithelial cells, but rarely in liver cells (Kato et al. 1963). Kato et al. (1955) called these structures A-type inclusion or Marchal bodies (ATIs). Virions may be occluded within the ATI, providing a means of environmental survival. Electron microscopic examination of infected L cells showed that with the Hampstead strain, all mature viral particles that developed in the B-type inclusion were occluded within the ATIs, whereas with the Ishibashi strain, the ATIs were devoid of virions (Matsumoto 1958; Ichihashi and Matsumoto 1966). The genetic basis for this observation likely resides in the presence or absence of an intact p4c gene product on the surface of the IMV particle (McKelvey et al. 2002). The Moscow strain
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Fig. 3-2 General orthopoxvirus replication cycle. IMV, intracellular mature virus; IEV, intracellular enveloped virus; CEV, cell-associated enveloped virus; and EEV, extracellular enveloped virus.
lacks an intact p4c and does not occlude virions (R. M. L. Buller, unpublished observations).
B. 1.
Growth in Tissue Culture and Eggs
Chick Embryo
Ectromelia virus infection of chick embryos was described simultaneously by Paschen (1936) and Burnet and Lush (1936a). Both grew the virus on the chorioallantoic membrane, and Burnet and Lush showed that inoculation of dilute virus suspensions facilitated development of discrete foci (pocks), which could be counted. This titration method was exploited by Fenner (1948a) for quantitative studies of mousepox infection. Chorioallantoic membrane inoculation with large doses of virus was usually followed by death of the embryo 4 or 5 days later, and there were often scattered areas of necrosis in the livers and spleens of these embryos.
Serial passage of ECTV on the chorioallantois sometimes modified its character. Paschen (1936) found that egg-passaged virus was more suitable for infection of the rabbit and guinea pig cornea and skin than was mouse live-passaged virus. Serial chorioallantoic passage of the Hampstead strain of ECTV (50–60 passages intermittently over a period of 10 years) resulted in greatly reduced virulence for mice (Fenner 1949c). No change in the high virulence of the Moscow strain of virus occurred after 20 consecutive passages on the chorioallantois. 2.
Tissue Culture
Ectromelia virus multiplies in cells from human (HeLa and human amnion), monkey (BSC-1), mouse (L and primary embryonic fibroblasts), and chicken (primary embryo fibroblasts) sources. Ichihashi and Matsumoto (1966) found that the Ishibashi strain of ECTV produced much larger plaques on chick embryo fibroblast monolayers than the Hampstead strain. Plaque production in mouse fibroblasts can be improved by the
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inclusion of DEAE-dextran in the overlay medium. The plaque assay in mouse L cells or fibroblasts or monkey BSC-1 cells is as sensitive as the chorioalloantoic membrane assay, but more reproducible. The tissue culture plaque assay has been adopted by most investigators as the standard assay method (Blanden 1974). Comparative studies determined the following relationship between virus particles (counted by electron microscopy) and various biological titration units: mouse (strain C57BL) footpad inoculation, 1 ID50 and 1 LD50 = 1.5 and 25 particles, respectively; chorioallantoic membrane, 1 pock-forming unit = 12 particles; chick embryo fibroblasts or mouse embryo fibroblasts, 1 plaque-forming unit = 19–20 particles (Schell 1960b).
VI.
CLINICAL DISEASE A.
Clinical Disease
Clinical disease differs among mouse strains and routes of inoculation both in natural epizootics and experimental infections. Briody (1966) observed that in natural epizootics A, BC, DBA/1, DBA/2, and CBA strains usually did not recover from infection whereas C57BL/6 mice were resistant to severe disease. Schell (1960b) showed that C57BL mice were resistant to lethal experimental infection when infected by the footpad route, but not other routes. ID50/LD50 ratios for footpad, intravenous, intranasal, intracerebral, and intraperitoneal inoculation routes were 5.4, 4.0, 3.7, 1.3, and 1.0, respectively. Early workers using outbred mice (Marchal 1930; McGaughey and Whitehead 1933; Schoen 1938) described two forms of the disease: a rapidly fatal form in which apparently healthy mice died within a few hours of the first signs of illness 6–7 days post-infection (PI) and had extensive necrosis of liver and spleen, and a chronic form characterized by ulcerating lesions of the feet, tail, and snout. Fenner (for review, see 1949a) showed that in every case there was a stage in which virus multiplied to high titer in the liver and spleen. Some mice died at this stage, but survivors almost invariably developed a generalized rash. Subsequently, it was shown that clinical signs were greatly affected by mouse genotype (see Section VIII, C). Clinical disease in inbred mouse strains occurred in two patterns (Wallace and Buller 1985; Bhatt and Jacoby 1987a,b). Ruffled fur, hunched posture, and prostration were observed in all dying mice following footpad inoculation, which mimics natural infections. For strains such as A, DBA/2, C3H, and BALB/c, death usually ranged from 6 to 14 days PI, depending on the infecting dose of virus. The ID50 and LD50 were similar. With C57BL/6 and AKR mice, over a range of viral doses,
clinical signs were minimal and the LD50 dose was 4 or 5 log10 higher than the ID50 dose.
B. 1.
Factors Affecting Clinical Disease
Age
Age affects the response of genetically susceptible mice (Fenner 1949d). Both the Moscow and Hampstead (egg) strains produced higher mortalities in suckling mice and in mice about a year old than in 8-week-old mice, the differences being more pronounced with the less virulent Hampstead virus. The increased severity was evident after footpad inoculation and in long-term experimental transmission experiments. In suckling mice there was a very short delay between peripheral inoculation of the virus and its appearance in the liver and spleen. In the year-old animals this interval, and the survival time of the mice, were the same as in 8-week-old animals. However, lethal titers of virus in the liver and spleen were attained only in occasional 8-weekold mice but occurred in most older animals. The causes of these differences were not elucidated, but may relate to a less effective immune system. 2.
Gender
Sexual dimorphism to disease has been observed in BALB/cJ and A/J mouse strains, and appears to have a partial hormonal basis (Buller et al. 1985). Day of death differences were noted between males and females, and day of death was modulated in males by castration or treatment with estrogen. This sex-related difference in severity of disease, although evident in the parental strains, was much more apparent in back-crossed populations (Wallace et al. 1985; Buller et al. 1986). The female mice had a longer time to death and higher proportion of survivors than males.
VII. A. 1.
PATHOGENESIS
The Mechanism of Infection
Intradermal Inoculation and Scarification
Infection by scarification, by footpad inoculation, or by instillation of virus into the cornea was followed by a disease indistinguishable from naturally acquired mousepox except for the localization of the primary lesion. Fenner (1948a) studied the pathogenesis of Hampstead and Moscow strains following footpad inoculation of small doses of virus. The incubation period, viral dissemination, and pathology all resembled that found in natural infections. The results with both strains of
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virus and in subsequent experiments with immunized mice indicated that the infection followed a constant pattern (Fenner 1949b) (see Section VII, B for a detailed description of virus spread within the animal). Other methods of inoculation (discussed below) caused lesions that differ considerably from those found in natural disease. 2.
Feeding
Fenner (1947c) found that infection by feeding virus occurred only if very large doses of virus were used, but the ensuing disease resembled that following natural infections. However Gledhill (1962a,b) and subsequently Horzinek and Höpken (1965) found that mice occasionally could be infected orally with much smaller doses, but that the infections were usually inapparent. Gledhill showed that chronic infection of Peyer’s patches occurred, and that small amounts of virus could be excreted in feces for as long as 119 days, sometimes associated with chronic tail lesions that also released small amounts of virus. In Gledhill’s experience, such carrier mice did not spread mousepox to uninfected mice by contact, nor was he able to “activate” acute mousepox in the carriers. Nevertheless, such mice clearly constitute a reservoir from which infection could be transferred by the inoculation of contaminated tissue suspensions. More recently, Wallace and Buller (1985) found that following intragastric inoculation, virus was isolated from feces of C57BL/6J mice for as long as 46 days and for up to 29 days from feces of BALB/cByJ mice. Transmission to cage mates from intragastrically infected C57BL/6J and BALB/cByJ occurred up to 36 and 30 days, respectively, after infection. 3.
Arthropod Vectors
Although this mode of transfer is important in several other poxvirus diseases, such as fowlpox and myxomatosis, Fenner was unable to demonstrate mechanical transmission of mousepox by mosquitoes (F. Fenner, unpublished experiments). Guillon (1970) has suggested another possible mode of transfer by arthropods. He found that the rat mite, Ornithonyssus bacoti, became infectious after a blood meal and suggested that it (and perhaps also cockroaches [Guillon 1975]) might act as a passive vector. 4.
Intraperitoneal Inoculation
This has been a commonly used way of passing ECTV. The lesions differ considerably from those observed in the natural disease, and it is obvious that some accounts of mousepox pathology are based upon the results of intraperitoneal inoculation (Fenner 1947b; Schell 1960b). There is, of course, no primary skin lesion, but in acutely fatal cases the necrosis of the liver and spleen resembles that found after natural infection. In addition, there is usually peritonitis with ascites, pancreatic edema, and considerable pleural fluid. In the rare animals that
Fig. 3-3 Postmortem appearances after intraperitoneal injection of ectromelia virus. Necrosis of the liver, hyperemia of the intestine, and peritoneal and pleural effusions are seen.
survive acute infection, general peritonitis is more pronounced (Fig. 3-3). There is a great excess of peritoneal and pleural fluid, the peritoneal surfaces of the liver and spleen are covered with a white exudate, the walls of the intestine are thickened and rigid, and there is often peritoneal fat necrosis. Extensive adhesions develop later. Survivors develop a characteristic rash. Survival times among mice inoculated intraperitoneally are usually 2 or 3 days shorter than those for mice given the same dose of virus inoculated into the foot. 5.
Intranasal Inoculation
Historically, mousepox has been induced unintentionally during mouse lung passage of influenza virus, illustrating that
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inoculation of the respiratory tract can result in generalized disease. Some workers (see, e.g., Kikuth and Gönnert 1940) suggested that ECTV acquires marked pneumotropic properties after serial lung passage, because deaths from pneumonia then occur with little macroscopic evidence of lesions in liver and spleen. When small doses of virus are inoculated intranasally, there is usually little change in the lungs except for the development of patchy congestion (Fenner 1947b; Jacoby and Bhatt 1987). The survival time of fatal cases is approximately the same after footpad inoculation with the same dose of virus, and lesions in liver and spleen are characteristic of acute, naturally acquired mousepox. With larger doses of virus, congestion of the lungs becomes more pronounced and pneumonia may occur. When very large doses are given, death occurs, with patchy or complete consolidation of the lungs and little change in the liver and spleen. Histologically, there is early exudation into the alveoli and bronchi and focal necrosis of bronchial epithelium. ATIs are seen in bronchial epithelium, histiocytes, pleural cells, and eventually in alveolar epithelium. When the lungs, liver, and spleen of fatal cases were examined virologically, titers in apparently normal liver and spleen were very high, just below the threshold at which demonstrable necrosis occurred (F. Fenner, unpublished observations; Ipsen 1945). Necrosis has been found in a few mice that survived for 6 (instead of the usual 4 or 5) days after the intranasal inoculation of a large dose of ECTV. Using fluorescent antibody staining, Roberts (1962a) showed that either macrophages or alveolar mucosal cells were initially infected, but it was the macrophages that carried virus to the pulmonary lymph nodes and thus to the bloodstream, from which it was taken up by the liver and spleen, in which multiplication then proceeded. The apparent pneumotropism is due to the fact that the local reaction, which occurs after the intranasal inoculation of very large doses of virus, kills the animal before there is time for the characteristic changes in the liver and spleen to occur. Similar findings were observed with inbred A/J and BALB/c mice (Jacoby and Bhatt 1987). Low doses of virus in small volumes (under 10 µl) resulted in local upper respiratory tract infections with little involvement of the lung. Death resulted from liver necrosis. A/NCr mice are highly susceptible to this route of infection and LD50 values of 0.3 PFU have been obtained. BALB/c mice are at least 100-fold more resistant (Bhatt and Jacoby 1987a; R. M. L. Buller, unpublished observations). 6.
Lower Respiratory Tract Inoculation
Mice can also be infected with small particle aerosols (mass median diameter of ~0.5 µm). In A/NCr mice, LD50 values have been achieved with presented doses as low as 36 PFU (Buller 2004; Schrewier et al. 2004). As with the intranasal route, infection with low doses of aerosolized virus results in death attributable to liver necrosis with only modest lung involvement.
7.
Intracerebral Inoculation
Kanazawa (1937), Jahn (1939), and Schoen (1938) described intracerebral passage of ECTV, and Kikuth and Gönnert (1940) confirmed their results. Jahn (1939) could not find ATIs in the brain, but Kikuth and Gönnert (1940) described them in neural cells and also in macrophages. It is apparent that after intracerebral inoculation, ECTV rapidly enters the systemic circulation, since virus titers in liver and spleen of fatal cases are always high (F. Fenner, unpublished observations), and except after large doses of virus, characteristic lesions develop (Kanazawa 1937). Jahn (1939) found no evidence of increased neurotropism after 20 brain-to-brain passages in mice. 8.
Intrauterine Infection
Mims (1969) showed that many pregnant mice infected intradermally with the attenuated Hampstead (egg) strain of ECTV survived, but there was extensive growth of virus in the placenta and infection of the fetuses. Infected fetuses died either in utero or soon after birth, and fluorescent antibody staining showed that there was widespread growth of virus throughout their bodies. Schwanzer et al. (1975) obtained similar results. In resistant strains of mice from enzootically infected colonies, intrauterine infection might present as a drop in reproductive success due to fetal deaths; however, in a largely immune population, this problem would probably not occur. Additional problems could arise from the use of mouse embryo cell cultures derived from infected embryos (Germer et al. 1961).
B.
Virus Spread in the Animal Following Footpad Infection
Fenner (1947b, 1949c) believed that the usual portal of entry of the ECTV in natural infections was through small abrasions of the skin. Using fluorescent antibody staining, Roberts (1962b) showed that after scarification, the first cells infected were dermal macrophages. Spread in the dermis initiated “island foci” of epidermal infection in advance of the more slowly spreading main dermal focus. Virus spread from the primary lesion through successive rounds of infection, with multiplication and liberation usually accompanied by cell necrosis. Spread to the lymphatics, often via infected macrophages, caused a sequential infection of local lymph nodes and internal organs (primary viremia; see Fig. 3-4). During the incubation period of 4 to 6 days PI, the virus multiplied locally and in the spleen and liver, and a primary lesion developed at the site of entry (Fig. 3-5). The subsequent disease course depended on the degree of virus replication in the liver and spleen. From the work of Mims (1964), it appeared that the phagocytic cells of the liver and spleen were important for the spread of the virus within the tissues. Virus from spleen and liver tissue released into circulation (secondary viremia) caused
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DAY 0
1
Peripheral routes of infection: lung and skin Regional lymph node: Multiplication
INCUBATION PERIOD
2 Blood-Stream: primary viraemia 3 Spleen and Liver: multiplication necrosis
4
5
Blood-stream: secondary: viraemia
6
Skin: focal infection: multiplication
7 Swelling of Foot: (primary lesion)
DISEASE
8
9
focal infections of the skin and sometimes of the kidneys, lungs, intestines, and other organs. If virus titers escalated, death occurred within 0–4 days of the appearance of the primary lesion (Fenner 1949b), and viral titers in the spleen and liver reached levels of l09 or 1010 pock-forming units per gram. The virus content of the skin was often very high, but not enough time had elapsed for skin lesions to develop. If the virus failed to reach a lethal concentration in the internal organs, virus deposited previously in the epidermis eventually caused multifocal necrosis, producing a generalized rash (Fig. 3-6). Healing of the skin lesions usually occurred in about a week, often leaving hairless scars. The severity of the rash depended upon the degree of viremia, which appeared to be directly related to virus concentration in liver and spleen. Virus can be detected in blood between 3 to 13 days PI, with peak titers of 104 PFU/ml detected on day 8 or 9 (Fenner 1949b). Viral multiplication in each organ followed a logarithmic course; however, titers fell steeply coincident with the onset of circulating antibody and cell-mediated immunity. Rash development was also affected by virus dose, route of inoculation, and mouse strain. It is important to note that all these observations refer to mousepox in young adult mice of a genetically susceptible Hall Institute outbred mouse strain; clinical signs are very different in disease-resistant mouse strains that have non-H-2 linked resistance genes (Section VIII, C).
Early eash Papules
C.
10 1.
11 Severe rash Ulceration Fig. 3-4
Diagram illustrating the pathogenesis of mousepox (Fenner 1948d).
Fig. 3-5
Gross and Microscopic Pathology
Susceptible Mouse Strains
Fenner described the gross and microscopic pathology of mousepox in susceptible outbred Hall Institute mice (Fenner 1948a). More recently, others have reported similar findings in
The primary lesion of mousepox on the left eyebrow of a naturally infected mouse, 8 days (left) and 14 days (right) after infection.
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A
B
C Fig. 3-6 The rash of mousepox as it appears 14 days after infection. (A) Normal mouse of a susceptible strain. (B) Normal mouse after depilation to reveal the rash. (C) In a naturally infected hairless mutant mouse (not athymic). (Courtesy of the Zentralinstitut für Versuchstiere, Hanover, Federal Republic of Germany.)
susceptible inbred mouse strains (BALB/c, DBA/2, and C3H) (Allen et al. 1981; Jacoby and Bhatt 1987). The following description is based mainly on Fenner’s work with susceptible outbred mouse strains. A. SKIN The primary gross lesion is localized swelling, usually surmounted by a minute breach of the surface (Fig. 3-5).
It rapidly increases in size, with pronounced edema of the surrounding tissues. Later, a hard, adherent scab forms and drops off after a week or two, leaving the site of the primary lesion marked by a deep, hairless scar that often persists for life (Fig. 3-7). The earliest histologic lesions precede the described gross change and may reflect viral replication that then had been in
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Fig. 3-7
Mouse that had recovered from mousepox, showing amputation of the foot (“ectromelia” of Marchal) and scars on the face.
progress for a week. The dermis and subcutaneous tissues may be edematous, and there may be lymphocytic infiltration of the dermis. ATIs may be seen in the overlying epidermal cells at the summit of the lesion (Fig. 3-8). Epidermal necrosis leads to ulceration and widespread dermal necrosis. Subsequent scab formation promotes healing. The secondary rash appears 2 or 3 days after development of the primary lesion as slightly raised, pale areas 2 or 3 mm in diameter. They may increase in number and size, ulcerate, and in animals that survive, heal by scarring 3 weeks after infection. Conjunctivitis and blepharitis may occur frequently during the secondary rash, and in severe cases ulcers can be found on the tongue and buccal mucous membrane. The first histologic changes of the secondary rash occur as localized areas of epidermal hyperplasia, with dark-staining nuclei surrounded by vacuoles; epidermal cells may contain intracytoplasmic ATIs. As areas of proliferation and edema increase in size, they become macroscopically visible as pale, slightly raised macules. Numerous ATIs are then present in epidermal cells. Fresh macules develop, and earlier lesions reveal epidermal necrosis of the superficial cells. Massive necrosis, which follows quickly, is accompanied by widespread inflammatory edema and lymphocytic infiltration of the dermis, and is expressed grossly as papules that convert to ulcers with closely adherent scabs. B. LIVER The liver is invariably affected during acute infection and is a prominent site of ECTV replication. The liver may remain macroscopically normal until within 24 hr of death, when it appears enlarged and studded with white foci representing hepatocytic necrosis (Fig. 3-9). Hepatic necrosis can extend rapidly into large semiconfluent zones. In animals that
survive, the liver usually returns to its normal macroscopic appearance, but occasionally numerous white necrotic foci are noted, especially along the anterior border of the median lobe. These foci represent the progression of viral lesion into abscesses. Histologically, little change is apparent until gross lesions also appear, that is, within a day or so of death in rapidly fatal cases; however, fluorescent antibody staining has shown that infection begins in littoral cells of the hepatic ducts, from which it spreads to contiguous parenchymal cells (Mims 1959). Numerous scattered foci of necrosis then develop and, in fatal cases, spread rapidly to semiconfluence (Fig. 3-9). Parenchyma at the margins of necrotic areas may display hepatocytic regeneration with many multinucleate cells, even in fatal cases. Liver regeneration commences early and is active, especially in nonfatal cases, and fibrosis does not occur. Portal tracts may be lightly infiltrated with lymphoid cells, but ATIs are rarely found in infected hepatocytes. The necrotic foci occasionally seen after recovery from infection consist of hyaline necrotic tissue. Survivors may have small accumulations of lymphocytes, usually in portal triads and occasionally in the liver parenchyma. C. SPLEEN The spleen often develops macroscopic changes at least a day earlier than the liver, and higher titers of virus can be found in the spleen each day until death, when virus titers in the spleen and liver are approximately the same. Fluorescent antibody studies (Mims 1964) revealed that virus probably reaches the spleen in infected lymphocytes, which initiate infection in splenic follicles. Infected follicles undergo necrosis, whereas neighboring follicles may show proliferative changes indicative of antibody production in the spleen.
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3. MOUSEPOX
A
B Fig. 3-8 Section of the skin of the foot of a mouse injected with ectromelia virus in the footpad 6 days earlier. (A) Low power. (B) High power. Mann’s stain. Almost every epithelial cell contains an eosinophilic ATI (arrows).
Initial gross lesions of splenic necrosis include pale, slightly depressed zones, which can become semiconfluent. In surviving mice, residual lesions are more common in spleen than liver, and vary from small raised plaques about 1 mm in diameter consisting of hyperplastic serosa to prominent parenchymal fibrosis. These changes constitute the most frequent and reliable gross evidence that a mouse has sustained and recovered from mousepox (Fig. 3-10). Histologically, the early splenic changes consist of hyperplastic lymphoid follicles, congestion of the red pulp, and focal necrosis of lymph follicles. Necrosis can extend rapidly to efface large segments of the parenchyma. D. OTHER LYMPHOID TISSUES Regional lymph nodes draining the primary lesions can be enlarged and show prominent to
confluent necrosis where pyknotic nuclear debris is observed in a featureless background. Numerous ATIs may be present. Lymphocytosis is characteristic of nonregional lymph nodes during recovery. Allen and colleagues (1981) reported thymic necrosis as a major finding in the 1979 National Institutes of Health mousepox outbreak. The necrosis was attributed to direct virus replication using immunohistochemistry, but the disintegration of the cortical thymocytes also pointed to the possibility of a secondary contribution from the stress of severe disease. E. INTESTINE The intestine is often engorged, and the Peyer’s patches can be enlarged in fatal cases. Greenwood et al. (1936) observed necrosis and ATIs in cells of the epithelium in about 65% of acutely fatal cases. Briody (1955, 1959) has commented
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A
B
Fig. 3-9 Acute hepatic necrosis in mousepox. A mouse sacrificed when moribund, 7 days after the injection of a large dose of ectromelia in the footpad. (A) Gross photograph of internal organs. (B) Higher power. There is extensive necrosis of the liver, with little inflammatory reaction.
upon the high frequency with which a hyperemic or bloodfilled small intestine was observed during epizootics of mousepox in the United States, especially in genetically highly susceptible strains of mice. No other organs are regularly affected in natural mousepox, but occasionally, especially in very young mice, necrotic, hemorrhagic foci can occur in the kidneys and in the urinary bladder. Necrosis may be especially prominent in the renal convoluted tubules, with ATIs in tubular epithelium, and sometimes free in the lumina. Renal fibrosis and lymphocytic infiltrations may occur among survivors. 2.
Resistant Mouse Strains
The pathology in genetically resistant C57BL/6 mice is distinctive. First, Jacoby and Bhatt (1987) and then Karupiah et al. (1993a) observed that pathology in ECTV-infected C57BL/6 mice was more restrictive than in susceptible mouse strains. Necrosis in spleen and liver was mild and accompanied by mononuclear cell inflammation and hyperplasia of
lymphoid tissues. Bone marrow, intestines, and oral tissues were relatively uninvolved.
VIII.
INNATE AND ADAPTIVE IMMUNE RESPONSES
The recovery of resistant C57BL6 mice from a primary peripheral (footpad) infection with ECTV is dependent on the seamless integration of innate defense mechanisms, including type I and II interferons, natural killer cells, and monocytes, as well as immune responses characterized by CD8+ cytotoxic T lymphocytes, CD4+ T cells, and antibody.
A.
Innate Resistance
The innate response is activated in the first few hours of infection. It is characterized by the production of soluble
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3. MOUSEPOX
the development of cytotoxic T cells, when natural killer cell lytic activity would be expected to restrict virus replication and spread (Müllbacher et al. 1999).
B.
Fig. 3-10 Scarring of the spleen after recovery from mousepox.
mediators, including interferons and the inflammatory cytokines: interleukin-1, tumor necrosis factor-α, IL-6, and IL-18. In addition, neutrophils, monocytes, and natural killer cells are activated and migrate toward the focus of infection. The importance of these early responses for recovery from infection was determined using genetic or pharmacological approaches to specifically block individual pathways. Karupiah et al. (1993a) found both interferon-α/β and interferon-γ were essential for recovery of resistant C57BL/6 mice. Further studies by Karupiah et al. (1993b) suggested that part of the antiviral activity of interferon-γ was mediated through nitric oxide production. Macrophages were also shown to be crucial for recovery of C57BL/6 mice, as lethal infections followed macrophage depletion by liposome-mediated intracellular delivery of dichloromethylene bisphosphonate (Karupiah et al. 1996). Natural killer cells are also necessary for recovery, as their depletion with rabbit anti-asialo-GM1 or mouse anti-NK1.1 monoclonal antibody resulted in lethal infections (Jacoby et al. 1989; R. M. L. Buller and G. Karupiah, unpublished observations). Consistent with these findings, perforin-deficient mice had enhanced virus replication early during infection, prior to
Adaptive Immunity
The importance of the adaptive response in recovery was initially determined by showing that recovered mice were immune to reinfection (Marchal 1930; Greenwood et al. 1936). Fenner (l949c) showed that 2 weeks after infection, mice were solidly immune to reinfection by footpad inoculation of the virus. This immunity declined slowly, but even after a year, multiplication of the virus was confined to the local skin lesion, and only in occasional animals could virus be isolated from the spleen. Local virus replication in the foot, with consequent swelling, was associated with elevated HI titers. Long-term observation of recovered mice illustrated the epidemiological importance of durable immunity. Only 3 out of 168 immune mice developed signs of reinfection, and this was restricted to a local lesion in the foot (Fenner 1948c). Cell-mediated immunity is critical for recovery from mousepox (for reviews, see Blanden 1974; Müllbacher, Hla, et al. 2003; Müllbacher and Blanden 2004). Pioneering experiments by Blanden (1970) showed that mice pretreated with antithymocyte serum died from otherwise sublethal doses of virus due to uncontrolled viral growth in target organs. These mice had impaired cytotoxic T cell responses but normal neutralizing antibody responses, elevated interferon levels in the spleen, and unchanged innate resistance in target organs. Anti-thymocyte serum–treated mice were protected from lethal infection by the transfer of immune spleen cells harvested 6 days after immunization of donor mice (Blanden 1971a). The transferred cells rapidly eliminated infection from the target organs of the recipients, while no anti-ECTV antibody or interferon was detected in the recipients. The active cells were CD8+ T cells. The kinetics of their generation and the requirement for sharing of H-2K or H-2D genes between donor and recipient identified them as cytotoxic T cells. The importance of CD8+ T cells for recovery from disease was confirmed using C57BL/6 mice depleted of CD8+ T cells by monoclonal antibody therapy or C57BL/6 β2m-/- mice lacking functional MHC class I surface molecules (Karupiah et al. 1996). As expected, C57BL/6 mice lacking granzyme A and B or perforin, which are necessary for FasLindependent CD8+ T cell killing of virus-infected cells, were also highly susceptible to lethal infection (Müllbacher et al. 1996; Müllbacher, Hla, et al. 1999; Müllbacher, Waring, et al. 1999). CD4+ T cells of the helper class, which recognize antigens dependent on the I region of the H-2 complex, are also important for recovery from ECTV infection. In C57BL/6 mice, CD4+ T cells were required for the optimal generation of cytotoxic T cells, although mice survived in their absence (Buller, Holmes, et al. 1987; Karupiah et al. 1996). C57BL/6 mice lacking CD4+ T cells sustained low levels of virus replication in most
82 organs and very high levels of replication in skin (Karupiah et al. 1996). The CD4+ T cells are likely important in facilitating immunoglobulin class switching in the production of anti-ECTV antibody, and in the activation of monocytes during hepatic infection (Blanden 1971b). B cell responses are important in protection from reinfection. Serum from recovered mice contained neutralizing (Burnet and Lush 1936a), HI (Burnet and Boake 1946), and precipitating (Horzinek 1965) antibodies. Serum-neutralizing antibody was not detected until day 8 PI, and virus-specific IgG was detected ~7–10 days PI of C57BL/6 mice (Buller et al. 1983; Chaudhri et al. 2004). Very large doses of interferon or immune serum were relatively ineffective against established infection in target organs, though high serum antibody titers could be demonstrated in the recipients of immune serum (Fenner 1949b; Blanden 1971a). These findings indicate that innate and adaptive responses are necessary for recovery from ECTV infection. Interferons help activate the innate response, including natural killer cells that slow the spread of virus until virus-specific immune CD8+ T cells can further retard viral spread by lysing infected cells before the maturation and assembly of progeny virions. In resistant C57BL/6 mice, cell-mediated immune responses by CD8+ and CD4+ T cells occur soon after infection; for example, virus-specific precursor cytotoxic T cells were detectable in the draining popliteal lymph node 2–3 days after footpad inoculation (O’Neill and Brenan 1987). Cytotoxic T cell activity reached peak levels in the spleen ~5 days later and returned to background levels as early as day 10 PI (Blanden and Gardner 1976). CD8+ T cells supplemented by virus-specific CD4+ T cells attract blood monocytes, which contribute to the elimination of the infection by phagocytosis and intracellular destruction of virus. Delayed type hypersensitivity was detectable by the footpad-swelling test 5–6 days after infection (Fenner 1948a; Owen et al. 1975). Activated macrophages locally produce interferon that may increase the efficiency of virus control and elimination. Recently, Karupiah and colleagues examined ECTV-infected susceptible (BALB/c and A/J) and resistant (C57BL/6) mice strains for cytokine production by CD4+ and CD8+ T cells, cytolytic activity of natural killer and T cells, and virus titers in various tissues, and concluded that polarized type 1 cytokine responses and cell-mediated immunity determined genetic resistance to mousepox (Chaudhri et al. 2004). Two features of the immunological response are important in explaining the natural history of mousepox. First, the effectiveness of cell-mediated immunity helps to explain the absence of clinical signs in some resistant genotypes. Second, lactogenic (humoral) immunity maternal antibody is very effective in protecting highly susceptible mouse pups from lethal infection (Fenner 1948b). Suckling mice attained a serum antibody titer of up to half that present in the mother, and following weaning on the 21st day, the antibody titer fell until the 6th week, when it was no longer detectable. Limited investigations on the spread of the disease in two breeding colonies suggested that
R. MARK, L. BULLER, AND FRANK FENNER
maternal antibody might play an important role in the persistence of the virus in laboratory colonies (see Section IX, D).
C.
Genetics of Resistance to Lethal Mousepox
Fenner’s characterizations of mousepox (Sections VI and VII) were carried out using highly susceptible non-inbred mice. The clinical picture and pathological findings are different if genetically resistant mice are studied. Some studies reported by Schell (1960a,b) in genetically resistant C57BL mice are pertinent. Schell found no difference in the infectivity endpoint of a viral suspension titrated in susceptible outbred mice and C57BL mice, but the titer of virus in the footpad of C57BL mice ceased to rise 6 days after infection, and the highest titers in blood, liver, and spleen were 2 to 3 logs lower than in outbred susceptible mice. Roberts (1964) showed that the rate of growth of ECTV in the littoral (reticuloendothelial) cells of the livers of mice depended upon the virulence of the strain of virus used (virulent Hampstead [mouse] versus attenuated Hampstead [egg]), whereas the subsequent growth of virus in the parenchymal cells of the liver depended upon the mouse strain. Additional studies by O’Neill and Blanden (1983b) suggested that C57BL mice have cells or factors resistant to radiation (950 rads) that impose an early barrier to viral spread from a peripheral site of infection via lymphocytes or blood to liver and spleen. Differences in immunological responses were also noted between mouse strains. Neutralizing antibody, delayed type hypersensitivity, and active immunity were demonstrable 1 or 2 days earlier in C57BL mice compared to outbred mice (Schell 1960a). Breeding experiments by Schell (1960b) and Ermolaeva et al. (1974) demonstrated that resistance was largely determined by a single dominant gene that was not linked to coat color or H-2 genotype (O’Neill et al. 1983a). Studies by Wallace et al. (1985) using additional susceptible DBA/2J, A/J, and BALB/cByJ mice and resistant C57BL/6J and AKR/J mice confirmed that resistance was determined by one or more independently assorting autosomal loci with dominant alleles for resistance in AKR/J and C57BL/6J mice and recessive alleles in A/J, BALB/cByJ, and DBA/2J mice. In the (B6 × A)F1 × A backcross, resistance to lethal infection appeared to be controlled by a single autosomal dominant gene, provisionally termed Rmp-1 (resistance to mousepox). Subsequent experiments by Brownstein and colleagues (1992) using a (C57BL/6 × DBA/2) F1 × DBA/2 backcross identified three additional C57BL/6 genes, Rmp-2, Rmp-3, and Rmp-4 and mapped the location of all four genes. Rmp-1 maps to an interval of chromosome 6 that includes the natural killer cell gene complex including the NKR-P1 and Ly-49 families of proteins (Delano and Brownstein 1995). Rmp-2 maps to an interval of chromosome 2 that includes the structural gene for the fifth component of complement (Brownstein et al. 1992). Rmp-3 and Rmp-4 map to chromosome 17 and chromosome 1, respectively (Brownstein
83
3. MOUSEPOX
et al. 1992; Brownstein and Gras 1995). Thus the genetics of resistance to mousepox are complex and can vary depending on which strain of ECTV is used (Brownstein et al. 1989).
IX.
EPIZOOTIOLOGY A.
1.
Host Range
Species
The mouse can be infected by all routes of inoculation with ECTV. Several other species of laboratory animals apart from Mus musculus have been tested for susceptibility to ECTV, as summarized below. A. OTHER SPECIES OF MUS There are two reports suggesting that mousepox might occur as a disease of wild rodents, other than of wild house mice associated with an outbreak in laboratory mice (e.g., McGaughey and Whitehead 1933). Gröppel (1962) examined wild mice belonging to three genera (Microtus, Apodemus, and Cletrionomys) that were captured in several rural localities in Germany, distant from possible contamination from laboratory mice. His observations may suggest that several of the Apodemus animals were infected with an agent that was probably ECTV. Kaplan et al. (1980) found that sera from several voles and woodmice captured in the wild in the United Kingdom contained complement-fixing antibodies against an orthopoxvirus that he proposed was ECTV. More recent studies suggest that the orthopoxvirus that is maintained in these British rodent populations is cowpox virus (Crouch et al. 1995; Bennett et al. 1997; Hazel et al. 2000). Resistance to severe mousepox varied across four genera of rodents (Buller et al. 1986). Experimental infections of representative species from the genus Mus (caroli, cookii, spretus, and cervicolor), and the genus Nannomys (minutoides) resulted in high mortality. Coelomys pahari and Pyromys platythrix could be infected, as indicated by seroconversion, but showed no signs of disease. B.
Mooser (1943) found that when peritoneal fluid from moribund mice, which were infected with both ECTV and Rickettsia prowaseki, was inoculated intraperitoneally into rats, there was a severe rickettsial peritonitis. Characteristic ectromelia ATIs were also seen in smears of the peritoneal fluid, suggesting that ECTV had replicated.
RAT Reports that rats can be naturally infected with ECTV and may indeed be a reservoir of the virus in nature (e.g., Iftimovici et al. 1976) are unconvincing. Burnet and Lush (1936a) showed that when large doses of virus were inoculated intranasally into the rat, an inapparent infection of the olfactory mucosa occurred, accompanied by the development of neutralizing antibody. This finding was confirmed by Reames (1940). Intradermal inoculation of large doses of ECTV produced seroconversion but no lesions, or at most a tiny papule at the site of injection (F. Fenner, unpublished observations). Intraperitoneal inoculation was also without effect except for seroconversion. Repeated ECTV inoculations into rats produced no further increase in antibody titer.
C. RABBIT Early workers found the rabbit was resistant to ECTV infection, although Paschen (1936) reported that virus that had been passed several times on the chorioallantois produced infection of the rabbit’s cornea, with the production of ATIs, and could then be passed to the rabbit skin. Burnet and Boake (1946) found that two rabbits inoculated intravenously with a large dose of virus died 6 and 10 days later, and hemagglutinin was present in suspensions of liver and spleen. Intradermal inoculation of ECTV produced indurated papules. Such papules regularly appeared about 4 days after intradermal inoculation of large doses of virus and sometimes ulcerated a few days later (F. Fenner, unpublished observations). In rabbits immunized by a prior infection, the papule formation and regression were accelerated, reaching their maximum size by the second day and fading by the sixth day. Such intradermal infections also resulted in seroconversion. Christensen et al. (1966) showed that ECTV could be used to immunize rabbits against rabbitpox, although it was preferable to use vaccinia virus for this purpose because of the danger to the mouse colony. D. GUINEA PIG Paschen (1936) found that inoculation of the guinea pig foot or cornea with egg-passaged ECTV but not with mouse liver virus produced typical ATIs. Intradermal inoculation of chick embryo membrane preparations of the virus regularly produced local indurated lesions, and HI antibodies were detected in the sera 14 days later (F. Fenner, unpublished observations). Intraperitoneal inoculation of large doses of ECTV caused no signs, but was followed by seroconversion. E. HAMSTER According to Flynn and Briody (1962), Syrian hamsters are not susceptible to infection with ECTV. F. MAN The only conscious attempt at infection of humans with ECTV was scarification of two men who had been vaccinated with vaccinia virus several times previously (F. Fenner, unpublished observations). In both a small papule appeared on the second day, became slightly vesicular in one on the fourth day, and disappeared by the eighth day. No significant increase in antibody titer was observed. Packalén (1947) showed the Laigret-Durand strain of “mousepathogenic murine typhus rickettsia,” like the epidemic typhus strain studied by Mooser (1943), owed its mouse pathogenicity to its ECTV content. Thus active ECTV, either alone or mixed with rickettsiae, was inoculated subcutaneously into hundreds of thousands of humans in doses of up to 1000 “mouse units” (Laigret and Durand 1939, 1941; Packalén 1945). No local or
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R. MARK, L. BULLER, AND FRANK FENNER
TABLE 3-3
TABLE 3-4
INFLUENCE OF GENOTYPE ON MORTALITY IN AN EPIZOOTIC OF MOUSEPOXa
COMPARISON OF INFECTIVITY AND MORTALITY OF ECTROMELIA VIRUS IN INBRED STRAINS OF MICEa
Case fatality rate Genotype
Stock mice (%)
DBA/1 A C3H MA/Nd C57BL/6 BALB/c AKR MA Sy N Sy D
84 84 71 2
